Paradigms of Neural Injury, Volume 30 (Methods in Neurosciences)

Paradigms of Neural Injury

Paradigms of Neural Injury

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Paradigms of Neural Injury

Edited by J. Regino Perez-Polo Department of Human Biological Chemistry and Genetics University of Texas Medical Branch Galveston, Texas

ACADEMIC PRESS San Diego New York Boston

London

Sydney Tokyo Toronto

Front cover photograph: Computer-enhanced picture of dissociated embryonic chick sensory neurons differentiated with 10 ng/ml nerve growth factor.

This book is printed on acid-free paper. Copyright 9 1996 by ACADEMIC PRESS, INC. All Rights Reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopy, recording, or any information storage and retrieval system, without permission in writing from the publisher.

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International Standard Serial Number: 1043-9471 International Standard Book Number: 0-12-185300-4

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Table of Contents

Contributors Preface Volumes in Series

vii xi xiii

1. Paradigms for Study of Neurotrophin Effects in Oxidant Injury George R. Jackson and J. Regino Perez-Polo

2. Nitric Oxide Toxicity in Central Nervous System Cultures

26

Valina L. Dawson and Ted M. Dawson

3. Development of in Vitro Injury Models for Oligodendroglia

44

A. Espinosa, P. Zhao, and J. de Vellis

4. Glia Models to Study Glial Cell Cytotoxicity

55

Antonia Vernadakis and M. Susan Kentroti

5. Rodent Glioma Models

81

William W. Maggio

6. Peripheral Lesioning of Olfactory System: Expression of Neurotrophin Receptors

97

Christopher P. Turner and J. Regino Perez-Polo

7. Basal Forebrain Cholinergic Lesions and Complete Transection of Septal-Hippocampal Pathway

106

Lawrence R. Williams ,

Neurochemical Lesions: Tools for Functional Assessment of Serotonin Neuronal Systems

115

Joan M. Lakoski, B. Jane Keck, and Ashish Dugar ,

Cerebral Glucose/Energy Metabolism: Valid Techniques in Humans and Animals

124

Siegfried Hoyer

10. Heavy Metal Effects on Glia

135

Evelyn Tiffany-Castiglioni, Marie E. Legare, Lora A. Schneider, Edward D. Harris, Rola Barhoumi, Jan Zmudzki, Yongchang Qian, and Robert C. Burghardt

11. Source, Metabolism, and Function of Cysteine and Glutathione in the Central Nervous System David K. Rassin

167

vi

TABLE OF CONTENTS 12. Magnetic Resonance Spectroscopy of Neural Tissue

178

Richard J. McClure, Kanagasabai Panchalingam, William E. Klunk, and Jay W. Pettegrew

13. Acute Stroke Diagnosis with Magnetic Resonance Imaging

209

Stephen C. Jones, Neng C. Huang, Michael J. Quast, Alejandro D. Perez-Trepechio, Gilbert R. Hillman, and Thomas A. Kent

14. Evaluation of Free Radical-Initiated Oxidant Events within the Nervous System

243

Stephen C. Bondy

15. Exogenous Administration of Cytokines into the Central Nervous System: Analysis of Alterations in Cell Morphology and Molecular Expression

260

M. A. Kahn and J. de Vellis

16. Animal Models to Produce Cortical Cholinergic Dysfunction

275

Reinhard Schliebs and Volker Bigl

17. In Vitro Studies of Liposome-Mediated Gene Transfection

290

K. Yang, J. Regino Perez-Polo, F. Faustinella, G. Taglialatela, and R. L. Hayes

18. Construction and Analysis of Transgenic Mice Expressing Amyloidogenic Fragments of Alzheimer Amyloid Protein Precursor

298

Rachael L. Neve and Frederick M. Boyce

19. Golgi Technique Used to Study Stress and Glucocorticoid Effects on Hippocampal Neuronal Morphology

315

Ana Maria Magarifzos, Eberhard Fuchs, Gabriele Fliigge, and Bruce S. McEwen

Index

327

Contributors

Article numbers are in parentheses following the names of contributors. Affiliations listed are current.

ROLA BARHOUMI (10), Department of Veterinary Anatomy and Public Health, Texas A & M University, College Station, Texas 77845 VOLKER BIGL (16), Paul Flechsig Institute for Brain Research, University of Leipzig, D-04109 Leipzig, Germany STEPHEN C. BONDY (14), Department of Community and Environmental Medicine, Center for Occupational and Environmental Health, University of California, Irvine, Irvine, California 92717 FREDERICK M. BOYCE (18), Department of Neurology, Harvard Medical School, Massachusetts General Hospital, Charlestown, Massachusetts 02129 ROBERT C. BURGHARDT(10), Department of Veterinary Anatomy and Public Health, Texas A & M University, College Station, Texas 77845 TED M. DAWSON (2), Departments of Neurology and Physiology, The Johns Hopkins University School of Medicine, Baltimore, Maryland 21287 VALINA L. DAWSON (2), Departments of Neurology and Physiology, The Johns Hopkins University School of Medicine, Baltimore, Maryland 21287 J. DE VELLIS (3, 15), Departments of Neurobiology and Psychiatry, Mental Retardation Research Center, University of California, Los Angeles, School of Medicine, Los Angeles, California 90024 ASHISH DUGAR (8), Departments of Pharmacology and Anesthesia, Pennsylvania State University, College of Medicine, Hershey, Pennsylvania 17033 A. ESPINOSA (3), Department of Neurobiochemistry, Mental Retardation Research Center, University of California, Los Angeles, School of Medicine, Los Angeles, California 90024 F. FAUSTINELLA(17), Department of Medicine, Baylor College of Medicine, Houston, Texas 77030 GABRIELE FLC)GGE(19), Division of Neurobiology, German Primate Center, D-37077 G6ttingen, Germany EBERHARD FUCHS (19), Division of Neurobiology, German Primate Center, D-37077 G6ttingen, Germany

vii

viii

CONTRIBUTORS

EDWARD D. HARRIS (10), Department of Biochemistry and Biophysics, Texas A & M University, College Station, Texas 77845 R. L. HAYES (17), Department of Neurosurgery, Health Science Center, University of Texas, Houston, Houston, Texas 77030 GILBERT R. HILLMAN (13), Departments of Pharmacology and Toxicology and Academic Computing, University of Texas Medical Branch, Galveston, Texas 77555 SIEGFRIED HOVER (9), Brain Metabolism Group, Departments of Pathochemistry and General Neurochemistry, University of Heidelberg, 69120 Heidelberg, Germany NENG C. HUANG (13), Departments of Pharmacology and Toxicology, and Marine Biomedical Institute, The University of Texas Medical Branch, Galveston, Texas 77555 GEORGE R. JACKSON(1), Department of Neurology, University of California, Los Angeles, School of Medicine, Los Angeles, California 90024 STEPHEN C. JONES (13), Cerebrovascular Research Laboratory, Department of Biomedical Engineering, Research Institute, Cleveland Clinic Foundation, Cleveland, Ohio 44195 M. A. KAHN (15), Departments of Neurobiology, Psychiatry, and Biobehavioral Sciences, Mental Retardation Research Center, University of California, Los Angeles, School of Medicine, Los Angeles, California 90024 B. JANE KECK (8), Department of Pharmacology, Pennsylvania State University College of Medicine, Hershey, Pennsylvania 17033 THOMAS A. KENT (13), Departments of Neurology and Pharmacology and Toxicology, University of Texas Medical Branch, Galveston, Texas 77555 M. SUSAN KENTROTI (4), Department of Pharmacology, University of Colorado Health Sciences Center, Denver, Colorado 80262 WILLIAM E. KLUNK (12), Department of Psychiatry, University of Pittsburgh, Pittsburgh, Pennsylvania 15213 JOAN M. LAKOSKI(8), Departments of Pharmacology and Anesthesia, Pennsylvania State University College of Medicine, Hershey, Pennsylvania 17033 MARIE E. LEGARE (10), Department of Veterinary Anatomy and Public Health, Texas A & M University, College Station, Texas 77845 ANA MARfA MAGARIlqOS(19), Neuroendocrinology Laboratory, Rockefeller University, New York, New York 10021

CONTRIBUTORS

ix

WILLIAM W. MAGGIO (5), Division of Neurosurgery, University of Texas Medical Branch, Galveston, Texas 77555 RICHARD J. MCCLURE (12), Department of Psychiatry, University of Pittsburgh, Pittsburgh, Pennsylvania 15213 BRUCE S. MCEWEN (19), Neuroendocrinology Laboratory, Rockefeller University, New York, New York 10021 RACHAEL L. NEVE (18), Department of Genetics, Harvard Medical School, McLean Hospital, Belmont, Massachusetts 02178 KANAGASABAI PANCHALINGAM(12), Department of Psychiatry, University of Pittsburgh, Pittsburgh, Pennsylvania 15213 J. REGINO PEREZ-POLO (1, 6, 17), Department of Human Biological Chemis-

try and Genetics, University of Texas Medical Branch, Galveston, Texas 77555 ALEJANDRO D. PEREZ-TREPECHIO (13), Cerebrovascular Research Laboratory, Department of Biomedical Engineering, Research Institute, Cleveland Clinic Foundation, Cleveland, Ohio 44195 JAY W. PETTEGREW(12), Departments of Psychiatry and Neurology, Health Service Administration, University of Pittsburgh, Pittsburgh, Pennsylvania 15213 YONGCHANGQIAN (10), Department of Biochemistry and Biophysics, Texas A & M University, College Station, Texas 77845 MICHAEL J. QUAST (13), Department of Radiology, and Marine Biomedical Institute, The University of Texas Medical Branch, Galveston, Texas 77555 DAVID K. RASSIN (11), Department of Pediatrics, The University of Texas Medical Branch, Galveston, Texas 77555 REINHARD SCHLIEBS(16), Paul Flechsig Institute for Brain Research, University of Leipzig, D-04109 Leipzig, Germany LORA A. SCHNEIDER (10), Department of Veterinary Anatomy and Public Health, Texas A & M University, College Station, Texas 77845 G. TAGLIALATELA(17), Department of Human Biochemistry and Genetics, University of Texas Medical Branch, Galveston, Texas 77555 EVELYN TIFFANY-CASTIGLIONI (10), Department of Veterinary Anatomy and Public Health, Texas A & M University, College Station, Texas 77845

CHRISTOPHER P. TURNER(6), Department of Neurology, Veterans Administration Medical Center, San Francisco, California 94121

X

CONTRIBUTORS ANTONIA VERNADAKIS (4), Departments of Psychiatry and Pharmacology, University of Colorado Health Sciences Center, Denver, Colorado 80262 LAWRENCE R. WILLIAMS (7), Amgen Neuroscience, Thousand Oaks, California 91320 K. YANG (17), Department of Neurosurgery, University of Texas, Houston, Health Science Center, Houston, Texas 77030 P. ZHAO (3), Department of Neurobiochemistry, Mental Retardation Research Center, University of California, Los Angeles, School of Medicine, Los Angeles, California 90024 JAN ZMUDZKI(10), Department of Pharmacology and Toxicology, Veterinary Research Institute, 24-100 Pulawy, Poland

Preface

The classic perturbation paradigms for the study of the nervous system date to the turn of the century. They relied on the augmentation of tissue present and the removal of tissues of interest, followed by careful microscopic work dependent on artfully conceived tissue fixation and staining protocols. Over the intervening decades, as neurotransmitters and their agonists and antagonists, growth factors and cytokines, and their antibodies and antisense oligonucleotides proliferated and newer and ever more discerning instruments and techniques were developed, the level of sophistication and physiological acuity of the models has increased, perhaps culminating with the most recent additions of transgenic mice, patch clamp measures of electrical activity and capture of unicellular mRNA samples, and noninvasive imaging of the nervous system. As research reports and interpretations of these paradigms become widespread, there is a tendency to focus on the results obtained and on their biological or clinical significance without a sufficient assessment of the actual paradigm being exploited. The result is that in some instances a technique may be adopted to other systems with sometimes confusing consequences. In the interest of providing a forum for the evaluation of some of the currently used paradigms, we offer in this volume an ecletic collection of a number of lesion and probing paradigms in use today. While it would be impractical to make such a list exhaustive, there is a sufficient variety to provide investigators with some insights into a technique being considered for adoption and, perhaps of more utility, some suggestions as to the power and limitations of such paradigms in general. We trust that the techniques presented will be useful to those selecting paradigms for perturbing neural structures or, perhaps, more wisely deter their inappropriate application. J. REGINO PEREZ-POLO

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Methods in Neurosciences

Editor-in-Chief P. Michael Conn

Volume 1 Gene Probes Edited by P. Michael Conn Volume 2 Cell Culture Edited by P. Michael Conn Volume 3 Quantitative and Qualitative Microscopy Edited by P. Michael Conn Volume 4 Electrophysiology and Microinjection Edited by P. Michael Conn Volume 5 Neuropeptide Technology: Gene Expression and Neuropeptide Receptors Edited by P. Michael Conn Volume 6 Neuropeptide Technology: Synthesis, Assay, Purification, and Processing Edited by P. Michael Conn Volume 7 Lesions and Transplantation Edited by P. Michael Conn Volume 8 Neurotoxins Edited by P. Michael Conn Volume 9 Gene Expression in Neural Tissues Edited by P. Michael Conn Volume 10 Computers and Computations in the Neurosciences Edited by P. Michael Conn Volume 11 Receptors: Model Systems and Specific Receptors Edited by P. Michael Conn Volume 12 Receptors: Molecular Biology, Receptor Subclasses, Localization, and Ligand Design Edited by P. Michael Conn Volume 13 Neuropeptide Analogs, Conjugates, and Fragments Edited by P. Michael Conn Volume 14 Paradigms for the Study of Behavior Edited by P. Michael Conn Volume 15 Photoreceptor Cells Edited by Paul A. Hargrave Volume 16 Neurobiology of Cytokines (Part A) Edited by Errol B. De Souza

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METHODS IN NEUROSCIENCES Volume 17 Neurobiology of Cytokines (Part B) Edited by Errol B. De Souza Volume 18 Lipid Metabolism in Signaling Systems Edited by John N. Fain Volume 19 Ion Channels of Excitable Membranes Edited by Toshio Narahashi Volume 20 Pulsatility in Neuroendocrine Systems Edited by Jon E. Levine Volume 21 Providing Pharmacological Access to the Brain: Alternate Approaches Edited by Thomas R. Flanagan, Dwaine F. Emerich, and Shelley R. Winn Volume 22 Neurobiology of Steroids Edited by E. Ronald de Kloet and Win Sutanto Volume 23 Peptidases and Neuropeptide Processing Edited by A. lan Smith Volume 24 Neuroimmunology Edited by M. lan Phillips and Dwight Evans Volume 25 Receptor Molecular Biology Edited by Stuart C. Sealfon Volume 26 PCR in Neuroscience Edited by Gobinda Sarkar Volume 27 Measurement and Manipulation of Intracellular Ions Edited by Jacob Kraicer and S. J. Dixon Volume 28 Quantitative Neuroendocrinology Edited by Michael L. Johnson and Johannes D. Veldhuis Volume 29 G Proteins Edited by Patrick C. Roche Volume 30 Paradigms of Neural Injury Edited by J. Regino Perez-Polo Volume 31

Nitric Oxide Synthase: Characterization and Functional Analysis Edited by Mahin D. Maines

[1]

Paradigms for Study of Neurotrophin Effects in Oxidant Injury G e o r g e R. J a c k s o n a n d J. R e g i n o P e r e z - P o l o

Neurotrophins

a n d Cell D e a t h

Naturally occurring cell death, or apoptosis, is a process critical to understanding neural development, injury, and regeneration. Apoptosis is sometimes referred to as "programmed" cell death, a description that unfortunately conveys a sense of preordained order not befitting the more ambiently regulated shaping and pruning of synaptic contacts that take place in the developing nervous system. Thus, neuronal cell death more appropriately may be considered as probabilistic or stochastic, rather than apoptotic (Hamburger and Oppenheim, 1982). Naturally occurring ontogenic cell death serves to establish specificity of synaptic connections. The attrition of neuronal populations to match fields of innervation, resulting in cell death and neurite pruning, may be considered positive regulatory factors in the establishment of specificity in the developing nervous system. (Cowan et al., 1984; Oppenheim, 1985; Hamburger and Oppenheim, 1982). Cell death following injury or during senescence of the nervous system, on the other hand, may have proved life threatening to the organism, particularly in the case of injury to the central nervous system, due to the limited capacity of these neurons to regenerate. Neurotrophins regulate naturally occurring and injury-induced neuronal cell death. In the former case, nerve growth factor (NGF) synthesized in target cells becomes available to some axonal endings and is retrogradely transported to neuronal somata during a critical developmental period (Hendry et al., 1974; Hendry, 1975; Johnson et al., 1987). In this fashion, the survival of neurons that have successfully synapsed with their targets is assured. Those neurons that fail to synapse and take up NGF die. In the case of the superior cervical ganglion, as much as 30% of the total neuronal population is lost during this period of attrition (Hendry and Campbell, 1976). Neurotrophins such as the nerve growth factor protein may also play a role in neuronal cell death that occurs as a consequence of injury. The injury-induced synthesis and secretion of neurotrophic factors are believed to facilitate functional recovery by stimulating axonal process formation (Nieto-Sampedro et al., 1983, 1984; Nieto-Sampedro and Cotman, 1985; Needels et al., 1985, 1986; Whittemore et al., 1985). The NGF-related family of neurotrophins is now known to include brain-derived neurotrophic factor, Methods in Neurosciences, Volume 30

Copyright 9 1996 by Academic Press, Inc. All rights of reproduction in any form reserved.

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PARADIGMS OF N E U R A L I N J U R Y

neurotrophin 3, and neurotrophin 4/5 (for reviews see Korsching, 1993; Lindsay et al., 1994; Maness et al., 1994). These factors achieve their neurotrophic effects via interaction with a low-affinity NGF receptor, referred to as p75 NCvR, and/or high-affinity tyrosine kinase receptors, referred to as Trk, TrkB, and TrkC (for reviews see Barbacid, 1993; Maness et al., 1994). Unrelated factors that have also been attributed neurotrophic effects include ciliary neurotrophic factor, leukemia inhibitory factor, and basic fibroblast growth factor (Ip and Yancopoulos, 1992; Baird, 1994; Murphy et al., 1993; Thaler et al., 1994). Despite these advances in understanding the physiology of neurotrophins and their receptors, the cellular and molecular mechanisms underlying the survival-promoting activity of neurotrophic factors remain poorly characterized. In contrast to better characterized effects of NGF on neurotransmitter metabolism and neurite formation, there is a dearth of knowledge about the mechanisms whereby NGF enhances cell survival. One approach to understanding cell death in mammals has been to study that occurring in more simple organisms. Genes regulating apoptosis have been characterized extensively in the nematode Caenorhabditis elegans (Ellis et al., 1991). Two C. elegans genes, ced-3 and ced-4, encode proteins that play an active role in cell death. A third gene, ced-9, encodes a protein that is a negative regulator of cell death. The mammalian homolog of ced-9 is the protooncogene bcl-2 (Hengartner and Horvitz, 1994). This gene, originally characterized at the breakpoint of a t(14: 18)-bearing B cell lymphoma, is now known to exert neurotrophic effects. High-level expression of bcl-2 in vitro prevents the death of NGF-deprived sympathetic neurons and of PC12 cells grown in serum-free medium (Garcia et al., 1992; Batistatou et al., 1993; Mah et al., 1993). The protein product of this gene has been ascribed an antioxidant function (Hockenberry et al., 1993; Kane et al., 1993). More recently, the mammalian homolog of ced-3, interleukin-1/~ converting enzyme (Yuan et al., 1993), has been shown to prevent NGF deprivation-induced apoptotic cell death in vitro (Gagliardini et al., 1994). Although a number of hypotheses have been advanced to explain neuronal cell death, definitive explanations for the survival-promoting effects of NGF remain elusive. The observation that inhibitors of transcription or translation can prevent naturally occurring neuronal cell death supports the existence of suicide genes encoding death proteins, or "thanatins" (Martin et al., 1988; Oppenheim et al., 1990; Scott and Davies, 1990); however, only one such protein, cyclin D~, has been demonstrated to be preferentially expressed in dying cells (Freeman et al., 1994). Another explanation for the observation that inhibitors of protein synthesis inhibit apoptosis is that such inhibitors preferentially shunt cysteine toward glutathione synthesis (Ratan et al., 1994).

[1] NEUROTROPHINEFFECTS IN OXIDANT INJURY Excitatory amino acid toxicity has also been implicated in ischemiareperfusion injury (Rothman, 1984; Simon et al., 1984; Drejer et al., 1985; Goldberg et al., 1987; Kochhar et al., 1988; Choi and Rothman, 1990; Lipton and Rosenberg, 1994). Recent evidence for the existence of a redox modulatory site on the NMDA receptor that affects excitotoxicity, as well as other relationships between glutamate and free radicals (Aizenman et al., 1990; Pelligrini-Giampetro et al., 1990), would suggest that interactions between excitotoxins and reactive oxygen species in the genesis of cell injury are more prevalent than previously assumed. The role of DNA fragmentation in apoptosis has been described in studies of glucocorticoid-induced effects on lymphocyte lysis, an event that may be relevant to the ontogeny of thymic autotolerance (Compton et al., 1987; Odaka et al., 1990). Internucleosomal DNA fragmentation has also been characterized in apoptosis associated with neuronal cell death, including that seen in NGF-deprived sympathetic neurons (Dipasquale et al., 1991; Deckwerth and Johnson, 1993; Diana et al., 1993).

Reactive

Oxygen

Species

and Cell Death

The generation of oxygen free radicals following traumatic or ischemic injury followed by reperfusion has been described in the nervous system and is held to be a biologically significant phenomenon (HalliweU and Gutteridge, 1985; McCord, 1987; Braughler and Hall, 1989; Hall and Braughler, 1989). An increasing amount of attention has been devoted to the role of oxygen free radicals in neuronal cell death following injury in a number of pathophysiologic conditions, including seizures, ischemia-reperfusion, and inflammation (Halliwell and Gutteridge, 1985; McCord, 1987; Braughler and Hall, 1989). Experimental investigations in the field of free radicals are fraught with technical difficulties, not the least of which are the instability and transient nature of many species of interest. Consequently, many studies seeking to link oxygen radicals to pathophysiology have employed indirect means, including analysis of end products of free radical-produced reactions, such as malondialdehyde (Dahle et al., 1962; Gutteridge, 1981). An alternative indirect approach has been to demonstrate inhibition of these reactions by known antioxidants (Simon et al., 1981; Hall and Braughler, 1989; Topinka et al., 1989). Neither kind of indirect approach, despite its utility in assessment of radical-induced injury, provides conclusive evidence linking free radical effects with pathology. Activated oxygen species, both endogenous and exogenous to the nervous system, may arise in a number of physiologic and pathologic states besides ischemia-reperfusion. The superoxide anion can be produced in mitochondria in several ways, including quinone autoxida-

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PARADIGMS OF NEURAL INJURY

tion (Misra and Fridovich, 1972; Boveris et al., 1976; Patole et al., 1986). Peroxisomes generate H202 directly (Masters and Holmes, 1977). The superoxide ion may be formed during arachidonate metabolism catalyzed by cyclooxygenase and lipoxygenase (Kukreja et al., 1986). Oxidation of catecholamines by monoamine oxidase may be an important endogenous source of H202 in the nervous system (Marker et al., 1981). Exogenous sources of reactive oxygen species may become increasingly important as the response to injury progresses. The generation of reactive oxygen metabolites by phagocytic enzyme systems such as NADPH oxidase and myeloperoxidase, as well as the microbicidal function of these mechanisms, have been documented (Klebanoff, 1968; Babior et al., 1976; Gabig and Lefker, 1984). The role of oxygen radicals in tissue destruction during inflammation has also been demonstrated (McCord, 1987); whether analogous processes occur in the late stages of injury to the nervous system, when secondary waves of neuronal cell death may emanate from primary traumatic or ischemic foci, is not known. Also, the generation of substantial amounts of reactive oxygen species, such as H202, by activated phagocytes may be a major source of such species following injury (Means and Anderson, 1983; Hallenbeck et al., 1986). Reactive oxygen metabolites considered to be of biologic importance include H202, the superoxide anion (O~), and the hydroxyl radical (. OH), and nitric oxide (NO). Lipid peroxidation in unsaturated fatty acids of membrane phospholipids may be initiated by 9OH (Raleigh et al., 1977; Gutteridge, 1982). Once initiated, membrane lipid peroxidation may be propagated, or terminated through the action of a number of cellular antioxidants, including glutathione and c~-tocopherol (Tappel, 1954; Lucy, 1972; Tsan et al., 1985). Hydroxyl radicals are one of the most reactive metabolites of oxygen. By interacting with ferrous iron, H202 is capable of forming 9OH in a Fentontype reaction [Eq. (1)]: F e 2+ + H202---> F e 3+ + O H - + . O H O~ + H202 -'--> 0 2 -}- O H - + 9O H

(1) (2)

Hemoglobin released from erythrocyte lysis following trauma may result in 9OH formation by the Fenton reaction (Gutteridge, 1986). The generation o f . OH can also occur by the Haber-Weiss reaction (2), in which iron catalyzes a one-electron transfer from O~ to H202 (Haber and Weiss, 1934). The ferrous iron then reacts with H202 to generate ferric iron, OH-, and 9OH. During ischemia, degradation of ATP may result in accumulation of hypoxanthine (Kleihues et al., 1974). Xanthine dehydrogenase may be converted proteolytically to xanthine oxidase during ischemia (Batelli, 1980).

[1] NEUROTROPHIN EFFECTS IN OXIDANT INJURY

H20 O2+H20 .= (R) l~k GSSG NADPH OH _ s ~1 /~. jz/X~ / \ Shunt 02 O~i > H202 / GSH ~ NADP R

-

--glutamy~ (~ t r ~ t i d a ~ ~'~-> Cys~Gly cycle

_

~'-Glu-Cys

~l;lutamylcysteine

3'-Glu-AA Cys ~ Glu ~, ~ ~+ f -glutarnyl~ I protein I / s-ox~o,,~=~ cyclotransferase ) ~.~ / AA ~-" 5-Oxoproline

FIG. 1 Overview of the metabolism of reactive oxygen species. For details, see text. From Jackson, G. R. Werrbach-Perez, K., Pan, Z., Sampath, D., and PerezPolo, J. R., Dev. Neurosci. (1995), with permission.

The action of this enzyme on hypoxanthine results in the formation of O~ (McCord and Fridovich, 1968). Under biologic conditions, the Haber-Weiss reaction occurs very slowly, but substantial 9OH formation by the Fenton reaction takes place (Freeman and Crapo, 1982). Nitric oxide is capable of interacting with superoxide to form the toxic peroxynitrile ion. On the other hand, nitric oxide may under some conditions exert a neuroprotective effect via interaction with a redox modulatory site of the NMDA receptor (Lipton et al., 1993). Figure 1 shows a simplified scheme of reactive oxygen metabolites and antioxidant defenses. Superoxide dismutase catalyzes the conversion of the superoxide anion to H202 (McCord and Fridovich, 1969). Catalase and glutathione peroxidase catalyze the degradation of H202 (Mills, 1957, 1960; Deisseroth and Dounce, 1970; Chance et al., 1979). The kinetic properties of the latter enzyme suggest that it may be of more importance under relatively low H202 concentrations (Cohen and Hochstein, 1963; Flohe and Brand, 1969). The importance of the glutathione peroxidase system as compared to catalase in oxidative injury has been emphasized by some investigators but disputed by others (Sadrzadeh et al., 1984; Suttorp et al., 1986; Gaetani et al., 1989; Giblin et al., 1990). On the other hand, the high capacity of catalase for H202, based on its rapid turnover number, would suggest that catalase activity is diffusion limited (Chance et al., 1979). Because the activity

6

PARADIGMS OF NEURAL INJURY

of the peroxidase system is closely linked to regeneration of GSH by GSH reductase and NADPH (Arrick et al., 1982; Harlan et al., 1984; Schrauffstatter et al., 1985), the availability of reducing equivalents for regeneration of GSH may become a critical factor regulating this pathway. The activity of enzymes capable of generating NADPH, such as malic enzyme and the hexose monophosphate shunt enzymes glucose-6-phosphate dehydrogenase and 6-phosphogluconate dehydrogenase, may also play a critical role in cellular responses to oxidative stress by supplying reducing equivalents (Harlan et al., 1984; Schrauffstatter et al., 1985; Suttorp et al., 1986; Meister et al., 1988; White et al., 1988). The coordinated operation of a number of cellular defense mechanisms is required for an integrated response to oxidative stress, the relative contribution of each element to the overall response depending on the severity of the injury sustained. Cellular effects of reactive oxygen species may be grouped into five categories: membrane damage, protein modifications, ion deregulation, DNA strand breakage, and effects on energy homeostasis. Interactions between effects in each category make these simplistic classifications artificial but do not limit their usefulness. The effects of lipid peroxidation on membrane integrity and the formation of cytotoxic by-products, such as aldehydes, have been documented (Benedetti et al., 1984; Demopoulos et al., 1979; Masaki et al., 1989). Despite a large body of evidence implicating lipid peroxidation in pathology, using indirect indices such as lactate dehydrogenase release or malondialdehyde formation, in most cases the demonstration that such changes are necessary and sufficient to elicit cytotoxic effects is lacking. Protein inactivation by thiol oxidation, cross-linking, and other modifications may rapidly begin to disrupt the normal functions of other systems, such as glycolysis due to effects on glyceraldehyde-3-phosphate dehydrogenase, or ion regulation due to effects on Na§247 (Fligiel et al., 1984; Kim et al., 1985; Girotti et al., 1986; Wolff and Dean, 1986; Hyslop et al., 1988; Richards et al., 1988; Kyle et al., 1989). Protein-phospholipid cross-linking may also impair cell function as a consequence of oxidative damage (Nielsen, 1981). Cellular ATP losses during oxidative stress may further compromise Na§247 activity and ion regulation, resulting in cellular swelling (Maridonneau et al., 1983). Elevation of intracellular free calcium, derived both extracellularly and from intracellular sources such as mitochondria, occurs as a consequence of membrane disruption and ion deregulation and activates enzymes such as phospholipases and endonucleases (Mallis and Bonventre, 1986; Hyslop et al., 1986; Cantoni et al., 1989a,b). Hydroxyl radicals formed from H202 in the Fenton reaction efficiently generate singlestrand breaks in DNA (Imlay and Linn, 1988; Olson, 1988). DNA damage activates the nuclear enzyme poly(ADP-ribose) polymerase, which catalyzes the ADP-ribosylation of chromatin proteins at the expense ofNAD § (Berger,

[1]

N E U R O T R O P H I N EFFECTS IN OXIDANT INJURY

1985; Berger et al., 1986; Schrauffstatter et al., 1986a,b; Olson, 1988). ADPribosylation of chromatin proteins such as histone H1 counteracts histone inhibition of DNA repair processes, such as DNA ligase activity (Durkacz et al., 1980; James and Lehmann, 1982; Morgan and Cleaver, 1983). One hypothesis of cell death following oxidative stress is that irreversible depletion of NAD + and hence ATP generation via oxidative phosphorylation occurs as a consequence of ADP-ribosylation in response to DNA damage (Spragg et al., 1985; Cantoni et al., 1986; Carson et al., 1986; Gille et al., 1989; Junod et al., 1989; Varani et al., 1990). Depletion of NAD + may result in decreased glycolytic flux, further perturbing energy homeostasis (Berger et al., 1986). Thus, the ability of cells to recover from depletion of pyridine nucleotides may be crucial to the final outcome of oxidative injury. Inhibition of poly(ADP-ribose) polymerase has been demonstrated to confer protection against chemically induced oxidative stress and against traumatic injury in hippocampal slices (Wallis et al., 1993). Another link between oxidative stress and the activation of poly(ADP-ribose) polymerase is the observation of nitric oxide-induced activation of the enzyme (Zhang et al., 1994). I n V i t r o P a r a d i g m s for S t u d y o f O x i d a n t I n j u r y

Various systems are available for study of neurotoxicity in vitro. Primary dispersed cultures of neurons, astrocytes, or oligodendrocytes provide a means of assessing effects of agents on a defined cell type. The use of transformed cells lines that exhibit neural properties has the advantage that large quantities of cells may be grown, permitting detailed biochemical analysis. Organotypic explants (Hauser and Stiene-Martin, 1991; Lyman et al., 1991, 1992; Newell et al., 1993), reaggregate cultures (Jones et al., 1993), synaptosomes (Pifl et al., 1993), and slice preparations (Schurr et al., 1993; Wallis et al., 1993) all provide ways of assessing neurotoxicity. Each of these techniques possesses its own inherent advantages and disadvantages (for review see Atterwill et al., 1992). Cell culture approaches to the study of injury have the advantage that the responses of isolated cell types can be analyzed in detail. The cell culture milieu may be precisely controlled and manipulated to study the effects of individual elements. The caveat that information gleaned from in vitro experiments with single cell types provides only a partial explanation for complex physiological processes does not negate the utility of such approaches. As indicated in Table I, a number of methods are available for generating oxidant injury experimentally. These include hypoxia, either chemically induced or generated by incubation in a low-oxygen atmosphere followed by return to normal oxygen content (Cazevieille et al., 1992; Zhang and

8

PARADIGMS OF NEURAL INJURY TABLE I

Methods for Inducing Oxidant Stress a

Hypoxia Rose bengal Glucose/glucose oxidase Xanthine/xanthine oxidase H202 Sodium nitroprusside/nitroglycerin/S-nitrosocysteine 6-Hydroxydopamine MPTP Amyloid fl-protein a For references, see text.

Piantadosi, 1992; Akaneya et al., 1994). Both the glucose/glucose oxidase and the xanthine/xanthine oxidase systems are capable of generating the superoxide anion, although singlet oxygen may also be a source of oxidative stress when these systems are employed (Fridovich and Handler, 1962; Griot et al., 1990; Pan and Perez-Polo, 1993; Hiraishi et al., 1994; Naveilhan et al., 1994). Rose bengal is another source of singlet oxygen (Fridovich, 1989; Van Reempts et al., 1993). Hydrogen peroxide can be added directly to culture medium, resulting in the production of the hydroxyl radical via the Fenton reaction (Jackson et al., 1990, 1992, 1994; Naveilhan et al., 1994). NO can be added to cultures directly or generated using nitroprusside or nitroglycerin (Chen et al., 1991; Lustig et al., 1992). Reactive oxygen species may also be generated by 6-hydroxydopamine, the toxicity of which depends in part on the generation of H 2 0 2 (Heikkila and Cohen, 1972; Cohen and Heikkila, 1974; Tiffany-Castiglioni and Perez-Polo, 1980, 1981; Spina et al., 1992; Cerutti et al., 1993). Another source of reactive oxygen species is 1methyl-4-phenyl-l,2,3,6-tetrahydropyridine (MPTP). The toxicity of MPP +, the biotoxic metabolite of MPTP, has been suggested to depend in part on the hydroxyl radical (Johannessen, 1991; Chiueh et al., 1992; Cleeter et al., 1992; Hartikka et al., 1992; Cerutti et al., 1993; Pifl et al., 1993; Hartley et al., 1994). Amyloid fl-protein toxicity has been suggested to depend on the generation of H 2 0 2 (Behl et al., 1994~. Studies of cell survival require reliable, reproducible techniques to assess cell viability. Table II lists a number of methods available for quantitating cell survival. A number of approaches have been taken to this end, including techniques measuring properties such as membrane integrity, protein or nucleic acid synthesis, or colony-forming ability. Each of these measures is perturbed by cell injury, but no single one is an entirely satisfactory index. Tests of membrane integrity, such as trypan blue exclusion, have long been

[1]

NEUROTROPHIN EFFECTS IN OXIDANT INJURY TABLE II

M e t h o d s of Assessing Cell Viability a

Radiolabeled precursor incorporation MTT reduction Trypan blue exclusion Rhodamine 123 fluorescence Propidium iodide fluorescence Morphology Acridine orange fluorescence 5~Cr release Cytosolic enzyme release Clonogenic survival a For references, see text.

accepted as measures of viability (Pappenheimer, 1917; Eaton et al., 1959; Tennant, 1964). Leakage of intracellular enzymes has been widely exploited both in vivo and in vitro (Smith et al., 1987; Loeb et al., 1988; Martin et al., 1988; Olson, 1988). Measurement of the release of cytoplasmic enzymes, such as lactate dehydrogenase (LDH) or adenylate kinase, is an effective index at late stages of cell injury but a poor measure of early homeostatic changes (Martin et al., 1988; Jackson et al., 1992; Cazevieille et al., 1993; Behl et al., 1994). Release of 51Cr may also be used as an index of cell viability (Sarafian and Verity, 1990). Dye-exclusion measurements fail to provide a sensitive index of cell viability because each cell must be classified as either alive or dead with no intermediate stages. Indices that rely on metabolic activities, such as tetrazolium salt reduction and incorporation of radioactive precursors, provide more sensitive measures of early perturbations in cell viability. Reduction of 3-(4,5-dimethylthiazol2-yl)-2,5-diphenyl-2H-tetrazolium bromide (MTT) was originally employed in mitogen studies of lymphoid cells, but has found application elsewhere (Mosmman, 1983; Hansen et al., 1989). This assay depends on mitochondrial activity, specifically succinate dehydrogenase, to reduce a tetrazolium salt to a colored product (Slater et al., 1963). Although the assay is highly sensitive, it may be inappropriate for measuring responses to injury when cells display impaired metabolic activity but nevertheless remain viable. Measurement of radioactive amino acid, uridine, or thymidine incorporation affords a sensitive index of activity in some cell types, but is useless in phases of the cell cycle when viable cells fail to incorporate precursors. Clonogenic survival may be particularly problematic in some cell types, such as neurons, that normally do not proliferate once they have reached maturity (Puck and Marcus, 1955; Tiffany-Castiglioni and Perez-Polo, 1980; Anderson, 1994;

10

PARADIGMS OF NEURAL INJURY

Van der Maazen et al., 1992). Fluorescence assays using agents such as rhodamine, propidium iodide, and acridine orange are convenient and may be applied to flow cytometry (Detta and Hitchcock, 1990; Gagliardinini et al., 1994; Deckwerth and Johnson, 1993; Ziv et al., 1994; Favit et al., 1992; Detta and Hitchcock, 1990; Darzynkiewicz et al., 1992; Sauer et al., 1992). Metabolic indices such as energy charge, which measures high-energy phosphate content (Atkinson, 1968), have been considered among the most definitive measures of metabolic capability. Nevertheless, even this technique may not be appropriate in all circumstances.

E f f e c t s o f N G F o n O x i d a n t I n j u r y in P C 1 2 Cells Our laboratory has used the PC12 rat pheochromocytoma cell line as a model for elucidating the effects of NGF on oxidant injury. Figure 2 shows the dose-response to HzO2 in control and NGF-treated cultures. The minimal concentration of NGF tested that was capable of enhancing survival above the control level was 1 ng/ml, a level effective only against 0.1 mM H202. Higher levels of NGF were effective against a wider range of H202 concentrations. The effects of NGF on cell viability under serum-free conditions were measured using protein synthesis and MTT reduction (Fig. 3). Under these conditions, the minimal dose of NGF tested that was capable of enhancing PC12 cell viability above the basal level was 1 ng/ml. Higher doses (10-100 ng/ml) had a more pronounced cell-sparing effect. The dose-response relationship between NGF concentration and survival in serum-free medium resembled that for survival following H202 treatment (Fig. 2). The time course for cell death following 0.5 mM H202 treatment is shown in Fig. 4, using dye exclusion, [35S]methionine incorporation, MTT reduction, and LDH release as viability assays. Decreases in all indices were observed following 0.5 mM H202 injury at the earliest time points measured, i.e., 2 hours postinjury for dye exclusion and LDH release and 4 hours for [35S]methionine incorporation and MTT reduction. NGF enhancement of survival was evident at the earliest time points obtained for dye exclusion and amino acid incorporation, but not for the other two indices. Enhanced survival of NGF-treated cells was not evident until 8 hours postinjury for MTT reduction and 4 hours for LDH release. No evidence for substantial recovery from initial viability losses was apparent except in the MTT reduction assay. This apparent recuperation from initial damage was most pronounced in NGFtreated samples. LDH release, in contrast, continued to increase such that the maximal value attained was at 24 hours. NGF-treated samples continued

[1]

NEUROTROPHIN EFFECTS IN OXIDANT INJURY

11

to release LDH after 4 hours, although these continued losses were less marked than those observed in control cultures. Previous studies of NGF effects on HzO 2 toxicity in PC12 cells and the SY5Y human neuroblastoma cell line used chronic NGF treatment regimens (6-11 days) before performing cytotoxicity tests (Tiffany-Castiglioni and Perez-Polo, 1981; Perez-Polo et al., 1986; Jackson et al., 1990). The data presented here indicate that NGF induction of a H202 refractory state does not require extended treatment. NGF treatment for 24 hours elicited a significant cytoprotective effect against oxidant injury. This effect was apparent prior to withdrawal of PC12 cells from the mitotic cycle, an effect requiring more prolonged NGF treatment (Greene and Tischler, 1976; Greene, 1978). Thus, cell cycle dependency of H202 injury, an important determinant of cellular ability to repair single-strand DNA breaks (Frankenberg-Schwager, 1990; Kleiman et al., 1990), does not appear to bear on enhanced survival of cells treated with NGF for 24 hours prior to injury. The minimal NGF concentration tested was capable of increasing PC12 cell survival above the control level was 1 ng/ml, although this NGF level was less effective than higher concentrations against high H202 levels. Similar doses of NGF were required for enhancement of survival following oxidant injury and serum deprivation. The ECs0 for NGF (i.e., the concentration necessary for a half-maximal effect at any given HzO 2 concentration) was estimated to be between 1 and 10 ng/ml. This dose-response relationship is similar to that previously described (Greene, 1978) and bears physiologic relevance in that the minimal effective dose tested (1 ng/ml or 3.7 nM) was similar to the Kd of the low-affinity NGF receptor (Sutter et al., 1979). Thus it appears from the dose-response characteristics that NGF induction of an HzOz-refractory state is a physiological rather than a pharmacological effect. Losses of viability following 0.5 mM HzO 2 treatment occurred at the earliest time points measured, regardless of the assay employed. Trypan blue exclusion decreased more rapidly in these studies than in those reported following a higher HzO 2 concentration (2.5 mM) in macrophage-like cells (Schrauffstatter et al., 1986b). The more rapid decreases in viability seen in PC12 cells were likely to reflect an enhanced susceptibility of neurally derived cell types to oxidant injury as compared to other cell types (Tiffany-Castiglioni et al., 1982). 6-Hydroxydopamine, a neurotoxin that generates H202 (Heikkila and Cohen 1972; Cohen and Heikkila, 1974), studied in the SY5Y human neuroblastoma line, revealed that dye exclusion was among the last viability index tested to be impaired (Tiffany-Castiglioni and Perez-Polo, 1980; TiffanyCastiglioni et al., 1982). The delayed changes in trypan blue exclusion reported in early studies may be a consequence of the prolonged generation of oxygen radicals by 6-hydroxydopamine as compared to delivery of a preformed HzO 2 concentration described here.

12

P A R A D I G M S OF N E U R A L I N J U R Y

110

o~ "-" z O_

~ 0 ~. nO o z__ bw "-~ U3

%.

100 90

' •

80

70

\"'-i

60

"*--*

%

50

.....

.....

~_ -e..

40 30

•

20 10 Illl

II, OH

,

i

~,. .

"i

-eZ~

-.

------_v.~" "e-

i i illl

i

i

i

i

0.10

i ilia

"k~

i

1.0

i

i

i i|

10.0

H20 2 (mM)

FIG. 2

Thus, the effect of NGF o n H 2 0 2 resistance was rapid and dose dependent. A direct comparison of four different viability assays demonstrated NGF protection from H 2 0 2 cytotoxicity. These findings refute the contention that NGF enhancement of cell viability as assessed by radiolabeled amino acid incorporation is an artifact secondary to NGF enhancement of protein synthesis. Homeostatic perturbation by oxidant injury has widespread repercussions on cellular function. The outcome of oxidant injury depends on a balance between the severity of the insult sustained and the capacity of cells to withstand and recover from such injury. Mechanisms of cellular recovery from the toxicity of reactive oxygen species may be as critical to the outcome of injury as are mechanisms for preventing such injury, such as antioxidant capabilities. The generation of single-strand DNA breaks in cultured cells during oxidant injury sets in motion a number of processes that may culminate in cell death (Olson, 1988; Schrauffstatter et al., 1986a,b). The activation of nuclear poly(ADP-ribose) polymerase (PADPRP) following DNA damage results in rapid depletion of NAD + (Sims et al., 1983; Berger, 1985; Schrauffstatter et al., 1985, 1986a; Berger et al., 1986; Junod et al., 1989). If extensive DNA damage causes irreversible NAD + depletion, cell death may ensue from inhibition of energy metabolism via effects on glycolysis and oxidative phosphorylation (Spragg et al., 1985; Berger et al., 1986; Carson et al., 1986; Schrauffstatter et al., 1986b). Inhibition of PADPRP using inhibitors such as 3-aminobenzamide (AB) has been reported to prevent NAD + depletion and cell death following oxidant exposure (Sims et al., 1983; Schrauffstatter et

13

[1] NEUROTROPHIN EFFECTS IN OXIDANT INJURY

FIG. 2 D o s e - r e s p o n s e effect of H20 2 on viability of PC12 cells treated with varying doses of N G F , using [35S]methionine incorporation as an index. Studies of N G F effects on H20 2 toxicity were performed as described previously (Jackson et al., 1990, 1992). PC12 cells (7.5 • 104) were plated in poly(D-lysine)-coated 6-mm-diameter wells and allowed to attach overnight in complete m e d i u m before addition of N G F . After 24 hours, these media were r e m o v e d and replaced with varying dilutions of H20 2 in RPMI 1640 medium. After 30 minutes o f H 2 0 2 t r e a t m e n t at room t e m p e r a t u r e , these solutions were r e m o v e d and replaced with complete medium. Following a 24hour r e c o v e r y period, viability was assessed. Incorporation of 35S]methionine was performed as previously described (Jackson et al., 1990, 1992). Each point represents the mean _+ S E M for quadruplicate samples. T , 0; O, 1; , , 10. II, 100 ng N G F / m l . Statistical analysis was carried out using A N O V A followed by F i s h e r ' s L S D test, comparing the values b e t w e e n N G F treatment groups at each H20 2 concentration. df, 29; F, 66.52. Statistics are summarized in the tabulations below, which give the most significant a level obtained for each comparison (NS, not significant, i.e., p -> 0.05) (note: at 5 m M H202, all values are NS):

NGF (ng/ml) NGF (ng/ml)

0

0 0.1 1.0 10

--

0.1

1.0

mM H202 NS NS -NS --

10

100

0.001 0.001 0.005

0.001 0.001 0.001 0.001

0.5

--

100 0.10

0 0.1 1.0 10

--

NS --

mM H20 z 0.01 NS ~

0.001 0.001 0.05 --

0.001 0.001 0.005 NS

100

mM HzO2 NS NS -NS

0.5

0 0.1

--

1.0

--

10

0.001 0.001

0.001 0.001

0.001

0.001

--

NS

100

0 0.1 1.0 10 100

--

1.0 mM HzOz NS NS -NS --

0.001 0.001 0.005 --

0.001 0.001 0.001 NS

14

PARADIGMS OF N E U R A L I N J U R Y 0.70 E tO

tO

Z

0.60

/

0.50

LU tO Z < rn n" O

0.20




25

~

s~-

~

9176

_.1

m < >

25

i

4

-_~---'F

8

12

l

16

i

20

TIME FOLLOWING H20 2 (HR)

24

0

0

a.

4

l

8

=

12

l

I6

20

24

TIME FOLLOWING H20 2 (HFI)

FIG. 4 Time course for cell death following 0.5 mM H20 2 treatment in control and NGF-pretreated cells as assessed by dye exclusion (A), [35S]methionine incorporation (B), MTT reduction (C), and LDH release (D). Experimental conditions were as described in the legend to Fig. 2. Viability assays were performed as described (Jackson et al., 1992). All points are mean _+ SEM for quadruplicate wells. For trypan blue exclusion, data were expressed as percentage of cells excluding dye. Values for NGF-treated samples (T) differed significantly from controls (0) at 2, 4, 8, and 24 hours postinjury (p < 0.05, 0.05, 0.005, and 0.05, respectively). For [35S]methionine incorporation, values were normalized to the initial value (100% for either control or NGF-treated wells). NGF-treated samples differed significantly from controls at 4, 8, and 24 hours postinjury (p < 0.001, 0.001, and 0.01, respectively). For MTT reduction, values were normalized to the initial value (100% for either control or NGF-treated wells). NGF-treated values differed from controls at 8 and 24 hours (p < 0.001). Values for LDH were expressed as a percentage of the total activity released into the medium at each time point. NGF-treated samples differed from controls at 0, 4, and 24 hours (p < 0.01, 0.005, and 0.001, respectively; Student's t-test). From Jackson, G. R., Werrbach-Perez, K., Ezell, E. L., Post, J. F. M., and Perez-Polo, J. R., Brain Res. 592, 239 (1992), with permission. 15

16

PARADIGMS OF NEURAL INJURY

required culture for longer times and with higher concentrations than those necessary to decrease HzO2 sensitivity (Jackson et al., 1990, 1994; Pan and Perez-Polo, 1993; Sampath et al., 1994. These efforts to elucidate effects of NGF on the degradation of activated oxygen species suggested that investigation of an aspect of oxidant injury other than the defense against initial injury might prove worthwhile, i.e., the recovery phase. NGF treatment rapidly decreases the content of PADPRP and its mRNA in PC12 cells (Taniguchi et al., 1988). Rapid (within 15 minutes) depletion of NAD + was observed following 0.5 mM H202 treatment in both control and NGF-pretreated cells (Fig. 5). By 1 hour following HzO2 treatment, intracellular NAD + concentrations had decreased to less than 20% of initial values. No evidence for NGF effects on initial depletion of NAD + was obtained; on the contrary, the mean NAD + concentration for NGF-treated cells (as a percentage of the NGF-treated control) was actually lower than the corresponding control value. At later time points, however, NGF-treated cells began to evidence a more extensive recovery from this initial depletion. Although there was an apparent enhancement of recovery from NAD + depletion in NGF-treated cells as compared to controls by 2 hours following injury, this increase did not attain statistical significance until 4 hours following H202 treatment. In control cells there was some evidence for enhanced recovery of NAD + from the minimum levels observed at 4 hours following injury, but the recovery was delayed and less complete than that observed in NGF-pretreated cells. By 24 hours following oxidant treatment, the NAD + concentration in NGF-treated cells had recovered to 80% of the initial value; in control cells, by contrast, the NAD + concentration was less than 50% of the initial value by this time. Analysis of LDH release confirmed that the more complete recovery of NAD + noted in NGF-treated as compared to control cells was matched by an enhanced viability, as indexed by greater enzyme retention (Jackson et al., 1992). These data indicated that NGF pretreatment did not prevent NAD + depletion occurring as a consequence of PADPRP activation; rather, the effect of the growth factor was to enhance recovery from initial NAD + depletion. Further experiments used a competitive inhibitor of PADPRP, AB, to examine further the role of this enzyme in H202 cytotoxicity. AB pretreatment results in a dose-dependent increase in cell survival following injury with H202 (Jackson et al., 1992). The effect of a single concentration of AB in control and NGF-pretreated cells, as assessed 24 hours following H202, is illustrated in Fig. 6. The protective effect of NGF in the absence of AB was qualitatively similar to that observed in experiments already described. In this particular experiment, incubation with AB had a protective effect on control PC12 cells only at 0.1 mM H202. The combination of AB with NGF

[1] NEUROTROPHIN EFFECTS IN OXIDANT INJURY

17

12o[

~. ~_ 0

110~ 100~ 90 80

60

//.L

50

,"

.~

40

z

30

" ~'

20 ~-~y,,~ 0

0

o..... - ~ ' ~ i 4

i 8

i 12

I 16

i 20

24

TIME FOLLOWING H20 2 (HR)

FIG. 5 Effect of treatment with 0.5 mM H202 on intracellular NAD + in control (0) and NGF-treated (!t) PC12 cells. Analysis of NAD + depletion was performed as described previously (Jackson et al., 1992). Briefly, cells (106) were plated on poly(Dlysine)-coated 60-mm-diameter dishes and pretreated 24 hours with NGF. NAD + concentrations were determined in perchloric acid extracts of cells harvested at various times following H202 treatment. Points are the mean + SEM for triplicate determinations. Values for NGF-treated cultures were significantly different from those for controls at 4, 8, and 24 hours following H202 treatment (p < 0.001, 0.001, and 0.05, respectively); the value for NGF-treated cells was significantly different from the control at 1 hour (p < 0.05, Student' s t-test). From Jackson, G. R., WerrbachPerez, K., Ezell, E. L., Post, J. F. M., and Perez-Polo, J. R., Brain Res. 592, 239 (1992), with permission.

pretreatment had a dramatic effect on survival. NGF-pretreated cells incubated with AB displayed an enhanced survival as compared to cells without AB at H202 concentrations between 0.1 and 1.0 mM. N G F treatment alone was insufficient to enhance survival above the control level at H202 concentrations above 0.1 mM. In cells treated with both N G F and AB, by contrast, robust tetrazolium salt reduction was observed at H202 levels as high as 1 mM. Thus, inhibition of PADPRP had an effect in NGF-treated cells that was more pronounced than that observed in undifferentiated cells. Consistent with studies of oxidant injury in other cell lines (Cantoni et al., 1989a; Gille et al., 1989; Junod et al., 1989; Schrauffstatter et al., 1986a,b), rapid depletion of N A D + was observed following 0.5 mM H202 treatment. Because N G F treatment diminished the PADPRP content of PC12 cells within the same time frame used in these experiments (Taniguchi et al.,

18

PARADIGMS OF N E U R A L INJURY 0.55 T 0.50 - ~I~, ~,

0.45

i,,.

0.40

t.u

0.30

z
Costar (Cambridge, MA) > Corning (Corning, NY) > Falcon (Lincoln Park, NJ). Primary cultures can also be grown in 35-mm dishes.

[2]

29

N I T R I C O X I D E T O X I C I T Y IN CNS C U L T U R E S

TABLE I

Stock Solutions Reagent

Eagle's minimal essential medium (MEM) with Earle's salts, without glutamine Fetal bovine serum ( F B S ) d heat inactivated at 56~ for 30 minutes then sterile filtered 5- Fluoro- 2'-deoxyuridine (5F2DU) b is highly toxic; wear gloves when handling Glucose b Glutamine a HEPES b Horse serum (HS), a heat inactivated at 56~ for 30 minutes then sterile filtered Polyornithine b Trypsin a

Source

Concentration

Storage

G I B C O - B R L (Gaithersburg, MD)

1x

500 ml/4~

GIBCO-BRL

1x

27 m l / - 2 0 ~

10 m M

10 m l / - 2 0 ~

Sigma (St. Louis, MO)

J. T. Baker (Phillipsburg, NJ) GIBCO-BRL Sigma GIBCO-BRL

Sigma GIBCO-BRL

2 M 200 m M 1M 1•

0.3 mg/ml 10•

100 ml/4~ 5.5 or 6.5 m l / - 2 0 ~ 100 ml/4~ 27 m l / - 2 0 ~

10 m l / - 2 0 ~ 1 ml/-20~

a Avoid repeated freeze/thaw cycles with these reagents. For best performance, aliquot once into sterile test tubes the volume needed for each preparation and thaw only the necessary aliquots. b Stock solutions made with autoclaved Milli-Q water and sterile filtered (0.2 txm) prior to aliquoting and freezing.

When neuronal cultures are grown in 96-well plates there is a nonuniform distribution of neuronal cell types in each well due to the small number of neurons plated in each well. For instance, the number of NOS neurons varies greatly in each well and some wells will have no NOS neurons. Thus, if the experiment is designed to examine activation of NOS there will be considerable variability among wells. Prior to plating the culture well with neurons, the well must be precoated with an attachment matrix. Stock polyornithine (0.3 mg/ml) (Table I) is diluted into autoclaved Milli-Q water at a l : 1 0 0 dilution (1 ml polyornithine/99 ml H 2 0 ). This solution is filtered through a 0.2-ixm filter to remove any crystals that may have formed, because the crystals are toxic to neurons. Using sterile technique, add 1 ml of polyornithine solution to each well, cover, and incubate in an oven for 60-90 minutes at 37-42~ Cultured neurons will lie flat and will not aggregate (clump) on a well-coated plate. Our optimal conditions are 42~ for 60 min-

30

PARADIGMS OF N E U R A L INJURY

TABLE II Culture Media Formulations a Reagents 10: 10:1 M E M MEM FBS HS Glutamine 5 : 1/5F2DU M E M MEM HS Glutamine 5F2DU 5:1 MEM MEM HS Glutamine M E M + 21 m M glucose MEM 2 M Glucose

V o l u m e (ml)

500 63.3 63.3 6.3 500 26.6 5.3 1.45 500 26.6 5.3 500 5.25

a Media containing glutamine should be used within 4 weeks of being made. Glutamine converts to glutamate with time in aqueous solution and may alter the cell culture conditions. If media cannot be used within 4 weeks, make a smaller volume.

utes. After incubation, aspirate off the polyornithine solution with a sterile Pasteur pipette. Gently rinse the well one time with autoclaved Milli-Q water and aspirate off the water. Apply water by placing the pipette on the side of the well and slowly releasing the water so as not to disturb the layer of polyornithine. Let the plates dry overnight in a clean hood. Store in a dustfree environment. For optimal results use the plates within the first week.

Culture Procedure At the beginning of the dissection, warm 10" 10" 1 Eagle's minimal essential medium (MEM) solution and the 1 x trypsin solution (Table II) in a 37~ H20 bath. Add 10-12 ml sterile Brooks-Logan solution (Table III) into a 15-ml test tube (one for each brain region to be dissected) and 15-25 ml of Brooks-Logan solution into several sterile 100-mm round tissue culture plates. Place dissecting tools into 70% ethanol. For cortex or caudate-puta-

[2]

31

N I T R I C O X I D E T O X I C I T Y IN CNS C U L T U R E S TABLE III Reagent Sucrose o-Glucose NaCl KC1 Na2HPO 4 KHzPO 4 1 M HEPES

Brooks-Logan

Solution a Concentration (mM) 44 25 137 2.7 10 1.8 10

a Dissolve all reagents in autoclaved Milli-Q water. Adjust pH to 7.4. Sterile filter through a 0.2-~m filter. This solution can be kept for several weeks at 4~ but should be sterile filtered before each use to prevent contamination of cultures by fungus or mold. The solution should be discarded if there are any visible particles or growth.

men cultures, decapitate a 14-day-old pregnant rat, or a 16-day-old pregnant rat for hippocampal or thalamic cultures. Place the rat supine, spray the abdomen with 70% ethanol, and expose the uterus by making an incision through the abdomen with a pair of scissors. Using forceps and a pair of scissors, remove the uterus and place in a sterile 100-mm culture plate. Make a longitudinal incision along the uterus, remove the fetuses, followed by removal of the heads to a sterile 100-mm culture plate containing 15-25 ml of sterile Brooks-Logan solution (enough to cover the heads completely). One at a time, remove a head to a different sterile 100-mm culture plate cantaining 10-15 ml Brooks-Logan solution. Utilizing a dissection microscope to assist with visualization, remove the brain from the cranial cavity by making a continuous longitudinal incision along the dorsal and ventral surface of the cranium. Take care to avoid cutting the underlying brain tissue. Remove the brain from the cranial cavity and remove the olfactory bulbs, cerebellum, midbrain, and spinal cord. Remove as much of the meninges as possible. Dissect out the brain region of interest (i.e., cortex, hippocampus, caudate-putamen, or thalamus) and put the dissected pieces of brain tissue into a 15-ml sterile conical test tube containing 10-12 ml of Brooks-Logan solution. Pool all the dissected tissue from one brain region from one pregnant rat in one test tube. Repeat for each fetal brain. After all the tissue has been dissected, draw off the Brooks-Logan solution from the 15-ml conical test tube with a sterile pipette. Be careful to avoid aspirating off the dissected tissue. Replace with 1.5-3 ml of a 1 x trypsin solution and place in a 37~ incubator for 20-25 min. Trypsin facilitates the dissociation of cells by digesting the protein contacts between cells; 37~ is the optimal temperature for

32

PARADIGMS OF NEURAL INJURY

trypsin activity. Do not overtreat the tissue, because trypsin will digest cell walls with time. The 1 x trypsin solution is made by adding 9 ml of Brooks-Logan solution to 1 ml of a 10x trypsin stock. After the trypsin digest, aspirate off as much of the trypsin solution as possible with a sterile pipette. Because trypsin-treated tissue is sticky, care must be used not to remove any tissue. This is followed by adding 5-7 ml of warm 10" 10" 1 MEM. (The serum in the MEM solution will inactivate the remaining trypsin.) Using sterile 9-inch Pasteur pipettes, with mouth openings of decreasing area (made by flaming the tips), triturate the cells and disperse them by gently moving the cells in and out of the pipette. The tissue should immediately form a cloudy suspension. Do not triturate more than 10 times. Let the tissue settle for a few minutes. Undissociated tissue will settle to the bottom of the test tube. Pipette 10/zl of cell suspension with a sterile pipette tip and eject between the coverslip and hemocytometer for cell counting and observation. There should not be any cell clumps, only individual cells. Count the number of cells in several squares and calculate the mean number of cells per square. Calculate the volume of media necessary to plate the cells at a particular density by the following equation: (mean number of cells per square x 16 • 10,000 x milliliters of dissociated cells) divided by desired density equals final volume (in milliliters). To dilute the dissociated cells to the desired density for plating, remove by pipette the suspended dissociated cells, leaving the bulk tissue in the test tube. Dilute the suspended cells in an appropriate volume of 10" 10" 1 MEM. Plate 1 ml of the diluted cells per well of a 24-well plate. Place in a 37~ 7% (v/v) CO2 humidified incubator. After 4-5 days change the media to 5 : 1/ 5F2DU MEM (Table II) to inhibit nonneuronal cell growth. The culture media are changed to 5"1 MEM 3-4 days later. The media are changed twice weekly by aspirating off one-half of the volume and replacing it with fresh 5:1 MEM (Table II). The cell cultures are maintained for 21 days.

NADPH-Diaphorase Stain for NOS in Primary Neuronal Cultures NOS is fully expressed by day 20 in culture and it is important to determine that the cultures are mature and express a full complement of NOS neurons. To determine the percentage of NOS neurons in the total cell population, stain the cultures for NADPH-diaphorase. In the CNS, NOS has been shown to account for the NADPH-diaphorase stain under conditions of paraformaldehyde fixation (3). To stain, fix the cell cultures in 4% freshly depolymerized paraformaldehyde/0.1 M phosphate-buffered solution (w/v) for 30 minutes at room temperature. Wash fixative off cells with control salt solution (CSS,

[2]

33

N I T R I C O X I D E T O X I C I T Y IN CNS C U L T U R E S

TABLE IV Control Salt Solution a Reagent

Concentration (mM)

NaC1 KC1 CaC12 D-Glucose Tris-HC1 MgCI2 b

120 5.4 1.8 15 25 0.8

a Dissolve all reagents in Milli-Q water and adjust pH to 7.4 at the temperature the experiment is to be performed (the pH of Tris is temperature sensitive). Sterile filter the solution through a 0.2-/~m filter and store at room temperature until use. All solutions should be sterile filtered before use and discarded if there is visible growth. b Except for NMDA experiments.

Table IV). Apply NADPH-diaphorase stain mix (see Appendix at end of chapter) to the cell cultures, approximately 500/xl per well. Float the dish in a 37~ water bath for 30 minutes. Check the development of the stain. The stain is complete when the neurons and projections are a deep purple/ black. If the glia begin to stain blue the cultures have been overstained. If the stain is not complete, return the plate to the water bath. Typically it takes between 30 and 90 minutes for the stain to develop in tissue culture. To stop the reaction wash the stain off with CSS. The cultures can be kept for a few days at 4~ if 0.05% sodium azide is added to the CSS. Greater than 100 NOS neurons are required per well of a 24-well plate to observe an NO component of glutamate neurotoxicity.

Toxicity Experiments

Exposure'Day 1 We routinely use cultures that are 21 days old because NOS is not fully expressed until day 20 of culture. Prepare exposure solutions that contain the pharmacologic agent of interest in sterile CSS (Table IV). Prior to exposing the neurons to excitatory amino acids (EAA) (see Appendix at end of chapter), the MEM must be removed because it contains salts and metals that may alter the outcome of toxicity experiments. Aspirate off growth media and replace with 1-1.5 ml CSS. Wash the wells with 1-1.5 ml CSS

34

PARADIGMS OF N E U R A L INJURY

2x to completely remove the MEM. Aspirate off the last rinse of CSS and replace with the exposure solution (0.5-1 ml). NO seems to play a major role in delayed neurotoxicity--a 5-minute exposure to glutamate or NMDA elicits cell death 24 hours later. Thus, exposure solutions should be applied to the neurons for exactly 5 minutes. Monitor intervals with a timer to deliver exposure solutions to each well. We have found that 15 seconds is the maximal time interval between wells that can safely be achieved without dislodging cells or losing time during the washes. Terminate the exposure by aspirating off the exposure solution and replacing with CSS (1-2 ml) and washing 2x. Replace the CSS with 1 ml MEM containing 21 mM glucose (Table II). Return the tissue culture plate to the incubator for 18-24 hours. Other forms of neurotoxicity require exposure times longer than 5 minutes. For exposure times greater than 30 minutes we use an exposure solution containing MEM + 21 mM glucose and perform the exposures in the incubator. Note: the buffering salt in CSS is Tris-HCl, which is temperature sensitive. Therefore, it is critical that the pH be adjusted to 7.4 at the same temperature at which the experiment will be performed.

Assay for Cell Death: Day 2 There are a variety of methods available to assay for cell death. We have found that the "gold standard" is cell counting and that other methods are not as sensitive for small but statistically significant changes. We routinely stain dead cells with 500 ~1 of 0.4% trypan blue for 20 minutes. Trypan blue is a vital stain that stains dead cells dark blue. Live cells exclude the die and remain phase bright. Very gently remove the trypan blue and wash with CSS until the cells can be visualized under a microscope. Photomicrographs or permanent images of several random fields of cells should be obtained for counting by an observer blinded to study design and treatment protocol. We routinely obtain between three and five images per cell well, representing approximately 20% of the total neuronal population. Viable versus nonviable cells should be counted. We usually perform at least two separate experiments utilizing four separate wells and routinely count a minimum of 4000-12,000 neurons for each data point. To determine whether cells are being "washed" away or are destroyed by either the treatment or staining protocol, photomicrographs should be made before and after treatment using a transparent grid etched with a diamond tip pen on the bottom of each culture plate. In our laboratory no appreciable loss of neurons has been identified when comparing "before" and "after" photomicrographs from cortical cultures. Other assays for cytotoxicity are based on biochemical parameters that

[2]

NITRIC OXIDE TOXICITY IN CNS C U L T U R E S

35

change in response to cell death. These assays work very well in homogeneous cell cultures. However, in mixed primary neuronal cell cultures that contain glia as well as neurons, the specificity of these methods can be clouded by background noise. The lactate dehydrogenase (LDH) assay is widely used for many cytotoxicity paradigms and has been found to be reliable in some cortical culture and retinal culture models (2, 6, 7). However, LDH activity is affected by the oxidative environment and appears to be a molecular target for transition metal-mediated radical attack in some cultured neuronal systems (8; R. Ratan, personal communication), limiting its use in these experimental paradigms. Additionally, a relatively large proportion of LDH activity resides in the small astrocytic population present in primary neuronal cultures. These astrocytes can become "leaky" under ischemic conditions and contribute nonspecifically and disproportionately to the measurement of LDH release (9). Therefore, LDH activity must be carefully evaluated before use as a measure of cell viability when the experimental paradigm is designed to examine certain models of oxidative stress or ischemia. Although the glia comprise less than 20% of the total cell population in primary neuronal culture, they contain more than 60% of the mitochondria. Cytotoxicity assays that measure mitochondrial function, such as rhodamine 123 and 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl-2H-tetrazolium bromide (MTT) dye assays, measure the response of the glia disproportionately to the response of neurons (9). The vital dye, neutral red, accumulates faster in glia than in neurons in primary neuronal cultures, again generating a larger glial than neuronal response. Therefore, these dye assays should be used in homogeneous neuronal cultures that contain minimal numbers of glia, such as cerebellar granule cell cultures or sympathetic ganglion cultures. An additional failing of these assays is the lack of assessment of live versus dead cells. In cultures that have survived for 3 weeks, the cell number can vary among wells. This is not a problem encountered in cultures that survive for only a few days before the experimental paradigm. Another method of cell counting that assesses both live and dead cells relies on fluorescent dyes. These are usually based on ethidium homodimer or fluorescein diacetate staining of dead cells (red) and calcienAM staining live cells (green) (Cytoprobe, Millipore, Bedford, MA; Live/Dead, Molecular Probes, Eugene, OR). These fluorescent vital dyes have the advantage that the incubation media do not need to be changed and they give graphic colorful photomicrographs. Additionally, the stained cultures can be analyzed by a fluorescent plate reader. The disadvantage to these dyes is that color photomicrographs must be made for cell counting, which is potentially cost prohibitive. If a plate reader is used to assess cell death by quantitating the amount of red versus green fluorescence, the center of each well must be uniform because the reader takes measurements from the center of each well.

36

PARADIGMS OF NEURAL INJURY

Criteria f o r N O S Activation A short (5 minutes) application of NMDA or glutamate to primary neuronal cultures clearly elicits neuronal cell death when determined 24 hours later. We and others have shown that NO mediates a component of neuronal cell death in these experimental paradigms (3). A number of criteria must be satisfied to establish that NO mediates cell death. A number ofNOS inhibitors with different mechanisms of action should be used (see Appendix). Potency of inhibition of toxicity should parallel their potencies as NOS inhibitors. The neuroprotection afforded by arginine analog inhibitors should be reversed by excess substrate L-arginine, and o-arginine should have no effect. Additionally, stereoisomers of the NOS inhibitors should be inactive. Depletion of arginine from the culture media should be neuroprotective, and hemoglobin, which complexes NO, should attenuate neurotoxicity. If the toxic insult occurs in part through NO, then increased levels of NO should be present during the exposure of the toxic agent. We routinely use cGMP as an indirect measure of NO formation in cultures (Fig. 2). Additional methods exist for measurement of NO formation, including the Greiss reaction and the NO electrode. The Greiss reaction is based on a colorimetric assay of nitrite (NO2) and nitrate (NO~-), reaction by-products of NO. Although the method is specific, it is usually not sensitive enough in primary neuronal cultures to access NO formation accurately. Commercially available NO electrodes allow the direct measurement of NO. Whether these electrodes are sensitive enough to reflect accurately changes in NO content in cultures is not known. Questions remain as to the specificity of the measurements as well as the calibration of the electrodes. This is an emerging technology and it may become the method of choice once the technical problems are resolved. Regardless of the choice of NO measurement, experiments should be performed to establish the specificity of NO formation. These experiments would be identical to those used in the toxicity experiments and include using a variety ofNOS inhibitors, stereoisomers, arginine-free media, and hemoglobin.

NO Chemistry NO can exist in three valence states: NO-, NO +, and NO .. The predominant species of NO formed in vivo is not known, but is probably dependent on the redox milieu of the local environment around NOS. NO- reacts with molecular oxygen to form NO2 and NO~- in the absence of metal ions. Several NO releasers are available, which preferentially release different valence states of NO. For instance, NO + is released by nitrosocysteine and organic

[2] NITRIC OXIDE TOXICITY IN CNS CULTURES

37

nitrates and reacts with tissue sulfhydryl groups such as those on the NMDA receptor, thus inactivating the NMDA receptor. In this way NO + is neuroprotective by decreasing NMDA receptor activity (10). Sodium nitroprusside (SNP) releases NO + or NO. depending on the redox environment of the experimental system, and 3-morpholinosydnonimine (SIN-l) preferentially releases NO .. The toxic valence state of NO appears to be N O . , which reacts with superoxide anion (O~) at a rate of 6.7 • 109 m -~ sec -~ to form the powerful oxidant ONOO- (11). There are several isoforms of superoxide dismutase (SOD) that convert O~ to hydrogen peroxide ( H 2 0 2 ) , which is then converted to H 2 0 by catalase or glutathione peroxidase. SOD is highly and ubiquitously expressed in all tissues as a first-line defense against oxidative stress. Of all identified enzymes, the copper-zinc isoform of SOD has the fastest rate constant for its substrate, O~ (2 • 109 M -~ sec-~), but the reaction between O~ and N O . is faster. Thus, NO. is the one biological molecule known that can outcompete SOD for O~. NO. is freely diffusible across lipid membranes; with a tl/2 < 1 second, it is a relatively "stable" free radical with a diffusion radius of approximately 40 tzm in tissue. In contrast, O~ has a diffusion radius of 1-3/zm intracellularly and can pass through cell membranes only at ion channels. Extracellularly, O~ has a diffusion radius of 20 /zm. Thus, O~ and N O . are ideally suited to react and form ONOO-. ONOOis somewhat membrane permeable and can diffuse several cell widths. It is estimated that approximately 10% of ONOO- exists in the trans form, which demonstrates hydroxyl-like chemistry and decomposes primarily to nitrate. The remainder of ONOO- exists in the cis form, which reacts with metals to nitrosylate amino acids such as tyrosine (12). Additionally, ONOO- oxidizes sulfhydryl groups, or ONOO- is protonated to peroxynitrous acid (ONOOH) (13), which decomposes to nitrogen dioxide and hydroxyl free radicals. The chemical state and the preferential reactions of NO and ONOO- are directly related to both the oxidative state and the pH of the tissue. Many of these conditions can be regulated in cell culture but clearly become quite complex in in vivo models of stroke and neurodegeneration.

Targets of NO The widespread role for NO in mammalian physiology and pathophysiology has just recently been recognized, and target molecules for NO that may play an active role in the genesis of neurotoxicity are still being discovered. In addition to reacting with O~ to form ONOO-, NO can activate hemecontaining enzymes, such as guanylate cyclase, through displacement of the heine moiety. NO can also induce the ADP-ribosylation and modification of several intracellular proteins, such as glyceraldehyde-3-phosphate dehydro-

38

PARADIGMS OF NEURAL INJURY

genase. Enzymes with iron-sulfur centers in both the respiratory cycle and in the pathway for DNA synthesis are also targets for NO. Target enzymes in this class so far identified include aconitase (aconitate hydratase; part of the Krebs cycle), reduced nicotinamide adenine dinucleotide phosphate" ubiquinone oxidoreductase [mitochondrial complex, NADH dehydrogenase (ubiquinone)], succinate:ubiquinone oxidoreductase [mitochondrial complex II, succinate dehydrogenase (ubiquinone)], and ribonucleotide reductase (a rate-limiting enzyme in DNA synthesis) (14, 15). Modulation of the function of any of these enzymes could obviously have a dramatic consequence to the target cells and possibly result in neurotoxicity. NO can cause deamination of DNA (16), which could result in sufficient DNA damage to be directly cytotoxic to the neurons. DNA damage resulting from deamination is different from DNA damage characterizing apoptotic, or "programmed," cell death in which DNA is degraded by endonucleases into specific fragments, resulting in a DNA ladder. One potential pathway leading toward neurotoxicity could be activation of poly(ADP-ribose) synthase in response to DNA damage. PARS has been extensively studied in eukaryotic cells such as thymocytes and in many secondary cell lines. PARS is a chromatin-bound enzyme that cleaves NAD and then transfers the ADP ribose moieties to various nuclear proteins, including histones, topoisomerase, DNA ligase II, and PARS itself. PARS can form lengthy ribose polymers of more than 80 residues to target proteins. Under normal physiologic conditions the biologic half-life of these polymers is less than 1 minute; therefore, nuclear enzymes can utilize NAD to create rapidly high molecular weight polyanions, which can have a drastic but transient effect on chromatin structure and enzyme function (for review, see Ref. 17). However, for every mole of NAD used as substrate by PARS, 4 moles of ATP are required to regenerate NAD. Therefore, when DNA is excessively damaged and PARS activation is sustained, both NAD and ATP intracellular levels drop precipitously. This depletion of NAD and ATP results in drastic reductions in energy-dependent processes, including synthesis of DNA, RNA, and protein. Application of PARS inhibitors such as benzamide blocks the depletion of NAD and ATP, resulting in restoration of energydependent processes and preservation of cellular integrity and neuroprotection against NMDA and NO donor-mediated neurotoxicity (4). It is possible that under certain conditions NO and other free radicals trigger massive DNA damage, overactivating PARS, resulting in depletion of the energy sources, NAD, and ATP, culminating in neuronal cell death. If this is the case, PARS inhibitors might have widespread therapeutic effects in diseases and neurodegenerative disorders that may involve glutamate neurotoxicity, such as stroke, Huntington's disease, and Alzheimer's disease.

[2]

NITRIC OXIDE TOXICITY IN CNS CULTURES

39

Appendix

NADPH-Diaphorase

Stain

FIXative: 4% Paraformaldehyde/O.1 M PB Use 8% paraformaldehyde (8 g/100 ml Milli-Q water); heat to 80~ for 30 minutes (do not boil). Clear with 1-2 drops of 10 N NaOH and replace volume of evaporated water. Add an equal volume of 0.2 M PB (100 ml 0.2 M NaHzPO4:400 ml 0.2 M NazHPO 4, pH 7.4) and pass through a 0.45tzm filter.

NADPH-Diaphorase Stain Solution Buffer 0.1 M Tris-HC1 (pH 7.2) 0.2% Triton X- 100 7.5 mg Sodium azide/100 ml buffer Mix 1 mM NADPH, reduced form (Sigma); 8.33 mg/5 ml 0.2 mM Nitroblue tetrazolium (NBT) (Sigma); 1.64 rag/5 ml Final volume is 10 ml; solubilize NADPH and NBT in separate test tubes to prevent precipitation. NBT may need to be sonicated to solubilize. Mix soluble NADPH and NBT to form stain mix.

E x p o s u r e Solutions Excitatory Amino Acids NMDA is readily soluble in water or buffer solutions up to 10 mM. Glycine is necessary for full activation of the NMDA receptor. Glycine is present in most water supplies, but to ensure consistent activation, a final concentration of 10/~M glycine should be added. Quisqualate is not readily soluble in water or buffer solutions. Make a 1 mM stock solution and heat to 52~ in a water bath. Vortex vigorously and heat until the fine crystals go into solution. Kainate is readily soluble in MEM. Add kainate to MEM and expose the cultures for 24 hours in the incubator to the kainate solution. Glutamate is readily soluble in water or buffer solutions.

40

PARADIGMS OF NEURAL INJURY

NOS Inhibitors NG-Nitro-L-arginine (N-Arg) competes with arginine for the catalytic site and is moderately soluble in water or buffer solutions up to a 1 mM concentration with vigorous vortexing and heating up to 42~ N-Methylarginine (NMA), N-iminoethyl-L-ornithine (NIO), NG-nitro-L arginine methyl ester (L-NAME), and NG-methyl-L-arginine acetate salt (L-NMMA) compete with arginine for the catalytic site and are readily soluble in water or buffer solutions. 7-Nitroindazole competes with arginine for the catalytic site and is more potent in inhibiting neuronal than endothelial NOS. Note: All of the arginine analog inhibitors of NOS inhibit all NOS isoforms. If more than one isoform is present in the preparation, care must be taken to assure that inhibition of all isoforms present does not confound the results. Diphenyleneiodonium (DPI) inhibits electron shuttling by flavoproteins. Dissolve in dimethyl sulfoxide (DMSO) and dilute in buffer solution. DMSO (Kodak, Rochester, NY) concentration cannot exceed 0.1%. GM1 ganglioside inhibits calmodulin, an essential cofactor for NOS activation. It is barely soluble in buffer solutions and requires a 2-hour preincubation before initiation of experiment. GTlb ganglioside inhibits calmodulin, an essential cofactor for NOS activation. It is barely soluble in buffer solutions and requires a 2hour preincubation before initiation of experiment. FK506, or immunophilin (Fujisawa Pharmaceuticals, Japan), binds to FKBP-12, and the FK506FKBP12 complex binds to calcineurin, inactivating the phosphatase. NOS is activated by calcineurin-mediated dephosphorylation.

Arginine Depletion Arginine is the precursor to NO formation by NOS. Cell cultures can readily be depleted of arginine by two methods. 1. Culture cells for 24 hours in arginine-free MEM (GIBCO, Grand Island, NY, special order) in the presence of 2 mM glutamine to inhibit argininosuccinate synthase. Arginine is a semi-essential amino acid and is synthesized by the cells unless the synthesis pathway is blocked by glutamine. After exposure of the cultures to experimental conditions the MEM + 21 mM glucose must also be arginine flee. 2. Treat both the media and the cell cultures with arginase, 10 units for 2 hours. Following exposure to experimental conditions place cells in arginase-treated MEM + 21 mM glucose.

[2]

N I T R I C O X I D E T O X I C I T Y IN CNS C U L T U R E S

41

NO Donor Reagents Sodium nitroprusside (SNP) is readily soluble in water or buffer solutions and releases NO and Fe(CNs) 2-. S-Nitroso-N-acetylpenicillamine (SNAP) (RBI, Natick, MA or Molecular Probes, Eugene, OR) is soluble in DMSO up to 100 mM and then can be diluted carefully into buffer solutions. Final DMSO concentrations cannot exceed 0.1% because DMSO is a free radical scavenger and is neuroprotective at concentrations greater than 0.1%. Additionally, penicillamine is a peroxynitrite (ONOO-) scavenger. In culture, a balance must be obtained between release of NO and scavenging of ONOO-. 3-Morpholinosydnonimine (SIN-l) (Molecular Probes) is soluble in water or buffer solutions. SIN-1 releases both NO and superoxide anion. Note: The NO donors should be weighed, kept in a foil-wrapped test tube, and solubilized immediately before use. In pure deoxygenated water, NO is stable for up to 30 minutes, but in buffer solutions the stability of NO is related to the degree of oxygenation and metal ions. For example, the stability of NO in Krebs buffer is less than 5 minutes. The NO rapidly converts to nitrite in the presence of metal ions. Choice of buffer solutions is therefore very important. Superoxide Anion Scavenger Superoxide dismutase is soluble in water and is a very stable protein. Hemoglobin Reduction Dissolve 2 mM hemoglobin (Sigma) in water. Dissolve 20 mM dithionite (Sigma) in water. When both reagents are in solution, mix them together in a 1:1 (v/v) ratio for a final concentration of 1 mM hemoglobin and 10 mM dithionite. The hemoglobin should turn from a brown color to a deep red color. Put the reduced hemoglobin in dialysis tubing with a pore size less than 60 kDa. Dialyze overnight in 1000-fold excess water at 4~ The dialysis vessel should be wrapped in foil. Replace the water with fresh water and store at 4~ wrapped in foil until use. Hemoglobin oxidizes very rapidly so prepare the cultures for the experiment and have all the solutions ready prior to diluting the hemoglobin to 500 tzM for the experiments. Long exposures to hemoglobin (in hours) should be avoided because hemoglobin can be toxic to neurons. Note: Hemoglobin does not exclusively scavenge NO, but other oxidants as well, including superoxide anion. PARS Inhibitors Benzamide (Aldrich) is soluble in DMSO; dilute (w/v) to less than 0.1% DMSO in final exposure solution. 3-Aminobenzamide (Aldrich) is soluble in DMSO; dilute (w/v) to less than 0.1% DMSO in final exposure solution. 1,5-

42

PARADIGMS OF NEURAL INJURY Dihydroxyisoquinoline (DHIQ), or 1,5-isoquinolinediol (Aldrich) is soluble in DMSO; dilute (w/v) to less than 0.1% DMSO in final exposure solution. Peroxynitrite

Peroxynitrite is not commercially available. O N O O - can be synthesized by the methods of Reed et al. (1974) and Beckman et al. (1994). O N O O - is relatively stable in alkaline solutions but rapidly decomposes in buffer solutions at physiologic pH. Both the volume of stock O N O O - added and the volume of exposure solution should be kept to a minimum to try to decrease the variability in concentration to which the cells will be exposed. O N O O decomposes and the concentration drops constantly while the reagent diffuses through the media to the cells. N o t e : Bicarbonate scavenges O N O O - ; therefore, bicarbonate-based buffers should be avoided when examining this chemical species in culture.

Acknowledgments VLD is supported by grants from National Alliance for Research on Schizophrenia and Depression, American Foundation for AIDS Research and the American Heart Association. TMD is supported by grants from the PHS, CIDA NSO 1578, the International Life Science Institute, and American Health Assistance Foundation. The authors own stock in and are entitled to royalty from Guilford Pharmaceuticals, Inc., which is developing technology related to the research described in this chapter. The stock has been placed in escrow and cannot be sold until a date that has been predetermined by the Johns Hopkins University.

References 1. M. A. Dichter, Brain Res. 149, 279 (1978). 2. R. R. Ratan, T. H. Murphy, and J. M. Baraban, J. Neurochem. 62, 376 (1994). 3. V. L. Dawson, T. M. Dawson, D. A. Bartley, G. R. Uhl, and S. H. Snyder, J. Neurosci. 13, 2651 (1993). 4. J. Zhang, V. L. Dawson, T. M. Dawson, and S. H. Snyder, Science 263, 687 (1994). 5. T. M. Dawson, D. S. Bredt, M. Fotuhi, P. M. Hwang, and S. H. Snyder, Proc. Natl. Acad. Sci. U.S.A. 88, 7797 (1991). 6. D. W. Choi, J. Koh, and S. Peters, J. Neurosci. 8, 185 (1988). 7. S.A. Lipton, N. J. Sucher, P. K. Kaiser, and E. B. Dreyer, Neuron 7, 111 (1991). 8. E. R. Stadman, Science 257, 1220 (1992). 9. B. H. J. Juurlink and L. Hetrz, Dev. Brain Res. 70, 239 (1993).

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10. S. Z. Lei, Z.-H. Pan, S. K. Aggarwal, H.-S. V. Chen, J. Hartman, N. J. Sucher, and S. A. Lipton, Neuron 8, 1087 (1992). 11. R. E. Huie and S. Padmaja, Free Rad. Res. Commun. 18, 195 (1993). 12. J. S. Beckman, H. Ischiropoulos, L. Zhu, M. van der Woerd, C. Smith, J. Chen, J. Harrison, J. C. Martin, and M. Tsai, Arch. Biochem. Biophys. 298, 438 (1992). 13. J. P. Crow, C. Spruell, J. Chen, C. Gunn, H. Ischiropoulos, M. Tsai, C. Smith, R. Radi, W. Koppenol, and J. S. Beckman, Free Rad. Biol. Med. 16, 331 (1994). 14. T. M. Dawson, V. L. Dawson, and S. H. Snyder, Ann. Neurol. 32, 297 (1992). 15. T. M. Dawson, V. L. Dawson, and S. H. Snyder, in "Neurobiology of NO. and 9OH." Annals of the New York Academy of Science. In press (1994). 16. D. A. Wink, K. S. Kasprzak, C. M. Maragos, R. K. Elespuru, M. Misra, T. M. Dunams, T. A. Cebula, W. H. Koch, A. W. Andrews, J. S. Allen, and L. K. Keefer, Science 254, 1001 (1991). 17. N. A. Berger, Rad. Res. 101, 4 (1985). 18. J. W. Reed, H. H. Ho, and W. L. Holly, J. Am. Chem. Soc. 96, 1248 (1974). 19. J. S. Beckman, J. Chen, H. Ischiropoulus, and J. P. Crow, in "Methods in Enzymology" (L. Packer, ed.), Vol. 233, p. 229. Academic Press, San Diego, 1994.

[3]

Development of in Vitro Injury Models for Oligodendroglia A. Espinosa, P. Zhao, and J. de Vellis

Introduction One of the major tasks of neural cultures has been to provide an understanding of the mechanism(s) involved in the development and function of the central nervous system. The role of environmental agents and naturally occurring substances that affect CNS activity has been elucidated to a large extent by using brain culture systems. Oligodendrocyte/astrocyte primary glial cultures are usually prepared in our laboratory from newborn rat brain (McCarthy and de Vellis, 1980). These cultures are the source of oligodendrocytes and their progenitors, which can be mechanically isolated from the astroglial monolayer (Cole and de Vellis, 1989). Both cell populations can be replated separately for further studies. These cultures have been extremely important for the characterization of oligodendroglial cell development (de Vellis and Espinosa, 1992). Recently we have focused our efforts to the establishment of various in vitro injury models for oligodendroglia. Such models facilitate the study of the serial changes suffered by these cells following injury. They allow evaluation of the either permanent or transient dysfunction and the potential of oligodendrocytes to recover. Here we describe a "freeze-thaw" method applied to two kinds of cultures: (a) primary 2-week-old glial cultures and (b) 3-day-old oligodendrocyte cultures prepared as described (Cole and de Vellis, 1989). The experimental design consists in the exposure of the cell cultures to severe hypothermia in an attempt to stop virtually all their metabolic functions. In order to preserve the cell structure and cytoplasmic membrane, the regular culture medium is substituted by a special freezing medium. Cultures are kept at -20~ and cells are reanimated 1 or 2 weeks later. Reanimation of the cells consists of placing them under regular culture conditions. We will discuss the phenotypic changes displayed by the cells as assessed by double-fluorescent immunostaining and the applications of this method for further studies on oligodendroglial cell injury.

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Methods in Neurosciences, Volume 30 Copyright 9 1996 by Academic Press, Inc. All rights of reproduction in any form reserved.

[3] In Vitro INJURY MODELS FOR OLIGODENDROGLIA

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Experimental Procedures

Preparation of Glial Cell Cultures Primary glial cultures are prepared from newborn rat brains (Cole and de Vellis, 1989). Cultures are maintained in the standard culture medium, Dulbecco's modified Eagle's medium/Ham's F12 (DMEM/F12). The DMEM/ F12 medium is supplemented with 10% calf serum. After 1 week in culture the flasks or multiwell plates are rinsed twice with phosphate balance salt (PBS) solution (NaC1, 120 mmol/liter, KC1, 2.7 mmol/ liter in 10 mM phosphate buffer at pH 7.4) at room temperature. The liquid is removed from the flasks by aspiration, leaving the cell layer virtually dry, and the freezing medium is slowly added to the cell layer. Observation of these cultures under phase-contrast microscopy reveals a confluent flat layer of astrocytes overlaid with oligodendrocytes. Secondary cultures of pure oligodendrocytes are similarly treated. For immunocytochemistry, glass coverslips are prepared in 24-well plates, coated with poly(D-lysine), and rinsed prior to cell plating.

Freezing Medium Preparation A mixture of dimethyl sulfoxide (DMSO) or glycerol, calf serum, and "glial development" culture medium is freshly prepared as follows: DMSO, 10%; calf serum, 89%; and GDM, 1% (v/v). This mixture is applied to the culture layer at room temperature (12 ml of freezing medium (FM) per 75 r The freezing medium should cover the total surface of the flask(s). For immunocytochemistry, 700 ~l to 1 ml per well of freezing medium is recommended.

Preparation of Cultures for Freezing Prior to freezing, the flasks or well plates are individually wrapped with a thick layer of cotton. A second layer should be prepared with dippers (to isolate the surfaces of the plate) followed by a third layer of thin foil. This step will allow a gradual decrease in temperature necessary to preserve plasma membrane integrity. Cultures should be placed in a -20~ freezer. The flasks are placed at -70~ 24 hours later. They can be kept indefinitely under these conditions. The minimum freezing period recommended is 48 hours.

46

PARADIGMS OF N E U R A L I N J U R Y TABLE I

Basal Medium DMEM/F12

Additives

GDM

Insulin Putrescine Sodium selenite a o(+)-Galactose Penicillin-streptomycin Transferrin Sodium bicarbonate L-Glutamine

5.0 mg/liter 16.1 mg/liter 10.0/A/liter 4.6 g/liter 1.0 ml/liter 50.0 mg/liter 2.2 g/liter

a Stock solution for sodium selenite is 0.8 mg/ml. Note: DF mix powder contains a large (4.5 g/liter) amount of glucose.

Culture Media Composition Our studies for the nutrient requirements of oligodendroglial cells indicate that, in a chemically defined medium, cells from the oligodendrocytelineage display an ability to proliferate and mature in a manner similar to the in vivo situation. The culture medium used is the glial developmental medium (GDM). The composition of the culture GDM is shown in Table I. Upon or after reanimation, oligodendroglial cells are fed with either GDM or with DMEM/F12 (1 : 1, v/v) mixture supplemented with 10% calf serum (DF-10).

Reanimation of Samples Regular culture medium is warmed at 37~ The frozen samples are totally unwrapped and placed for 10 to 12 minutes in a 37~ incubator. As soon as the freezing medium is liquid it is aspirated, leaving the cell layer as dry as possible. Then the flasks are rinsed twice with fresh culture medium, leaving the second rinse at least 3 minutes in each flask. After the last rinse add 10 ml of the same medium or 1 ml/well for the twenty-four or four well plates and keep the cultures in the incubator, at least 24 hours. Cultures can be studied any time (beyond the initial 24 hours) and it is recommended that the medium be changed 24 hours after thawing. These cultures can be used to address a variety of questions. For the present study we used mixed glial cultures kept frozen for 2 weeks, followed by a 2-week incubation at 37~ or we used pure oligodendrocyte cultures reanimated in either DF-10 or GDM. Samples were fixed 3 days after reanimation.

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Immunocytochemical

47

Characterization

Characterization of the phenotype of the cells present in the cultures after the freeze-thaw treatment is essential. Double immunofluorescence has been very useful to assess the expression of specific markers by a given cell population (Espinosa and de Vellis, 1988, 1990). It is well known that the morphological features of a cell together with the expression of specific cell markers define not only the cell type but in many cases also indicate its developmental or functional stage (Espinosa and de Vellis, 1995). Many techniques have been described for the characterization of cells in culture. All these techniques offer advantages and disadvantages that need to be considered in order to select the most adequate method for the examination of the changes produced by the freeze-thaw injury model. The most current technique for visualization of a single antigen is the avidin-biotin complex (ABC) method. This procedure allows a very sharp detection of even very slight changes of antigen expression and/or its distribution within the cell. Such small changes would be undetectable by using the peroxidase-antiperoxide method or fluorescent immunocytochemistry. Another advantage of the ABC technique is that it often requires the use of much less antibody than do other methods. At the present time, many products are commercially available for the ABC method. To study a small number of samples, a kit is convenient and cost effective. If the technique is to be used on a regular basis for a large number of samples, however, it is more cost effective to purchase the reagents separately. For the study presented here, we selected double and triple immunofluorescence for the characterization of the cell populations present in reanimated cultures. We used the following markers: sulfatides detected with the monoclonal antibody 04, galactocerebroside (GC) and myelin basic protein (MBP) for oligodendrocyte/myelin markers, and the glial fibrillary acidic protein (GFAP) for identification of astrocytes. Transferrin (Tf) has been described as an early marker for oligodendroglia (Espinosa et al., 1988) and appears in these cells prior to any myelin component, including GC. We also studied Tf expression in our samples. For visualization of these markers, a secondary goat antimouse immunoglobulin G (IgG) tagged with fluorescein, a goat antirabbit IgG tagged with Texas Red, and a goat antimouse IgM are combined. All the antibodies are purchased from Boehringer Mannheim (Indianapolis, IN). The dilutions for primary and secondary antibodies are predetermined by titration. The method for immunostaining used in the present study has been described in detail for freshly dissociated cells (Espinosa et al., 1989). It has been successfully used for the characterization of well-established cell cultures such as glial cells from normal rat brains (Espinosa and

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PARADIGMS OF NEURAL INJURY

de Vellis, 1988) or from myelin-deficient (md) rat mutant (Espinosa et al., 1989) cell lines and other cell cultures. Briefly, the method consists of permeabilization of the cells with Triton X-100 treatment and blockage of nonspecific binding sites with normal goat serum. Preparation of the cocktail of primary antibodies and their incubation overnight are carried at 4~ The next day, primary antibodies are rinsed and secondary antibodies are prepared. Samples are incubated at room temperature with a mixture of secondary antibodies for 1 hour, rinsed, and mounted. Samples are observed under a micro Fx fluorescent Nikon microscope equipped with the appropriate filters and barriers.

Results

Observation of Samples Primary glial cultures are characterized by the presence of a layer of astrocytes and oligodendroglial progenitors that appear within the first week postplating. Oligodendrocyte progenitors display a dark and flat appearance, and very short (or no) processes. Initially, the majority are found as single cells (Espinosa et al., 1985). During the second week in culture dark cells change to round bright cells with thin long cell processes either in clusters or as single cells surrounding such clusters (Espinosa et al., 1985; Cole and de Vellis, 1989). When primary glial cultures that were frozen are reanimated at 37~ in the incubator, the freezing medium melts within 10 to 12 minutes; at this time cells are observed under the microscope. The layer of astrocytes and small clusters of oligodendrocytes present in these cultures appear normal when compared to nonfrozen cultures. However, large oligodendrocyte clusters, as well as some single oligodendrocytes, detach and float in the culture medium. At this time, the freezing medium is substituted for fresh culture medium at 37~ Cultures are placed in the incubator and subsequently observed daily. Observation of the cultures reveals no morphological differences in frozen versus nonfrozen cells. Cultures are fed every fourth day. When oligodendroglial cultures are reanimated with the same procedure used for primary glial cultures, a large number of cells detach from the coverslip, leaving approximately 40% of viable cells on the coverslips. Floating cells are eliminated by the replacement of freezing medium for either GDM or DF-10 fresh culture medium. Cultures reanimated and fed with either one of the media look very similar under phase-contrast microscopy. No difference is detected morphologically in reanimated cultures fed with

[3] In Vitro INJURY MODELS FOR OLIGODENDROGLIA

49

FIG. 1 Phase-contrast views of oligodendroglial cells cultured for 10 days after their isolation from primary glial cultures. (A) The oligodendroglial cells display multiple cell processes; some of the cells have formed clusters and some single cells interconnect with neighboring cells, forming a network with their processes. All cells have a small cell body with similar morphology. (B) View of a reanimated culture. Cells were grown for 1 week, as was their sister culture (A), then were frozen for 1 week. Cells were reanimated as described in the text and cultured for 3 more days. These cultures differ from their sister culture counterparts in cell numbers, the almost total absence of cell clusters, the large amount of debris, and vacuolated cells. However, the most remarkable finding is the morphological diversity of the cell populations. None of these oligodendrocytes have long cell processes; many cells lack cell processes, their cell bodies appear larger, and some are vacuolated. A subpopulation of astrocyte-like cells is present. These GFAP § cells have multiple long cell processes and are not found in normal glial cultures. Magnification: x200.

DF-10 (Fig. 1) or G M D (now shown). Despite the c o n s i d e r a b l e loss of cells during r e a n i m a t i o n , a diversity of cell types is still p r e s e n t in these cultures (Fig. 1B). This h e t e r o g e n e i t y in r e a n i m a t e d cultures s h o w s that the t r e a t m e n t affects to a different e x t e n t cells from the same lineage t r e a t e d identically

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PARADIGMS OF NEURAL INJURY

through the duration of the experiment. All the different cell types survive and remain healthy in these cultures until fixation time (in this case 3 days after reanimation). In contrast, oligodendrocyte cultures are composed of a homogeneous cell population with similar morphology. These cells present a healthy appearance with their characteristic small round and bright cell body and multiple thin cell processes. These cells are either single or are in small clusters homogeneously distributed on the surface of the coverslip (Fig. 1A).

Fixation and Characterization of Samples After 3 days or 2 weeks after reanimation, primary glial cells or oligodendroglial cultures are fixed. Nonfrozen sister samples are fixed either 3 days or 2 weeks after their counterparts are frozen and stored at 4~ until reanimated samples would be fixed. Previous studies have led us to use formaldehyde (3.7% v/v) as a routine fixative for most of our studies instead of other methods, such as acetone-ethanol, paraformaldehyde, or cold air flow, which have also been used to study mature oligodendroglia in culture (Espinosa et al., 1986a,b; Espinosa, 1987). Every fixative offers advantages and disadvantages with respect to others; for instance, paraformaldehyde offers an excellent preservation of cell morphology, whereas cold air flow or acetoneethanol enhances the visualization of membrane components, but in some cases soluble antigens are poorly detected or even undetectable due to their partial or total extraction during fixation procedure. Formaldehyde allows the detection of a large panel of cell markers independently of whether they are in a soluble form in the cytoplasm, membrane associated, membrane components, or cytoskeleton proteins. Formaldehyde fixation provides a good preservation of cell morphology and of the antigens we studied. The appearance of reanimated primary glial cultures under phase-contrast after fixation is very similar to that of nonfixed cells. At the time of fixation, cells do not show any changes and remain attached to the culture dish. However, when fixative is added to secondary cultures of oligodendrocytes, 10-12% of cells lift from the coverslip; these cells are eliminated from the coverslip. The rest of the cells remain attached throughout the procedure.

Immunocytochemical Analysis of the Cultures After fixation, double or triple immunofluorescence is performed as described in the methods section. Reanimated primary glial cultures are examined by double immunofluorescence for GFAP and GC or MBP. Figure 2 (color

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51

plate) shows the comparison of reanimated samples (Fig. 2A and B) to nonfrozen sister age-matched cultures (Fig. 2C and D). Reanimated cultures are characterized by the presence of GC + cells in clusters or as single cells. However, they do not have any cell processes. GFAP + cells with round cell body, eccentric nucleus, and multiple long cell processes are frequently observed. These cells are also GC +, giving an orange-yellow color by the colocalization of rhodamine and fluorescein-tagged antibodies in the same cell (Fig. 2A). In some cases, these cells have extremely long GFAP + cell processes (in green), whereas GC expression is restricted to the cell body (in orange-yellow, Fig. 2B). In sister cultures that were never frozen, all the cells either in clusters or single are GC +, with branches forming a GC + network (Fig. 2C). All the cells in these cultures (100%) express the myelin marker MBP, of the cells in the cell bodies, their processes, and extensive layers of membrane, as seen in Fig. 2D. These cultures are virtually devoid of GFAP immunoreactivity (data not shown). The most remarkable finding is the presence of MBP + membrane sheaths in control cultures and the absence of both membrane sheath and cell processes in reanimated cultures. Reanimated oligodendrocytes fed with either one of the culture media (DF-10 or GDM) are analyzed by triple immunofluorescence. The reason for this is the diversity of cell types found in treated samples. Phase contrast does not reveal a major difference between DF-10 and GDM reanimated cultures, because only the cell bodies are visible. The primary antibodies are prepared in combination, Tf/O4/GFAP. Both oligodendrocyte markers Tf and 04 appear early in the lineage. Tf is expressed only by 5% of mature oligodendrocytes, but 100% by young oligodendrocytes. Sulfatides (detected by 04) appear later than Tf but persist in the mature cell and its membrane(s). The pattern of expression of 04 and GC is very similar (04 is used here as an alternative to GC because it is a monoclonal IgM, allowing its combination with mGFAP, IgG, and polyclonal TfIgG). Immunocytochemistry reveals more information about the effects of the treatment. Besides the expression of the antigens studied, the cell morphology is more distinct when viewed with epifluorescence, instead of phase contrast. The appearance of cultures fed with DF-10 after reanimation is shown in Fig. 3 (color plate); the cells are flat and have either no processes or irregular processes. These cells have a weak expression for GFAP (Fig. 3A) as well as for 04, they have retracted all their processes, and their morphology differs from what is expected for a normal 04 + cell (Fig. 3C). These cells are extremely immunoreactive for Tf (Fig. 3B), which appears soluble or in vesicles within the cell cytoplasm. Some spheres of Tf + are also frequently outside the cells (on the substrate). In some cases these vesicles are in the inner side of the plasma membrane. Tf + vesicles are

52

PARADIGMS OF NEURAL INJURY

negative for 04 and GFAP. Occasionally a short display of cytoplasmic extension is found but no cell processes or myelin-like sheaths are visible. When sister reanimated samples are cultured in GDM, the findings are very different. Most of the cells have cytoplasmic extensions in a perinuclear symmetric manner. Various intensities of 04 are observed, at the level of the cell body, with a smooth appearance as a soluble product. In a few cells 04 is within intracellular vesicles (Fig. 3F). However, the most remarkable finding is the preservation of extensive myelin-like sheaths that are also O4+, as shown in Fig. 3D. All of the cells have very strong expression of Tf (Fig. 3F) and more than 50% of them display a weak GFAP immunoreactivity (Fig. 3D). It is important to note that the cell bodies are smaller than those from cells maintained in DF-10.

Interpretation of Data and Concluding Remarks The elucidation of the mechanisms elicited in oligodendrocytes by injury is necessary in order to be able to provide support for recovery of function in these cells. Several in vitro approaches have been described to study chemical damage focusing mainly on a nutrient deficit or cytotoxicity. The experimental injury model for oligodendrocytes that we have described here demonstrates that injury is not necessarily accompanied by cell death as a result of injury. However, cells suffer major phenotypic changes as a result of severe hypothermia. This model allows direct observation of the changes induced by the freeze-thaw treatment. The model offers a simple yet effective way of evaluating the extent of injury. Furthermore, this system can be used as a working model to test potential factors that may promote recovery of the cells. Examination of reanimated cultures can be done not only by immunocytochemistry but samples can also be prepared in culture dishes or flasks for Northern blot analysis of mRNA expression of the mRNA markers of interest. Western blots can also be prepared, yielding not only qualitative results, but also quantitative information. The application of this method can be very extensive, going from time course studies to different cell types or culture conditions. Furthermore, this method can be used to study interspecies response to injury and plasticity going from fish to mammals. Using the model described in this review, we have learned that the freeze and thaw treatment appears to affect mature oligodendrocytes differently when cultured as mixed glia versus purified oligodendroglia. We observed that in mixed glial cultures cell loss was relatively low, but oligodendrocytes had lost their cell processes and myelin sheaths. All of these cells were GC + both in clusters or single cells. The expression of GFAP by a subpopulation of cells in these cultures was frequent, approximately 30% of the total number

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In Vitro INJURY MODELS FOR OLIGODENDROGLIA

53

of cells. GFAP + cells displayed multiple long cell processes; some of them were also GC +, indicating their oligodendroglial origin and suggesting that oligodendrocytes, just like astrocytes, can be reactive under certain conditions. A large number of studies of oligodendrocytes in culture have led to the finding of GFAP expression by some or most of these cells, depending on the culture conditions (Raft et al., 1983). When these GFAP + cells were first found in cultures prepared from optic nerve or brains of newborn rat pups, they were considered an alternative phenotype to oligodendrocytes, derived from the same progenitor, termed a "bipotential progenitor," thus the GFAP + cell was named a type II astrocyte. The culture conditions wherein type II astrocytes were present have been considered as the optimal conditions for the appearance and maintenance of this type II phenotype. To our knowledge, the present study is the first of its kind to show the changes resulting from injury to mature oligodendroglia. Extensive studies have shown that reactive gliosis occurs as a result of trauma or pathology. Initially, this phenomenon was assessed in the mature CNS and the term "gliosis" was and is still used for astroglial cells. Among the features characteristic of a reactive astrocyte are cell proliferation, hypertrophy of the cell body and cell processes, as well as overexpression of the intermediate filament protein GFAP (Duffy, 1983). Our preliminary findings using the freeze-thaw model indicate that injury induces the expression of the GFAP gene in oligodendroglia to a different extent. The colocalization of 04 and GFAP confirms that GFAP + cells had synthesized 04 before freezing and that they are oligodendrocytes. On this basis, GFAP expression by these cells indicates a "reactive" stage(s) of oligodendrocytes. The overexpression of Tf experienced by reanimated cells indicates that injury up-regulates this gene as well. The colocalization of GFAP, 04, and Tf in oligodendrocytes is not an unusual finding. In other experiments performed in our laboratory whereby cultured oligodendrocytes were exposed to abnormal conditions, such as hyperoxia, colocalization of oligodendroglial markers with GFAP was observed (Espinosa and de Vellis, 1995). On the basis of these observations it is clear that plasticity of oligodendrocytes may also include a reactive stage under nonphysiological conditions, e.g., the so-called type II astrocyte arising from oligodendroglial progenitors in culture exposed to fetal calf serum. It remains to be determined whether the reactive oligodendrocyte described here is a transient or permanent phenotype. The use of different culture media resulted in both protection of performed myelin sheaths and their maintenance after reanimation. This last finding strongly suggests that oligodendroglia can support their myelin sheath after injury if they are provided with adequate conditions for their recovery.

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PARADIGMS OF NEURAL INJURY

Acknowledgments We thank Dennis Espejo, Danny Vu, and Nancy Wainwright for help in the preparation of this manuscript. Research was supported by NIH Grant HD-06576.

References Cole, R., and de Vellis, J., in "A Dissection and Tissue Culture Manual of the Nervous System," pp. 121-133. Alan R. Liss, New York, 1989. de Vellis, J., and Espinosa de los Monteros, A., Neuromethods 23, 323-352 (1992). Duffy, P. E., "Astrocytes; Normal, Reactive and Neoplastic." Raven Press, New York, 1983. Espinosa de los Monteros, A., Ph.D. Thesis. Universit Louis Pasteur de Strasbourg (1987). Espinosa de los Monteros, A., and de Vellis, J.,J. Neurosci. Res. 21, 181-187 (1988). Espinosa de los Monteros, A., and de Vellis, J., in "Cellular and Molecular Biology of Myelination (H. Althaus, ed.). Springer-Verlag, Berlin and New York, 1990. Espinosa de los Monteros, A., and de Vellis, J., "Vulnerability of Oligodendrocytes to Environmental Insults: Potential for Recovery (M. Aschner, ed.). CRC Press, Boca Raton, Florida, 1995. Espinosa de los Monteros, A., Roussel, and Nussbaum, J. L., and Labourdette, G., Dev. Biol. 108, 474-480 (1985). Espinosa de los Monteros, A., Roussel, G., and Nussbaum, J. L., Biochem. Soc. Trans. 14, 648-650 (1986a). Espinosa de los Monteros, A., Roussel, G., and Nussbaum, J. L., Dev. Brain Res. 24, 117-125 (1986b). Espinosa de los Monteros, A., Chiappelli, F., Fisher, R. S., and de Vellis, J., Int. J. Dev. Neurosci. 6, 167-175 (1988). Espinosa de los Monteros, A., Pena, L. A., and de Vellis, J., J. Neurosci. Res. 24, 125-136 (1989). Levison, S., and McCarthy K., pp. 103-104. Alan R. Liss, New York, 1990. Raft, M. C., Miller, R. H., and Noble, M., Nature (London) 303, 390-396 (1983).

[4]

Glia Models to Study Glial Cell Cytotoxicity Antonia Vernadakis and M. Susan Kentroti

Introduction The neuron-glia functional partnership first proposed by Hyden in 1961 is now generally accepted, and several reviews have been written describing the cellular events in neuron-glia interactions [see three volumes, "Astrocytes," edited by Fedoroff and Vernadakis (1986a-c); see also the reviews by Vernadakis (1988) and by Abbott (1991)]. The role of cell-to-cell interactions involved in neurogenesis and neuronal growth and differentiation has been a major theme of developmental neurobiology in the past two decades. The coexistence of neurons and glial cells during early neuroembryogenesis places these cells in a strategic position to interact with each other and thus to influence their individual growth and differentiation. In vivo and in vitro studies described in the above-mentioned reviews demonstrate (1) the influence of glial cells on neuronal growth and differentiation and (2) the influence of neurons on glial growth and differentiation. Such interactions appear to be mediated through cell surface components and cell-secreted factors in the microenvironment. The second volume of "Astrocytes" edited by Fedoroff and Vernadakis is dedicated to the physiological and pharmacological aspects of astrocytes. Specific attention is given to intracellular metabolic activity; to membrane components and functions, including receptors, uptake, and transport; to responses to neurotransmitters and other intrinsic factors; and to neuron-glia interactions. Hertz and Schousboe (1984) eloquently describe the role of astrocytes in compartmentation of amino acid energy and metabolism. They position the astrocyte functionally between the y-aminobutyric acid (GABA)ergic and glutamatergic neuron, thus regulating the amount of both glutamate and GABA in the extracellular environment (Fig. 1). One can, therefore, deduct from this interrelationship that any changes occurring in the fucntion of astrocytes would be reflected in both the glutamatergic and GABAergic neuronal function. Such changes can be produced in astrocytes both by exogenous administration of cytotoxins and by cytotoxic substances produced endogenously; the latter are implicated in aging and in neuropathological conditions. A re vie w of Aschner and LoPachin ( ! 993) describes astroc ytes as targets and mediators of chemically induced injury. Several glial cell models have been used to study glia cell cytotoxicity and include both glioma Methods in Neurosciences, Volume 30

Copyright 9 1996 by Academic Press, Inc. All rights of reproduction in any form reserved.

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PARADIGMS OF NEURAL INJURY

Astrocyte

GABAergic Neuron

GABA

GIu

Glutamatergic Neuron

FIG. 1 Schematic drawing of evoked release and uptake of glutamate and GABA in GABAergic or glutamatergic neurons and in astrocytes. The sizes of the arrows give an estimate of the relative magnitudes of the respective fluxes. It can be seen that neuronally released glutamate, to a major extent, is accumulated in astrocytes, whereas most of the released GABA is reaccumulated in neurons. Reproduced with permission from Hertz, L. and Schousboe, A., in "Model Systems of Development and Aging of the Nervous System" (A. Vernadakis, A. Privat, J. M. Lander, P. S. Timiras, E. Giacobina, eds.), pp. 19-32. Martinus Nijhoff, Boston, 1984. cell lines and primary glial cells. In this chapter method of studying cytotoxicity using C6 glial cells and primary astrocytes derived from chick embryos will be described. Methodology for isolation of astrocytes from rodents is described in other chapters in this book and thus will be omitted here.

Glial Cell Models

Primary Glial Cell Cultures Glial-enriched cultures can be prepared from various animal species, but the most commonly used animals are rodents (e.g., mouse or rat) or birds (e.g., chicken). In this chapter we will describe glial-enriched cultures obtained from chick brain.

Glial-Enriched Cultures Derived from Chick Brain: Mixed Astrocyte-Oligodendrocyte Cultures Glial-enriched cultures can be prepared from cerebral hemispheres (or other brain areas) of 15-day-old chick embryos or from the telencephalons of 3-day-old chick embryos. In cultures derived from 15-day-old chick embryo cerebral hemispheres, neuronal cells do not survive after 5 days in culture, and by 15 days the cultures consist predominantly of glial cells and about 10% fibroblasts (after removal of meninges). In cultures derived from the

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MODELS TO STUDY G L I A L C E L L CYTOTOXICITY

57

3-day-old chick embryonic telencephalon, neuronal elements disappear within 1 week. The uniqueness of these early embryonic cultures is that they remain glioblastic until they are subcultured at least once. Thus these cultures can be used to study the influence of cytotoxins on early gliogenesis and glial phenotypic differentiation.

Preparation of Glial-Enriched Cultures Glia Cultures Derived from 15-Day-Old Chick Embryo (E15CC) The cerebral cortices from 15-day-old chick embryos are dissected under sterile conditions and collected on a piece of 73-~m mesh placed on a 100mm petri dish containing 2 ml Dulbecco's modified Eagle's medium (DMEM) + 20% fetal bovine serum (FBS). Tissue is mechanically dissociated through the mesh and the cell suspension counted. Each paired cerebral cortex yields approximately 1.0 x | 0 6 cells. Cells are plated directly on 60-mm plastic dishes at a concentration of 2.0 x 10 6 cells/dish in 3 ml DMEM + 20% FBS. Cells are grown at 37~ in 7.5% (v/v) CO2 in air. After 24 hours, the medium is changed to remove floating cells. Cultures are further grown with media changes every 4-6 days until neuronal elements disappear and cells form confluent cultures. After approximately 3 weeks, cultures are ready to be subcultured. Cultures grown in 20% FBS exhibit a high percentage of astrocytes. If one desires a higher population of oligodendrocytes, cultures are first plated in DMEM + 20% FBS and after 24 hours medium is changed to medium containing 2.5-5% FBS. Glia Cultures Derived from 3-Day-Old Chick Embryo (E3H) The telencephalic region from 3-day-old chick embryos is dissected under sterile conditions and collected on a piece of 48-~m mesh placed on a 100mm petri dish in 2 ml DMEM + 20% FBS. Tissue is mechanically dissociated through the mesh and the cell suspension counted. Each telencephalic region yields approximately 0.385 x 10 6 cells. Cells are plated directly on 60-mm plastic dishes at a concentration of 1.0 x l 0 6 cells/dish in 3 ml DMEM + 20% FBS. After 24 hours, the medium is changed to remove floating cells. Cultures are further grown with media that have been changed every 4-6 days until neuronal elements disappear and cells form confluent cultures. After approximately 3 weeks, cultures are ready to be subcultured.

Biochemical Profiles of Glial Marker Enzymes in Culture For biochemical studies, culture medium is aspirated and a 2-ml solution containing 0.1% trypsin is added to each 100-mm dish, or ! ml for a 60-mm dish, and immediately aspirated. This is to neutralize any peptidases left

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PARADIGMS OF NEURAL INJURY

from the serum in the medium and to minimize the effectiveness of trypsin in detaching the cells from the dish surface. In order to detach the cells from the dish, cultures are incubated at 37~ with 0.1% trypsin (5 ml for a 100mm dish or 3 ml for a 60-mm dish) for 3-5 minutes. The cell suspension from each dish is transferred with a Pasteur pipette to a conical tube containing an equal volume of DMEM + 10% FBS, centrifuged at 1000 rpm, 4~ for 10 minutes. Medium is aspirated and cell pellets are resuspended in 2-5 ml Earle's balanced salt solution (EBSS) and centrifuged as above. This procedure is repeated twice in order to remove all traces of FBS, which will interfere with protein analysis. The final pellet is frozen at -20~ until enzyme assays are performed. Pilot studies have shown that the enzyme activity does not decrease during this time. Two biochemical markers are used to identify astrocytes and oligodendrocytes. The activity of glutamine synthetase (GS; glutamate-ammonia ligase) is used for astrocytes (Norenberg and Martinez-Hernandez, 1979) and 3',5'cyclic-nucleotide phosphohydrolase (CNP) is used for oligodendrocytes (Podulso, 1975; Podulso and Norton, 1972). The biochemical assays used to determine the activities of GS and CNP are described in detail in Sakellaridis et al. (1983). Figures 2 and 3 illustrate the enzyme profiles of glial-enriched cultures prepared from 15-day-old chick embryo cerebral hemisphers (Sakellaridis et al., 1983). The decrease in CNP with days in culture is attributed to the lack of neurons in the cultures. To test this hypothesis, conditioned medium prepared from neuronal cultures can be added to the medium. For such a study the reader is referred to the report by Sakellaridis et al. (1984). Figure 4 illustrates the GS activity profile in E3H cultures, passage 7. I m m u n o c y t o c h e m i c a l Characterization o f E 3 H and E15CC Cultures Glial-enriched cultures are prepared from telecephalons of 3-day-old chick embryos or 15-day-old chick embryo cerebral hemispheres, as described above. An aliquot of 4000 cells is plated in four chamber slides. At various days in culture, slides are immunostained for glial markers. Markers used are galactocerebroside (GalC) for labeling oligodendrocytes (Raft et al., 1978); A2B5 (Eisenbarth et al., 1979) for labeling neurons, bipotential progenitors, type 2 astrocytes, and immature oligodendrocytes (Abney et al., 1983; Raft et al., 1978; 1984); glial fibrillary acidic protein (GFAP) for labeling type 1 and type 2 astrocytes (Bignami and Dahl, 1973; Raft et al., 1984); and vimentin, a major cytoskeletal component of immature glia (Dahl et al., 1981). These cell markers are identified using specific monoclonal or polyclonal antibodies directed against the appropriate antigen. Cells to be stained for GFAP or double-stained for GFAP and A2B5 are first fixed in 100 acetone at -20~ for 5 minutes. Cells to be stained for GalC or double-stained for GalC and A2B5 are first fixed in 3.7% formaldehyde in PBS for 30 minutes at room temperature. Cells are incubated in the primary antibody for 30

[4]

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MODELS TO STUDY GLIAL CELL CYTOTOXICITY 1.200 E "0

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Days in Culture FIG. 2 Changes with days in culture in glutamine synthetase activity in glial-enriched cultures dissociated from cerebral hemispheres of 15-day-old chick embryos. Activity is expressed in mircomoles of y-glutamylhydroxamic acid formed in 15 minutes/mg protein and plotted vs. days in culture. Points represent means +- SE of 3-4 separate culture dishes. From Sakellaridis, N., Bau, D., Mangoura, D., and Vernadakis, A., Neurochem. Int. 5, 685-689 (1983), with permission. minutes at room temperature, washed with PBS, and incubated an additional 30 minutes with secondary antibody conjugated to either fluorescein (GFAP; GalC) or rhodamine (A2B5). After a single passage, glia from E3H exhibit a glioblastic phenotype (Vim +, GFAP-) early in culture (C1) and progress to mature astrocytes (Vim +, GFAP +) later. Cells follow this same transition with each successive passage. In E15CC cultures, glial cells exhibit multiple phenotypes, with cells representing astroblasts (Vim+, GFAP-), intermediate astrocytes (Vim +, GFAP+), mature astrocytes (GFAP+), and oligodendrocytes (GalC+), and some cells express the extracellular matrix protein, fibronectin. Data in Figs. 5 and 6 illustrate the glia phenotypes exhibited in E 15C and E3H cultures.

C6 Glial Cells as M o d e l s C6 glial cells from a rat glioma cell line have been used to study glial cell properties and function. This cell line has generally been designated as an astrocytoma (Benda et al., 1968). C6 glioma cells can be obtained from the

60

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FIG. 3 Changes with days in culture in 2',3'-cyclic-nucleotide 3'-phosphohydrolase activity in glial-enriched cultures dissociated from cerebral hemispheres of 15-day-old chick embryos. Activity is expressed in micromoles of 2'-adenosine monophosphate formed in 20 minutes/mg protein and plotted vs. days in culture. Points represent means -+SE of 3-4 separate culture dishes. From Sakellaridis, N., Bau, D., Mangoura, D, and Vernadakis, A., Neurochem. Int. 5, 685-689 (1983), with permission. American Tissue Culture Collection (Rockville, MD). We use C6 glioma cells, 2B clone obtained courtesy of Dr. Jean deVellis from the University of California at Los Angeles. This cell line was given to us at passage 11 and we currently have passages ranging from 11 to 80 frozen in liquid nitrogen. Cell Passage Procedure Cells are plated at a density of 0.5 x 106 for a 100-ram petri dish or 0.1 x 10 6 for a 60-mm dish (Falcon, Lux, respectively). The growth medium is Dulbecoo's modified Eagle's medium (GIBCO, Grand Island, NY) supplemented with 10% fetal bovine serum (GIBCO). For a 100-mm dish, 6 ml of medium is used and for a 60-mm dish, 3 ml of medium is used. Cells are grown at 37~ in 7.5% (v/v) CO2 in air for 3 days, at which point medium is aspirated and replaced with new medium. At culture day 7 (C7), medium is aspirated and a solution containing 2 ml 0.1% trypsin is added to the 100mm dishes or 1 ml is added to the 60-mm dishes and again aspirated. In order to detach the cells, cultures are incubated at 37~ with 0.1% trypsin (5 ml for a 100-mm dish and 3 ml for a 60-mm dish) for 3-5 minutes. The cell suspension is transferred with a sterile pipette to a beaker containing 10-20 ml DMEM + 10% FBS. The cell suspension is mixed using a syringe with a large needle (14-gauge) or a pipette attached to a pipettor moved "up

[4]

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M O D E L S TO S T U D Y G L I A L C E L L C Y T O T O X I C I T Y "o (1)

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FIG. 4 Changes with days in culture in glutamine synthetase (glutamate-ammonia ligase) activity in glial-enriched cultures dissociated from 3-day-old chick embryo telencephalon (head) at passage 7. Activity is expressed in micromoles of y-glutamylhydroxamic acid formed in 15 minutes/mg protein and plotted vs. days in culture. Barograms with bracketed lines represent means +_ SE of 4-5 separate culture dishes per point.

and down" about l0 times. An aliquot is counted using a hemocytometer, which gives the amount of cells in 1 ml. This number is then multiplied by the total volume in the beaker to give a total number of cells. From this cell suspension new cultures can be plated. Cell passages up to passage 30 are characterized as early passage, whereas passages 70 and above are considered late passage. Figure 7 shows the morphology of early and late passage C6 glial cells (Lee et al., 1992).

Freezing Cells The procedure for freezing cells is similar to passing cells, up to the point when cells are counted. Instead of plating, the cell suspension is divided into sterile, screw-cap centrifuge tubes (15-ml volume) and centrifuged at room temperature for 5 minutes at low speed. The supernatant is decanted and the cell pellet resuspended in a solution containing 90% FBS and 10% glycerol. The volume to be used should be adjusted to contain 4-3 million cells/ml (labeled with cell passage, number, and date). An aliquot of 1 ml is transferred to sterile cryotubes, which are then capped. The above procedure, except for the centrifugation, is performed under a sterile laminar

62

PARADIGMS OF NEURAL INJURY

FIG. 5 Double-staining immunofluorescence labeling of glial cells derived from 15day-old chick embryo cerebral cortex. Cultures were double stained for GFAP and vimentin at various passages (P) and days in culture (C). (A and B) Cells from P1 at C11. At this early passage, the presence of intermediate astrocytes (GFAP § Vim§ large arrowhead) and of mature astrocytes (GFAP+; small arrowhead) is observed. (C and D) Cells from P4 at C 1; (E-H) cells from P4 at C 13. At C 1 cells are predominantly immature glioblasts exhibiting Vim +, GFAP- immunostaining (occasional cells are also GFAP +, Vim +, as shown by the arrowheads in (C and D). By C13 of the same passage 9P4), many more cells exhibit the intermediate astrocyte phenotype [GFAP § Vim§ small arrowheads (F)] as well as the phenotype of immature glioblasts [Vim§ GFAP-; large arrowheads (E)]. (G) Cells immunoreactive for the oligodendrocyte marker, GalC (outlined arrowhead); (H) same frame with a large, epitholioid cell exhibiting positive staining for the basement membrane protein, fibronectin.

[4]

MODELS TO STUDY GLIAL CELL CYTOTOXICITY

FIG. 5

63

(continued)

flow hood. The cryotubes are transferred to a bucket containing ice and refrigerated for 30 minutes. The cryotubes are then transferred to a bucket containing dry ice for 4 hour at -20~ Last, the cryotubes are placed in labeled racks and immersed in liquid nitrogen. Liquid nitrogen is replenished weekly. This method has resulted in our successfully keeping frozen cells for over 10 years.

Thawing Cells To retrieve cells for use from liquid nitrogen, cryotubes are removed and rapidly thawed in a 37~ water bath. Under a sterile hood, cryotubes are wiped with 70% (v/v) ethanol and the caps carefully removed. Cells are aspirated into a sterile, cotton-plugged Pasteur pipette and transferred to a tissue culture flask (75 cm) containing 30 ml DMEM + 10% FBS. This volume

64

PARADIGMS OF NEURAL INJURY

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Double-staining immunofluorescence labeling of glial cells derived from 3-day-old chick embryo telencephalon (head) at various passages and days in culture. (A-D) Cells from passage 0 (primary cultures) at C 17. Cultures consist predominantly of immature glioblasts [Vim+, GFAP-; large arrowheads (B)] with occasional intermediate astrocytes [Vim+, GFAP+; small arrowheads (A)]. (C) Cluster of oligodendrocytes, which stain positive for GalC (outlined arrowhead). A light micrographic representation of cultures is shown (D). (E-J) Cells at P1, C1, and C13. Whereas at C1 most cells are Vim § glioblasts (large arrows) with few cells exhibiting GFAP § Vim § immunostaining, by C13, cultures continue to contain Vim § glioglasts as well as cells that exhibit exclusively GFAP § immunostaining, considered mature astrocytes (G; small arrowhead). (I) Numberous GalC § oligodendrocytes. It should be noted here that this morphological pattern also agrees with biochemical expression of glutamine synthetase in Fig. 4. FIG. 6

[4]

MODELS TO STUDY GLIAL CELL CYTOTOXICITY

FIG. 6

(continued)

65

66

PARADIGMS OF NEURAL INJURY

FIG. 7 Photomicrographs of early (P23-P24) and late (P73) passage glial cells grown in DMEM + 10% FBS (A, early passage; B, late passage) or CDM + TIPS (C, early passage; D, late passage). (A) Cells are primarily glioblastic (large arrows), with some stellate cells representing mature-like glial cells (small arrows). (C) In contrast, cells are differentiated, mature stellate-typema mixture of astrocyte (large arrow) and oligodendrocyte-like (small arrow) cells. Morphology of late passage cells in B and D is not strikingly different except for a decrease in proliferation and apparent increase in cell size. Also, in D, the appearance of oligodendrocytic cells is evident (small arrows) Magnification: x590. From Lee, K., Kentroti, S., Billie, H., Bruce, C., and Vernadakis, A., Glia 6, 245-257 (1992), with permission.

is necessary to remove the glycerol from the cells. After 24 hours the medium is replaced with 10 ml fresh D M E M + 10% FBS. Culture medium is again changed (C3). Cells can be passed at C5 for stocks.

Characterization of Glial Phenotypes in C6 Glial Cell Cultures Biochemical Markers Using CNP and GS as biochemical markers in an early study, Parker et al. (1980) found that early passage (20-26) C6 glial cells (2B clone) exhibit high activity levels of CNP with low levels of GS and that late passage (80-88)

[4]

MODELS TO STUDY GLIAL C E L L CYTOTOXICITY

67

cells exhibit high activity levels of GS and low levels of CNP. From these studies it appears that early passage cells exhibit oligodendrocytic properties, whereas late passage cells exhibit astrocytic properties. Immunocytochemical Characterization of C6 Glial Cells C6 glial cells from passage 23 (early) or 111 (late) are plated on four-chamber plastic slides at a concentration of 20,000 cells/chamber in a volume of 0.3 ml DMEM + 10% FBS. Figure 8 shows early passage C6 glial cells grown in the presence (Fig. 8A-D) or absence (Fig. 8 E - H ) of serum (Lee et al., 1992). As noted in Fig. 8, we only considered cells that exhibited intense immunoreactivity, because faintly stained cells may express nonspecific fluorescein staining. In the presence of serum, many cells are A2B5 + and are assumed to be glioblastic progenitor cells. A few cells were A2B5 + GFAP +, indicating type 2 astrocytes (Fig. 8A and B), and, as expected, several cells were A2B5 + GalC + (Fig. 8C and D), indicating oligodendrocytes. In the absence of serum, most cells were A2B5 + GFAP + (Fig. 8E and F), indicating the differentiation to type 2 astrocyte. A2B5 + GalC + oligodendrocytic cells were very rare (Fig. 8G and H). In late passage C6 glial cells grown in either the presence or absence of serum, the predominant cells are A2B5 + GFAP +, indicating type 2 astrocytes (Fig. 9A, B, E, and F). Of importance was the presence of several A2B5 + GalC + in cultures grown in the absence of serum (Fig. 9G and H), suggesting that possible progenitor cells still present in the late passage cells differentiate into oligodendrocytes in the absence of serum, as has been reported by others (Raft et al., 1984).

Usefulness o f Early and Late Passage C6 Glial Cells Based on both biochemical and immunocytochemical characterizations, early passage C6 glial cells can serve as a model to study various conditions that shift phenotypic expression, oligodendrocytic versus astrocytic. The late passage cells can be used to study effects on a predominantly astrocytic population. As an example, a study by Mangoura et al. (1989) has shown similarities of early (20-22) and late (78-82) passage C6 glial cells with primary chick brain glial cells in culture. The conditions tested were culture substrata [collagen, poly(e-lysine), plastic] or supplements for the culture medium, i.e., DMEM (fetal bovine serum, heat-inactivated fetal bovine serum, or media conditioned from mouse neuroblastoma cells or primary chick embryo cultured neurons). GS and CNP were the biochemical glial markers used. The

68

PARADIGMS OF N E U R A L INJURY

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[4] MODELS TO STUDY GLIAL CELL CYTOTOXICITY

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findings showed that C6 glial cells are pluripotential and have the plasticity to express both astrocytic and oligodendrocytic properties, whereas the late passage cells are more committed to astrocytic expression.

Paradigm of Astrogliosis Using C6 Glial Cells as Models The potential role of glial cells in neuroinflammation and reactive gliosis was examined by testing the response of C6 glial cells to platelet-activating factor (PAF) (Kentroti et al., 1991). P A F (1-O-alkyl-2-acetyl-sn-glycero-3-phosphocholine) is a potent biologically active phospholipid found in various cells, including endothelial, mast, and kidney cells, as well as platelets and macrophages. It is normally released during inflammation and immune reactions to modulate cellular functions. In the central nervous system, PAF exerts modulatory effects on neuronal differentiation and calcium fluxes and its synthesis is enhanced by certain neurotransmitters. In this study, cells from passages 18-25 were designated early passage whereas cells from passages 152-167 were designated late passage. Cells were plated in D M E M + 10% fetal bovine serum on uncoated 100-mm polystyrene petri dishes (Lux) at a density of 0.5 x 106 cells/dish. Cultures were incubated at 37~ in an atmosphere of 7.5% CO2 in air, saturated with H20. Because PAF is rapidly metabolized in the presence of serum, and in order to assess the direct effect of test substances without the interaction of serum factors, media were replaced with chemically defined media (CDM) supplemented with transfer-

FIG. 8 Double-staining immunofluorescence labeling of C6 glial cells, 2B clone, early passage (P23), grown in chamber slides in DMEM + 10% FBS (A-D) or CDM + TIPS without serum (E-H), 2 days in culture. (A and B) Cells were stained with anti-GFAP (fluorescein optics in A) and anti-A2B5 (rhodamine optics in B). Note two A2B5 + cells (B), which also express GFAP (A) and are considered type 2 astrocytes. The remaining cells in A cannot be considered positive for GFAP when compared to the two cells that express both A2B5 and GFAP intense immunoreactivity. All other cells in B are exclusively A2B5 +. (C and D) Cells were stained with anti-GalC (fluorescein optics in C) and anti-A2B5 (rhodamine optics in D). Note three A2B5 + cells (D) that are also GalC + (C, arrows) and are considered oligodendrocytes; most other cells in D are exclusively A2B5+. Again, the faint stain in the remaining cells of C is considered nonspecific fluorescein staining. (E and F) Cells were stained with anti-GFAP (fluorescein optics in E) and anti-A2B5 (rhodamine optics in F). Note that marked difference, when compared to A and B, that most, if not all, A2B5 + cells also express GFAP and are considered type 2 astrocytes. (G and H) Cells were stained with anti-GalC (fluorescein optics in G) and anti A2B5 (rhodamine optics in H). Note three A2B5 + cells (H), which are also GalC + (G, arrows) and are considered oligodendrocytes. All other cells are exclusively A2B5 +. From Lee, K., Kentroti, S., Billie, H., Bruce, C., and Vernadakis, A., Glia 6,245-257 (1992), with permission.

70

PARADIGMS OF N E U R A L INJURY

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[4] MODELSTO STUDY GLIAL CELL CYTOTOXICITY

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rin, insulin, and penicillin/streptomycin (TIPS) after 24 hours in culture (C 1). At that time, cells were gently washed twice with D M E M and the media replaced with 6 ml CDM + TIPS. Cells were initially administered test substances at this time in culture. Groups of cultures (four to six/group) from early or late passages were treated at C1 with PAF (2-200 nM), lyso-PAF (3-300 nM), dBcAMP (1.0 mM), or RO20-1724 (0.1 mM). Cells were harvested at C4 with 0.2% trypsin, centrifuged for 5 minutes at 250 g and 4~ washed twice with Earle's balanced salt solution (EBS), and the pellet frozen at - 2 0 ~ for assay of GS activity. PAF increases GS activity in early passage (glioblastic) cells and, more importantly, it increases GS activity in late passage cells already committed to the astrocytic phenotype. Furthermore, cells from both passages fail to respond to addition of lyso-PAF, the nonbiologically active analog of PAF, to the medium. We compared PAF effects with that of dibutyryl cyclic AMP (dBcAMP) and RO20-1724, a phosphodiesterase inhibitor. Cells from the early passage responded to both dBcAMP and RO20-1724 treatments with a significant increase in GS activity, whereas late passage cells showed no significant change, confirming earlier reports from this laboratory. These findings indicate that the response of C6 glioma cells to PAF (at least in the late passage) is not mediated via cyclic AMP. We suggest that in early passage cells PAF promotes expression of the astrocytic phenotype, and in the late passage cells PAF mediates a gliosis-type response.

FIG. 9 Double-staining immunofluorescence labeling of C6 glial cells, 2B clone, late passage (P73), grown in chamber slides in DMEM + 10% FBS (A-D) or CDM + TIPS without serum (E-H), 2 days in culture. (A and B) Cells were stained with anti-GFAP (fluorescein optics in A) and anti-A2B5 (rhodamine optics in B). All A2B5 + cells (B) also express GFAP (A) and are considered type 2 astrocytes. (C and D) Cells were stained with anti-GalC (fluorescein optics in C) and anti-A2B5 (rhodamine optics in D). Note three A2B5 + cells (D) that are also GalC + (C, arrows) and are considered oligodendrocytes. One cell (D) appears to be A2B5 + and GalC + and is considered to be a precusor (p?). The remaining cells in panel C are faintly stained, probably nonspecific fluorescein staining. (E and F) Cells are stained with anti-GFAP (fluorescein optics in E) and anti-A2B5 (rhodamine optics in F). Note that all A2B5 + cells are also GFAP + and are considered type 2 astrocytes. Cells grown in CDM + TIPS (E, F) have a more differentiated stellate appearance than those grown in DMEM + 10% FBS (A, B). (G and H) Cells were stained with anti-GalC (fluorescein optics in G) and anti A2B5 (rhodamine optics in H). Note three A2B5 + cells (H) that also express GalC + (G, arrows) and are considered oligodendrocytes. From Lee, K., Kentroti, S., Billie, H., Bruce, C., and Vernadakis, A., Glia 6, 245-257 (1992), with permission.

72

PARADIGMS OF N E U R A L INJURY

Glial C y t o t o x i c i t y P a r a d i g m s

Ethanol Gliotoxicity and Relation to Glutamate Neurotoxicity There is now abundant evidence of the neuropathologic effects of ethanol both in the brains of chronic alcoholics and in children afflicted with fetal alcohol syndrome, the preponderance of these studies center on the structure, physiology, and biochemistry in neurons. The extent to which glial cells are vulnerable to ethanol has been less thoroughly examined [see review by Davies (1992)]. Studies from our laboratory using the chick embryo as an animal model have established that the critical period for ethanol neuroembryotoxicity is between embryonic days 1 and 3 (Brodie and Vernadakis, 1990; Kentroti and Vernadakis, 1992). Neuronotoxic effects of ethanol include decline in neuronal survival, shifts in neuronal phenotypic expression, neuronal migration, and neuronal maturation. In view of the role that astrocytes play in neuronal homeostasis from development to aging (Vernadakis, 1988; Muller, 1992; Abbott, 1991), the cytotoxicity of ethanol to glial cells may be an important factor on ethanol neuronotoxicity. One can investigate ethanol gliotoxicity using various models, including C6 glial cell cultures. We have reported ethanol gliotoxicity (Davies and Vernadakis, 1986) in late passage C6 glial cells because, as discussed previously, these late passage cells consist predominantly of an astrocytic phenotype. C6 glial cells at passage 76 (late passage) were plated at 1 x 10 6 cells per 100-mm dish in DMEM + 10% FBS. On culture day 3 (logarithmic growth) or on culture day 10 (postconfluency) the cultures were divided into control and ethanol-treated groups (five to six cultures per group). Medium is aspirated and replaced with DMEM + 5% FBS containing either 0.2, 0.5, or 1% ethanol. Ethanol-treated cultures are placed over a pan containing water plus ethanol up to 2% and the entire pan is enclosed in a thick plastic bag and sealed. We have found that this procedure maintains constant levels of ethanol in the culture medium for several days (Kentroti and Vernadakis, 1990). Control and ethanol-treated cultures can be removed after 24-72 hours and cultures are harvested with 0.1% trypsin for cell counts using a hemocytometer and also for biochemical analysis for GS activity. In this study, cultures treated with ethanol at C3 and C4 (logarithmic growth phase) were harvested at C5 whereas cultures treated with ethanol at C10 and C11 (confluent growth phase) were harvested at C12. The results obtained have shown that glial cells exposed to ethanol at postconfluency growth phase are more vulnerable to ethanol and exhibit a marked decrease in GS activity.

[4]

MODELS TO STUDY G L I A L C E L L CYTOTOXICITY

73

We have also studied ethanol gliotoxicity using glial cultures prepared from 15-day-old chick embryo cerebral hemispheres as described above (Davies and Vernadakis, 1984). Cultures are prepared as described above. On day 6, in vitro cultures are assigned to one of five groups: experimental cultures treated with DMEM + 10% FBS containing ethanol at one of four dosages [0.1, 0.5, 1.0, or 2.05 (w/v) (i.e., 21.7, 108.5, 217, or 434 mM)]. Control cultures receive the same culture medium without ethanol. Ethanoltreated cultures are placed over a pan containing water with 500 mM ethanol and the pan with cultures is placed in a plastic bag and sealed. Cellular maturation of cultures is followed by phase microscopy. The proliferation of cells is assessed on culture days 5, 7, 8, and 10 in the control group and the 0.5% and 1.0% ethanol groups. At each designated time point, four or five petri dishes from each group are harvested with 0.1% trypsin as described above. Viable cells are counted in a hemocytometer; the trypan blue exclusion method is employed to evaluate cell death. Cultures are harvested on day l0 in vitro for determination of GS activity and protein content (GS is expressed per mg/protein). Although the doses of ethanol appear high, they represent neurotoxicity doses in vivo. The striking finding in this study is the marked effect of 1% ethanol on cell n u m b e r ~ a n almost 50% decrease by day 10. Also, the surviving cells exhibit immature characteristics, i.e., appearing as flat, epitheloid cells. Of interest is the finding that GS activity is also markedly decreased, which is reflected by the decline in cell number. Another significant observation is the presence of reactive astrocytes exhibiting enlarged somata extending numerous branching processes. For comparison of the ethanol response of glial cells from various species, we present here the results of a study by Bass and Volpe (1988) using glia derived from newborn rat brain. Newborn Sprague-Dawley rats,

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pipetted over the cell layer, 2 ml/dish. After a 10-minute incubation at 37~ the radioactive medium is removed and cells are washed with 2 ml of 150 mM NaC1 (adjusted to pH 4.0) at room temperature for 30 seconds. Copper67 uptake for each concentration of 67Cu is expressed as picomoles Cu/mg protein/minute. A double reciprocal plot of velocity v e r s u s 67Cu concentration is determined to asses the apparent K m and gmax parameters. These data are shown in Fig. 1. A straight-line relationship is obtained by plotting 1/pmol Cu/mg protein/minute v e r s u s 1/[67Cu]. Values for apparent K m and gma x ar e calculated on the basis of the straight-line relationship of the Michaelis-Menten equation: llv-

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which graphically is equivalent to y = b + m x , where b is the intersecting point on the y axis (1/Vmax), and m is the slope, equal to K m / V m a x . The calculation of x when y equals zero sets x equal to - 1 / K m, which is an alternative and perhaps more convenient method for determining the K m parameter. Using the equations, one is able to derive values of K m -- 512 nM and Vmax = 3.57 pmol Cu/mg protein/minute for Cu transport into astroglial cells. Moreover, the linearity of the plot suggests the cells contain either

142

PARADIGMS OF N E U R A L I N J U R Y

a single binding site for copper in the membrane or multiple sites with identical binding affinities.

Time Dependence of 67Cu(II) Efflux Cultures are prepared as described, and washed with 2 ml DPBS, after which 2 ml of fresh DMEM/F12 serum-free medium containing 50 nM 67CUC12 is carefully pipetted over the cells. After a 50-minute incubation at 37~ the radioactive medium is removed and the cells are rinsed gently with 2 ml DPBS at room temperature for 30 seconds. Fresh DMEM/F12 medium (2 ml) is then pipetted over the cells and incubation is allowed to continue. After 5, 15, or 30 minutes of incubation at 37~ the cells are collected as above with 0.5 M NaOH and a cell scraper, and 67Cu retained in the cells is determined and expressed as picomoles Cu/mg protein. Copper-67 appearing in the medium with time is also monitored. Effluxing ceases in about 30 minutes. Any 67Cu remaining within the cells at 50 minutes is, therefore, regarded as nonexportable. The difference between cell-retained 67Cu at 50 minutes and time zero is used to calculate the exported fraction.

Determination of Kinetic Parameters (K m and Vmax):Efflux of 6ZCu(II) Plated cells are preloaded with 10, 20, 50, 100, or 200 nM of 67CUC12 in fresh DMEM/F12 serum-free medium at 37~ for 50 minutes. According to the Darwish assumption (14), the concentration of copper in the cell at the efflux site is equivalent to the concentration of copper in the medium once equilibrium has been established. After the incubation, the radioactive medium is removed and the cells are washed with 2 ml DPBS at room temperature for 30 seconds. Fresh DMEM/F12 medium (2 ml) is layered over the cells as before. Cell-retained 67Cu is determined at time zero and after a 5minute incubation at 37~ The efflux velocity is obtained from the difference between the two readings for each 67Cu load and expressed as picomoles Cu/mg protein/minute. Graphic analysis of the velocity versus 67Cu concentrations in the medium gives the kinetic curve of 67Cu eff~ux. The values for Km and Vmaxof 67Cu efflux are determined with a double-reciprocal plot (see preceding). The results of the kinetic analysis are shown in Fig. 2. Based on the values shown, astroglial cells have an efflux Km = 67.7 nM and an efflux gmax = 0.677 pmol Cu/mg protein/minute.

Interpretation and Limitations o f Kinetics Constants Determinations The above procedure estimates Km and Vmax transport constants by standard kinetic analysis and employs standard assumptions. Rates of uptake must be performed rapidly, preferably when only unidirectional flow of 67Cu into

[10] HEAVY METAL EFFECTS ON GLIA

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FIG. 2 Double-reciprocal plot of copper efflux in cultured C6 rat glioma cells. For this plot, y = 1.47 + 100.15x; r 2 = 1.00. Kinetics constants for copper efflux are K m = 67.7 nM and Vmax = 0.677 pmol Cu/mg protein/minute. Each point represents the mean of duplicate determinations. the cell is occurring. Binding of 67Cu to the membrane must also occur rapidly, but depends on the concentration of 67CHin the medium, i.e., below the level that would saturate the membrane binding sites. Transport is viewed as a two-step process with binding preceding the actual activation of the mobilizing factor. The method gives no insight into how the transport mechanism or carrier is functioning. It is essential that the 67Cu in the medium have free and open access to the membrane carrier and that other competing ligands be eliminated from the medium. Because serum contains albumin, and albumin has at least one high-affinity site for copper on the protein, the albumin must be removed. Cells, therefore, are suspended in serum-free medium for the duration of exposure to 67CH. Our interpretation of the data presented in Figs. 1 and 2 is that the import and export of copper by astroglia require two systems. The efflux system appears eight times more sensitive. This conclusion can be interpreted to mean that cells export copper with a greater affinity than they take it up. The imbalance is met by having Vmax for efflux only one-fifth as rapid (at saturation) as input. Because of the strong binding affinity of the efflux system, internal systems that require copper for metabolic function must compete with a highly efficient efflux system designed to keep the cell in homeostasis and to protect the cell from transient high environmental exposure. The remainder of the chapter will address methods for analyzing disruptions in cell homeostasis that result from exposure to metals in culture.

144

PARADIGMS OF NEURAL INJURY

A n a l y s i s o f Cell F u n c t i o n : C e l l u l a r F l u o r e s c e n c e I m a g i n g

Rationale for Use of Imaging Techniques Over the past several years, the convergence of several technologies has led to the development of new experimental approaches for investigating mechanisms of chemical toxicity. Innovations in cell culture strategies and a renaissance in light microscopy centered around fluorescence techniques have led to the development of a new generation of in vitro toxicity assays based on the quantification of fluorescent probes in living cells. The authors have been exploiting the powerful new technology of microscopic image analysis with vital fluorescent probes and interactive laser cytometry for the development of highly sensitive, mechanistically relevant assays for neurotoxicants. This technology has many potential advantages over traditional in vitro toxicity assays. First, the assays, which are based on the quantitation of fluorescent probes in individual cells and cultures, are noninvasive and are typically carried out with living cells in the culture dish. Second, the assays are highly sensitive and quantitative. Third, data can be collected from both individual cells and cell populations. This feature is valuable for screening out variant cells (such as those in mitosis) that may respond differently from the majority of cells to a toxin. Fourth, in many cases repeated measurements can be taken from individual cells as well as from cultures. Appropriate controls may also be run in the same culture dish because of the amenability of the system to repeated measurements. Fifth, the types of measurements obtained are highly mechanistic, and thus provide not only a screening device for the presence of a toxicant, but also information on the mechanisms of toxic action. Sixth, once assays have been developed and validated with an instrument such as the Meridian ACAS interactive laser cytometer (as described further in the next section), they should be readily adaptable to automated cytofluorometric assay systems.

Fluorescent Probes and Analytical Instrumentation Fluorescent probes have been created (and continue to evolve) that are designed to react with biomolecules under conditions of relatively low temperature (i.e., temperatures compatible with living cells) and near-neutral pH (15). Many fluorescent indicators of cellular function are uniquely suitable for probing living cells because of a combination of five properties (16): specificity, the ability to detect the probe in a complex mixture of biomolecules; sensitivity, the potential for detection of few molecules in a given volume; spectroscopy, differences in absorption or emission caused by sensi-

[10] HEAVY METAL EFFECTS ON GLIA

145

tivity to their immediate physicochemical environment; temporal resolution, the potential for fluorescence measurements over time points longer than the excitation/emission of the fluorescent probe, i.e., longer than 10-s seconds; and spatial resolution, defined by the resolving power of the microscope lens used to image the fluorescent probe. Many of the probes, such as the acetoxymethyl (AM) ester derivatives of a number of fluorescent indicators, may be noninvasively delivered (without microinjection or damage to the plasma membrane) into cells because of their membrane permeability properties. After entry into the cell the ester derivatives are subsequently cleaved by nonspecific cytosolic esterase activity to yield charged, membrane-impermeant probes. Other noninvasively delivered probes are membrane permeable but partition into cells based on charge, or are nonfluorescent until covalently bound to molecules within the cytoplasm. Coinciding with advances in fluorescent probe technology are developments in analytical microscopy instrumentation that permit the analysis of diverse molecular targets on or within cells. Technical improvements in microscopy include the development of lasers with emission lines suitable for use with many fluorescent probes, computer automation and control over the intensity and duration of sample illumination, and digital imaging photometric systems coupled with computer-controUed stage positioning, all of which combine to permit spatially resolved fluorescence signals. Modern fluorescence detection systems provide a means to balance the need for sufficient illumination with sensitivity, permitting excitation irradiance levels low enough to avoid damage to the sample while providing detectability equivalent to micromolar concentrations of fluorophores. There are a number of commercially available instruments capable of exciting and detecting fluorescent signals from living cells. The following description is restricted to our experience with an instrument manufactured by Meridian Instruments, Inc. (Okemos, MI), referred to as an ACAS 570 Interactive Laser Cytometer (ACAS is an acronym referring to the Adherent Cell Analysis and Sorting capabilities of the instrument). Relevant features of the instrument have been previously described (17) and include a Coherent Innova 90-5 5 W argon ion laser that produces ultraviolet (UV) illumination in the range of 351.1-363.8 nm and several visible lines throughout the range of 457.9-528.7 nm. The selected laser illumination line is directed through the rear illumination port of an Olympus IMT-2 inverted microscope equipped for epifluorescence. Fluorescence excitation with minimal destruction of fluorophores is optimized by regulation of the intensity and duration of the laser spot. Attenuation of the laser beam is controlled with two separate components. Neutral-density filters are inserted into the optical path to reduce beam intensity. The second component of beam attenuation employs an acoustooptic modulator (AOM). The AOM is used to separate the zero

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PARADIGMS OF NEURAL INJURY

and first diffracted order of light. Only the first diffracted order light is admitted to the optical system and the amount of first-order light is regulated by activation of the AOM. By application of a high-frequency, high-voltage signal to the AOM, changes in the Bragg angle of the crystal diverts the firstorder beam from the optical axis to attenuate the beam with potentially rapid (_ o O tD

rr

30

20

10 -

=

Fe4 Fe5 control

0

Time (rain) FIG. 12 Gap FRAP analysis of astroglial gap junctional communication after exposure to 0, l0 (FeS), or 100/~M (Fe4) FeC12 . In this experiment, cells were exposed daily for 7 days to the Fe dose indicated. For each datum 15-19 cells were measured. Error bars represent standard errors. At each time point measured (1-4 minutes) the mean percentage of recovery was significantly higher in the Fe4 group than in the other two groups. The rate constant (k) for diffusion of CFDA into the photobleached cell was calculated, which is proportional to the permeability of the channel. The k value was calculated to be 0.3425, 0.3784, and 0.5275 for the 0, FeS, and Fe4 groups, respectively. Thus the permeability of the gap junctional channels was increased by over 40% in cells treated with 100/zM Fe (Fe4) compared to the other treatment groups (p < 0.01).

Fe up-regulates gap junctional communication, as shown in Fig. 12. The dose of Fe (100/~M) selected was that which produces the same amount of metabolic injury in astroglia as 1 /~M Pb [as measured by reduction of glutamine synthetase (glutamate-ammonia ligase) specific activity]. The finding that Pb and Fe affect GJIC differently supports the concept that metals damage cells by distinct pathways, leaving a unique set of molecular end points, which we term their "signatures." It should be possible to develop a battery of sensitive assays to detect and characterize these unique signatures, particularly by the use of cellular fluorescence imaging techniques.

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Acknowledgments Work in the laboratories of E.T.-C., R.C.B., and E.D.H. has been supported by grants from the NIH (R01-ES05871, R01-HD26182, and R01-HD29959) and an NIH Superfund Grant to Dr. Stephen Safe (P42-ES04917). M.E.L. and L.A.S. are recipients of Physician Scientist Awards (K11-ES00251 and K11-ES00279) from the NIH. We thank Jeff Bowen and Michelle Nyberg for excellent assistance in the preparation of the manuscript.

References

.

9. 10.

11. 12. 13. 14. 15.

16.

D. M. Danks, in "The Metabolic Basis of Inherited Disease" (C. R. Scriver, A. I. Beaudet, W. S. Sly, and D. Valle, eds.), 6th Ed., p. 1411. McGraw-Hill, New York, 1989. M. J. Davis, NeuroToxicology 11, 285 (1990). E. Tiffany-Castiglioni, NeuroToxicology 14, 513 (1993). S. W. Levison and K. D. McCarthy, in "Culturing Nerve Cells" (G. Banker and K. Goslin, eds.), p. 309. MIT Press, Cambridge, Massachusetts, 1991. K. D. McCarthy, J. Pharmacol. Exp. Therap. 226, 282 (1983). E. Tiffany-Castiglioni, J. Zmudzki, J. N. Wu, and G. R. Bratton, Metab. Brain Dis. 2, 61 (1987). D. Holtzman, C. DeVries, H. Nguyen, J. I. Olson, and K. Bensch, NeuroToxicology 5, 97 (1984). J. A. Thomas, F. D. Dallenbeck, and M. Thomas, J. Pathol. 109, 45 (1973). C. Vandecasteele and C. B. Block (eds.), "Modern Methods for Trace Element Determination." Wiley, New York, 1993. T. J. Kneip and L. Friberg, "Handbook On The Toxicology of Metals" (L. Friberg, G. F. Nordberg, and V. B. Vouk, eds.), 2nd Ed., Vol. I, p. 44. Elsevier, New York, 1986. Thermo Jarrel Ash Corp., "Methods Manual for Furnace Operation," Vol. II, 1993. T. J. Goka, R. E. Stevenson, P. M. Hefferan, and R. R. Howell, Proc. Natl. Acad. Sci. U.S.A. 73, 604 (1976). H. Kodama, Y. Meguro, T. Abe, M. H. Rayner, K. Y. Suzuki, and S. Kobayashi, J. Inherit. Metab. Dis. 14, 896 (1991). H. Y. Darwish, R. C. Schmitt, J. C. Cheney, and M. J. Ettinger, Am. J. Physiol. 246, 648 (1984). R. P. Haugland, "Molecular Probes: Handbook of Fluorescent Probes and Research Chemicals" (K. D. Larison, ed.). Molecular Probes Inc., Eugene, Oregon, 1992. D. L. Taylor and E. D. Salmon, in "Fluorescence Microscopy of Living Cells in Culture" (Y. Wang and D. L. Taylor, eds.), Part A, p. 207. Academic Press, San Diego, 1989.

166

PARADIGMSOF NEURAL INJURY 17. M. Schindler, M. H. Allen, M. R. Olinger, and J. F. Holland, Cytometry 6, 368 (1985). 18. R. C. Burghardt, R. Barhoumi, D. Doolittle, and T. D. Phillips, in "Principles and Methods of Toxicology" (A. W. Hayes, ed.), 3rd Ed., p. 1231. Raven Press, New York, 1994. 19. A. Meister, Pharmacol. Therap. 51, 155 (1991). 20. S. P. Raps, J. C. K. Lai, L. Hertz, and A. J. L. Cooper, Brain Res. 545, 312 (1989). 21. A. Jain, J. Martensson, E. Stole, P. A. M. Auld, and A. Meister, Proc. Natl. Acad. Sci. U.S.A. 88, 1913 (1991). 22. D. Holtzman, Toxicol. Appl. Pharmacol. 89, 211 (1987). 23. P. Goering, NeuroToxicology 14, 45 (1993). 24. P. S. Rabinovitch, C. H. June, and T. J. Kavanagh, in "Clinical Flow Cytometry: Principles and Application" (U. D. Bauer, R. E. Dunque, and T. V. Shanley, eds.), p. 505. Williams and Wilkins, Baltimore, Maryland, 1993. 25. R. Barhoumi, J. A. Bowen, L. S. Stein, J. Echols, and R. C. Burghardt, Cytometry 14, 747 (1993). 26. M. E. Legare, R. Barhoumi, R. C. Burghardt, and E. Tiffany-Castiglioni, NeuroToxicology 14, 267 (1993). 27. B. Ehrenberg, V. Montanta, M. D. Wei, J. P. Wuskell, and L. M. Loew, Biophys. J. 53, 785 (1988). 28. J. R. Bunting, T. V. Phan, E. Kamali, and R. M. Dowben, Biophys. J. 56, 979 (1989). 29. M. Poot, T. J. Kavanagh, H. C. Kang, R. P. Haugland, and P. S. Rabinovitch, Cytometry 12, 184 (1991). 30. T. J. B. Simons, NeuroToxicology 14, 77 (1993). 31. R. Y. Tsien, Rev. Neurosci. 12, 227 (1989). 32. R. C. Burghardt, R. Barhoumi, E. Lewis, R. H. Bailey, K. Pyle, B. Clement, and T. D. Phillips, Toxicol. Appl. Pharmacol. 112, 235 (1992). 33. M. E. Legate, R. Barhoumi, E. Hebert, R. C. Burghardt, and E. Tiffany-Castiglioni, in preparation. 34. F. A. X. Schanne, T. L. Dowd, R. K. Gupta, and J. F. Rosen, Proc. Natl. Acad. Sci. U.S.A. 86, 51 (1989). 35. J. L. Tomsig and J. B. Suszkiw, Am. J. Physiol. 259, C762 (1990). 36. J. J. Anders, Glia 1, 371 (1988). 37. E. L. Hertzberg and R. G. Johnson (eds.), "Gap Junctions of Modern Cell Biology." Alan R. Liss, New York, 1988. 38. A. H. Cornell-Bell, S. M. Finkbeiner, M. S. Cooper, and S. J. Smith, Science 247, 470 (1990). 39. W. R. Loewenstein, Am. Rev. Respir. Dis. 142, $48 (1990). 40. M. H. Wade, J. E. Trosko, and M. Schindler, Science 232, 525 (1986). 41. L. S. Stein, J. G. Boonstra, and R. C. Burghardt, In Vitro Cell Dev. Biol. 28A, 436 (1992). 42. S. O. Mikalsen, Carcinogenesis 11, 1621 (1990).

[11]

Source, Metabolism, and Function of Cysteine and Glutathione in the Central Nervous System David K. Rassin

Introduction In recent years there has been an explosion of interest in the role of glutathione; this tripeptide (y-glutamylcysteinylglycine) has been associated with antioxidant function (reviewed in Refs. 1-3), immunologic functions (4, 5), the disease process associated with human immunodeficiency syndrome (HIV) infection (6, 7), and modulation of neurotransmitter function in the central nervous system (CNS) (8, 9). Of particular interest with respect to neural injury are the antioxidant and excitatory amino acid modulatory roles of glutathione. Glutathione may directly modify oxidant stress or it may modulate release of excitatory amino acids by influencing the function of the N-methyl-D-aspartate (NMDA) receptor (8, 9). In addition, glutathione and cyst(e)ine may prevent excitatory amino acid-induced apoptosis by protecting against intracellular glutathione depletion (10, 11). Glutathione is readily synthesized in mammalian cells from its three precursor amino acids, glutamate, cysteine, and glycine. However, glutathione does appear to be most dependent on the availability of cysteine in order to maintain organ pools (12, 13). Cysteine may fill many of the functions supported by glutathione, reflecting the common structural feature of these two compounds, the sulfhydryl group. Cysteine is usually synthesized from the essential amino acid methionine via the transsulfuration pathway (Fig. 1); however, there are several circumstances, particularly during early development, wherein cysteine may have to be supplied in the diet due to the low hepatic activity of cystathionase, the enzyme responsible for catalyzing cysteine synthesis (12, 13). In the following presentation the origin and metabolism of cysteine and glutathione will be discussed with attention to some of the methods that have been particularly useful in investigating their characteristics.

Cysteine Cysteine is a sulfur-containing amino acid that is synthesized from the essential amino acid methionine via the transmethylation and transsulfuration pathways (Fig. 1). Cysteine may be made available to the central nervous Methods in Neurosciences, Volume 30 Copyright 9 1996 by Academic Press, Inc. All rights of reproduction in any form reserved.

167

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PARADIGMS OF N E U R A L INJURY

PROTEIN


Giu ~

4

FIG. 3 Some aspects of the metabolism of glutathione. 1, y-Glutamyltransferase; 2, y-glutamylcyclotransferase; 3, 5-oxoprolinase; 4, y-glutamylcysteine synthetase (glutamate-cysteine ligase); 5, glutathione synthetase; 6, dipeptidase; 7, glutathione reductase; 8, peroxidases.

(2), as a modulator of the N-methyl-D-aspartate receptors (8, 9), and as a component of the 7-glutamylamino acid transport cycle in the CNS (32) (see Fig. 3). The role of glutathione in protecting against antioxidant damage has become more appreciated with the implication of oxidant stress in the neuropathology associated with hypoxia, hyperoxia, Alzheimer's disease, Parkinson's disease, and aging (33-39). The relative amounts of reduced (GSH) and oxidized (GSSG) glutathione may serve as markers of oxidant stress, thus in situations of oxidative attack the GSSG/GSH ratio increases as an index of such stress. Protection against oxidative stress depends on maintenance of intracellular GSH concentrations, and such maintenance depends on the availability of precursor cysteine (21). Glutathione may be protective both as a function of its role in modulating the excitotoxic effects of glutamate (10-11) and as a function of its role in protecting against the metabolic continuum, from oxidant attack to energy depletion to neurotrophin disruption to cell death in the CNS (40). These considerations have emphasized the need for measures of glutathione, oxidized and reduced, in the CNS. Such methodology has been reviewed

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(41-42), and the two most useful techniques are the spectrophotometric method developed by Tietze (43) as modified by Griffith (44), and highperformance liquid chromatographic methodology using dual-cell electrochemical detection (45).

Spectrophotometric Assay of Glutathione Total and oxidized glutathione are measured using an enzymatic recycling method (based on glutathione reductase) and the GSH blocking agent, 2vinylpyridine (43, 44). Tissues are homogenized directly in 3% sulfosalicylic acid (w/v) (preferable technique) or first in a phosphate buffer and then treated with sulfosalicylic acid. Rapid preparation is necessary to minimize autoxidation of glutathione to the disulfide form. The protein precipitate is removed by centrifugation (17,000 g for 15 minutes) at 5~ and the supernatant solution is used for the glutathione assay. Four tubes are prepared for each sample; 200/zl of unknown is added to each tube. Two tubes have 4/zl of 2-vinylpyridine added (this should be a clear liquid; obtain fresh material if an amber color is present). Add 12/~1 of triethanolamine to all tubes to raise the pH to 7-7.5. Mix for 1 minute and let stand at room temperature for 60 minutes. Place 200 ~1 of sample into a 1.5-ml cuvette, add 700/zl of NADPH (0.3 mM in EDTA-phosphate buffer, pH 7.5), 100/zl of DTNB (6.0 mM in EDTA-phosphate buffer, pH 7.5), and 10/zl of glutathione reductase (50 units/ml), invert cuvette two or three times to mix, place immediately into a recording spectrophotometer set at a wavelength of 412 nm, and measure change in absorbance for 120 seconds. The slope of the line is calculated and compared to a standard curve prepared from slopes given by different known amounts of glutathione. Glutathione measured in the tubes with added 2-vinylpyridine represents the oxidized form and is subtracted from tubes without 2-vinylpyridine (total glutathione) to calculate the amount of reduced glutathione.

HPLC Assay of Glutathione In cases in which direct measurement of oxidized and reduced glutathione in the presence of cysteine and cystine is desired at high sensitivity, highperformance liquid chromatography with dual-cell electrochemical detection is useful. Tissue samples may be prepared as described above in sulfosalicylic acid. These samples are then injected onto a 5-~m C18 reversed-phase cartridge column (22 cm • 4.6 mm; Pierce, Rockford, IL) with a 3-cm guard column packed with the same stationary phase. Compounds are eluted with

Ill]

173

CSH A N D G S H IN T H E CNS

A

B

=

-lto

to co

,+ -r to

cO to to

!

03

UPSTREAM ELECTRODE OFF

to to

-r" to (.9 r

, 9

. L'~

UPSTREAM ELECTRODE ON

F I G . 4 The separation and detection of glutathione and cyst(e)ine by HPLC with dual-cell electrochemical detection. (A) Upstream electrode is off, so only the already reduced (sulfhydryl) compounds are detected; (B) upstream electrode is on, and all four forms of the two compounds are detected.

96% monochloroacetic acid at pH 3.0 and 4.0% methanol containing 1.0 mM sodium octyl sulfate as an ion-pairing agent. Dual, serially assembled mercury-gold electrodes are prepared (46). Cysteine, cystine, oxidized glutathione, and reduced glutathione separate on the column and then are all modified to their reduced forms by the first (upstream) electrode set at - 1 . 0 V. The detector (downstream) electrode, set at 0.15 V, then is used for quantitation based on the oxidation potential of the reduced compounds (Fig. 4).

Inhibitory Agents Agents that inhibit the synthesis of glutathione have been particularly useful in elucidating the metabolism of this tripeptide (47). The most widely used inhibitor has been buthionine sulfoximine (BSO), an analog of the glutamine synthetase (glutamate-ammonia ligase) inhibitor methionine sulfoximine. Administration of BSO effectively inhibits y-glutamylcysteine synthetase by binding to the enzyme. BSO effectively reduces GSH synthesis in brain of newborn animals, but is not as effective in adult animals, presumably due

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PARADIGMS OF N E U R A L INJURY

to poor transport of the agent across the blood-brain barrier (48). Administration of BSO directly to the brain and in solutions of dimethyl sulfoxide (DMSO) represent alternative techniques to bypass the problem of transport into the CNS, and are effective in reducing glutathione synthesis (49, 50). A second pharmacologic agent that has been useful in the manipulation of both cysteine and glutathione is the cystathionase inhibitor, propargylglycine (PPG). PPG inhibits cystathionase noncompetitively and irreversibly, limiting the conversion of cystathionine to cysteine (51, 52). Adult rats treated with PPG (40/xmol/day) for 15 days had about a 50% reduction in plasma cystine with a 200-fold increase in plasma cystathionine, the immediate cysteine precursor (53). Brain glutathione was reduced by approximately 20% in these studies; unfortunately the cystathionine and cysteine in brain were not reported. Other studies have shown that animals treated with increased dietary methionine accumulate cystathionine in the central nervous system, reflecting a possible metabolic block and indicating that peripheral cysteine synthesis may be required to support transport of cysteine into the CNS followed by in situ synthesis of glutathione in the CNS (27, 28, 54). Several repletion agents are available to assess the specificity of the inhibitory agents noted above. Poor absorption of glutathione and cysteine oxidation to the more insoluble cystine are factors that complicate direct use of these metabolites. However, glutathione esters, particularly the monoester, appear to be readily absorbed and efficiently replete glutathione when it is depleted by BSO treatment (55). In like manner the cysteine analog, Nacetylcysteine, is readily absorbed and converted to cysteine, in which form it also can further promote glutathione synthesis (56-58). Thus, the tools to deplete and replete cysteine and glutathione are available for further investigation of the roles of these compounds in the central nervous system.

Conclusion Glutathione and cysteine have important roles as antioxidant protective agents in the CNS. The increasing evidence that protection of CNS tissue from oxidative stress is an important mechanism in preventing neuronal cell death is leading to better understanding of some of the degenerative processes that attack the CNS. Two studies have investigated mechanisms of oxidatively induced cell death or apoptosis in cultured embryonic cortical neurons (10, 11). Hypothesizing that oxidant cell death is mediated through excitatory amino acids, these studies explored the role of glutathione and cysteine in these cells on treatment with homocysteate or glutamate (11). The protective effect of N-acetylcysteine and depletion of glutathione by homocysteate led

[11] CSH AND GSH IN THE CNS

175

to the suggestion that cells are protected by cystine uptake into the cell and conversion to glutathione (11). The above described protective mechanisms suggest that if indeed diseases such as Alzheimer's and amyotrophic lateral sclerosis involve oxidative stress and cell death, antioxidant administration may offer some hope of ameliorating the damage. Antioxidant maintenance in the CNS appears to require cyst(e)ine transport followed by glutathione synthesis in situ. Thus, agents that promote CNS glutathione synthesis may be important in altering the cascade of events that is initiated by any form of oxidative stress, and maintaining homeostasis requires an appropriate supply of cyst(e)ine.

References 1. A. Meister, in "Functions of Glutathione: Biochemical, Physiological, Toxicological, and Clinical Aspects" (A. Larsson, S. Orrenius, A. Holmgren, and B. Mannervik, eds.), pp. 1-22. Raven Press, New York, 1983. 2. A. Meister, in "Glutathione: Chemical, Biochemical, and Medical Aspects" (D. Dolphin, O. Avramovi6, and R. Poulson, eds.), Part A, pp. 1-48. Wiley, New York, 1989. 3. M. Taniguchi, K. Hirayama, K. Yamaguchi, N. Tateishi, and M. Suzuki, in "Glutathione: Chemical, Biochemical, and Medical Aspects" (D. Dolphin, R. Poulson, and O. Avramovi6, eds.), Part B, pp. 645-727. Wiley, New York, 1989. 4. W. Dr6ge, H. P. Eck, H. GmOnder, and S. Mihm, Am. J. Med. 91, 140S144S (1991). 5. M. K. Robinson, M. L. Rodrick, D. O. Jacobs, J. D. Rounds, K. H. Collins, I. B. Saporoschetz, J. A. Mannick, and D. W. Wilmore, Arch. Surg. 128, 29-35 (1993). 6. R. Buhl, K. J. Holroyd, A. Mastrangeli, A. M. Cantin, H. A. Jaffe, F. B. Wells, C. Saltini, and R. G. Crystal, Lancet 2, 1294-1297 (1989). 7. S. Mihm, J. Ennen, U. Pessara, R. Kurth, and W. Dr6ge,AIDS 5,497-503 (1991). 8. D. I. Levy, N. J. Sucher, and S. A. Lipton, NeuroReport 2, 345-347 (1991). 9. N. J. Sucher and S. A. Lipton, J. Neurosci. Res. 30, 582-591 (1991). 10. R. R. Ratan, T. H. Murphy, and J. M. Baraban, J. Neurosci. 14, 4385-4392 (1994). 11. R. R. Ratan, T. H. Murphy, and J. M. Baraban, J. Neurochem. 62, 376-379 (1994). 12. A. Meister, Nutr. Rev. 42, 397-410 (1984). 13. A. Meister, in "Glutathione: Chemical, Biochemical, and Medical Aspects" (D. Dolphin, O. Avramovi6, and R. Poulson, eds.), Part A, pp. 367-474. Wiley, New York, 1989. 14. J. A. Sturman, G. E. Gaull, and N. C. R~iih~i, Science 169, 74-76 (1970). 15. G. E. Gaull, J. A. Sturman, and N. C. R. R~iih~i,Pediatr. Res. 6, 538-547 (1972). 16. H. H. Tallan, S. Moore, and W. H. Stein, J. Biol. Chem. 230, 707-716 (1958). 17. T. L. Perry, K. Berry, S. Hansen, S. Diamond, and C. Mok, J. Neurochem. 18, 513-519 (1971).

176

PARADIGMSOF NEURAL INJURY 18. S. H. Mudd, J. D. Finkelstein, F. Irreverre, and L. Laster, J. Biol. Chem. 240, 4382-4392 (1965). 19. J. A. Sturman, D. K. Rassin, and G. E. Gaull, Int. J. Biochem. 1, 251-253 (1970). 20. R. K. Shaw and J. D. Heine, J. Neurochem. 12, 151-155 (1965). 21. N. Tateishi, T. Higashi, A. Naruse, K. Nakashima, H. Shiozake, and Y. Sakamoto, J. Nutr. 107, 51-60 (1977). 22. E. S. Cho, N. Sahyoun, and L. D. Stegink, J. Nutr. 111, 914-922 (1981). 23. M. H. Malloy, D. K. Rassin, and G. E. Gaull, Anal. Biochem. 113,407-415 (1981). 24. T. L. Perry, S. Hansen, K. Berry, C. Mok, and D. Lesk, J. Neurochem. 18, 521-528 (1971). 25. T. L. Perry, H. D. Sanders, S. Hansen, D. Lesk, M. Kloster, and L. Gravlin, J. Neurochem. 19, 2651-2656 (1972). 26. H. H. Tallan, D. K. Rassin, J. A. Sturman, and G. E. Gaull, in "Handbook of Neurochemistry" (A. Lajtha, ed.), 2nd Ed., Vol. 3, pp. 535-558. Plenum, New York, 1983. 27. M. H. Malloy and D. K. Rassin, Pediatr. Res. 18, 747-751 (1984). 28. M. H. Malloy, D. K. Rassin, W. C. Heird, and G. E. Gaull, Am. J. Clin. Nutr. 34, 1520-1525 (1981). 29. M. K. Gaitonde, Biochem. J. 104, 627-633 (1967). 30. G. E. Gaull, D. K. Rassin, and J. A. Sturman, Neuropdediatrie 1, 199-266 (1969). 31. M. Das, R. Dixit, P. K. Seth, and H. Mukhtar, J. Neurochem. 36, 1439-1442 (1981). 32. A. Meister, Science 180, 33-39 (1973). 33. J. D. Adams, Jr., L. K. Klaidman, I. N. Odunze, H. C. Shen, and C. A. Miller, Mol. Chem. Neuropathol. 14, 213-226 (1991). 34. G. Benzi, F. Marzatico, O. Pastoris, and R. F. Villa, J. Neurosci. Res. 26, 120-128 (1990). 35. A. Meister, Biochem. Pharmacol. 44, 1905-1915 (1992). 36. K. Kramer, H.-P. Voss, J. A. Grimbergen, C. Smink, H. Timmerman, and A. Bast, Gen. Pharmacol. 23, 105-108 (1992). 37. A. Pean, G. B. Steventon, R. H. Waring, H. Foster, S. Sturman, and A. C. Williams, J. Neurol. Sci. 124, 59-61 (1994). 38. M. A. Verity, NeuroToxicol. 15, 81-92 (1994). 39. S. Fahn and G. Cohen, Ann. Neurol. 32, 804-812 (1992). 40. Z. Pan and R. Perez-Polo, J. Neurochem. 61, 1713-1721 (1993). 41. R. C. Fahey, in "Glutathione: Chemical, Biochemical, and Medical Aspects" (D. Dolphin, O. Avramovi6, and R. Poulson, eds.), Part A, pp. 303-307. Wiley, New York, 1989. 42. M. E. Anderson, in Glutathione: Chemical, Biochemical, and Medical Aspects" (D. Dolphin, O. Avramovi6, and R. Poulson, eds.), Part A, pp. 339-365. New York, 1989. 43. F. Tietze, Anal. Biochem. 27, 502-522 (1969). 44. O. W. Griffith, Anal. Biochem. 106, 207-212 (1980). 45. L. A. Allison, J. Keddington, and R. E. Shoup, J. Liq. Chromatog. 6, 17851798 (1983). 46. Anonymous, LCEC Application Note No. 53, pp. 1-3 Bioanalytical Systems Inc., West Lafayette, Indiana, 1983.

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47. A. Meister, Pharmacol. Therap. 51, 155-194 (1991). 48. O. W. Griffith and A. Meister, Proc. Natl. Acad. Sci. U.S.A. 76, 5606-5610 (1979). 49. E. Pileblad and T. Magnusson, Neurosci. Lett. 95, 302-306 (1988). 50. R. Steinherz, J. M~trtensson, J. Wellner, D. and A. Meister, Brain Res. 518, 115-119 (1990). 51. W. Washtien and R. H. Abeles, Biochemistry 16, 2485-2491 (1977). 52. R. H. Abeles and C. T. Walsh, J. Am. Chem. Soc. 95, 6124-6125 (1973). 53. E. S. Cho, J. Hovanec-Brown, R. J. Tomanek, and L. D. Stegink, J. Nutr. 121, 785-794 ( 1991). 54. D. K. Rassin, in "Absorption and Utilization of Amino Acids" (M. Friedman, ed.), Vol. II, pp. 71-85. CRC Press, Boca Raton, Florida, 1989. 55. J. M~trtensson, R. Steinherz, A. Jain, and A. Meister, Proc. Natl. Acad. Sci. U.S.A. 86, 8727-8731 (1989). 56. R. J. Flanagan and T. J. Meredith, Am. J. Med. 91, 131S-139S (1991). 57. R. Ruffmann and A. Wendel, Klin. Wochenschr. 69, 857-862 (1991). 58. M. F. Banks and M. H. Stipanuk, J. Nutr. 124, 378-387 (1994).

[12]

Magnetic Resonance Spectroscopy of Neural Tissue R i c h a r d J. M c C l u r e , K a n a g a s a b a i P a n c h a l i n g a m , William E. K l u n k , a n d J a y W. P e t t e g r e w *

Introduction Magnetic resonance spectroscopy (MRS) is a powerful physical technique for examination of the quantity and structure of organic molecules in solution and in vivo to monitor metabolites noninvasively in intact animals and humans. MRS provides useful information about high-energy and phospholipid metabolism from observations of their characteristic metabolites. In this chapter we will (1) cover the key topics necessary to understand the MRS procedures used to examine neural tissue both in vitro and in vivo, (2) present 1H and 31p MRS methods to examine metabolite levels in autopsy brain tissue, and (3) discuss in vivo 31p MRS methods to monitor metabolite levels in Alzheimer's disease (AD) brain. The examples given to illustrate these procedures have been taken from studies done in this laboratory.

Terms and Theory This section provides an introduction to MRS terms and theory. More detailed explanations can be found in several reviews (1-3).

Nuclear Magnetic Moment The property of the atomic nucleus that allows it to produce an MRS signal in a magnetic field is its magnetic moment, a property arising from the intrinsic nuclear spin caused by an uneven number of protons and neutrons in the nucleus. In the nucleus, the proton and neutron spins combine to give a net spin (angular momentum) to the nucleus. In a number of nuclei, such as those in 1H, 13C, 19F, and 31p, the charges are distributed uniformly and

* T o w h o m c o r r e s p o n d e n c e s h o u l d be a d d r e s s e d .

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Spherical Non-Spinning Nucleus: !~ = 0 eQ = 0

Spherical Spinning Nucleus: I~ • 0 eQ = 0

Ellipsoidal (Prolate) Spinning Nucleus: I~ ~ 0 eQ>0

Ellipsoidal (Oblate) Spinning Nucleus: !~ • 0 eQ 120) or moderate (60-119). The results of this study demonstrate that PME levels of the mildly demented group are increased compared to the moderately demented group. Also, PME levels correlate negatively with the clinical rating, the Mattis

205

[12] MRS OF NEURAL TISSUE I

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Fro. 12 Distribution of phosphomonoester (PME) levels (mole%) by Mattis score for Alzheimer's disease patients. The p and r values represent significance level and goodness of fit, respectively. Reprinted from Neurobiol. Aging 15, Pettegrew, Panchalingam, Klunk, McClure, and Muenz. Alterations of cerebral metabolism in probable Alzheimer's Disease: A preliminary study, 117-132, Copyright 1994, with kind permission from Elsevier Science Ltd, The Boulevard, Langford Lane, Kidlington 0X5 1GB, UK. score (Fig. 12). Taken together, these results indicate that the milder the dementia the greater the PME levels. This suggests that alterations in membrane phospholipid metabolism could be an early "molecular trigger" in AD, perhaps resulting in alterations to mitochondrial membranes as well as to the plasma membrane. Levels of PCr were decreased in the mildly demented AD subjects and increased as the dementia worsened. The data indicate that both of the immediate precursors of ATP (PCr and ADP) are diminished early in AD. This decrease in energy availability and reserve may lead directly to neuronal dysfunction and possibly place neurons at more risk of neurotoxic insult from glutamate (32).

In Vivo 31p M R S Studies of Acetyl-L-Carnitine Treatment of AD Brain In vivo MRS is a technique well suited to monitor the course of treatment of AD brain. We have monitored the effect of the administration of acetyl-Lcarnitine to AD patients by in vivo 31p MRS using the depth-pulse localization technique. In a double-blind, placebo study, acetyl-L-carnitine was administered to 7 probable Alzheimer's disease patients who were then compared by clinical and 3~p magnetic resonance spectroscopic measures to 5 placebotreated probable AD patients and 21 age-matched healthy controls over the course of 1 year (7). Compared to AD patients on placebo, acetyl-L-carnitinetreated patients showed significantly less deterioration in their Mini-Mental

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Status (MMS) and Alzheimer's Disease Assessment Scale test scores. Furthermore, the decrease in phosphomonoester levels observed in both the acetyl-L-carnitine and placebo AD groups at entry was normalized in the acetyl-L-carnitine-treated but not the placebo group. Similar normalization of high-energy phosphate levels were observed in the acetyl-L-carnitinetreated but not placebo-treated patients. This is the first direct in vivo demonstration of a beneficial effect of a drug on both clinical and CNS neurochemical parameters in AD. Acetyl-L-carnitine-treated patients showed significantly less deterioration in their MMS scores compared to AD patients on placebo. Although MMS scores were equivalent at entry in the two AD groups (p = 0.97), acetyl-Lcarnitine-treated patients had significantly higher scores than placebo-treated patients at 6 (p = 0.01) and 12 months (p = 0.01). This is the first direct in vivo demonstration of a beneficial effect of a drug on both clinical and CNS neurochemical parameters in AD.

Conclusion MRS spectroscopy provides useful information about the status of highenergy and phospholipid metabolism from observations of characteristic metabolites. MRS is useful for studying AD brain, both in vivo and in vitro. MRS findings support the suggestion that there is derangement of both the membrane structure and the metabolism of characteristic membrane lipids, which would contribute to deranged cellular functions. In vivo MRS is uniquely suited to identify the molecular underpinnings of neural injury and to monitor the efficacy of treatment protocols.

Acknowledgments This work was supported in part by NIA Grants AG08371, AG08974, AG50133, and AG9017.

References 1. J. W. Pettegrew, in "Handbook of Neuropsychology" (F. Boller and J. Grafman, eds.), pp. 39-56. Elsevier, Amsterdam, 1991. 2. "NMR: Principles and Applications to Biomedical Research" (J. W. Pettegrew, ed.) Springer-Verlag, Berlin and New York, 1990.

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,

10.

11. 12. 13. 14. 15. 16. 17. 18. 19. 20.

21. 22.

23. 24.

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"Spectrometric Identification of Organic Compounds." (R. M. Silverstein, G. C. Bassler, and T. C. Morril, eds.), 5th Ed. Wiley, New York, 1991. K. Panchalingam, S. Sachedina, J. W. Pettegrew, and T. Glonek, Int. J. Biochem. 23, 1453-1469 (1991). R. Vink, Mol. Chem. Neuropathol. 18, 279-297 (1993). M. R. Bendall, Bull. Magn. Reson. 8, 17-44 (1986). A. A. Maudsley, D. B. Tweig, D. Sappey-Marinier, B. Hubesch, J. W. Hugg, G. B. Matson, and M. W. Weiner, Magn. Reson. Med. 14, 415-422 (1990). K. O. Lim, J. Pauly, P. Webb, R. Hurd, and A. Macovski, Magn. Reson. Med. 32, 98-103 (1994). W. P. Aue, Magn. Reson. Med. 1, 21-72 (1986). L. Bolinger and R. E. Lenkinski, in "Biological Magnetic Resonance 11, In Vivo Spectroscopy" (L. J. Berliner and J. Reuben, eds.), pp. 1-53. Plenum, New York, 1992. T. Glonek, S. J. Kopp, E. Kot, J. W. Pettegrew, W. H. Harrison, and M. M. Cohen, J. Neurochem. 39, 1210-1219 (1982). J. W. Pettegrew, K. Panchalingam, G. Withers, D. McKeag, and S. Strychor, J. Neuropathol. Exp. Neurol. 49, 237-249 (1990). W. E. Klunk, K. Panchalingam, J. Moossy, R. J. McClure, and J. W. Pettegrew, Neurology 42, 1578-1585 (1992). J. W. Pettegrew, G. Withers, K. Panchalingam, and J. F. Post, Magn. Reson. Imaging 6, 135-142 (1988). J. H. Ackerman, T. H. Grove, G. G. Wong, D. G. Gadian, and G. K. Radda, Nature (London) 283, 167-170 (1980). H. Bruhn, J. Frahm, M. L. Gyngell, K. D. Merboldt, W. Hanicke, and R. Sauter, Magn. Reson. Med. 9, 126-131 (1989). J. W. Pettegrew, M. S. Keshavan, K. Panchalingam, S. Strychor, D. B. Kaplan, M. G. Tretta, and M. Allen, Arch. Gen. Psychiatry 48, 563-568 (1991). G. G. Brown, S. R. Levine, J. M. Gorell, J. W. Pettegrew, J. W. Gdowski, J. A. Bueri, J. A. Helpern, and K. M. Welch, Neurology 39, 1423-1427 (1989). J. W. Pettegrew, K. Panchalingam, W. E. Klunk, R. J. McClure, and L. R. Muenz, Neurobiol. Aging 15, 117-132 (1994). W. E. Klunk, M. Keshavan, K. Panchalingam, and J. W. Pettegrew, in "American Psychiatric Press Review of Psychiatry" (J. M. Oldham, M. B. Riba, and A. Tasman, eds.), Vol. 12, pp. 383-419. American Psychiatric Press, Washington, D.C., 1993. W. E. Klunk, C. J. Xu, K. Panchalingam, R. J. McClure, and J. W. Pettegrew, Neurobiol. Aging 15, 133-140 (1994). M. Baramy and T. Glonek, in "Methods of Enzymology" (D. L. Frederiksen and L. W. Cunningham, eds.), Part B, Vol. 85, pp. 624-676. Academic Press, New York, 1982. M. M. Cohen, J. W. Pettegrew, S. J. Kopp, N. Minshew, and T. Glonek, Neurochem. Res. 9, 785-801 (1984). P. A. Bottomley, J. P. Cousins, D. L. Pendrey, W. A. Wagle, C. J. Hardy, F. A. Eames, R. J. McCaffrey, and D. A. Thompson, Radiology 183, 695-699 (1992).

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PARADIGMS OF NEURAL INJURY 25. R. S. Lara, G. B. Matson, J. W. Hugg, A. A. Maudsley, and M. W. Weiner, Magn. Reson. Imaging 11, 273-278 (1993). 26. O. A. C. Petroff, J. W. Prichard, K. L. Behar, J. R. Alger, J. A. den Hollander, and R. G. Shulman, Neurology 35, 781-788 (1985). 27. J. W. Pettegrew, J. Moossy, G. Withers, D. McKeag, and K. Panchalingam, J. Neuropathol. Exp. Neurol. 47, 235-248 (1988). 28. J. W. Pettegrew, K. Panchalingam, J. Moossy, J. Martinez, G. Rao, and F. Boller, Arch. Neurol. 45, 1093-1096 (1988). 29. D. M. A. Mann, Mech. Ageing Dev. 31, 213-255 (1985). 30. E. Masliah, M. Ellisman, B. Carragher, M. Mallory, S. Young, L. Hansen, R. DeTeresa, and R. D. Terry, J. Neuropathol. Exp. Neurol. 51, 404-414 (1992). 31. E. Masliah, A. Miller, and R. D. Terry, Med. Hypotheses 41, 334-340 (1993). 32. R. C. Henneberry, Neurobiol. Aging 10, 611-613 (1989). 33. E. L. Harris, "NMR and Periodic Table." Academic Press, New York, 1978. 34. R. J. McClure, J. N. Kanfer, K. Panchalingam, W. E. Klunk, and J. W. Pettegrew, Neuroprotocols 5, 80-90 (1994).

[13]

Acute Stroke Diagnosis with Magnetic Resonance Imaging S t e p h e n C. J o n e s , N e n g C. H u a n g , M i c h a e l J. Q u a s t , A l e j a n d r o D. P e r e z - T r e p e c h i o , G i l b e r t R. H i l l m a n , a n d T h o m a s A. K e n t

Introduction The diagnosis of early ischemic stroke is primarily clinical within the first 8 hours, the time period during which therapeutic measures should be administered in order to be maximally successful. After 6 hours, the damage to the brain and the blood-brain barrier is generally throught to be irreversible (1) and the most recent evidence indicates that therapy must start before 4 hours to be effective (2). Thus we are left with a dilemma: How can early therapy be administered if early diagnosis cannot be made? We will review the use of three new magnetic resonance imaging (MRI) techniques that might alleviate, if not solve, the dilemma: first, diffusion-weighted imaging (DWI) is sensitive to proton diffusion and to the very early stages of ischemia; second, magnetic resonance spectroscopy (MRS) produces lactate and Nacetylaspartate (NAA) images using chemical shift imaging (CSI); and third, MR perfusion imaging yields on-line circulatory status. This review is focused on the immediate clinical application of these new MR techniques by the cerebrovascular community. The question that this review poses is whether the ischemic penumbra, that region that is salvageable in the early stages of ischemic stroke, can be identified by these new MR imaging methodologies.

E a r l y D i a g n o s i s of S t r o k e The early diagnosis and recognition of ischemic stroke is complicated by its symptomatology and pathophysiology. The changing pathophysiological situation during the early stages of ischemia has made predictions concerning ultimate outcome based on clinical findings alone difficult. Angiography, computed tomography (CT) (3), single-photon emission computed tomography (SPECT), positron emission tomography (PET), or conventional MRI Methods in Neurosciences, Volume 30 Copyright 9 1996 by Academic Press, Inc. All rights of reproduction in any form reserved.

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(4, 5) all have limited utility or applicability for the very early diagnosis of ischemic stroke. The early diagnosis of arterial occlusion that precedes infarction is possible using angiography, an invasive and risky procedure. However, early angiography has been used safely in one tissue plasminogen activator (tPA) trial to confirm clot position and lysis (6). Cerebral blood flow (CBF) techniques such as SPECT (7), PET, or stable xenon CT are neither highly available, accepted, nor proved. Noninvasive vascular imaging techniques such as transcranial Doppler ultrasound or MR angiography have not yet been utilized for clinical trials. Conventional CT can be used to exclude intracranial hemorrhage, and has been used for this purpose in another tPA trial for the early treatment of stroke (8), but is typically normal during the first 24 hours and cannot provide a diagnosis of early stroke. Currently available MRI techniques are not sensitive to early stroke. Tz-weighted MRI may be abnormal 12 hours after stroke onset, but is not useful for the very early diagnosis of stroke (9).

Early Treatment of Stroke: Entry Time, Therapeutic Window, and Ischemic Penumbra Much evidence exists that early treatment is important for stroke. Early entry times (or pretreatment) in experimental studies are associated with successful outcomes (10-13). Clinical studies that have been relatively successful tend to have earlier entry times (14-16) than those that were unsuccessful (17-20). Entry times of 3-4 hours after stroke have been suggested as necessary for clinical trials to take advantage of the therapeutic window (2, 21), that period during which some cells are at risk of death and can be saved by the restoration of blood flow. The limit of entry time is characterized by the progression of cerebral ischemic edema (22) from cytotoxic edema to vasogenic edema (23), the subsequent leakage of the blood-brain barrier to macromolecules in plasma, and the permanent disruption of cellular integrity 6-24 hours after stroke onset. Evidence from the comparison of H 2 clearance CBF and neuropathology suggests that 3 hours is the longest time that ischemia can be tolerated during which the "therapeutic window" still exists (1). Using a rat model of ischemic stroke, Kaplan et al. (2) have come to the same conclusion and defined the therapeutic window in terms of the ischemic penumbra. Clinical trials have disregarded this time factor until now, because of results of trials of the thrombolytic agent, tPA; treatment within 90 minutes has been achieved using emergency medical personnel to recognize stroke (8, 24). Another tPA trial that uses the much lengthier process of angiography

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for screening has reduced the entry time to 6 hours (6). However, one of the weak links in clinical trials is the lack of quick and noninvasive diagnostic techniques that are specific for very early stroke and that can identify and characterize evolving brain infarction, because treatment based solely on neurologic findings and time parameters is clearly suboptimal.

Diffusion-Weighted Imaging Diffusion-weighted imaging is a relatively new MR modality that is sensitive to the translational motion of water protons and was first introduced, implemented, and extended by Le Bihan et al. (25-27). Its role in ischemia is based on the observation that the diffusional rate of water in ischemia is much lower than that of normal brain. For diffusion-weighted imaging, gradient coils capable of generating high magnetic field gradients are required. The diffusion-weighted image is acquired by introducing a pair of gradient pulses with respect to the refocus 180~radio frequency pulse in the spin-echo sequence to increase the effect of motion on the echo signal (25). The resulting echo signal is dependent on proton motion (26). Therefore only the a p p a r e n t diffusion coefficient (ADC) can be obtained. The ADC image is a quantitative measure of the amount of diffusion and is inversely proportional to the DW image. It is calculated, on a pixel-by-pixel basis, from two or more images with different diffusion weighting (26). Values in normal and ischemic brain have been estimated to be 700-800 ~m2/sec and 200-400/xmZ/sec, respectively (28). Several theories have been advanced to explain the sensitivity of DWI to cerebral ischemia. Although it is reasonable that the increased DWI is not due to the magnitude of total water increase in cerebral ischemia, shown to be less than 5% (29), it is possible that the shift of water into the intracellular compartment (30), where its diffusion is restricted, or the associated decrease in membrane permeability due to the inactivation of the Na+/K + pump, could well be the cause of the decreased ADC in early ischemia (31). In either case, the decrease in ADC during early cerebral ischemia is due to the presence of cytotoxic edema. In normal brain, the Na+/K + pump maintains a large space of extracellular water, which has a higher diffusional constant than intracellular water. In ischemia, the pump is disabled and the extracellular space is decreased (22). The evidence supporting this conjecture is based on an experiment in which ouabain, which disables the Na+/K + pump, was administered intraparenchymally, decreasing the ADC from 840 to 460/xm2/ sec (30). The initial decrease of ADC after ischemia (28, 32) certainly is consistent with the initial decrease in extracellular space after arterial occlu-

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sion (22). Thus the decrease in ADC during early cerebral ischemia is most probably due to some characteristic of cytotoxic edema. In the late phase of cerebral ischemic edema characterized by vasogenic edema, in which the blood-brain barrier becomes dysfunctional, evidence concerning DWI is not conclusive. Using a model ofphotochemically induced intracerebral thrombosis (33), which has an uncertain relationship to classical vasogenic edema, DWI was increased in the periphery of the lesion, but not in the central area that suffered immediate endothelial barrier dysfunction (34). In several other reports, high DWI intensities at 12 hours (35) or low ADC values up to 24 hours (32) were noted after permanent occlusion, indicating that DWI is sensitive to vasogenic edema, as well as cytotoxic edema. Although DWI changes after middle cerebral artery (MCA) occlusion in the rat have been described by Mintorovitch et al. (28), the data lack certain features that limit our understanding of DWI for the diagnosis of ischemic stroke. First, the data are reported in terms of the ratio to contralateral cortex. Second, DW intensity is used, not ADC values. Third, the data are collected at only one gradient factor (b = 1413 sec/mm2). Fourth, ischemia was produced for only 33 minutes, so no information beyond this period is available. Finally, quantitative regional comparison to CBF or histology was not performed, although infarct was confirmed histologically at the end of the experiment. Although this study does not provide necessary CBF or histology data, it is extremely interesting because the initial DWI intensity at approximately 20 minutes after occlusion is 1.38 times the preocclusion control. It should be noted that this work used the intraluminal suture model of focal cerebral ischemia of Zea-Longa et al. (36), which produces large ischemic regions. Thus the sensitivity reported is based on only large ischemic regions and does not provide knowledge on the detectability of small ischemic regions. The time course of DWI changes after MCA occlusion in the rat has been described by Knight et al. (32) at 1.5-4, 4-8, 18-24, and 48-72 hours, in addition to longer periods. ADC remained at 50% of control values until the 18- to 24-hour time point, then increased to control thereafter. The time resolution is coarse, especially for the early and more interesting times, and although regions of interest were placed over "ischemic" cortex, no confirmation of ischemia was provided. In a further publication in which only ipsilateral/contralateral differences are given, Knight et al. (37) showed ADC changes until 24 hours in core and bordering regions. DWI has been proposed as a neuropathological marker at 30 minutes after ischemic onset using the suture model of MCA occlusion (38), but interestingly a subsequent work from the same group suggests that the ADC threshold changes during the first 2 hours (39). Both of these studies use

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neuropathology at 24 hours to assess ADC changes, without regard to the changing pathophysiology of CBF during the first hours after ischemia. Although ADC alone will undoubtedly have a role in stroke detection, we are unsure of exactly how ADC will be used for penumbra localization. Most probably, the addition either of an MR perfusion technique (40) or of MRS with NAA and lactate (41) as additional pointers will be necessary for clinical utility.

Chemical Shift Imaging Spectroscopy and Lactate Spectroscopy and Imaging MR proton spectroscopy was first performed with a surface coil placed over a region from which the spectra were to be obtained. This limitation made it difficult, if not impossible, to use the technique in either experimental or human focal cerebral ischemia, because the location of the ischemic core is often unknown, making the placement of the surface coil problematic. The combination of diffusion-weighted imaging and surface coil lactate spectroscopy has been applied to experimental stroke (42, 43). Recently, spectra have been obtained in a specified position by choosing a volume of interest using a saddle or slotted tube resonator (44). Localized proton spectroscopy with a vowel size of 2 x 2 x 2 cm has been used in normal humans (45) and a stroke patient (44). A technique to obtain a voxel size of 12/xl (3 x 2 x 2 mm) has been described, making this method useful for the study of experimental stroke in rats (41).

Cerebral Tissue Lactate in Stroke: Time Course and Levels Lactate plays a central role in ischemia. Figure 1 shows the results of combining cerebral tissue lactate determinations from different ischemia models and species (46-55). These data from different sources agree well, showing an immediate elevation of brain tissue lactate to 10 mM within minutes of ischemia, with a linear rise to 20 mM at 6 hours, dropping to 10 mM at 24 hours. The increasing lactate up until 6 hours, and perhaps beyond, could be a factor in the loss of tissue vitality that occurs when the last stages of cytotoxic edema end at 6-10 hours after onset. The only points that are not consistent with this pattern are those above 30 mM, after complete (53) or focal (54) ischemia in the cat. These values are two to three times the other values, because in the global ischemia model (53) there presumably was just enough residual flow to supply glucose for the anaerobic production of lac-

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FIG. 1 /3-Galactosidase transfection efficiency was calculated from X-Gal staining in septohippocampal cultures at four time points after transfection with pCMV//3gal using 1/xg DNA/3/xl liposomes/well. Cell counts were conducted independently by two investigators. The values represent the mean _+ SEM of the average number of X-Gal-stained cells in each well at 1, 2, 7, and 14 days following transfection (n = 4)./3-Galactosidase expression was maximal 2 days after transfection and persisted for 2 weeks.

concentrations, we did not see any increase in transfection efficiency by increasing DNA concentrations from 1 to 2/zg/well. To further investigate the temporal profile of liposome-mediated gene transfection,/3-gal transfection efficiency was calculated from X-Gal staining in septohippocampal cultures at 1, 2, 7, and 14 days after incubation with a transfection concentration of 1 /zg DNA/3 /zl liposomes/well. Maximal X-Gal staining (>1000 cells/well) was detected 2 days after transfection (Fig. 1). We also found that/3-gal-transfected cells could continue expressing /3-gal for at least 2 weeks, although at markedly lower levels (Fig. 1). Decreased levels of/3-gal expression were partially due to cell loss caused by prolonged incubation of primary septohippocampal cell cultures. Because of the limited life span of primary septohippocampal cell cultures, we were unable to study gene expression for longer periods.

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Our studies employing the fl-gal reporter gene in CNS cell cultures further confirmed observations of other researchers that the concentrations of liposomes can influence the efficiency of transfection (5). The differential transfection efficiencies in CNS cell cultures associated with varying concentrations of liposomes are consistent with the view that the higher the net positive charge of DNA-liposome complexes is, the better the interaction with the negatively charged cell membrane will be. However, overly high levels of liposomes can cause cell lysis. Without increasing liposome concentrations, increased amounts of DNA did not improve transfection efficiency. The efficiency of the DOTMA- and DOPE-mediated pCMV/fl-gal transfection observed by us in septohippocampal cell cultures (> 1000 transfected cells per 16 mm well) exceeds previously reported transfection efficiency for fl-ga! in hippocampa! cultures employing the transfection reagents. Transfectam and DOTAP (40-200 per 35-mm well) (16). The sustained expression of fl-gal for at least 2 weeks longer suggests the potential therapeutic utility of liposomal-mediated gene transfection in CNS injury and degeneration.

Sustained Expression of Nerve Growth Factor by Liposome-Mediated Gene Transfer Exogenous supplementation of nerve growth factor (NGF) has been reported to spare neurons from death and degeneration following injury (9, 18, 24, 36) and to increase choline acetyl transferase (CHAT) activity (31, 35). Furthermore, long-term NGF administration also increases the activity of protective antioxidant enzymes in rat brain (27). The rodent hippocampus is preferentially vulnerable to a variety of central nervous system insults, including traumatic brain injury and ischemia (8). Thus, enhancing the availability of NGF following CNS injury may have significant therapeutic potential. Many different approaches, such as continuous infusion, have been developed to deliver exogenous NGF to the nervous system of mammals (9, 18, 24, 36). Although these methods have generated important information and have therapeutic potential, significant limitations imposed by protein degradation and by the blood-brain barrier restrict the clinical utility of these approaches (1). Gene transfer is another way to introduce NGF into CNS cells and tissues. As pointed out above, a number of viral vector systems have been used to introduce genes into localized regions of the nervous system. Each of these has its advantages. However, each is compromised by limitations, including concerns regarding safety and toxicity. Cationic liposomes have also been used to carry various agents into CNS cells, including plasmid DNA for gene transfer (13, 33, 40). Roessler and Davidson report liposome-mediated fl-gal gene transfection and expression in adult mouse

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brain (29). Optimal concentrations of liposomes for transfection of the/3-gal gene in primary septohippocampal cultures have been determined by our laboratory (39). The transient expression produced by liposome-mediated gene transfection may limit its application in diseases caused by genetic defects. However, liposomal transfection of trophic factors may prove useful for treatment of central nervous system injury by blunting transient pathological processes and/or facilitating recovery. Because of the simplicity, reproducibility, safety, and efficiency of cationic liposome-mediated gene transfection (5, 12, 26), we have examined liposome-mediated NGF gene transfection in vitro (38). We used a pUC 19-based plasmid containing a CMV promoter as an expression vector for NGF transfection (23, 39). The rat NGF DNA was subcloned into a unique NotI site under the conrol of the CMV promoter. We used the commercially available DOTMA and DOPE (GIBCO-BRL) liposome formulated from a 1 : 1 (w/w) mixture of the cationic lipid N-[1-(2,3-dioleyloxy)propyl]-N,N,N-trimethylammonium chloride and dioleoylphosphatidylethanolamine in membrane-filtered water. Because of our interest in injury mechanisms in the hippocampus and the preferential vulnerability of the hippocampus to traumatic or ischemic brain injury (14, 22), we used mixed primary septohippocampal cell cultures for in vitro studies of liposome-mediated gene transfection. Cultures were incubated for 1 week prior to transfection. By that time, astrocytes reached confluence and were no longer actively multiplying, whereas neurons were well differentiated and stable. Based on our previous studies of liposome-mediated gene transfection in septohippocampal cultures (39), we employed a concentration of 1 /xg DNA/3/zl liposomes/well (16-mm well) to transfer the NGF gene to primary septohippocampal cell cultures. The purpose of these experiments was to examine systematically if liposome-mediated NGF gene transfection could produce increased expression of NGF mRNA and protein. We also sought to confirm the biological activity of the NGF protein produced following transfection. Reverse transcription-polymerase chain reaction (RT-PCR) analyses of NGF mRNA were conducted 1 day after liposome-mediated NGF transfection of septohippocampal cultures. Increased NGF mRNA was observed in pCMV/NGF-transfected cells as compared to sham transfections. To check for possible DNA contamination during RNA preparation, we included RNA samples without performing reverse transcription. These control studies confirmed the absence of DNA contamination. NGF protein levels were examined 2 days after liposome-mediated NGF transfection using an antibody enzyme-linked immunosorbent assay sandwich (ELISA). NGF concentrations were quantified against a standard con-

295

[17] LIPOSOME-MEDIATED GENE TRANSFECTION 10000 9000 8000

[~]

Control NGF

-

*** 7 0 0 0

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-

6000 Q.

5000 -

u.. Q~

4000 -

v

Z 3000 -

2000 1000

3 days

7 days

14 d a y s

Time after Transfection Fro. 2 ELISA analysis of NGF protein in culture medium: Three days after NGF gene transfection, NGF protein was increased 10-fold in the medium from NGF DNA-transfected cultures. Increased secreted NGF could be detected in the medium 2 weeks after NGF DNA transfection (values represent means +_ SEM; n = 4). The medium was exchanged three times a week after gene transfection.

centration curve of pure isolated murine NGF. E L I S A studies detected dramatic increases in N G F protein in cell pallets from transfected septohippocampal cultures. Three days after N G F gene transfection, robust increases of N G F protein were detected by E L I S A in the cell culture medium. The secreted form of N G F in the medium could still be detected 2 weeks after p C M V / N G F transfection (Fig. 2). Because we routinely exchange the medium three times a week after gene transfection, the consistent detection of the secreted form of N G F in the medium suggests that septohippocampal cells express and secrete N G F for at least 2 weeks after liposome-mediated gene transfection. Rat pheochromocytoma (PC12) cells were used to confirm the specific biological activity of N G F in medium conditioned by cell cultures transfected with N G F DNA. PC12 cell medium was removed 3 hours after plating, a sufficient amount of time for cells to attach to wells, and replaced with 0.5 ml of conditioned medium collected from cultures 3 days following liposomemediated N G F DNA transfection. N G F (20 ng/ml) was added to sister wells

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PARADIGMS OF NEURAL INJURY to assay the response of the cells to exogenous NGF. Cells were observed after 33 hours for the presence of neurite outgrowth. The secreted form of N G F in the N G F DNA-transfected cell medium produced biological effects similar to those of N G F isolated from mouse submaxillary gland. However, medium from control cells incubated only with liposomes did not produce neurotrophic effects. These results represent the first reported use of liposome-mediated N G F transfection in postmitotic central nervous system cell cultures. The levels of N G F protein expressed in our transfection system are particularly high, persist for at least 14 days, and elicit prominent neurotrophic effects such as neurite growth and growth cone formation. The persistent secretion of large amounts of N G F in media after p C M V / N G F transfection suggests the potential utility of neurotrophin gene transfection for treatment of neuronal injury or degenerative disorders.

References 1. M. Barinaga, Science 264, 773 (1994). 2. X. O. Breakefield and N. A. DeLuca, New Biol. 3, 203-218 (1991). 3. K. W. Culver, Z. Ram, S. Wallbridge, H. Ishii, E. H. Oldfield, and R. M. Blaese, Science 256, 1550-1552 (1992). 4. X. J. Fang, A. Keating, J. deVilliers, and M. Sherman, Hepatology 10, 78i787 (1989). 5. P. L. Felgner, T. R. Gadek, M. Holm, R. Roman, H. W. Chan, M. Wenz, J. P. Northrop, G. M. Ringold, and M. Danielsen, Proc. Natl. Acad. Sci. U.S.A. 84, 7413-7417 (1987). 6. F. H. Gage, J. A. Wolff, M. B. Rosenberg, L. Xu, J. L. Yee, C. Shults, and T. Friedmann, Neuroscience 23, 795-807 (1987). 7. T. Giordano, T. H. Howard, J. Coleman, K. Sakamoto, and B. H. Howard, Exp. Cell Res. 992, 993-997 (1991). 8. R. L. Hayes, L. W. Jenkins, and B. G. Lyeth, J. Neurotrauma, 9, S173-S178 (1991). F. Hefti, J. Neurosci. 8, 2155-2162 (1986). lO. M. S. Horwitz, in "Fields Neurology" (B. N. Fields and D. N. Knipe, eds.), 2nd Ed., pp. 1679-1721. Raven Press, New York, 1990. ll. Q. Huang, J. P. Vonsattel, P. A. Schaffer, R. L. Martuza, X. O. Breakefield, and M. DiFiglia, Exp. Neurol. 115, 303-316 (1992). 12. P. Hug and R. G. Sleight, Biochim. Biophys. Acta 1097, 1-17 (1991). 13. S. Imaizumi, V. Woolworth, R. A. Fishman, and P. K. Chan, Stroke 21, 13121317 (1990). 14. L. W. Jenkins, K. Moszynski, B. G. Lyeth, W. Lewelt, D. S. DeWitt, A. Allen, C. E. Dixon, J. T. Povlishock, T. J. Majewski, G. L. Clifton, H. F. Young, D. P. Becker, and R. L. Hayes, Brain Res. 477, 211-224 (1989). ,

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15. P. A. Johnson, K. Yoshida, F. H. Gage, and T. Friedmann, Mol. Brain Res. 12, 95-102 (1992). 16. S. Kaech, J. A. Drazba, and E. Ralston, Soc. Neurosci. Abstr. 19, 1746 (1993). 17. M. D. Kawaja, M. B. Rosenberg, K. Yohida, and F. H. Gage, J. Neurosci. 12, 2849-2864 (1992). 18. L. F. Kromer, Science 235, 214-216 (1987). 19. G. leGalLaSalle, J. J. Robert, S. Berrard, V. Ridoux, L. D. Stratford-Perricaudet, M. Perricaudet, and J. Mallet, Science 259, 988-990 (1993). 20. A. P. Li, C. A. Myer, and D. L. Kaminski, Cell Dev. Biol. 28A, 373-375 (1992). 21. E. Lycke, B. Hamark, M. Johansson, A. Krotochwil, J. Lycke, and B. Svennerholm, Arch. Virol. 101, 87-104 (1988). 22. B. G. Lyeth, L. W. Jenkins, R. J. Hamm, C. E. Dixon, L. L. Phillips, G. L. Clifton, H. F. Young, and R. L. Hayes, Brain Res. 526, 249-258 (1990). 23. G. R. MacGregor and C. T. Caskey, Nucl. Acids Res. 17, 2365 (1989). 24. C. N. Montero and F. Hefti, J. Neurosci. 8, 2986-2999 (1988). 25. N. Mori and T. Fukatsu, Epilepsia 33, 994-1000 (1992). 26. E. G. Nabel, D. Gordon, Z. Y. Yang, L. Xu, H. San, G. E. Plautz, B. Y. Wu, X. Gao, L. Huang, and G. J. Nabel, Human Gene Therap. 3, 649-656 (1992). 27. G. Nistico, M. R. Ciriolo, K. Fiskin, M. Iannone, A. Demantino, and G. Rotilio, Free Rad. Biol. Med. 12, 171-181 (1992). 28. Z. Ram, K. W. Culver, S. Walbridge, R. M. Blaese, and E. H. Oldfield, Cancer Res. 53, 83-88 (1993). 29. B. J. Roessler and B. L. Davidson, Neurosci. Lett. 167, 5-10 (1994). 30. M. B. Rosenberg, T. Friedmann, R. C. Robertson, M. Tuszynski, J. A. Wolff, X. O. Breakefield, and F. H. Gage, Science 242, 1575-1578 (1988). 31. R. J. Rylett, S. Goddard, B. M. Schmidt, and L. R. Williams, J. Neurosci. 13, 3956-3963 (1993). 32. H. San, Z. Y. Yang, V. J. Prompili, M. L. Jaffe, G. E. Plautz, L. Xu, J. H. Felgner, C. J. Wheeler, P. L. Felgner, X. Gao, L. Huang, D. Gordon, G. J. Nabel, and E. G. Nabel, Human Gene Therap. 4, 781-788 (1993). 33. M. J. Stewart, G. E. Plautz, L. Del-Buono, Z. Y. Yang, L. Xu, X. Gao, L. Huang, E. G. Nabel, and G. J. Nabel, Human Gene Therap. 3,267-275 (1992). 34. T. A. Thompson, M. N. Gould, J. K. Burkholder, and N. S. Yang, Cell Dev. Biol. 29A, 165-170 (1993). 35. L. R. Williams and R. J. Rylett J. Neurochem. 55, 1042-1049 (1990). 36. L. R. Williams, S. Varon, G. M. Peterson, K. Wictorin, W. Fischer, A. Bjorklund, and F. H. Gage, Proc. Natl. Acad. Sci. U.S.A. 83, 9231-9235 (1986). 37. D. Wolf, C. Richter-Landsberg, M. P. Short, C. Cepko, and X. O. Breakefield, Mol. Biol. Med. 5, 43-49 (1988). 38. K. Yang, F. Faustinella, J. J. Xue, J. Whitson, A. Kampfl, X. S. Mu, X. Zhao, G. Taglialatela, J. R. Perez-Polo, G. Clifton, and R. L. Hayes, Neurosci. Lett. 182, 291-294 (1994). 39. K. Yang, F. Faustinella, J. J. Xue, J. Whitson, A. Kampfl, X. S. Mu, X. Zhao, G. Taglialatela, J. R. Perez-Polo, G. L. Clifton, and R. L. Hayes, Neurosci. Lett. 182, 287-290 (1994). 40. N. Zhu, D. Liggitt, Y. Liu, and R. Debs, Science 261, 209-211 (1993).

[18]

Construction and Analysis of Transgenic Mice Expressing Amyloidogenic Fragments of Alzheimer Amyloid Protein Precursor Rachael L. Neve and Frederick M. Boyce

Introduction Two pathological aspects of Alzheimer's disease (AD) that may be related, but which are not fully understood, are the accumulation of amyloid in the brain and the destruction of brain cells. Even though the relationship between these two phenomena remains to be defined with precision, it is reasonable to suspect that any information we can obtain about/3-amyloid (A4) and its derivation from the/3-amyloid protein precursor (/3-APP) will yield insights into the mechanisms by which nerve cells degenerate in the disease. One of the earliest pieces of evidence linking AD neurodegeneration and/3-APP and/or its/3/A4-containing derivatives was the finding that the/3-APP gene is on chromosome 21: virtually all individuals trisomic for this chromosome will show AD-like neurodegeneration by the age of 40. More recently, the discovery that specific point mutations in the/3-APP gene are associated tightly with some forms of familial Alzheimer's disease has contributed to the increased interest in the role of/3-APP in the disease. Our laboratory has focused on a specific aspect of/3-APP and its connection with the neuronal destruction of AD. This work evolved from our observation several years ago that the carboxy-terminal 100 amino acids of the amyloid precursor protein (/3-APP-C100, or simply C100; previously termed AB1 or /3-APP-C104) were neurotoxic (1). Other laboratories subsequently revealed that C100 was amyloidogenic (2, 3). The neurotoxicity of C100 has been confirmed by other laboratories (4, 5). We hypothesized on the basis of these data that C100 or a similar/3-A4containing fragment of/3-APP may be centrally involved in the amyloidogenesis and neurodegeneration of Alzheimer' s disease. To test this latter hypothesis, we designed and generated an in vivo model for the action of C 100, in the form of transgenic mice expressing C100 in the brain (6). These animals display neuropathology that resembles some features of Alzheimer' s disease neuropathology, lending strength to our hypothesis that C100, or a/3-APP fragment very much like it, may be the perpetrator of neurodegeneration in Alzheimer's disease. In the following pages, using our C 100 transgenic mice

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Copyright 9 1996 by Academic Press, Inc. All rights of reproduction in any form reserved.

[181

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T R A N S G E N I C M I C E E X P R E S S fl-APP

RNA Splice & cDNA Polyadenylation

Promoter

r FIG. 1

A m

Diagram of a prototypic transgenic construct.

as a prototype, we describe the methodology involved in generating transgenic mice for the study of the pathological effects of amyloidogenic fragments of the Alzheimer amyloid protein precursor.

Practical Considerations The construction of transgenic mice for the study of amyloidogenesis and neurodegeneration in Alzheimer's disease entails the following pragmatic considerations: (1) How will the construct be designed and prepared for microinjection? (2) How will the resultant transgenic mice be analyzed for the presence of the transgene? (3) How will the mice be bred and maintained? (4) How will the RNA and protein products of the transgene be detected? (5) What types of histological analyses of the mouse brains should be carried out, and what are some of the technical and procedural issues involved? In the following sections, we describe methodologies that address these practical considerations for creating transgenic mice to study the neuropathological effects of amyloidogenic portions of the amyloid protein precursor.

Design of Transgenic D N A C o n s t r u c t The DNA construction for a transgenic animal requires three elements: (1) a promoter to drive expression of the transgene, (2) the transgene, and (3) RNA splicing and polyadenylation signals (Fig. 1). The choice of promoter is crucial, particularly when expression of the transgene in the brain is desired. Selection of the appropriate promoter can drive expression of the transgene in cells throughout the brain, or confine its expression to neurons or glia, or target expression to cells of a particular transmitter phenotype,

300

PARADIGMS OF NEURAL INJURY TABLE I

Neural Promoters Tested in Transgenic Mice

Promoter

Specificity

Ref.

Neuron-specific enolase SCG10 L7 Rhodopsin Dopamine fl-hydroxylase Preproenkephalin Tyrosine hydroxylase S 100 gene Olfactory marker protein Dopamine-/3-hydroxylase Acetylcholine receptor a2

Panneuronal Neural Cerebellar Purkinje and retinal bipolar neurons Photoreceptor cells Sympathetic and other neurons Brain and some peripheral tissues TH-immunopositive cells Astrocytes, some neurons Olfactory neurons Noradrenergic and adrenergic cells Cholinergic subregions of CNS Neurons and astrocytes Neurons

30 31 32 33 34 35 7 36 37 38 39 40 41

c-fos Calmodulin gene II

for example. It is most economical to choose a promoter whose specificity in directing expression of a reporter gene in transgenic mice has previously been verified, given the uncertainties of predicting in vivo specificity of expression of a promoter from in vitro studies (see Ref. 7). A partial listing of neural promoters that have been tested in transgenic mice is given in Table I. For the construction of our C100 transgenic mice, we used the human dystrophin brain promoter (8), which had been shown to confer expression preferentially in the brain in transgenic mice (unpublished data of F. M. Boyce and M. Rosenberg). We elected to place the transgene under the control of the dystrophin brain promoter because dystrophin is widely expressed in the brain at relatively low levels (9) and because its expression in the cortex peaks postnatally (unpublished data of F. M. Boyce and R. L. Neve). These quantitative and qualitative features of transcripts controlled by the dystrophin brain promoter may have been important in allowing survival of the C100 transgenic mice beyond the embryonic stage. In contrast, Quon et al. (10), who overexpressed the full-length fl-APP751 cDNA in transgenic mice, used the neuron-specific enolase promoter. In their case, a promoter conferring robust expression in the brain was desirable. At least three other groups have created transgenic mice overexpressing human fl-APP under the control of its own promoter by introducing the entire fl-APP gene into the mice (11-13). Such use of an entire gene in transgenic mice often gives optimal expression of the transgene; however, most transgenic mice studies employ cDNAs or partial cDNAs. The advantages of cDNA transgenes are that they are usually easier to clone and to manipulate than are entire genes, and that cDNAs

[18]

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TRANSGENIC MICE EXPRESS/3-APP

Dystrophin Neuronal Promoter

Mlul

Hindll,

~

C100 SV40 Splice & eDNA Polyadenylation

Hindlll ~

r

~

/N

BamHI

AAAAAA

FIG. 2 Diagram of the C100 transgenic construct.

can, with more facility, be mutated or truncated for comparison of parallel transgenics possessing the wild-type cDNA transgene with those possessing genetically altered versions of the cDNA. In our case, we wished to express only the carboxy-terminal 100 amino acids of/3-APP (which fortuitously possessed its own translation start methionine), and we had already established that it could be exogenously expressed from an amino-terminal-truncated /3-APP cDNA in cell lines in vitro. It is important, when making constructs for transgenic mice, to include not only a methionine that will be a translational start signal, but also a stop codon (if a carboxy-terminal truncation of the cDNA is planned). The cDNA transgene must be fused to an RNA cleavage and polyadenylation signal sequence at the 3' end. Many of these 3' cassettes [most of which have been derived from simian virus 40 (SV40)] include an intron as well. Inclusion of introns has been shown to enhance expression of transgenes (14). When we created our C100 transgenic construct, we added the SV40 mRNA processing signals that comprise the 800-bp BglII-BamHI fragment of pRSV-/3-globin (15). The complete C100 construct, then, in which the Rous sarcoma virus (RSV) promoter was replaced with the 3.0-kb dystrophin brain promoter, and/3-globin was replaced with C100 (0.85 kb), followed by the SV40 mRNA processing signals, was approximately 4.65 kb (Fig. 2). An additional consideration in planning the transgenic construct is to design unique restriction enzyme sites flanking the transgene. This allows the transgene to be excised from the bacterial plasmid vector sequences, which could interfere with expression. To prepare the C100 DNA for microinjection, 20txg CsCl-pure plasmid was cleaved with MluI and BamHI to release the 4.65-kb C100 transgenic construct. The digested DNA was preparatively electrophoresed on a 14-cm 0.6% Seakem GTG (FMC, Rockland, ME) agarose gel in standard Trisacetate-EDTA (TAE) buffer until the 4.0-kb vector and the 4.65-kb transgene-containing fragment were well separated. The latter band was cut out

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of the gel (after being revealed by ethidium bromide UV fluorescence) and placed into dialysis tubing with approximately 500/~1 of TAE, after which the DNA was electroeluted at 80 V for 45 minutes. A brief (1 minute) reversal of the voltage gradient ensured removal of the DNA from the walls of the dialysis tubing. The TAE buffer (containing the now-eluted DNA fragment) was removed from the dialysis tubing and the DNA fragment precipitated by the addition of 1/10 volume of 3 M sodium acetate (pH 5.0) and 2.0 volumes of 95% (v/v) ethanol. The DNA was pelleted by spinning it at 14,000 rpm for 30 minutes in a microcentrifuge, and the resultant pellet was resuspended in 50/~1 of TE (10 mM Tris, pH 7.5, 0.25 mM EDTA).

Analysis of Mice for Presence of Transgene The two mostly commonly used methods for detecting mice that possess the transgene are Southern blot analysis and polymerase chain reaction (PCR) analysis. At the time of weaning, when the approximately 3-week-old mice are tagged and males are separated from females, approximately 1 cm is clipped from the end of the tail of each mouse in a given litter (if PCR will be used to detect the transgene, care must be taken not to transfer any tissue from one tail to another, e.g., by using a pair of scissors consecutively on the mice). The tail fragments are kept on ice (if DNA preparation is to occur immediately) or frozen. DNA is then prepared from this tissue according to the following simplified protocol obtained from Dr. M. Rosenberg (personal communication, 1991). Add the tail to 600/~1 of tail buffer (500 mM Tris, pH 8.0; 100 mM EDTA, pH 8.0; 100 mM NaC1; 1% SDS) in a 1.5-ml centrifuge tube. Add 35 p.l of 10 mg/ml proteinase K, and invert the tube or vortex it briefly to mix the contents. Incubate at 55~ for 5 hours to overnight. Then add 202/~1 of 5 M NaC1 (to bring the final concentration of salt to 1.5 M) while the tube is still warm. Add 600 /~1 of chloroform : isoamyl alcohol (24:1, v/v) and vortex each tube for 20-30 seconds. Separate the aqueous and organic layer by spinning at 14,000 rpm in a microcentrifuge for 10 minutes. A very large layer of protein and SDS will form between the aqueous and organic phases. Carefully remove the upper, aqueous layer to a new tube containing 1.5 ml of 95% (v/v) ethanol. Invert the tube two or three times, and remove the precipitated DNA with a capillary micropipette, one end of which has been heated over a flame to create a small hook, which can then be used to retrieve the DNA. Allow the ethanol to drain for 1 minute, then place the micropipette into 150/~1 of TE, pH 8.0. Allow the DNA to resuspend overnight at 4~ before subjecting it to further manipulations. The yield will be 50-100 p.g of DNA. If Southern blots are used to identify transgene-containing DNAs, the

[18] TRANSGENIC MICE EXPRESS/3-APP

303

DNA should be cleaved with an enzyme(s) that will release the transgene or some portion of the construct intact. We digested our mouse tail DNAs with HindIII, which we knew should release the dystrophin brain promoter as an intact 3-kb band (see Fig. 2). The HindIII ends were put onto the brain promoter fragment artificially during the original cloning of the promoter, so that while the transgene promoter would be revealed as a 3-kb fragment on Southern blots, the endogenous promoter would appear on a DNA fragment(s) of a different size. Thus, the endogenous and transgenic dystrophin brain promoters could be distinguished by size on Southern blots. Tail DNA (30/zl, or approximately 15/xg) was digested to completion with HindIII and electrophoresed at 50 V for 18 hours. The DNA in the gel was transferred to 1.2/zm Biotrans (ICN, Costa Mesa, CA) in the presence of 20 x SSC (1 x SSC: 0.15 M sodium chloride, 0.015 M sodium citrate), and the blot probed with a dystrophin brain promoter DNA fragment radiolabeled by the random hexanucleotide priming method. Blots were washed to a maximum stringency of 0.2x SSC at 65~ with 0.1% SDS. An overnight exposure of the washed blot was usually sufficient to reveal which DNAs contained the transgene. Approximate copy number of the transgene in each line can be determined by densitometric analysis of the blots, in which the intensity of the transgene band is compared to the intensity of the endogenous dystrophin promoter band. If the polymerase chain reaction is utilized to detect the presence of the transgene, the primers must not both be internal to the promoter or to the cDNA unless they span introns, for otherwise the PCR product from the endogenous gene cannot be distinguished from the PCR product from the transgene. PCR tends to be most consistent with primers that define an approximately 100- to 300-bp fragment. The primers should be tested beforehand on control mouse DNA, and on dilutions (down to one-tenth genome copy) in control mouse DNA of the plasmid carrying the transgene. The 5' primer that we used was P6 in the dystrophin promoter (8); the 3' primer (which was within the C 100-encoding region) represented the reverse complement of base pairs 2040-2062 in/3-APP695 (see Fig. 2). PCR was carried out in 50-/xl reactions containing 50 ng of each primer and 0.5/zl (approximately 200 ng) of each tail DNA, for 32 cycles (94~ 1 minute; 50~ 1 minute; 72~ 3 minutes), followed by a 10-minute extension at 72~

Maintenance of Transgenic Mouse Colony Because of the effort and expense involved in the creation of transgenic mice, it is crucial that each transgenic line is properly maintained. Sufficient numbers of animals must be generated both for propagation of the transgenic

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line and for analysis of any neuropathology in each line. In addition, transgenic animals that are models for the effects of amyloidogenic fragments of fl-APP and for other aspects of Alzheimer's disease will almost certainly need to be aged. For these reasons, it is important that the animals are kept in a barrier or viral antibody-flee facility to avoid loss of animals to infectious agents. Our animals are housed in positive individually vented caging systems to minimize the transmission of airborne diseases, and manipulations of the mice are carried out in a laminar flow hood. We have observed neuropathology in C100 transgenic mice as early as 4.5 months postnatally, but the pathology may advance with age. Interestingly, normal aged mice do not show plaques or tangles, which are characteristic of aged human brains. The founder transgenic mice, derived from F 2 C57BL/6 • SJL/J hybrid mice, are mated with C57BL/6J mice to give rise to F~ progeny. All subsequent generations (F2, F3, etc.) are also backcrossed to C57BL/6J. Weaning is done at approximately 3 weeks of age and consists of separating each litter into separate cages of males and females. Ear tagging and typing of each offspring for the presence of the transgene are also performed at the time of weaning. Transgene-negative siblings are used as age-matched controls. Because the founder mice are derived from both C57BL/6J and SJL/J strains, it is also important to include animals from each of these strains as controls for strain variations. We determine which founder mice are carrying the transgene, and then use F1 litters to determine whether each line is expressing the transgene at the RNA and protein levels (see following). Failure to detect expression of the transgene in several F1 mice carrying the transgene suggests that the line will not be useful for further analyses. However, because expression may depend on integration site, other lines carrying the same transgene may yield expression. Thus it is important to screen founder animals for the presence of the transgene as well as F 1 animals for expression in order to minimize the number of transgenic animals that must be maintained. It is customary to maintain several expressing lines for each transgene. The presence of the transgene is monitored in each generation by PCR or Southern blotting. For long-term propagation of each transgenic line, siblings can be bred to yield offspring that are homozygous for the transgene. Because both heterozygotes and homozygotes are positive for the transgene, homozygotes are usually detected by test breeding with C57BL/6J mice. Homozygotes will yield litters with all progeny carrying the transgene, whereas heterozygotes will yield mixed litters. When a homozygous male and female have been identified, they may be bred to yield homozygous offspring. Creation of a homozygous line reduces the need for further transgene typing. However, it is wise to check for the presence and expression of the transgene periodically, because errors in breeding may occur, and because expression of the transgene may be extinguished with time.

[18] TRANSGENIC MICE EXPRESS/3-APP

305

Detection of RNA and Protein Products of Transgene To assess the tissue specificity of the expression of C100 under the control of the dystrophin brain promoter, we first examined the transgenic mice for the presence of RNA transcribed from the transgene. Total RNA was prepared using 100-500 mg of tissue by the guanidinium thiocyanate procedure (with adaptations) (16). The final step involved precipitation of the RNA with 1/2 volume of ethanol, which preferentially precipitates RNA but not DNA. Because the dystrophin brain promoter drives the expression of relatively low levels of dystrophin in the brain, we anticipated that similarly low expression of the transgene might make detection of the transgene RNA by Northern blots difficult. We chose to use reverse transcription coupled with PCR (RT-PCR), using the same primers that were used to analyze the tail DNAs. It is important to confirm that the 5' primer, if part of the promoter region, is also represented in the expected RNA transcript. RNA (1 ~g) from each tissue was treated with DNase I (0.3 U//A in a volume of 13.1 ~1) at 37~ for 20 minutes to remove possible contaminating DNA, and then at 65~ to inactivate the DNase. CH3HgOH (3 ~1, 0.1 M) was added to remove RNA secondary structure; 1.55/~1 of 0.7 M 2-mercaptoethanol was then added, after which cDNA was synthesized from each RNA sample in a 25-~1 reverse transcription reaction that consisted of 50 mM Tris (pH 8.2 at 42~ 50 mM KC1, 6 mM MgCI 2, 10 mM dithiothreitol, 1000 U/ ml Promega (Madison, WI) Biotec RNasin, 1 /~g of 3' primer (identical ~o the 3' primer used in the tail DNA PCR reactions), 400 ~M dNTPs, and 350 U/ml Life Sciences avian myeloblastosis virus reverse transcriptase. After incubation of these reactions at 41.5~ for 2 hours, the reactions were placed at 65~ for 10 minutes to inactivate the reverse transcriptase; 1 /~1 of each cDNA reaction was used as template in a 50-/.d PCR reaction mix containing 200 ng of each of the primers (identical to the primers used for PCR analysis of tail DNAs) and 0.25 ~1 Taq polymerase (Perkin-Elmer, Norwalk, CT). The reactions were subjected to 40 cycles of PCR (94~ 1 minute; 50~ 2 minutes; 72~ 3 minutes) and 40 ~1 of each reaction was electrophoresed on a 1% agarose gel in TAE buffer and transferred to 1.2-~m Biotrans (ICN) in the presence of 20• SSC. The filter was then hybridized with an internal oligonucleotide probe that was 32p-labeled using T4 polynucleotide kinase (U.S. Biochemicals, Cleveland, OH). We used RT-PCR to assess expression of the C100 transgene in brain, muscle, heart, and liver of animals from nine different founder lines. The expected 320-bp RT-PCR product was seen at highest levels in the brain, in all lines evaluated. Although the transgene was transcribed at low levels in other tissues in some of the lines, in all transgenic animals its expression was - 10 times higher in brain than in any other tissue examined. As expected, we were unable to detect the transgene transcript on RNA blots.

306

PARADIGMS OF NEURAL INJURY

46K

----

3 0 K "-'-p,-

14.3K

----

FIG. 3 Detection of C100 protein in the brains of transgenic mice. Immunoblot of the antibody 10D5 against total brain homogenates of three nontransgenic (-) and two transgenic (+) 9-month-old mice in lines 2 and 3 is shown.

Attempts to detect the C100 transgene protein product in the brains of the transgenic mice highlighted one of the difficulties often encountered during the characterization of a transgene product, which is the inability to distinguish between endogenous and transgene protein products by immunocytochemical or immunoblot analysis. In this instance, our efforts to detect the C100 were hampered by the fact that available antibodies to human C100 (expressed by the transgene) also reacted with endogenous mouse flAPP. Among the endogenous processing products of mouse fl-APP, as of human fl-APP, are amyloidogenic carboxyterminal fragments of fl-APP that have a similar size to the product of the C100 transgene and are detected by most antibodies to human C100. However, we were able to obtain a monoclonal antibody to made human ~/A4 1-15 (10D5, gift ofD. Schenk, Athena Neurosciences, Inc.) that does not react with mouse fl-APP. Brains of 9-month-old mice were rapidly homogenized and 20/~g of each homogenate was electrophoresed and transferred to PVDF membranes. The membranes were immunostained with the monoclonal antibody 10D5, and enhanced chemiluminescence was used to detect positive immunoreactivity at a band of approximately 28 kDa only in lanes containing homogenates of positive transgenic mouse brains (Fig. 3). We believe that this band represents a C 100 dimer, largely because the predominant portion of the C 100 that we generate in bacterial or baculovirus systems migrates at 28 kDa (about 10% migrates at 15 kDda; unpublished results of R.L.N.).

307

[18] TRANSGENIC MICE EXPRESS fl-APP TABLE II

Epitope Tags Tag

Sequence

Ref.

FLAG Streptavidin-affinity tag Influenza virus hemagglutinin a-Tubulin epitope HSV-Tag

AspTyrLysAspAspAspAspLys SerAlaTrpArgHisProGlnPheGlyGly TyrProTyrAspValProAspTyrAlaSerLeu GluGluPhe GlnProGluLeuAlaProGluAspProGluAsp GluGlnLysLeulleSerGluGluAspLeu

17 42 43 44 45 46

c-myc

To improve our ability to detect the C100 transgene protein product in mice, we have constructed a second transgenic mouse, expressing a cDNA that we term FLAG/3-APP-C100. We have fused the coding sequence for a hydrophilic 10-amino acid sequence termed FLAG (17) onto the aminoterminus of C100. We have successfully expressed this fusion fragment in cells, and have shown that it retains its neurotoxicity in culture, and that it binds to the specific C 100 receptor (18). We will be able to distinguish the transgene product from the endogenous/3-APP gene product in the transgenic mice expressing FLAG/3-APP-C 100 by using monoclonal antibodies to FLAG (M2 and M5, available commercially from VWR, Greenbelt, MD) or polyclonal antibodies to FLAG that we have made to detect FLAG/3-APP-C 100 and its metabolic derivatives in vivo. We have tested the FLAG antibodies immunocytochemically, and they demonstrate specificity, with the polyclonal antibodies showing particularly robust immunoreactivity to the FLAG epitope, both immunocytochemically and on immunoblots. We will use these antibodies to determine the location and state of aggregation of FLAG/3-APP-C100 in the transgenic mice. We have illustrated the use of epitope tags by describing the rationale for, and the design of, our FLAG/3-APP-C 100 transgenic mice. Additional epitope tags have been described, however, which are useful for the same purpose. A partial listing of such tags is given in Table II. Selection of the appropriate tag will depend on whether the investigator wishes to use it at the amino terminus or the carboxy terminus of the transgene protein (or even internal to the protein), and on availability and characteristics of the antibody.

Histological Analyses of Transgenic Mouse Brains Expressing Amyloidogenic Fragments of/3-APP in Brain Classic histological stains for Alzheimer's disease neuropathology have historically been those that detect amyloid plaques and neurofibrillary tangles, the two major features of Alzheimer's disease neuropathology that form the

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basis for postmortem diagnoses. During the past decade, hov,ever, we have developed antibodies and cloned genes that enable increasingly sophisticated analyses of features of Alzheimer's disease that precede the end-stage plaques and tangles. As a result, a broader definition of the neuropathology characteristic of this disease has emerged, as exemplified by the analyses that we carried out on our C 100 mice (6). We used an affinity-purified antibody, E1-42 (19), raised against a peptide representing the 42-amino acid/3/A4 fragment, to detect amyloid deposition in the transgenic mouse brains. This antibody does not recognize normal human/3-APP, but distinctively immunoreacts with pathologic structures specific to Alzheimer's disease and (to a lesser extent) normal aged brain. These include the amyloid cores of neuritic plaques, as well as the so-called diffuse amyloid deposits that are not revealed by conventional histological stains for amyloid. Immunostaining of control mouse brains with E1-42 showed that in mouse brain, in contrast to human brain, this antibody recognizes normal structures to some degree in that it displayed light homogeneous staining of cell bodies. Both quantitative and qualitative differences in the pattern of E1-42 (/3/A4) immunoreactivity were exhibited in transgenic mouse brains. Intensely E1-42 immunoreactive material was seen in neuronal cells throughout the brains of all transgenic mice tested. In most cases the/3/A4 immunoreactivity occurred as punctate deposits within neurons that have a rounded, compact appearance. Such staining was clearly visible in Ammon's horn of the hippocampus and in the stratum oriens. The intracellular accumulation of/3/A4 was particularly prominent within the hilus, where the immunoreactivity was seen not only in the cell bodies but also in abnormal processes. The emergence of/3/A4 immunoreactivity in the neuropil may represent a stage of amyloid deposition later than that observed in the cell soma, because it was seen only in the three lines expressing highest levels of the transgene. Punctate accumulations of/3/A4 immunoreactivity in dystrophicappearing fibers were dramatically apparent in the stratum radiatum of the CA2/3 region of the hippocampus in these lines. When we analyzed these same mice with additional antibodies to/3/A4 and its subdomains, we discovered that they exhibited patterns of immunoreactivity somewhat different from those displayed by the antibody E1-42. For some antibodies, the difference was merely quantitative, whereas with others we detected no differences between transgenic and control animal brain. When these antibodies were used to probe immunoblots of E1-42 peptide, we discovered that only those antibodies recognizing aggregated forms of E1-42 revealed/3/A4 deposits in the transgenic mice. The antibodies that immunodetected only the E 1-42 monomer on immunoblots did not show abnormal immunoreactive structures in the brains of the mice. Interestingly,

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most antibodies that recognized only the E1-42 monomer were made to subdomains of the j3-amyloid fragment (e.g., 1-28) or to the coupled 1-42 peptide. Those detecting aggregates of/~/A4 comprised primarily antibodies generated with uncoupled 1-42 peptide. We had previously shown that staining of Alzheimer's disease brains with F5, an antibody to the carboxy-terminal nine amino acids of/~-APP, exposed intracellular aggregates of this epitope in secondary lysosomes in pathologically afflicted regions of Alzheimer's disease brain, such as in the CA1 neurons of Sommer's sector (20). Hence, we might expect to detect a similar phenomenon in the transgenic mice. And indeed, staining of the brain sections with the antibody F5 showed a striking change in the subcellular localization of the F5 epitope that was particularly evident in the CA2/3 region of the hippocampus in transgenic mice. Whereas control mice showed homogeneous light F5 immunoreactivity predominantly of the neuronal somata in this region, the F5 immunoreactivity in the transgenic mice presented as dark punctate accumulations in subcellular compartments that extended markedly into the neuronal processes. Adjacent Nissl stained sections did not reveal detectable gross abnormalities in the area of altered F5 staining, suggesting that the disorganization evident in the immunostained sections mainly involves the neuropil in transgenic mice of this age. In transgenic mice from the three lines expressing highest levels of the transgene, the cells in the CA2/3 region showed particularly dense reaction product in the neuropil, and the reactivity in the soma took the form of larger accumulations, as if the punctate vesicular immunoreactive material had fused or aggregated. The appearance of the F5 immunoreactivity in these cells was very similar to what we observed in Alzheimer's disease brains (20). F8, an independent antibody made to the carboxy terminus of/3-APP (gift of D. Schenk, Athena Neurosciences), exhibited the same abnormal pattern of immunoreactivity revealed by the F5 antibody. The immunocytochemistry is carried out as follows. F1 backcross progeny from six different founder lines (ages 3 weeks to 4 months) and founders only from three additional lines (6 months of age) are analyzed histologically. We also analyzed eight age-matched C57BL/6 and SJL controls. Mouse brains are either immersion fixed (in cases in which we use half of the brain for RNA analysis) or are perfused with freshly prepared 4% paraformaldehyde (w/v) buffered with 0.1 M sodium phosphate, pH 7.2, and postfixed overnight at 4~ before cryoprotection in 30% sucrose in 0.1 M sodium phosphate, pH 7.2. The brains are frozen in a dry ice-acetone bath maintained at -40~ then processed immediately or stored at -70~ The tissue is cut at 50-~m intervals on a sliding microtome into ice-cold Tris-buffered saline (TBS: 50 mM Tris, pH 7.5,300 mM sodium chloride). Human brain tissue is obtained in formalin and is cryoprotected and cut as described for

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the mouse tissue except that the TBS used for the human tissue contains only 145 mM sodium chloride. E 1-42 immunocytochemistry on mouse and human sections, and F5 immunocytochemistry on human sections, are carried out as follows. Sections are pretreated with 1% H202 in TBS (100 mM Tris, pH 7.5, 145 mM sodium chloride) for 20 minutes, rinsed for 5 minutes in TBS and 15 minutes in TBSA (TBS plus 0.1% Triton X-100), blocked for 30 minutes in TBS-B (TBS-A plus 2% BSA), and incubated overnight at room temperature in primary antibody diluted 1" 750 in TBS-B. Sections are then rinsed for 15 minutes in TBS-A and 15 minutes in TBS-B, reacted with the secondary antibody (biotinylated goat antirabbit IgG, 1:250 in TBS-B according to manufacturer's specifications for the Elite kit (Vector Labs, Burlingame, CA), rinsed for 15 minutes in TBS-A and 15 minutes in TBS-B, reacted with horseradish peroxidase (HRP)-conjugated avidin-biotin complex (as specified for the Elite kit; Vector Labs), rinsed 3x 5 minutes in TBS, and visualized using Vector Labs HRP substrate (diaminobenzidine) kit. Nickel chloride is included with the chromogen for F5. F5 and F8 immunocytochemistry on mouse brains is carried out as follows. The brain sections are pretreated with 3% H202 in TBS (50 mM Tris, pH 7.5, 300 mM sodium chloride) for 20 minutes, blocked for 1 hour in 0.3% Triton X-100, 20% goat serum in TBS, washed 2• 15 minutes in TBS, and incubated for 40 hours at 4~ in primary antibody diluted 1 : 1000 in 0.1% Triton X-100, 20% goat serum (TBST). Sections are then rinsed 2x 15 minutes in TBS, reacted with the secondary antibody (biotinylated goat antirabbit IgG, 1"250 in TBST according to manufacturer's specifications; Vector Labs), rinsed 2x 15 minutes in TBS, reacted with HRP-conjugated avidin-biotin complex (as specified; Vector Labs), rinsed 2x 15 minutes in TBS, and the reaction is visualized using 0.05% diaminobenzidine (Sigma, St. Louis, MO) plus 0.8% nickel chloride as a chromogen. For all antibodies used, sections incubated in parallel without primary antibody fail to develop any staining. Sections are mounted onto chrom-alum subbed slides, covered, and photographed with Kodak (Rochester, NY) T-MAX film. We do not know whether the /3/A4 immunoreactive aggregates or the abnormal subcellular buildup of the carboxy-terminal portion of/3-APP as revealed by staining with the F5 antibody derive from the endogenous/3APP or from the product of the transgene. This question can begin to be answered if the product of the transgene is tagged, as described previously. With regard to our new FLAG/3-APP-C100 transgenic mice, double staining with the FLAG antibody and either the E 1-42 or the F5 antibody may reveal whether the abnormal immunoreactivity is generated from the product of the transgene.

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We showed previously that the enlarged intracellular organelles into which the F5 immunoreactivity segregates in the disease are probably fused lysosomes (20). The dense F5 immunostaining in these enlarged organelles was particularly prominent in regions of the hippocampus that were heavily invested with pathology, such as Sommer's sector (CA1), in which the heavy staining of the pyramidal cells was accompanied by atrophy of many of these cells (20). Although Nissl and Bielschowsky stains did not reveal gross neuronal death in any of the transgenic brains at 4-6 months of age, it is possible that aggregation of the F5 epitope into enlarged lysosomes may presage neuronal degeneration. To determine whether the lysosomal system is indeed involved in the pathology in our transgenic mice, it is necessary to carry out immunoelectron microscopy to confirm that the F5 aggregates in the transgenic mice, as in AD brain, are fused lysosomes. Enzymatically active lysosomal proteases have been detected in association with senile plaques in Alzheimer's disease, and immunoreactivity for certain of these enzymes is abnormally increased in degenerating neurons in the disease (21). It will be of interest to look, in transgenic mouse models for Alzheimer's disease, for abnormalities of the lysosomal system that resemble those seen in the disease. Thioflavin S histochemistry was employed for identification of amyloid deposits in the mouse and human brains. Sections were incubated for 20 minutes in 1"1 100% ethanol'chloroform, and then rinsed 3x 1 minute in 95% (v/v) ethanol, 3 minutes in 70% (v/v) ethanol, 3 minutes in 50% (v/v) ethanol, 3 minutes in H20. Sections were then incubated for 4 minutes in 1% thioflavin S (w/v) (Sigma) in H20 and differentiated by rinsing in 80% ethanol. The thioflavin S histochemistry showed no abnormal fluorescence of structures in the control mice, or in six of the transgenic mouse lines. However, the mice from the three lines with highest brain expression of the transgene displayed prominent thioflavin S fluorescence around blood vessels, which we also observed in Alzheimer's disease brains stained with thioflavin S. This fluorescence suggests that amyloid has accumulated in the cerebral blood vessels of these transgenic animals. Cytoskeletal alterations, particularly with regard to the phosphorylation state of the microtubule-associated protein tau (r), should be evaluated in any animal model for the effects of amyloidogenic fragments of/3-APP. We have used the monoclonal antibody Alz50 to reveal cytoskeletal pathology in the brains of mice that had received transplants of C100-producing PC12 cells (22). The Alz50 antibody was originally identified on the basis of its selective immunoreactivity with a neuronal antigen in the brains of Alzheimer disease patients (23). A modified form of the microtubule-associated r protein is present in Alzheimer' s disease brain, (24), and Alz50 has been hypothesized to recognize a modification of r that occurs early in the sequence of events

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PARADIGMS OF NEURAL INJURY leading to neurofibrillary degeneration (25, 26). Caution should be exercised, however, when using Alz50 to assess animal models of Alzheimer's disease (27). This antibody stains normal structures in both human and animal brains, so that careful controls must be carried out to show that a particular pattern of Alz50 staining that is suspected to be abnormal is indeed not observed in control brains. Moreover, because it is a mouse monoclonal antibody, an antirodent secondary antibody is normally used in conjunction with it. This antibody will, of course, nonspecifically react with endogenous rodent immunoglobulins, necessitating careful controls for such an artifact. During the past several years, numerous additional antibodies that define pathological phosphorylated epitopes of r and that reciprocally mark the normal lack of phosphorylation at these epitopes have been defined, and are reviewed in two articles (28, 29).

Acknowledgment This work was supported by NIH Grant NS28965 (R.L.N.).

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12. B. E. Pearson and T. K. Choi, Proc. Natl. Acad. Sci. U.S.A. 90, 10578 (1993). 13. J. D. Buxbaum, J. L. Christensen, A. A. Ruefli, P. Greengard, and J. F. Loring, Biochem. Biophys. Res. Commun. 1997, 639 (1993). 14. R. D. Palmiter, E. P. Sandgren, M. R. Avarbock, and D. D. Allen, Proc. Natl. Acad. Sci. U.S.A. 88, 478 (1991). 15. C. Gorman, R. Padmanabhan, and B. H. Howard, Science 221, 551 (1983). 16. R. L. Neve, P. Harris, K. S. Kosik, D. M. Kurnit, and T. D. Donlon, Mol. Brain Res. 1, 271 (1986). 17. K. S. Prickett, D. C. Amberg, and T. P. Hopp, Biotechniques 7, 580 (1989). 18. M. R. Kozlowski, S. F. Spanoyannis, S. P. Manly, S. A. Fidel, and R. L. Neve, J. Neurosci. 12, 1679 (1992). 19. B. J. Cummings, J. H. Su, J. W. Geddes, W. E. Van Nostrand, S. L. Wagner, D. D. Cunningham, and C. W. Cotman, Neuroscience 48, 763 (1992). 20. L. I. Benowitz, W. Rodriguez, P. Paskevich, E. J. Mufson, D. Schenk, and R. L. Neve, Exp. Neurol. 106, 237 (1989). 21. A. M. Cataldo and R. A. Nixon, Proc. Natl. Acad. Sci. U.S.A. 87, 3861 (1990). 22. R. L. Neve, A. Kammesheidt, and C. F. Hohmann, Proc. Natl. Acad. Sci. U.S.A. 89, 3448 1.1992). 23. B. L. Wolozin, A. Pruchnicki, D. W. Dickson, and P. Davies, Science 232, 648 (1986). 24. A. Nieto, I. Correas, E. Montejo de Garcini, and J. Avila, Biochem. Biophys. Res. Commun. 154, 660 (1988). 25. K. Ueda, E. Masliah, T. Saitoh, S. L. Bakalis, H. Scoble, and K. S. Kosik, J. Neurosci. 10, 3295 (1990). 26. V. M.-Y. Lee, B. J. Valin, L. Otvos, and J. Q. Trojanowski, Science 251, 675 (1991). 27. P. Davies, Neurobiol Aging 13, 613 (1992). 28. M. Goedert, Trends Neurosci. 16, 460 (1993). 29. E.-M. Mandelkow and E. Mandelkow, Trends Biochem. Sci. 18, 480 (1993). 30. C. W. Wuenschell, N. Mori, and D. J. Anderson, Neuron 4, 595 (1990). 31. S. Forss-Petter, P. E. Danielson, S. Catsicas, E. Battenberg, J. Price, M. Nerenberg, and J. G. Sutcliffe, Neuron 5, 187 (1990). 32. J. Oberdick, R. J. Smeyne, J. R. Mann, S. Zackson, and J. I. Morgan, Science 248, 223 (1990). 33. D. J. Zack, J. Bennett, Y. Wang, C. Davenport, B. Klaunberg, J. Gearhart, and J. Nathans, Neuron 6, 187 (1991). 34. E. H. Mercer, G. W. Hoyle, R. P. Kaput, R. L. Brinster, and R. D. Palmiter, Neuron 7, 703 (1991). 35. D. M. Donovan, M. Takemura, B. F. O'Hara, and M. T. Brannock, Proc. Natl. Acad. Sci. U.S.A. 89, 2345 (1992). 36. W. C. Friend, S. Clapoff, C. Landry, L. E. Becket, D. O. Hanlon, R. J. Allore, I. R. Brown, A. Marks, J. Roder, and R. J. Dunn, J. Neurosci. 12, 4337 (1992). 37. B. L. Largent, R. G. Sosnowski, and R. R. Reed, J. Neurosci. 13, 300 (1993). 38. S. Morita, K. Kobayashi, T. Mizuguchi, K. Yamada, I. Nagatsu, K. Titani, K. Fujita, H. Hidaka, and T. Nagatsu, Mol. Brain Res. 17, 239 (1993).

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[19]

Golgi Technique Used to Study Stress and Glucocorticoid Effects on Hippocampal Neuronal Morphology Ana Marfa Magarifios, Eberhard Fuchs, Gabriele Fltigge, and Bruce S. McEwen

Introduction More than a century ago, Camilo Golgi introduced the "reazione nera," the black reaction, or silver impregnation technique (12), which has become an invaluable tool in helping to understand the morphology of the nervous system. This technique is used to study isolated neurons in order to visualize their dendritic arbor structures, branching patterns, and density of spines. Given the high quality of morphological detail that can be obtained with this method, it also lends itself to being combined with electron microscopic procedures. In this way, it is possible to study the synaptic organization of neurons with great detail (2, 3, 29, 30). For a long time, Golgi's approach faced serious criticism; its unpredictability was the main concern. Today, modified versions of the original technique allow us to obtain much better results. Not only can the Golgi technique visualize normal neuronal structures, but it also permits the identification of atrophic neurons or neurons undergoing plastic change in response to natural stimuli or hormonal manipulations or as a result of certain diseases. Basically, the Golgi technique involves treatment of fixed neural tissue with potassium dichromate, followed by exposure to silver nitrate. The resulting impregnation consists of a reduced silver salt. In contrast, damaged and degenerating neurons are visualized by another technique that is based on the fact that tissue elements in traumatized neurons will catalyze the reduction of silver salts. This method, the silver degeneration technique, takes advantage of this property and labels only damaged cells. This brief review summarizes the uses of the Golgi technique for studies of stress-induced alterations in hippocampal neuronal morphology, but only after first considering the distinction between the modern version of the Golgi method and the methods for studying degenerating neurons and glial cells. Methods in Neurosciences, Volume 30 Copyright 9 1996 by Academic Press, Inc. All rights of reproduction in any form reserved.

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Degeneration Staining Methods for Neural Tisuse Because the classical Golgi method and the silver degeneration stain provide different information and yet are similar in some respects, it is important to consider how to discriminate between neurons damaged by primary injury in vivo and those damaged by secondary tissue changes induced by alterations in blood flow, oxygen supply, or uneven fixation. Even removing the brain immediately after perfusion can lead to artifacts. Thus, close attention should be paid in order to avoid distortion of the brain as well as to rule out the fact that the injury-induced reactions that produce the affinity for silver may take place in insufficiently fixed brain tissue (4). A recently developed method for silver degeneration staining has demonstrated how to minimize the "dark" neuron reaction as an artifact of brain manipulations at the time of perfusion (11, 31). Using appropriate controls and carefully regulated experimental conditions, it was shown that in normal, untreated brain tissue no cell body axons or dendrites of any neuron type are stained. Furthermore, the silver impregnation method can be potentially useful in detecting and assessing different stages of progressive neuronal damage. Another advantage of this method is the visualization of dendritic arborizations, which provides information about the different neuron types undergoing degeneration. It should be noted that other common histological techniques, such as cresyl violet staining, have been used to demonstrate traumatized neurons. Indeed, these cells are usually characterized by their darkly stained condensed chromatin and light or absent cytoplasm. The fact that the various processes are unable to be observed makes it difficult to recognize different neuronal types. Both cresyl violet and silver degeneration techniques have been used successfully in studying adrenalectomy-induced cell death in the dentate gyrus of the hippocampus in rats (13, 15, 26).

Applications of Single-Section Golgi Method to Investigate Effects of Stress and Glucocorticoids In the remainder of this chapter, we describe our studies on hippocampal morphology using a modification of the single-section Golgi impregnation technique (10). In particular, we examine the effect of chronic stress and glucocorticoid hormones on the structure of neurons in the rat hippocampus. Furthermore, using a model of psychosocial stress, we applied the same histological studies to examine hippocampal neuronal morphology in the tree shrew (Tupaia belangeri). Tree shrews are placental mammals whose brains

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combine primitive as well as primate-like features (18, 20). Because part of the central nervous system of tree shrews resemble primates, this species was formerly considered to be phylogenetically related to primates (14, 17). However, according to more recent evidence, tree shrews are classified as a separate order (Scandetia or Tupaiidae) (27). Confrontations between dominant and subordinate animals proved to induce effectively atrophy of dendrites of hippocampal neurons that are similar to those produced by glucocorticoid administration.

Materials and Methods

Experimental Animals For the chronic-resistant stress studies, male Sprague-Dawley rats (CD strain, Charles River, Kingston, NY), weighing 290-300 g at the beginning of the experiments, are housed in groups of three per hanging metallic cage with ad libitum access to food and tap water. Animals are maintained in a temperature- and light-controlled environment (12/12 hours, light/dark cycle; lights on from 0700 to 1900 hours). After handling the animals daily during 1 week, they are randomly assigned to experimental groups. For the psychosocial stress studies, male tree shrews (German Primate Center, G6ttingen, Germany) are housed individually on a regular day-night cycle (lights on from 0800 to 2000) with water and food ad libitum. Experimental Treatment Groups Restraint Stress in Rats Experiments are performed during the light period of the light-dark cycle. Rats are assigned to the following experimental groups: 1. Unstressed control groupmthe rats remain in their cages except for body weight recordings. 2. Chronic-restraint stress control groupnconsists of 6 hours/day (from 10 a.m. to 4 p.m.) restraint of the rats for 3 weeks in wire mesh restrainers, secured at the head and tail ends with clips. During the restraint sessions, the rats are placed again in their home cages. 3. Drug-treated chronic-restraint stress group--the rats receive daily restraint stress for 21 days as described above. Three different sets of rats receive daily injections of one of the following treatments: (1) 10 mg of corticosterone (CORT) in 250 ~1 of sesame oil, subcutaneously, (2) phenytoin (40 mg/kg), dissolved in propylene glycol, or (3) tianeptine (15 mg/kg), dis-

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solved in propylene glycol. Treatments (2) and (3), are applied prior to the restraint sessions and the injections are given intraperitoneally. Psychosocial Stress in Tree Shrews Removal of the opaque partition between the cages of two males unknown to one another results in an active competition for control of the enlarged territory. After establishment of a stable dominant/subordinate relationship, the two males are separated by a transparent wire mesh. The wire mesh is removed every day for 1-2 hours. Morning urine samples are collected and the animals are weighed daily. This period of psychosocial stress lasts for 28 days. Control animals are housed in separate quarters in the animal facility. [For details see Fuchs et al. (9).]

Single-Section Golgi Staining Procedure Fixation of Brains At the end of the treatment period, the rats are deeply anesthetized with Metofane (Pitman-Moore, Mundelein, IL) and transcardially perfused with 150 ml of 4% paraformaldehyde in 0.1 M phosphate buffer with 1.5% (v/v) picric acid. The brains are then postfixed in the perfusate overnight at 4~

Potassium Dichromate Treatment Using a Vibratome, 100-/zm-thick sections are cut into a bath of 3% w/v potassium dichromate dissolved in distilled water. Sections are then processed according to a modified version of the single-section Golgi impregnation procedure (10). Brain sections are then incubated in an aqueous solution of 3% w/v potassium dichromate overnight, at room temperature. The following day, the tissues are rinsed twice--but only for a few s e c o n d s ~ i n distilled water and mounted onto plain slides.

Construction of Assemblies and Silver Nitrate Treatment After mounting tissue sections onto slides, the excess of potassium dichromate is carefully wiped off the tissue edges. A coverslip is glued over the section at the four corners and these assemblies are then incubated in a 1.5% w/v silver nitrate solution dissolved in distilled water overnight and in the dark. The construction of the assemblies is a critical step. If the sections are too wet, the undesired formation of crystals on top of the tissue sections is likely. If, on the other hand, the tissue is dried excessively, not enough potassium dichromate will be available to react with the silver nitrate solution and no impregnation will be evident. Thus, a critical degree of humidity is

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essential in order for the reaction to occur. Another crucial concern is to avoid trapping air bubbles between the tissue and the glass of the assemblies. If this happens, the silver nitrate solution will not diffuse evenly and certain areas will not be impregnated.

Initial Analysis of Impregnation It is possible to follow the progress of the labeling method with a light microscope. A quick examination will give information about the number and density of Golgi-impregnated cells in areas of interest. If the impregnation is not satisfactory, it is still feasible to recycle the sections: the crystals should be removed from the tissue by treating them with 1% sodium thiosulfate three separate times for 15 minutes each with constant agitation, followed by three separate washes in distilled water for 5 minutes each. From our experience, this recycling procedure produces adequate results but the quality of the tissue impregnation is not comparable with that obtained initially. In other words, in the case of quantitative evaluation of dendritic spines we do not recommend the recycling procedure because many structural details are lost and the results can be seriously underestimated. After 2 days of incubation in silver nitrate, the assemblies are carefully dismantled. Then the tissue sections are rinsed in distilled water and dehydrated through an ascending series of ethanol concentrations (95 and 100%). The sections are then defatted in Histoclear (Americlear), mounted onto gelatinized slides, and coverslipped under Permount (Fisher Scientific).

Data Analysis Slides containing brain sections are coded prior to quantitative analysis. This renders the experiment completely blind and the code is not broken until the analysis is completed. In order to be selected for analysis, Golgi-impregnated neurons must possess the following characteristics: (1) a location in the appropriate subregion of the rostral hippocampusunamely, within the region of the area CA3 extending from a point just below the apex of the lateral bend in the pyramidal cell layer to a point directly ventral to the most lateral extension of the upper limb of the dentate granule cell layer; (2) a dark and consistent impregnation throughout the entire length of the dendrites; and (3) relative isolation from neighboring impregnated cells, which could interfere with the analysis. Moreover, in order to avoid a partial picture, only cells located in the middle third of the tissue section are analyzed. For each brain, six to eight pyramidal cells from CA3c are selected. Each selected neuron is traced at 400 x magification using a light microscope with a camera lucida drawing tube attachment. From each pyramidal cell drawing, the number of dendritic branch points of each dendritic tree is determined.

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In addition, the length of the dendrites is determined for both apical and basal dendritic trees using a Zeiss interactive digitizing analysis system. Means are determined for each variable and for each brain. The resulting values are analyzed using a one-way ANOVA with Tukey HSD posthoc comparisons.

Results

Effects of Chronic-Restraint Stress and Glucocorticoids We have used Golgi impregnated tissue to assess the effects of chronicrestraint stress on the structure of the hippocampus. Close attention was paid to the selection of an equal number of the various cell types within each treatment group in order to validate the subsequent comparisons (7). All cells being analyzed exhibited optimal impregnation quality, regardless of the subtype they belonged to, or the degree of atrophy they displayed. CORT treatment during 21 days resulted in significantly fewer apical dendritic branch points in CA3 pyramidal neurons as contrasted with their control counterparts (36). In contrast, no differences in the number of CA3 pyramidal cell basal dendritic branch points were detected with CORT treatment. Quantitative analysis of CA1, CA2, and dentate gyrus neurons revealed no significant differences between treatment groups in their branching patterns. CORT treatment also resulted in a significantly reduced total apical dendritic length in CA3 pyramidal neurons (Fig. 1). Daily restraint stress for 3 weeks induced an identical atrophic effect of hippocampal CA3 pyramidal neurons (33). Once again, this morphologic change was not observed in neurons belonging to other areas of the hippocampus, namely, CA1, CA2, or dentate gyrus. Furthermore, the atrophy was restricted to the apical dendritic tree of CA3 neurons (Fig. 2).

Effects of Pharmacological Treatmets Watanabe et al. (34) showed that daily administration of phenytoin, an antiepileptic drug that interferes with excitatory amino acid release and excitatory synaptic transmission (25), prevented the atrophic branching pattern and the shrinkage in apical dendrites of CA3 area induced by both chronic-restraint stress and CORT treatment (Figs. 1 and 2). Another drug, tianeptine, led to the same result (35); this atypical antidepressant, known for facilitating serotonin uptake (21), was able to interfere with the atrophic effect. Recent results showed that a third drug, the steroid synthesis inhibitor,

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~

t~

.=

~9 0.001

.8 o

fi

0.000 control

cort

cort+ phenytoin

FIG. 1 Effects of corticosterone or corticosterone plus phenytoin on (A) dendritic branch points in CA3c pyramidal neurons of the rat hippocampus and (B) thymus/ body weight ratios (mean +_ SEM); ,, significant difference from control; p < 0.05. Reprinted with permission from Ref. 34. Y. Watanabe, E. Gould, H. A. Cameron, D. C. Daniels, and B. S. McEwen. Phenytoin prevents stress- and corticosteroneinduced atrophy of CA3 pyramidal neurons. Hippocampus 2(4), 431 (1992). Copyright 1992, Churchill Livingstone, New York.

322

PARADIGMS OF NEURAL INJURY

A ~-~ 2000 1 , 1500 ,

~9 1000" O

500 "

B

9

control

Stress

stress + phenytoin

control

stress

stress + phenytoin

2o] 15

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FIG. 2 Effect of a 21-day daily restraint stress and phenytoin plus stress on dendritic length (A) and dendritic branch points (B) in CA3c pyramidal neurons of the rat hippocampus (mean +_ SEM); ,, significant difference from control; p < 0.05. Reprinted with permission from Ref. 34. Y. Watanabe, E. Gould, H. A. Cameron, D. C. Daniels, and B. S. McEwen. Phenytoin prevents stress- and corticosterone-induced atrophy of CA3 pyramidal neurons. Hippocampus 2(4), 431 (1992). Copyright 1992, Churchill Livingstone, New York.

[19] GOLGI METHOD: STRESS AND GLUCOCORTICOIDS

323

cyanoketone, also blocked completely the chronic restraint-induced CA3 atrophy, and the apical branching pattern did not differ from control animals (19). Under these conditions basal levels of CORT were not affected but the stress-induced CORT response was blocked.

Effects of Psychosocial Stress After 28 days of daily confrontations, the Golgi impregnation studies showed that the apical trees of CA3 neurons in subordinate tree shrews were atrophic compared to control ones, as judged by the lower number of branch points and the decreased total dendritic length observed (8). In this study we also quantified the spine density in both basal and apical CA3 dendrites but no significant differences were detected between groups.

Discussion The atrophy observed in the hippocampus of either chronically stressed or CORT-treated rats might be considered as a very early pathophysiological transformation of these neurons, because adjacent sections stained with cresyl violet showed no significant number of pyknotic neurons or other signals of neuronal degradation. It could be argued that, because the Golgi technique does not label all neurons contained in a given area and the criteria followed to choose representative cells might rule out neurons that are poorly impregnated and perhaps atrophic, the results might reflect characteristics of a subgroup of nonrepresentative cells. This seems n o t to be the case in view of the Nissl stain results mentioned previously. Moreover, we have evidence employing the same impregnation technique that demonstrates the reversibility of the atrophy following the termination of stress. It is interesting to note that beyond the intrinsic differences in the nature of the stresses applied, two phylogenetically distant species showed the same specific atrophic pattern in response to two qualitatively different types of chronic stressors, namely, restraint and psychosocial confrontation. Yet the same altered morphological change was observed in spite of the fact that the restraint stress led to habituation of CORT secretion whereas the social confrontation seems not to induce tolerance in the cortisol response. In order to understand the physiological significance of this phenomena, we should note that CA3 apical dendrites are the main postsynaptic target for the mossy fibers (MF), which originate in the granule cells of the dentate gyrus (DG). The DG, in turn, receives excitatory amino acid (EAA) input from the entorhinal cortex (EC) (1, 5). Thus, CA3 neurons play a crucial role in mediating excitatory transmission between the EC, the DG, and the

324

PARADIGMS OF NEURAL INJURY CA1 pyramidal cells of the hippocampus. Moreover, numerous studies show that stress and CORT induce excitatory amino acid release and other EAAmediated metabolic actions (16, 22-24, 28). These pathways are known to be involved in aspects of learning and memory (6). In this respect, the morphological changes were correlated with altered performance of rats in solving maze tasks (32). However, not only do CA3 neurons receive EAA transmission through mossy fibers, but hippocampal interneurons are known to be heavily innervated too. Thus, an excess of EAA stimulation inducing atrophy of the postsynaptic elements in CA3 neurons is a p o s s i b l e ~ b u t not sufficient--explanation. In summary, it has yet to be established whether these morphological effects are a consequence of a potentiating role of glucocorticoids on detrimental effects of EAA neurotransmission, a result of the lack of integrity of inhibitory counterbalances, an effect of cortisol/corticosterone on other neurotransmitter systems (GABA, serotonin, etc.), or a protective response, given that a smaller dendritic surface would prevent CA3 pyramidal cells from being excessively stimulated by EAA. Nevertheless, our strategy to investigate and understand how the adult brain deals with stress is increasingly focusing on dendritic remodeling effects and their correlation with behavioral responses.

References 1. S. A. Bayer, in "The Rat Nervous System" (G. Paxinos, ed.), Vol. 1, p. 335. Academic Press, New York, 1985. 2. T. W. Blackstad, Z. Zellforsch 67, 819 (1965). 3. T. W. Blackstad, "The Interneuron" (M. Brazier, ed.), p. 391. UCLA Forum in Medical Sciences, Los Angeles, 1969. 4. J. Cammemeyer, Histochemistry 56, 97 (1978). 5. B. J. Clairbone, D. G. Amaral, and W. H. Cowan, J. Comp. Neurol. 246, 435 (1986). 6. H. Eichenbaum and T. Otto, Behav. Neural Biol. 57, 2 (1992). 7. J. M. Fitch, J. M. Juraska, and L. W. Washington, Brain Res. 479, 105 (1989). 8. E. Fuchs, G. Fl~igge, H. Uno, B. S. McEwen, and A. M. Magarifios, N. Y. Acad. Sci. Abstr. (1994). .

10. 11. 12. 13. 14. 15.

E. Fichs, O. J6hren, and G. FliJgge, Psychoneuroendocrinology 18 (1993). P. L. Gabbott and J. Somogy, J. Neurosci. Methods 11, 221,557 (1984). F. Gallyas, M. Tsu, and G. Buszaki, J. Neurosci. Methods 10, 159 (1994). C. Golgi, Arch. Ital. Biol. 3, 285. (Reprinted in ibid. 97) (1883). E. Gould, C. S. Woolley, and B. S. McEwen, Neuroscience 37, 367 (1990). D. E. Haines and D. R. Swinder, J. Hum. Evol. 1, 407 (1972). D. Jaarsma, F. Postema, and J. Korf, Hippocampus 2(2), 143 (1992).

[19] GOLGI METHOD: STRESS AND GLUCOCORTICOIDS

325

16. M. T. Lowy, L. Gault, and B. K. Yamamoto, J. Neurochem. 61, 1957 (1993). 17. W. P. Lucket, "Comparative Biology and Evolutionary Relationship of Tree Shrews." Plenum, New York, 1980. 18. J. S. Lund, D. Fitzpatrick, and A. L. Humphrey, in "Cerebral Cortex" (A. Peters and E. G. Jones, eds.), Vol. 3, p. 157. Plenum, New York, 1985. 19. A. M. Magarifios and B. S. McEwen, Neuroscience 69, 89 (1995). 20. R. B. Martin, in "Primate Origins and Evolution" (R. B. Martin, ed.), p. 191. Chapman and Hall, London, 1990. 21. T. Mennini, E. Mocaer, and S. Garratini, Naunyn-Schmiebergs, Arch. Pharmachol. 336, 478 (1987). 22. D. R. Packan and R. M. Sapolsky Neuroendocrinology 51, 613 (1990). 23. R. M. Sapolsky, "Stress, the Aging Brain and the Mechanisms of Neuron Death." MIT Press, Cambridge, Massachusetts, 1992. 24. R. E. M. C. Schasfoort, L. A. De Bruin, and J. Korf, Brain Res. 475, 58, 1988. 25. J. Skerrit and G. Johnston, Clin. Exp. Pharmacol. Physiol. 10, 527 (1983). 26. R. S. Sloviter, G. Valiquette, G. M. Abrams, C. Ronk, A. I. Sollas, L. A. Paul, and S. A. Neubort, Science 243, 535 (1989). 27. D. Starck, "Vergleichende Anatomie der Wirbeltiere auf Evolutionsbiologischer." (Grundlage, ed.), Vol. 1. Springer-Verlag, Berlin and New York, 1978. 28. B. A. Stein-Behrens, E. M. Elliot, C. A. Miller, J. W. Schilling, R. Newcombe, and R. M. Sapolsky, J. Neurochem. 58, 1730 (1992). 29. W. K. Stell, Anat. Rec. 153, 389 (1965). 30. W. K. Stell, Am. J. Anat. 121, 401 (1967). 31. A. Van den Pol and F. Gallyas, J. Comp. Neurol. 296, 654 (1990). 32. M. Villegas, C. Martinez, V. N. Luine, and B. S. McEwen, Brain Res. 639, 167 (1994). 33. Y. Watanabe, E. Gould, and B. S. McEwen, Brain Res. 588, 341 (1992). 34. Y. Watanabe, E. Gould, H. A. Cameron, D. C. Daniels, and B. S. McEwen, Hippocampus 2(4), 431 (1992). 35. Y. Watanabe, E. Gould, H. A. Cameron, D. C. Daniels, and B. S. McEwen, Eur. J. Pharmacol. 222, 157 (1992). 36. C. S. Woolley, E. Gould, and B. S. McEwen, Brain Res. 531, 225 (1990).

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Index

ACAS 570 interactive laser cytometer, 145-146 N-Acetylaspartate, in normal brain and cerebral ischemia, 215 Acetyl-L-carnitine, effects on Alzheimer's disease brain, in vivo 31p MRS, 205-206 Acidity, cytosolic, and reactive oxygen species generation, 245 Adenylate kinase release, cell injury assessment, 9 Age effects on cerebral glucose/energy metabolism, 132-133 on serotonin neuronal system, 116-117, 121-122 Aluminum chloride, neurotoxicity, astrocyte role, 75-76 Alzheimer's disease /3-APP expression in transgenic mouse brain, 307-312 1H MRS in vitro studies, 203 31p MRS studies acetyl-L-carnitine treatment effects, 205-206 in vitro, 203 in vivo, 204-205 3-Aminobenzamide, effects on NAD + depletion, 14-18 a-Aminohydroxy-5-methyl-4-isoxazoleproionic acid, lesioning of basal forebrain nuclei, 280-282 /3-Amyloid protein precursor, expression in transgenic mice animal maintenance, 303-304 brain amyloidogenic fragments, histological analysis, 307-312 C 100 neurotoxicity, 298 DNA construction, 299-302 polymerase chain reaction, 302-303 protein product detection, 305-307 RNA detection, 305-307 Southern blot analysis, 302-303 Anesthesia, for studies of cerebral glucose/energy metabolism, 132 Antibodies, to/3/A4 fragment, detection of amyloid deposition, 308-309, 308-312 Antioxidants, intracellular, determination, 252-254 J3-APP, see j3-Amyloid protein precursor Apparent diffusion coefficient vs. cerebral blood flow in ischemic cortex, 219-222

changes after ischemia, 223-224 characteristics, 211 comparison with blood flow autoradiography for early stroke assessment, 217-224 threshold determination, 218-219 in vasogenic edema, 224 Arginine, depletion, methods, 40 Ascorbic acid, intracellular, and oxidative status, 253 Astrocytes heavy metal accumulation in cultured astroglia atomic absorption spectroscopy, 136-139 cellular fluorescence imaging, 144-146 gap junctional intercellular communication, 159-164 glutathione content, 146-153 intracellular Ca 2+ measurements, 153-159 kinetics constants, 139-143 mitochondrial membrane potential, 146-153 lead gliotoxicity, 74-75 1-methyl-4-phenyl- 1,2,3,6-tetrahydropyridine effects, 76-77 role in aluminum chloride neurotoxicity, 7576 Astrogliosis, C6 glial cell models, 69-71 Autoradiography blood flow, comparison with apparent diffusion coefficient, 217-224 for local cerebral glucose utilization, 129 Avian sarcoma virus-induced glioma model (rodent), 85-87 Avidin-biotin technique, for cultured cells, 47

Basal forebrain, cholinergic lesions cholinergic immunotoxins, 284-286 ethylcholine aziridinium, 282-284 excitotoxins, 280-282 general procedures, 278-279 mechanical damage, 279-280 Benzoic acid, hydroxylation, for reactive oxygen species determination, 249-250 Biochemical markers A2B5, 58 galactocerebroside, 47, 58-59 glial fibrillary acidic protein, 47, 58-59

327

328

INDEX

glutamine synthetase, 58, 66-67 myelin basic protein, 47 3',5'-cyclic-nucleotide phosphohydrolase, 58, 66-67 transferrin, 47 vimentin, 58 Blood flow, cerebral vs. apparent diffusion coefficient in ischemic cortex, 219-222 autoradiography, comparison with apparent diffusion coefficient, 217-224 correlation with diffusion-weighted imaging, 231-235 and diffusion-weighted imaging hyperintensity, 231-235 flow indices, calculation, 228-227 mismatch with diffusion-weighted imaging, 222 Bulbectomy, olfactory effects on p75 NGFRexpression in olfactory bulb, 102-104

procedure, 98 Buthionine sulfoximine, inhibition of glutamine synthesis, 173-174, 254

genes regulating apoptosis, 2 Calcium, intracellular, measurement interpretation, 156-159 limitations, 156-159 method, 155-156 rationale for, 153-155 0-Carotene, intracellular, and oxidative status, 253 Catalase, role in reactive oxygen species detoxification, 253 Cell cultures, primary neurons nitric-oxide synthase expression, 32-33 procedure, 30-32 tissue culture plate preparation, 28 Cell death assays, cell counting, 34-35 reactive oxygen species in, 3-7 Cell lines developed from nitrosourea-induced glioma (rodent), 88 PC12, nerve growth factor effects on oxidant injury (rat pheochromocytoma), 10-19 Cell survival, assessment methods, 8-9 Cell viability, assessment methods, 8-9 Cerebral blood flow, see Blood flow, cerebral C a e n o r h a b d i t i s elegans,

Cerebral glucose/energy metabolism, see Glucose/ energy metabolism, cerebral Cerebral perfusion pressure, control of cerebral blood flow, 124-125 Chemiluminescence method, Western analysis, 270 Cholinergic lesions, basal forebrain cholinergic immunotoxins, 284-286 ethylcholine aziridinium, 282-284 excitotoxins, 280-282 general procedures, 278-279 mechanical damage, 279-280 Cholinergic neurons, aspirative transection of timbria fornix materials, 109-110 surgical procedure, 110-112 Chronic-restraint stress, effects on hippocampal morphology, 320 Ciliary neurotrophic factor, induction of astrogliosis, 272-273 Copper, 67Cu(II) efflux from glial cells kinetic parameters, 142 time dependence, 142 uptake by glial cells kinetic parameters, 140-141 time dependence, 140 Correlation time, in magnetic resonance spectroscopy, 186 Corticosterone, effects on hippocampal morphology, 320-321 Cresyl violet staining, 316 Crosslinking, protein-phospholipid, 6 Cyanoketone, effects on hippocampal morphology, 323 Cyclic GMP, for nitric oxide formation in cultures, 36 Cyclin D1. expression in dying cells, 2 Cystathionase, assay, 170 Cysteine bound, assay, 169-170 in central nervous system, 167-168 chemical forms in vivo, 169 metabolic pathway, 168 total, assay, 169-170 Cytokines, injection into CNS animal preparation, 262 immunocytochemistry, 266-269 model system, 261-262 procedure, 262-264 tissue fixation, 265-266

INDEX tracer dyes, 262 Western analysis, 269-272 Cytomegalovirus promoter, for/3-gal gene transfection, 291 Cytometer, interactive laser, ACAS 570, 145-146 Cytotoxicity, glial cell aluminum chloride effects, 75-76 C6 cells, 59-71 ethanol effects, 72-74 lead effects, 74-75 methylmercuric chloride effects, 77-78 1-methyl-4-phenyl- 1,2,3,6-tetrahydropyridine effects, 76-77 primary cultures, 56-59 Dichlorofluorescein, assay of reactive oxygen, 249 Diffusion-weighted imaging, s e e Magnetic resonance imaging, diffusion-weighted 5,7-Dihydroxytryptamine, serotonergic lesions applications, 121-122 experimental design, 117-118 functional impact of denervation, 121 intracerebral, 120-121 intraventricular, 118-120 3-(4,5-Dimethylthiazol-2-yl)-2,5-diphenyl-2Htetrazolium bromide reduction applications, 9 in PC12 cells, 10 DNA damage fragmentation internucleosomal, 3 during oxidant injury in PC12 cells, 13-14 poly(ADP-ribose) synthase activation, 38 strand breakage, 6-7 Dye-exclusion measurements, of cell viability, 9 Dyes, tracer, for cytokine injections, 262 Edema, vasogenic, 224 Eicosanoids, production, 245-247 Electron spin resonance detection of free radical species, 248-250 low molecular weight iron compounds, 254 Energy homeostasis, reactive oxygen species effects, 6-7, 13 Energy metabolism, cerebral, s e e Glucose/energy metabolism, cerebral Enzyme-linked immunosorbent assay, nerve growth factor after liposome-mediated transfection, 294-295 Enzymes, mitigating oxidative damage, 252-254

329 Ethanol, gliotoxicity, 72-74 Ethylcholine aziridinium, lesioning of basal forebrain nuclei, 282-284 N-Ethylnitrosourea, glioma model (rodent), 81-85 Excitotoxins interactions with reactive oxygen species, 3 lesioning of basal forebrain nuclei, 280-282 Fenton reaction, OH (hydroxyl radical) formation, 4 Fimbria fornix, aspirative transection, 108-109 materials, 109-110 surgical procedure, 110-112 Fixation, tissue cryoprotection, 265-266 perfusion technique, 265 sectioning, 265-266 Fluorescence imaging ACAS 570 interactive laser cytometer, 145-146 for cell viability, 9-10, 35 rational for, 144 Fluorometry glutathione assay, 253-254 tryptophan content of solubilized proteins, 252 Free induction decay, 181-182 Freeze-thaw methods C6 glial cells, 61-66 oligodendroglia data interpretation, 52-53 experimental procedures, 45-48 immunocytochemical analysis of cultures, 50-52 observation of samples, 48-50 Galactocerebroside markers, 47 in reanimated glial cultures, immunofluorescence, 50-53 Gap junctional intercellular communication, measurement interpretation, 162-164 method, 159-161 rationale for, 159-160 Genes, C a e n o r h a b d i t i s e l e g a n s , role in apoptosis, 2 Gene transfer, liposome-mediated cytomegalovirus promoters, 291 nerve growth factor expression characteristics, 293-296 transfection efficiency, 291-293

330

INDEX

Glial cells biochemical markers, 57-59 C6 glioma cell passage procedure, 60-61 early passage, 67-69 freezing, 61-63 glial phenotype characterization, 66-67 late passage, 67-69 thawing, 63-66 oligodendrocytes, freeze-thaw method data interpretation, 52-53 freezing medium, 45 immunocytochemical characterization, 47-48, 50-52 medium, 46 preparation, 45 preparation for freezing, 45 reanimation, 46 sample fixation, 50 sample observation, 48-50 primary cultures E3H-derived immunocytochemical characterization, 58-59, 62-63 preparation, 57 E15CC-derived immunocytochemical characterization, 58-59, 62-63 preparation, 57 mixed astrocyte-oligodendrocyte, 56-57 Glial fibrillary acidic protein expression after cytokine injection into CNS immunocytochemistry, 267-269 Western analysis, 269-272 markers, 47 in reanimated glial cultures, immunofluorescence, 50-53 Glioma models, rodent avian sarcoma virus, 85-87 cell lines developed from, 88 nitrosourea compounds, 81-85 transplantation, 87-92 Glucocorticoids, effects on hippocampal morphology, 320 Glucose/energy metabolism, cerebral age effects, 132-133 anesthetics, 132 biochemical analyses, 130-131 blood flow control mechanisms, 124-125 enzyme activities, 131-132

in experimental animals, 129-132 in humans, 125-129 in vivo tissue studies, 129-130 Kety-Schmidt technique, 126-127 nuclear magnetic resonance in experimental animals, 131 in humans, 128 physiological steady state, 125 positron emission tomographic techniques, 127-128 postmortem studies, 128-129 substrate utilization, 129 Glutamate neurotoxicity, NO-initiated, model for, 28 y-Glutamyltransferase, assay, 252 Glutathione chemical forms in vivo, 169 cytosolic, analysis, 146-150, 152-153 high-performance liquid chromatography assay, 172-173 inhibitors, 173-174 intracellular, and oxidative status, 253 metabolism, 171 role in protection, 171 sources, 170-171 spectrophotometric assay, 172 tissue levels, determination, 254 Glutathione peroxidase in oxidative injury, 5 role in reactive oxygen species detoxification, 253 Golgi method, single-section staining procedure, 318-320 Greiss reaction, for nitric oxide formation in cultures, 36 Heavy metals, effects on cultured astrogial cells atomic absorption spectroscopy, 136-139 cytosolic glutathione content, 146-150, 152-153 fluorescence imaging, 144-146 gap junctional intercellular communication, 159-164 intracellular Ca 2§ content, 153-159 mitochondrial membrane potential, 146-148, 150-152 transport kinetics, 139-143 Hemoglobin, reduction, 41 High-performance liquid chromatography biochemical analysis of cerebral glucose/energy metabolism, 130-131

INDEX cysteine, 173 glutathione, 172-173 Hippocampus, neuronal morphology chronic-restraint stress effects, 320 glucocorticoid effects, 320 pharmacological treatment effects, 320-323 psychosocial stress effects, 323 single-section Golgi staining, 318-320 Hydrogen peroxide, toxicity in PC12 cells, nerve growth factor effects, 11-18 Ibotenic acid, lesioning of basal forebrain nuclei, 280-282 Immunocytochemistry amyloidogenic fragments of fl-amyloid protein precursor expression in transgenic mouse brains, 307-312 C6 glial cells, 67 characterization of reanimated oligodendroglia, 47-52 ED1 monoclonal antibody expression after cytokine injection, 267-269 E3H- and E15CC-derived cultures, 58-59, 62-63 glial fibrillary acidic protein expression after cytokine injection, 267-269 Immunofluorescence, characterization of reanimated oligodendroglia, 47-52 Immunohistochemistry, p75 NGFRexpression in olfactory system after bulbectomy, 98-101, 103-104 Immunotoxins, 192IgG-saporin conjugate, lesioning of basal forebrain nuclei, 284-286 Indicator-dilution method, for mean transit time measurement, 226-227 In vitro injury models, oligodendroglia freeze-thaw method, 45-47 immunocytochemical characterization, 47-53 Ion regulation, reactive oxygen species effects, 6 Iron compounds, low molecular weight assay, 254 in neurological disorders, 244 Ischemic penumbra and chemical shift imaging spectroscopy, 215-217 therapeutic window, 210-211 Kety-Schmidt technique, 126-127 Kinetics constants interpretations, 142-143 limitations, 142-143

331 measurement methods, 140-142 rationale for measurement of, 139-140 k-space substitution method, 227-228 Lactate cerebral tissue, in stroke, 183-184 proton resonances, 183 Lactate dehydrogenase, release for cell injury assessment, 9 in PC12 cells, 10 Larmor frequency expression for, 180 field dependence, 182 Lead accumulation in cultured astroglia atomic absorption spectroscopy, 136-139 cellular fluorescence imaging, 144-146 gap junctional intercellular communication, 159-164 glutathione content, 146-153 mitochondrial membrane potential, 146-153 gliotoxicity, 74-75 Lipid peroxidation, cytotoxic effects, 6 Lipofusin, formation, 251 Magnetic moment, nuclear, 178-179 Magnetic resonance imaging contrast agents, 227 diffusion-weighted, 211-213, 218 correlation with cerebral blood flow, 231-235 mismatch with cerebral blood flow, 222-223 sensitivity and specificity, 218 in vivo glioma metabolism (human), 91 perfusion techniques, 226 cerebral blood flow-diffusion-weighted imaging hyperintensity, 231-235 contrast agents, 227 diffusion-weighted spin-echo image, 226 flow indices, 228-227 k-space substitution, 227-228 relative mean transit time and cerebral blood flow images, 226-228 region of interest analysis, 230-231 Magnetic resonance spectroscopy applications Alzheimer's disease brain, 203-206 freeze-clamped Fischer 344 rat brain, 201-203 chemical shifts N-acetylaspartate in cerebral ischemia, 215 applications, 183

332

INDEX

and ischemic penumbra, 215-216 lactate in stroke, 213-215 lactate resonances, 183-184 correlation time, 186 free induction decay, 181-182 information contained in spectrum, 186-187 in vitro

quantitation of spectra, 194-195 signal identification, 193-194 in vivo

quantitation of spectra, 195-197 signal identification, 194 spatial localization depth-pulse technique, 190-191 spin-echo technique, 191-193 Larmor frequency, 180 line broadening chemical exchange, 188-189 correlation time effects, 190 field inhomogeneity, 188 magnetic field effects, 179-180 nuclear magnetic moment, 178-179 nuclear spin types, 179 radio frequency pulse application, 179-180 resonance frequency of a nucleus, 185 sample preparation, 197 saturation of nuclear magnetization, 181 sensitivity, 187 signal identification, in vitro spectra, 193-194 spin coupling, 187 spin relaxation, 185-186 Magnetic resonance spectroscopy, 1H instrumental conditions, 198 in vitro, Alzheimer's disease brain, 203 perchloric acid of brain tissue, 183 Magnetic resonance spectroscopy, 31p freeze-clamped Fischer 344 rat brain, 201-203 instrumental conditions, 197-198 in vitro

Alzheimer's disease brain, 203 perchloric acid of brain tissue, 184 in vivo

acetyl-L-carnitine treatment of Alzheimer's disease brain, 205-206 Alzheimer's disease brain, 204-205 with depth-pulse localization technique, 198-199 with spin-echo localization technique, 199-200 with spin-echo pulse sequence, 191-193

Mean arterial blood pressure, control of cerebral blood flow, 124-125 Mean transit time, blood through brain, indicatordilution method, 226-227 Membrane potential, mitochondrial, analysis, 150-153 Messenger RNA, nerve growth factor, after liposome-mediated transfection, 294 Metal ions, multivalent, 245 N-Methyl-D-aspartate, neurotoxicity, attenuation, 27-30 N-Methyl-D-aspartic acid, lesioning of basal forebrain nuclei, 280-282 Methylmercuric chloride, gliotoxicity, 77-78 N-Methylnitrosourea, glioma model (rodent), 81-85 1-Methyl-4-phenyl- 1,2,3,6-tetrahydropyridine, Parkinsonism-inducing effects, astrocyte role, 76-77 Mitochondria function, cytotoxicity assays for, 35 membrane potential, analysis, 150-153 superoxide anion production, 3-4 Monochlorobimane, reaction with glutathione, 253-254 Monoclonal antibodies 192IgG-saporin conjugate, lesioning of basal forebrain nuclei, 284-286 ED1, immunoreactivity after cytokine injection into CNS, 267-269 04, 47, 51-52 Myelin basic protein markers, 47 in reanimated glial cultures, immunofluorescence, 50-53

NAD + depletion nerve growth factor effects, 13-18 in NO-initiated glutamate neurotoxicity, 28 poly(ADP-ribose) polymerase inhibitor effects, 13-18 NADPH-diphorase stain, for nitric-oxide synthase in primary neuronal cultures, 32-33, 39 Nerve growth factor effects on oxidant injury in PC12 cells, 10-19 expression after liposome-mediated gene transfer, 293-296 liposome-mediated gene transfer in septohippocampal cultures, 290-293

INDEX Nerve growth factor receptor p75 NGFRexpression, unilateral bulbectomy effects bulbectomy procedure, 98, 102 immunohistochemistry, 98-101, 103-104 intranasal irrigation, 98, 101-102 P75 NGFRinteraction with neurotrophins, 1-2 Neurotoxicity aluminum chloride, astrocyte role, 75-76 fl-amyloid protein precursor C100, 298 glutamate, NO-initiated, 28 N-methyl-D-aspartate, attenuation, 27-30 and peroxynitrite formation, 28 Nitric oxide chemistry, 36-37 donor reagents, 41 formation, levels of regulation, 27 targets, 37-38 toxicity, in primary neuronal cultures cell death assay, 34-35 exposure, 33-34, 39-40 nitric-oxide synthase activation, 36 valence states, 36-37 Nitric-oxide synthase activation characterization, 27 criteria for, 36 in primary neuronal cultures, NADPH-diphorase stain for, 32-33, 39 Nitrosourea glioma models (rodent), 81-85 Nuclear magnetic resonance, techniques for cerebral glucose/energy metabolism in experimental animals, 131-132 ex vivo in vitro technique, 131 in humans, 128 Nuclei, properties, magnetic resonance spectroscopy-related, 180 Nucleus basalis magnocellularis, lesioning cholinergic immunotoxins, 284-286 ethylcholine aziridinium, 282-284 excitotoxins, 280-282 general procedures, 278-279 mechanical damage, 279-280 Oligodendrocytes glial cultures, freeze-thaw method data interpretation, 52-53 freezing medium, 45 immunocytochemical characterization, 47-48, 50-52

333 medium, 46 preparation, 45 preparation for freezing, 45 reanimation, 46 sample fixation, 50 sample observation, 48-50 lead gliotoxicity, 75 Oxidant injury experimental induction methods, 8 PC12 cells, nerve growth factor effects, 10-19 Oxidases, role in reactive oxygen species generation, 247 Oxidative stress, quantifiable parameters, 245-247 Partition pressure of carbon dioxide in arterial blood (paCO2), 124-125 Perchloric acid extract 1H MRS spectrum, 183 lap MRS in vitro spectrum, 184 31p MRS in vivo spectrum, 184 Peroxynitrite formation, and neurotoxicity, 28 synthesis, 42 Phagocytosis, role in reactive oxygen species generation, 247 Phenytoin, effects on hippocampal morphology, 320 pH levels, and reactive oxygen species generation, 245 Phosphorylation, oxidative, 245 Platelet-activating factor, C6 glial cell response, 69-71 Poly(ADP-ribose) polymerase activation, DNA damage-related, 38 inhibitors, effects on NAD + depletion, 13-18 Polyclonal antibodies, for Western analysis, 270-271 Polymerase chain reaction reverse transcription, s e e Reverse transcriptionpolymerase chain reaction transgene detection, 302-303 Positron emission tomography, techniques for cerebral glucose/energy metabolism, 127-128 Promoters cytomegalovirus, for fl-gal gene transfection, 291 neural, tested in transgenic mice, 300 Prooxidant status direct assays with electron spin resonance, 248-250

334

INDEX

dyes for, 244 indirect indices, 252-254 oxidation products of cellular constituents, 250-252 Propargylglycine, inhibition of glutamine synthesis, 174 Psychosocial stress, effects on hippocampal morphology, 323 Quinolinic acid, lesioning of basal forebrain nuclei, 280-282 Quisqualic acid, lesioning of basal forebrain nuclei, 280-282 Radio frequency pulse, application in magnetic resonance spectroscopy, 179-180 Radiolabeled precursors, for cell viability, 9-10 Reactive oxygen species cellular effects, 6-7 direct assay, 248-249 benzoate hydroxylation, 250 with dichlorofluorescein, 249 endogenous sources, 3-4, 245-247 cytosolic activity, 245 eicosanoid production, 245-247 metal ions with multivalence potential, 245 oxidases, 247 oxidative phosphorylation, 245 phagocytosis, 247 enzymes mitigating oxidative damage, 252-254 estimation using oxidation products of cellular constituents, 250-252 glutamine synthase assay, 252 protein tryptophan residue degradation, 252 exogenous sources, 4 interactions with excitotoxins, 3 intracellular antioxidant determination, 252-254 metabolism, 4-5 Resonance, defined, 180 Reverse transcription-polymerase chain reaction C100 transgene RNA and protein products, 305-307 nerve growth factor mRNA, 294 Ribonucleic acid, C 100 transgene, RT-PCT detection, 305-307 RT-PCR, s e e Reverse transcription-polymerase chain reaction Saporin, 192IgG-saporin conjugate, lesioning of basal forebrain nuclei, 284-286

Saturation of nuclear magnetization, 181 Septohippocampal cell cultures liposome-mediated gene transfer in, 290-293 nerve growth factor expression after transfection, 293-296 Serotonergic neurons, 5,7-dihydroxytryptamineinduced lesioning experimental design, 117-118 functional impact of denervation, 121 intracerebral, 120-121 intraventricular, 118-120 Silver degeneration staining, 316 Singlet oxygen, sources, 8 Southern blotting, transgene detection, 302-303 Spectrophotometry for biochemical analysis of cerebral glucose/energy metabolism, 130-131 glutathione assay, 172 Spin, nuclear, 179 Spin coupling, characteristics, 187 SPINECHO research pulse sequence, 192 Spin-lattice relaxation, characterization, 185 Spin-spin relaxation, characterization, 186 Stroke, ischemic early diagnosis/assessment apparent diffusion coefficient and blood flow autoradiography, 217-224 factors affecting, 209-210 early treatment, 210-211 entry time, 210-211 ischemic penumbra, 210-211 magnetic resonance imaging techniques animal model, 225-226 diffusion-weighted imaging, 211-213 magnetic resonance spectroscopy, 213-217 perfusion imaging, 224-235 therapeutic window, 210-211 Superoxide anion, production in mitochondria, 3-4 Superoxide dismutase, role in reactive oxygen species detoxification, 253 Superparamagnetic iron oxide contrast agent, 227

Tetrazolium salt reduction, for cell viability assessment, 9 Thanatins, characterization, 2 Therapeutic window, in ischemic stroke, 210-211 Thiol oxidation, protein inactivation by, 6 Tianeptine, effects on hippocampal morphology, 320

INDEX a-Tocopherol, intracellular, and oxidative status, 253 Transection, aspirative, fimbria fornix, 108-109 materials, 109-110 surgical procedure, 110-112 Transferrin markers, 47 in reanimated oligodendroglial cells, immunofluorescence, 50-53 Transgenic mice,/3-APP expression animal maintenance, 303-304 DNA construction, 299-302 polymerase chain reaction, 302-303 protein product detection, 305-307

335 RNA detection, 305-307 Southern blot analysis, 302-303 Transplantation glioma models (rodent), 87-92 Transverse relaxation, 186 Triton X-100, intranasal irrigation with, 98, 101-102 Tryptophan residues, determination, 252 Valence states, of nitric oxide, 36-37 Vasogenic edema, apparent diffusion coefficient, 224 Western blotting, glial fibrillary acidic protein expression after cytokine injection, 269-272

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FIG. 2 Double immunofluorescence of primary glial cultures. (A and B) Cultures 2 weeks after reanimation; (C and D) nonfrozen 4-week-old sister cultures. Reanimated cultures were characterized by the presence of GC + cells, the majority in clusters (A); these cells had cell processes. Single GC + cells are seen in B (again no processes observed). The presence of GFAP + cells with long processes was very frequent (45% approximately). Magnification: x200.

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FIG. 3 Triple immunofluorescence for the expression of GFAP (green), Tf (red), and 0 4 (blue) in oligodendroglial cultures 3 days after reanimation. (A-C) Cultures fed with DF-10; (D-F) sister cultures fed with GDM. Cells in DF-10 were flat and without distinct cell processes. 100% of the cells in the cultures expressed GFAP, Tf, and 0 4 at different intensities. Diffuse staining was observed for both GFAP and 0 4 whereas Tf + vesicles were frequently seen on the cell membrane or extracellularly (arrows). Cells fed with the glial development medium also shared the expression of the three antigens, GFAP, Tf, and 04. However, the cell morphology was different from DF-10 cells. (D) Weak expression of GFAP at the level of the cell body. Interestingly, 0 4 and Tf staining revealed the preservation of myelinlike sheaths. Both antigens were expressed in the cell by all the cells as well as in membranous cell processes (arrowheads). Magnification: x200. Other cells had modest cytoplasmic extensions (small arrows).

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