Current Protocols in Toxicology

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Wiley InterScience: Reference Work: Current Protocols in Toxicology

Current Protocols in Toxicology Copyright © 2005 by John Wiley & Sons, Inc. All Rights Reserved. ISBN: 0-471-24106-7 Last time updated: September 2005

Current Protocols in Toxicology is a "best-practices" collection of lab protocols for accurate, efficient assessments of toxicity in whole organisms, organs and tissues, cells, and biochemical pathways. Continuously updated since its initial publication in May 1999, this quarterly-updated two-volume set... - provides the latest models and methods from molecular biology, cell biology, biochemistry, and genetics, plus sophisticated toxicological procedures from leading laboratories. - offers expert guidelines for evaluating the effects of substances on human physiology and metabolism. - provides valuable reference information in three appendices, including stock solutions and equipment, commonly used techniques, and using information.

Edited by: Lucio G. Costa (University of Washington); Ernest Hodgson (North Carolina State University); David A. Lawrence (Wadsworth Center); Terence R. Ozolins (Pfizer, Inc.); Donald J. Reed (Oregon State University); William F. Greenlee, Advisory Editor (CIIT); Past Editor-in-Chief: Mahin Maines; Past Editors: I. Glenn Sipes, Shigeru Sassa Series Editor: Kathy Morgan

Editors EDITORIAL BOARD Lucio G. Costa University of Washington Seattle, Washington Ernest Hodgson North Carolina State University Raleigh, North Carolina David A. Lawrence Wadsworth Center Albany, New York Donald J. Reed Oregon State University Corvallis, Oregon ADVISORY EDITOR William F. Greenlee CIIT Centers for Health Research Research Triangle Park, North Carolina PAST EDITOR-IN-CHIEF Mahin D. Maines University of Rochester School of Medicine Rochester, New York PAST EDITORS Shigeru Sassa Rockefeller University New York, New York I. Glenn Sipes University of Arizona Tucson, Arizona

Chapter 1 Toxicological Models Introduction Unit 1.1 Nonhuman Primates as Animal Models for Toxicology Research Unit 1.2 Statistical Approaches to the Design of Toxicology Studies Unit 1.3 Transgenic Animals in Toxicology Unit 1.4 DNA Microarrays: An Overview of Technologies and Applications to Toxicology Unit 1.5 The Use of Fish-Derived Cell Lines for Investigation of Environmental Contaminants Unit 1.6 Sea Urchin Embryos and Larvae as Biosensors for Neurotoxicants Unit 1.7 Zebrafish: An Animal Model for Toxicological Studies Unit 1.8 Preclinical Models of Parkinson's Disease Chapter 2 Assessment of Cell Toxicity Introduction Unit 2.1 Current Concepts in Cell Toxicity Unit 2.2 Determination of Apoptosis and Necrosis Unit 2.3 Detection of Covalent Binding Unit 2.4 Measurement of Lipid Peroxidation Unit 2.5 Measurements of Intracellular Free Calcium Concentration in Biological Systems Unit 2.6 In Vitro Methods for Detecting Cytotoxicity Unit 2.7 In Situ Hybridization Histochemistry Unit 2.8 Confocal Microscopy Unit 2.9 Measurement of Expression of the HSP70 Protein Family Unit 2.10 Analysis of Mitochondrial Dysfunction During Cell Death Chapter 3 Genetic Toxicology: Mutagenesis and Adduct Formation Introduction Unit 3.1 The Salmonella (Ames) Test for Mutagenicity Unit 3.2 Measurement of a Malondialdehyde-DNA Adduct Unit 3.3 Mutagenesis Assays in Mammalian Cells Unit 3.4 Cell Transformation Assays Unit 3.5 Assays for DNA Damage Unit 3.6 Detecting Epigenetic Changes: DNA Methylation Unit 3.7 Assays for Detecting Chromosomal Aberrations Unit 3.8 Methods for Measuring DNA Adducts and Abasic Sites I: Isolation, Purification, and Analysis of DNA Adducts in Intact DNA Unit 3.9 Methods for Measuring DNA Adducts and Abasic Sites II: Methods for Measurement of DNA Adducts Chapter 4 Techniques for Analysis of Chemical Biotransformation Introduction Unit 4.1 Measurement of Cytochrome P-450 Unit 4.2 Purification of Cytochrome P-450 Enzymes Unit 4.3 Measurements of UDP- Glucuronosyltransferase (UGT) Activities Unit 4.4 Detection of Metabolites Using High-Performance Liquid Chromatography and Mass Spectrometry Unit 4.5 Measurement of Aryl and Alcohol Sulfotransferase Activity Unit 4.6 Measuring the Activity of Arylamine N-Acetyltransferase (NAT) Unit 4.7 Measurement of Carboxylesterase (CES) Activities Unit 4.8 Analysis of the Aryl Hydrocarbon Receptor (AhR) Signal Transduction Pathway Unit 4.9 Measurements of Flavin-Containing Monooxygenase (FMO) Activities Unit 4.10 Assays for the Classification of Two Types of Esterases: Carboxylic Ester Hydrolases and Phosphoric Triester Hydrolases Unit 4.11 Techniques for Measuring the Activity of Carboxylic Acid:CoA Ligase and Acyl-CoA:Amino Acid N-Acyltransferase: The Amino Acid Conjugation Pathway Unit 4.12 Determination of Paraoxonase 1 Status and Genotypes at Specific Polymorphic Sites Unit 4.13 Human Cytochrome P450: Metabolism of Testosterone by CYP3A4 and Inhibition by Ketoconazole Unit 4.14 Biotransformation Studies Using Rat Proximal Tubule Cells Unit 4.15 TaqMan Real Time—Polymerase Chain Reaction Methods for Determination of Nucleotide Polymorphisms in Human N-Acetyltransferase-1 (NAT1) and -2 (NAT2) Unit 4.16 Evaluation of the Cytochrome b5/Cytochrome b5 Reductase Pathway Unit 4.17 Measurement of Xenobiotic Carbonyl Reduction in Human Liver Fractions

Chapter 5 Toxicokinetics Introduction Unit 5.1 Measurement of Bioavailability: Measurement of Absorption Through Skin In Vitro Unit 5.2 Measurement of Bioavailability: Measuring Absorption Through Skin In Vivo in Rats and Humans Unit 5.3 Measurement of Disposition Half-Life, Clearance, and Residence Times Unit 5.4 Isolated Perfused Porcine Skin Flap Unit 5.5 Porcine Skin Flow-Through Diffusion Cell System Unit 5.6 Toxicant Transport by P-Glycoprotein Unit 5.7 Collection of Bile and Urine Samples for Determining the Urinary and Hepatobiliary Disposition of Xenobiotics in Mice Chapter 6 The Glutathione Pathway Introduction Unit 6.1 Overview of Glutathione Function and Metabolism Unit 6.2 Measurement of Glutathione and Glutathione Disulfide Unit 6.3 Measurement of Glutathione Transport Unit 6.4 Measurement of Glutathione Transferases Unit 6.5 HPLC-Based Assays for Enzymes of Glutathione Biosynthesis Unit 6.6 -Glutamyl Transpeptidase Activity Assay Unit 6.7 Oxidant-Induced Regulation of Glutathione Synthesis Unit 6.8 Measurement of Glutathione Conjugates Unit 6.9 Coenzyme A and Coenzyme A-Glutathione Mixed Disulfide Measurements by HPLC Chapter 7 Assessment of the Activity of Antioxidant Enzymes Introduction Unit 7.1 Analysis of Glutathione-Related Enzymes Unit 7.2 Measurement of Glutathione Reductase Activity Unit 7.3 Analysis of Superoxide Dismutase Activity Unit 7.4 Measurement of Thioredoxin and Thioredoxin Reductase Unit 7.5 Measurement of MnSOD and CuZnSOD Activity in Mammalian Tissue Homogenates Unit 7.6 Measurement of Ascorbic Acid and Dehydroascorbic Acid in Biological Samples Chapter 8 Heme Synthesis Pathway Introduction Unit 8.1 The Heme Biosynthesis Pathway and Clinical Manifestations of Abnormal Function Unit 8.2 Measurement of ALA Synthase Activity Unit 8.3 Measurement of Heme Concentration Unit 8.4 Measurement of Uroporphyrinogen Decarboxylase Activity Unit 8.5 Measurement of Protoporphyrinogen Oxidase Activity Unit 8.6 Measurement of -Aminolevulinate Dehydratase Activity Unit 8.7 Measurement of Ferrochelatase Activity Unit 8.8 Measurement of Erythrocyte Protoporphyrin Concentration by Double Extraction and Spectrofluorometry Unit 8.9 HPLC Methods for Analysis of Porphyrins in Biological Media Chapter 9 Heme Degradation Pathway Introduction Unit 9.1 Overview of Heme Degradation Pathway Unit 9.2 Detection of Heme Oxygenase Activity by Measurement of CO Unit 9.3 Detection of Heme Oxygenase 1 and 2 Proteins and Bilirubin Formation Unit 9.4 Detection of Biliverdin Reductase Activity Unit 9.5 Histochemical Analysis of Heme Degradation Enzymes Unit 9.6 An HPLC Method to Detect Heme Oxygenase Activity Unit 9.7 Functional Analysis of the Heme Oxygenase-1 Gene Promoter Unit 9.8 Quantitation of Human Heme Oxygenase (HO-1) Copies by Competitive RT-PCR Unit 9.9 Purification and Characterization of Heme Oxygenase Chapter 10 The Nitric Oxide/Guanylate Cyclase Pathway Introduction Unit 10.1 Overview of the Pathway and Functions of Nitric Oxide Unit 10.2 Assay of Tissue Activity of Nitric Oxide Synthase Unit 10.3 Detection of Nitrosated Proteins Unit 10.4 Fluorometric Techniques for the Detection of Nitric Oxide and Metabolites Unit 10.5 Measurement of cGMP and Soluble Guanylyl Cyclase Activity Unit 10.6 Histochemical Analysis of Nitric Oxide Synthase by NADPH Diaphorase Staining Unit 10.7 Immunocytochemical Analysis of Cyclic Nucleotides Unit 10.8 Methods for Distinguishing Nitrosative and Oxidative Chemistry of Reactive Nitrogen Oxide Species Derived from Nitric Oxide Unit 10.9 Inducible Nitric Oxide Synthase Expression

Chapter 11 Neurotoxicology Introduction Unit 11.1 Overview of Neurotoxicology Unit 11.2 Neurobehavioral Screening in Rodents Unit 11.3 Assessment of Spatial Memory Unit 11.4 Advanced Behavioral Testing in Rodents: Assessment of Cognitive Function in Animals Unit 11.5 Testing for Organophosphate-Induced Delayed Polyneuropathy Unit 11.6 Risk Assessment and Neurotoxicology Unit 11.7 Neurobehavioral Testing in Humans Unit 11.8 Mouse Models of Global Cerebral Ischemia Unit 11.9 Mouse Models of Focal Cerebral Ischemia Unit 11.10 Principles of Electrophysiology: An Overview Unit 11.11 Electrophysiological Studies of Neurotoxicants on Central Synaptic Transmission in Acutely Isolated Brain Slices Unit 11.12 Whole-Cell Patch-Clamp Electrophysiology of Voltage-Sensitive Channels Unit 11.13 Detection and Assessment of Xenobiotic-Induced Sensory Neuropathy Unit 11.14 Methods to Produce Brain Hyperthermia Chapter 12 Biochemical and Molecular Neurotoxicology Introduction Unit 12.1 Biochemical Approaches to Studying Neurotoxicity Unit 12.2 Development of an In Vitro Blood-Brain Barrier Unit 12.3 Culturing Rat Hippocampal Neurons Unit 12.4 Isolation of Neonatal Rat Cortical Astrocytes for Primary Cultures Unit 12.5 Analytical Cytology: Applications to Neurotoxicology Unit 12.6 Estimating Cell Number in the Central Nervous System by Stereological Methods: The Optical Disector and Fractionator Unit 12.7 Isolation of Cerebellar Granule Cells from Neonatal Rats Unit 12.8 Measurement of Glial Fibrillary Acidic Protein Unit 12.9 Aggregating Neural Cell Cultures Unit 12.10 Coculturing Neurons and Glial Cells Unit 12.11 Determining the Ability of Xenobiotic Metals to Bind a Specific Protein Domain by Electrophoresis Unit 12.12 Morphological Measurement of Neurotoxic Injury in the Peripheral Nervous System: Preparation of Material for Light and Transmission Electron Microscopic Evaluation

Chapter 13 Teratology Introduction Unit 13.1 Overview of Teratology Unit 13.2 Rat Embryo Cultures for In Vitro Teratology Unit 13.3 Micromass Cultures in Teratology Unit 13.4 Using Chicken Embryos for Teratology Studies Unit 13.5 In Vivo Assessment of Prenatal Developmental Toxicity in Rodents Unit 13.6 Organ Culture of Midfacial Tissue and Secondary Palate Unit 13.7 Overview of Behavioral Teratology Unit 13.8 Statistical Analysis of Behavioral Data

Chapter 14 Hepatotoxicology Introduction Unit 14.1 Overview of Hepatotoxicity Unit 14.2 Preparation of Hepatocytes Unit 14.3 Small Animal Models of Hemorrhagic Shock—Induced Liver Dysfunction Unit 14.4 Isolation of Liver Kupffer Cells Unit 14.5 Measurement of Hepatobiliary Transport Chapter 15 Gene Targeting Introduction Unit 15.1 Embryonic Stem (ES) Cell Culture Basics Unit 15.2 Genotyping Embryonic Stem (ES) Cells Unit 15.3 Aggregation Chimeras (ES Cell—Embryo) Unit 15.4 Reporter Genes to Detect Cre Excision in Mice

Chapter 16 Male Reproductive Toxicology Introduction Unit 16.1 In Vivo Models for Male Reproductive Toxicology Unit 16.2 Guidelines for Mating Rodents Unit 16.3 Histopathology of the Male Reproductive System I: Techniques Unit 16.4 Histopathology of the Male Reproductive System II: Interpretation Unit 16.5 Monitoring Endocrine Function in Males: Using Intra-Atrial Cannulas to Monitor Plasma Hormonal Dynamics in Toxicology Experiments Unit 16.6 Epididymal Sperm Count Unit 16.7 Performing a Testicular Spermatid Head Count Unit 16.8 Transgenerational (In Utero/Lactational) Exposure to Investigate the Effects of Endocrine Disrupting Compounds (EDCs) in Rats Chapter 17 Oxidative Stress Introduction Unit 17.1 Formation and Functions of Protein Sulfenic Acids Unit 17.2 Measurement of Protein Sulfenic Acid Content Unit 17.3 Fluorescence Microplate Reader Measurement of Tissue Susceptibility to Lipid Peroxidation Unit 17.4 In Situ Localization of Nonenzymatic Peroxidase-Like Activity of Tissue-Bound Transition Metals Unit 17.5 F2-Isoprostanes as Markers of Oxidant Stress: An Overview Unit 17.6 Quantification of F2-Isoprostanes by Gas Chromatography/Mass Spectrometry as a Measure of Oxidant Stress Unit 17.7 Immuno-Spin Trapping: Detection of Protein-Centered Radicals

Chapter 18 Immunotoxicology Introduction Unit 18.1 Associating Changes in the Immune System with Clinical Diseases for Interpretation in Risk Assessment Unit 18.2 Local Lymph Node Assays Unit 18.3 Murine Asthma Models Unit 18.4 Use of Bronchoalveolar Lavage to Detect Lung Injury Unit 18.5 Measuring Lymphocyte Transcription Factor Activity by ELISA Unit 18.6 Measuring the Activity of Cytolytic Lymphocytes Unit 18.7 Solid-Phase Immunoassays Unit 18.8 Immune Cell Phenotyping Using Flow Cytometry Unit 18.9 In Vitro Model for Modulation of Helper T Cell Differentiation and Activation Appendix 1 Using Information 1A Safe Use of Radioisotopes 1B Transgenic and Gene-Targeted Mouse Lines for Toxicology Studies Appendix 2 Laboratory Stock Solutions and Equipment 2A Common Stock Solutions and Buffers 2B Standard Laboratory Equipment Appendix 3 Commonly Used Techniques 3A Molecular Biology Techniques 3B Techniques for Mammalian Cell Tissue Culture 3C Enzymatic Amplification of DNA by PCR: Standard Procedures and Optimization 3D Detection and Quantitation of Radiolabeled Proteins in Gels and Blots 3E Northern Blot Analysis of RNA 3F One-Dimensional SDS Gel Electrophoresis of Proteins 3G Spectrophotometric Determination of Protein Concentration 3H Dialysis and Concentration of Protein Solutions 3I The Colorimetric Detection and Quantitation of Total Protein Appendix Suppliers Selected Suppliers of Reagents and Equipment

FOREWORD

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oxicological research is driven by the need to understand and assess the human and ecological risks of exposure to chemicals and other toxicants as well as by interest in using toxic agents to elucidate basic biological and pathobiological processes. The level of research activity in this field is higher, the rate of change in knowledge more rapid, and interest in applying scientific information to societally important issues is greater than ever before. These are exciting and challenging times to be working in toxicology. The ongoing ferment builds on the extraordinary advances being made in the understanding of biological systems at the molecular level. This fundamental knowledge provides the opportunity for greatly enhanced insight into how chemicals and other stressors may damage biological structures and processes, influence the rate of biological repair, and lead to reversible or irreversible diseases or to a return to health. Society increasingly calls on the scientific community for the knowledge needed both to reevaluate the health hazards of existing products and technologies and to evaluate the prospective hazards of new ones. Such information is used to develop guidelines and regulations designed to ensure that these new products and technologies do not harm people or the environment. Acquiring sound, reproducible scientific data that can be integrated with existing information to advance the knowledge of toxicants and living systems requires rigorous adherence to the scientific method. This means intelligent, thoughtful individuals identifying important needs, formulating testable hypotheses, designing experiments to test them, meticulously conducting these experiments, carefully reviewing and interpreting data, and ultimately presenting this information to scientific peers, including publishing it in the peer-reviewed literature. Current Protocols in Toxicology is a clear and well-documented compendium of the most important methods in the field—proven approaches developed by leading researchers—for the benefit of other experimentalists, from students to seasoned investigators. Since toxicology by its nature is multidisciplinary, other titles in the Current Protocols series may also provide relevant methods. Although review of the literature cited for each procedure can give added insight into the underlying theory and breadth of applications, the protocols have been carefully designed to provide clear, step-by-step descriptions that can easily be followed even by the relatively inexperienced. Regular updates to Current Protocols in Toxicology manual will help ensure an awareness of changes in previously documented methods and of methods newly developed. Use of these protocols will avoid unnecessary duplication of effort in development and validation when the methods are applied without modification, and will speed up the development of more refined methods that will further advance the field of toxicology and, in turn, may have a place in future updates. Roger O. McClellan Chemical Industry Institute of Toxicology Research Triangle Park, North Carolina

Current Protocols in Toxicology Contributed by Roger O. McClellan Current Protocols in Toxicology (1999) Copyright © 1999 by John Wiley & Sons, Inc.

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PREFACE

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he span of research in toxicology has been expanding and diversifying precipitously in recent years. One cause for this is the ongoing increase in industrial activity and in the generation of toxic compounds that then find their way into the environment. Another is the intensifying public awareness of the health effects of chemical exposure. The expansion of the field can be observed by attending any major scientific event dedicated to toxicology—such as the annual meetings of the Society of Toxicology, whose attendance has tripled in the course of the 1990s. Examining the meeting program for one of these events provides a very good feel for the broad scope of toxicology. For those who have attended such meetings periodically over the past few years, the dynamic nature of the field and its explosive growth is obvious: there is simply more in-depth research going on every year. This is in contrast to toxicology’s early years, when the field was dominated by research involving gross assessment of organisms’ responses to toxic chemicals. More recent times have witnessed the emergence of applications of state-of-the-art technology to the study of toxicity responses in organisms and living cells, along with phenomenal advancement in molecular and biochemical techniques, which increasingly are finding their way into toxicology research laboratories. A growing number of presentations at toxicology meetings constitute bridges between basic toxicology research and approaches to improving human health and environmental quality. It is this changing and expanding face of toxicology and its methodologies that represented the greatest challenge in assembling Current Protocols in Toxicology. We have attempted to include those methods that are presently central to modern toxicology and that we expect will remain valuable tomorrow. Like the field of toxicology, with its quarterly supplements this book will continue to expand in scope, to include more topics and methods as the field advances.

Because toxicological questions may be addressed using methods deriving from a wide variety of disciplines, other titles in the Current Protocols series may also provide methods that can be applied in your research. Molecular biology techniques, in particular, are integral to toxicological investigation. Such techniques are included where appropriate within units in this book; however, where these protocols are located may not be readily apparent from the table of contents. To help you find them, Table A.3A.1 in APPENDIX 3A provides a listing of specific techniques and where they can be found, either in this book or in related Current Protocols manuals. In addition, protocols for a number of basic techniques will be added to APPENDIX 3 in future supplements. Although mastery of the techniques in this manual will enable readers to pursue research in toxicology, the manual is not intended to be a substitute for graduate-level courses or a comprehensive textbook in the field. An inevitable hazard of manual writing is that protocols may become obsolete as the field expands and new techniques are developed. To safeguard this manual from inexorable obsolescence (and perhaps pleasantly surprise the users of the manual!), we provide quarterly supplements to provide protocols that utilize new innovations and technologies in the field. The updatable formats—looseleaf binder, CD-ROM, Intranet, and online Internet—easily accommodate the addition of this new material.

Current Protocols in Toxicology Contributed by Mahin D. Maines, Lucio G. Costa, Donald J. Reed, Shigeru Sassa, and I. Glenn Sipes Current Protocols in Toxicology (1999) iii-vi Copyright © 1999 by John Wiley & Sons, Inc.

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HOW TO USE THIS MANUAL Format and Organization This publication is available in both looseleaf and CD-ROM format. For looseleaf purchasers, a binder is provided to accommodate the growth of the manual via the quarterly update service. This format allows easy insertion of new pages, units, and chapters that are added. The index and table of contents are updated with each supplement. CD-ROM purchasers receive a completely new disc every quarter and should dispose of their outdated discs. The material covered in the two versions is identical. Subjects in this manual are organized by chapters, and protocols are contained in units. Protocol units, which constitute the bulk of the book, generally describe a method and include one or more protocols with listings of materials, steps and annotations, recipes for unique reagents and solutions, and commentaries on the “hows” and “whys” of the method. Other units present more general information in the form of explanatory text with no protocols. Overview units contain theoretical discussions that lay the foundation for subsequent protocols. Other discussion units present more general information. Page numbering in the looseleaf version reflects the modular arrangement by unit; for example, page 1.2.3 refers to Chapter 1 (Toxicological Models), UNIT 1.2 (Statistical Methods in Toxicology), page 3 of that particular unit. Many reagents and procedures are employed repeatedly throughout the manual. Instead of duplicating this information, cross-references among units are used and recipes for common reagents are supplied in APPENDIX 2A. Cross-referencing helps to ensure that lengthy and complex protocols are not overburdened with steps describing auxiliary procedures needed to prepare raw materials and analyze results. Introductory and Explanatory Information Because this publication is first and foremost a compilation of laboratory techniques in toxicology, we have included explanatory information where required to help readers gain an intuitive grasp of the procedures. Some chapters begin with special overview units that describe the state of the art of the topic matter and provide a context for the procedures that follow. Chapter and unit introductions describe how the protocols that follow connect to one another, and annotations to the actual protocol steps describe what is happening as a procedure is carried out. Finally, the Commentary that closes each protocol unit describes background information regarding the historical and theoretical development of the method, as well as alternative approaches, critical parameters, troubleshooting guidelines, anticipated results, and time considerations. All units contain cited references and many indicate key references to inform users of particularly useful background reading, original descriptions, or applications of a technique. Protocols

Preface

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Many units in the manual contain groups of protocols, each presented with a series of steps. One or more basic protocols are presented first in each unit and generally cover the recommended or most universally applicable approaches. Alternate protocols are provided where different equipment or reagents can be employed to achieve similar ends, where the starting material requires a variation in approach, or where requirements for the end product differ from those in the basic protocol. Support protocols describe additional steps that are required to perform the basic or alternate protocols; these steps are separated from the core protocol because they might be applicable to other uses in the manual, or because they are performed in a time frame separate from the basic protocol steps.

Reagents and Solutions Reagents required for a protocol are itemized in the materials list before the procedure begins. Many are common stock solutions, others are commonly used buffers or media, while others are solutions unique to a particular protocol. Recipes for the latter solutions are provided in each unit, following the protocols (and before the commentary) under the heading Reagents and Solutions. It is important to note that the names of some of these special solutions might be similar from unit to unit (e.g., electrophoresis buffer) while the recipes differ; thus, make certain that reagents are prepared from the proper recipes. On the other hand, recipes for commonly used stock solutions and buffers are provided once in APPENDIX 2A. These universal recipes are cross-referenced parenthetically in the materials lists rather than repeated with every usage. Commercial Suppliers Throughout the manual, we have recommended commercial suppliers of chemicals, biological materials, and equipment. In some cases, the noted brand has been found to be of superior quality or it is the only suitable product available in the marketplace. In other cases, the experience of the author of that protocol is limited to that brand. In the latter situation, recommendations are offered as an aid to the novice in obtaining the tools of the trade. Experienced investigators are therefore encouraged to experiment with substituting their own favorite brands. Addresses, phone numbers, and facsimile numbers of all suppliers mentioned in this manual are provided in the SUPPLIERS APPENDIX. Safety Considerations Anyone carrying out these protocols may encounter the following hazardous or potentially hazardous materials: (1) radioactive substances, (2) toxic chemicals and carcinogenic or teratogenic reagents, and (3) pathogenic and infectious biological agents. Check the guidelines of your particular institution with regard to use and disposal of these hazardous materials. Although cautionary statements are included in the appropriate units, we emphasize that users must proceed with the prudence and precaution associated with good laboratory practice, and that all materials must be used in strict accordance with local and national regulations. Animal Handling Many protocols call for use of live animals (usually rats or mice) for experiments. Prior to conducting any laboratory procedures with live subjects, the experimental approach must be submitted in writing to the appropriate Institutional Animal Care and Use Committee (IACUC) or must conform to appropriate governmental regulations regarding the care and use of laboratory animals. Written approval from the IACUC (or equivalent) committee is absolutely required prior to undertaking any live-animal studies. Some specific animal care and handling guidelines are provided in the protocols where live subjects are used, but check with your IACUC or governmental guidelines to obtain more extensive information. Reader Response Most of the protocols included in this manual are used routinely in the authors’ laboratories. These protocols work for them; to make them work for you they have annotated critical steps and included critical parameters and troubleshooting guides in the commentaries to most units. However, the successful evolution of this manual depends upon readers’ observations and suggestions. Consequently, a self-mailing reader-response

Current Protocols in Toxicology

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survey can be found at the back of the manual (and is included with each supplement); we encourage readers to send in their comments. ACKNOWLEDGMENTS This manual is the product of dedicated efforts by many of our scientific colleagues who are acknowledged in each unit and by the hard work by the Current Protocols editorial staff at John Wiley and Sons. We are extremely grateful for the critical contributions by Kathy Morgan (Series Editor), who kept the editors and the contributors on track and played a key role in bringing the entire project to completion, and by Gwen Crooks and Virginia Chanda, who provided developmental support in the early stages of the project. Other skilled members of the Current Protocols staff who contributed to the project include Joseph White, Kathy Wisch, Michael Gates, Demetra Kagdis, Alice Ro, Scott Holmes, Tom Cannon, and Alda Trabucchi. The extensive copyediting required to produce an accurate protocols manual was ably handled by Rebecca Barr, Allen Ranz, Elizabeth Harkins, Ben Gutman, Karen Hopkin, Monte Kendrick, Caroline Lee, Candace Levy, and Cathy Lundmark, and electronic illustrations were prepared by Gae Xavier Studios.

Mahin D. Maines, Lucio G. Costa, Donald J. Reed, Shigeru Sassa, and I. Glenn Sipes

Preface

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CHAPTER 1 Toxicological Models INTRODUCTION

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his chapter illustrates a variety of general models and approaches that can be used in toxicological studies. As such, it is considerably broader and more diverse than other chapters in Current Protocols in Toxicology, presenting a broad group of methodological approaches, in vivo models, and in vitro systems. As well as established toxicological protocols, the chapter will cover both traditional and novel methods developed in other disciplines that have potential application to toxicology. examines the role of nonhuman primates as animal models in toxicology research. While not used extensively, for certain obvious reasons (such as cost and the necessity for special facilities), their similarity animals to humans makes nonhuman primates invaluable in certain aspects of toxicology. The unit discusses several areas of investigation (including reproductive toxicology, neurobehavioral toxicology, and immunotoxicology) where studies in nonhuman primates have provided important data relating to understanding mechanisms of toxicity and setting safe levels of exposure to toxicants.

UNIT 1.1

Because the end-points of toxicity used in in vivo or in vitro studies are so diverse, use of the appropriate statistical approach is of the utmost importance. UNIT 1.2 reviews statistical methods in toxicology, with an emphasis on the approaches that should be used with different toxicological tests. The ability to generate transgenic animals, most often mice, that overexpress or lack a certain protein (such as an enzyme or receptor) has been one of the major achievements in life science research over the past several years. The availability of transgenic animals allows a much better understanding of the physiological functions of proteins of interest and of their potential role in chemical toxicity. UNIT 1.3 discusses strategies and applications of transgenic animals in toxicology, as well as methods currently used to generate transgenic mice. DNA microarrays, also known as DNA “chips,” allow detection of expression of RNA for thousands of genes that can be modified by toxic chemicals. In addition, they can be used to detect DNA sequence polymorphisms, thus providing a powerful method to assess genetic variations. An overview of the technologies of DNA microarrays and their applications to toxicology is presented in UNIT 1.4. UNIT 1.5 describes a series of methods for the preparation and use of fish-derived cell lines for cytotoxicity testing of environmental contaminants. Another interesting model system for toxicity testing is represented by sea urchin embryos and larvae. The model, described in UNIT 1.6, appears to be particularly promising for studies of the effects of developmental neurotoxicants.

describes yet another rather novel test system—the zebrafish—which has potential for a number of applications. UNIT 1.7

Toxicological Models Contributed by Lucio G. Costa Current Protocols in Toxicology (2003) 1.0.1-1.0.2 Copyright © 2003 by John Wiley & Sons, Inc.

1.0.1 Supplement 18

Two important in vivo models for studying Parkinson’s disease are discussed in UNIT they utilize treatments with MPTP in mice and non-human primates and with 6-OHDA in rats.

1.8;

Upcoming units will discuss in vitro methods to assess toxicity and genotoxicity in mammalian cells among other topics. Lucio G. Costa

Introduction

1.0.2 Supplement 18


Nonhuman Primates as Animal Models for Toxicology Research The use of nonhuman primates in biomedical research has a long and distinguished history (Bennett et al., 1995). An integral part of this history is biomedical research in the area of toxicology. The purpose of this unit is to present an overview of the use of nonhuman primate models in toxicological research. The unit is organized into five sections. The first section provides an overview of the extensive work with nonhuman primate models in the areas of reproductive toxicology and teratology (birth defects). The second section focuses on neurotoxicology research, including a brief discussion of nonhuman primate models for Parkinson’s disease and methanol-induced ocular toxicity. This section also offers an overview of studies that used infant nonhuman primate models to investigate the neurobehavioral toxicology of early exposure to environmental pollutants (lead, methylmercury, polychlorobiphenyls) and drugs of abuse (ethanol, cocain e) . Section three focuses on immunotoxicology. Recent studies that used nonhuman primate models to examine the effects of polychlorobiphenyls (PCBs) and early ethanol exposure are provided as examples. The fourth section discusses research in respiratory or lung toxicology and highlights the use of nonhuman primate models in studies of inhaled particles. The final section provides an overview of the use of nonhuman primate models for research in chemical carcinogenesis. This section also discusses long-term National Cancer Institute studies that used nonhuman primates in tumor-incidence research. More recent uses of nonhuman primates in studies of the role of diet in the development of cancer are also presented. The unit closes with a few comments on other important uses of nonhuman primates in toxicological research. Although this unit describes the numerous contributions of nonhuman primate models in toxicology, it is important to keep in mind that the majority of toxicology research is conducted using rodent animal models. Rodents have more diverse behavioral repertoires, are less expensive to purchase (thus allowing larger sample sizes), and are easier to care for than nonhuman primates. Rodents also develop quickly, so adult physical stature and sexual maturity are reached in months instead of years.

Mazue and Richez (1982) delineated the benefits and problems associated with using nonhuman primates in toxicological research. Issues such as phylogenetic proximity and physiologic, metabolic, and behavioral similarity were listed as benefits, whereas supply, small sample sizes, the potential for disease transmission to humans, and cost were listed as problems. In addition to the above, the ethical use and treatment of nonhuman primates is an issue of great importance in toxicology research. Although the ethical issues are not specific to nonhuman primate research or research in toxicology (Dennis, 1997), researchers in toxicology must weigh these issues—such as why nonhuman primates are necessary in the investigation of toxic effects and how many animals are required to define potential toxicity—carefully when the use of nonhuman primates is considered. The nonhuman primates most frequently used in toxicology are members of the Macaca genus and include the crab-eating macaque (M. fascicularis), the rhesus macaque (M. mulatta), and the pig-tailed macaque (M. nemestrina). Less widely used are the baboon, squirrel monkey, and chimpanzee. The specific requirements for housing and maintenance of these animals are described in the congressional Animal Welfare Act (AWA). (To obtain a copy of the AWA, call the USDA at 916-857-6205.) Administered by the US Department of Agriculture (USDA), this act covers all warmblooded animals, with the exception of rats, mice, and birds. The US Public Health Service (USPHS) requires that all institutions supported by the National Institutes of Health (NIH) meet or exceed the regulations published in the AWA. Briefly, the minimum space (cage size) that must be provided to nonhuman primates is based on the animal’s weight, except for brachiating species (those that rely on an overhead arm swing for locomotion) and the great apes (i.e., chimpanzees, orangutans, and gorillas). The AWA contains a table for calculating appropriate cage size. Animals are typically fed twice a day to support natural foraging behavior and to minimize the potential of clinical disorders, such as bloating. Purina High Fiber Monkey Chow (#5049) provides all the basic nutritional requirements, although diets are typically supplemented with vegetables and

Contributed by Thomas M. Burbacher and Kimberly S. Grant Current Protocols in Toxicology (1999) 1.1.1-1.1.9 Copyright © 1999 by John Wiley & Sons, Inc.

UNIT 1.1

Toxicological Models

1.1.1

fruits, such as grapes, apples, green peppers, cherry tomatoes, onions, potatoes, and yams. Water is typically available ad libitum. Recently, the AWA was amended to include environmental enrichment programs for nonhuman primates and dogs. Environmental enrichment for primates typically includes regular opportunities for social contact for grooming and play, chew toys, at least one perch in each cage, food treats, positive interaction with a caregiver or another familiar person, and daily visual and auditory contact with at least one animal of the same or a compatible species. When working with nonhuman primates, all laboratory personnel must follow special precautions for minimizing the transmission of zoonoses. These precautions include, but are not limited to, gloves, protective eyewear, shoe covers, and laboratory coats. Many of these steps have been adopted in response to the potential lethal nature of the herpes B virus. Because small research facilities find it difficult to meet all the necessary requirements, most nonhuman primate research in the United States is carried out at one of the seven NIHsponsored Regional Primate Research Centers (www.ncrr.nih.gov/compmed/cmrprc.htm).

REPRODUCTIVE TOXICOLOGY AND TERATOLOGY Reproductive Toxicology

Nonhuman Primates as Animal Models for Toxicology Research

The value of the nonhuman primate model in reproductive toxicology is largely based on similarities of the hypothalamic-pituitaryovarian-uterine axis in monkeys and humans. The general reproductive parameters shared between many nonhuman primates and humans include the plasma hormone patterns that support menstruation, the length of the menstrual cycle, the onset of chorionic gonadotropin secretion, placental structure, and length of gestation (Hendrickx and Cukierski, 1987). Macaque menstrual cycles are typically 28 days long with 3 to 5 days of actual menses, closely matching the human menstrual cycle. Conception rates vary among species and laboratories, ranging from 25% to 50% after a single mating. In addition, similarities in embryonic and fetal development are evident, beginning with the timing and length of organogenesis. As in humans, successful reproduction in primates requires a fertilized oocyte to implant in the endometrial lining of the uterus and complex hormonal interactions to successfully maintain the pregnancy. Once conception has taken place, both humans and macaques show similar

early pregnancy plasma hormone patterns. Organogenesis begins on day 21 in the macaque and day 18 in the human, ending on day 50 in the macaque and day 60 in the human. Although human placentas are monodiscoid (single lobed) and macaque placentas are bidiscoid (double lobed), placental function is virtually identical. The gestation of a full-term infant macaque ranges from 165 to 175 days (23 to 25 weeks), whereas human gestation is, on the average, ~280 days (40 weeks). Ovarian function and pregnancy are generally well understood in many nonhuman primate species and thus provide an opportunity to examine the relationship between toxicant exposure and reproductive dysfunction (e.g., alterations in menses and fertility). Studies can be designed to evaluate changes in the production of steroids after toxicant exposure and the role these changes may play in adverse reproductive outcomes. In addition to more immediate outcome measures, such as menses, ovulation, and pregnancy, reproductive processes at the opposite end of the reproductive continuum, such as menopause, can also be examined in lifespan studies (Sakai and Hodgen, 1988). Within the context of reproductive toxicology, Sakai and Hodgen (1988) pointed out the importance of minimally invasive experimental procedures. Examples of such procedures are the collection of blood during the menstrual cycle for analysis of gonadotropin and steroid concentrations, the collection of urine for analysis of steroids and pregnancy gonadotropins, and minor surgical procedures (e.g., laparoscopy and laparotomy). Behavioral methods have also been developed to minimize the handling and subsequent stress of adult female monkeys used for reproductive toxicology studies. Monkeys can be trained to present their perineum to human observers so menstrual bleeding can be detected, and early pregnancies can be reliably palpated by 3 to 4 weeks postconception (Burbacher et al., 1988). Various macaque species have been used as models in studies investigating the reproductive effects of exposure to environmental pollutants (lead, methylmercury) and of drug abuse (ethanol, cocaine). Procedures for evaluating the characteristics of the menstrual cycle (length, hormone status), breeding status (number of timed matings to conception, conception rate, live-birth delivery rate), and offspring viability (gestation length, birth size, perinatal mortality) are typically included in these studies. Results indicated that lead exposure suppresses circulating levels of luteinizing hormone (LH),

1.1.2 Current Protocols in Toxicology

follicle-stimulating hormone (FSH), and 17-βestradiol (E2) during the menstrual cycle in crab-eating macaques (M. fascicularis). The length of the menstrual cycle, the length of menses, and the circulating levels of progesterone were unaffected (Foster, 1992). Chronic methylmercury exposure was associated with a decrease in the number of live-born offspring in crab-eating macaques, but the menstrual cycle and menses lengths were again unaffected (Burbacher et al., 1988). Reproductive effects associated with ethanol exposure in pig-tailed macaques (M. nemestrina) also included a significant decrease in the number of live-born offspring (Clarren and Astley, 1992). This effect was primarily the result of an increase in the number of abortions. A study of the effects of maternal cocaine exposure in rhesus monkeys (M. mulatta) did not reveal significant effects on reproductive parameters (Morris et al., 1996a). The number of females investigated in this study (n = 3/group) may, however, have been too small to detect such effects.

Teratology One of the most important uses of nonhuman primate models has been in research aimed at identifying toxicants that cause birth defects. Compared to adults, embryos and fetuses exhibit an increased sensitivity to the structural and functional effects of many chemical compounds. Studies designed specifically to address the risk of birth defects are required to evaluate the health risks from exposure to certain environmental compounds or drugs. For drugs that are likely to be taken by pregnant women and for a widespread environmental pollutant, such as lead or methylmercury, studies using nonhuman primate models may be appropriate. The antinausea drug thalidomide provides a good example of the importance of using nonhuman primate models in teratology research. Limb malformations documented in infants born to women who used this drug during pregnancy were not observed in routine teratology tests using rodent models. Parallel effects were observed in nonhuman primates, including timing (sensitive period), type of malformation (limb defects), and the dose required to produce the teratogenic response (Hendrickx, 1973). The thalidomide episode established nonhuman primates as an important animal model for specific malformation syndromes seen in human infants. However, as Hendrickx and Binkerd (1990) noted, although nonhuman primates provided an excellent model for the

effects of thalidomide, this was not the case for the developmental effects after exposure to the rubella virus in early pregnancy. In a study evaluating fifteen known human teratogens, rodent models predicted a human teratogenic response ∼70% of the time, but nonhuman primate models predicted such a response only ∼50% of the time (Schardein et al., 1985). Although several factors may account for this—e.g., metabolic differences and the small sample size typical in nonhuman primate work—it is prudent to note that nonhuman primates do not always mimic humans in their teratogenic responses. Nonhuman primate models have been used in a wide range of studies aimed at assessing the safety of pharmaceutical agents, environmental pollutants, physical agents (e.g., X-rays), and drugs of abuse as well as the effects of infectious diseases. Hendrickx and Binkerd (1990) provided a comprehensive listing of the compounds that have been tested using nonhuman primate models and the corresponding indices of toxicity in the offspring (e.g., fetal death, structural malformations, growth retardation, and functional deficits). In addition to thalidomide, nonhuman primate models are particularly well recognized for helping elucidate the dysmorphology associated with prenatal exposure to vitamin A and its derivatives (retinoids), the anticonvulsant valproic acid, and triamcinolone acetonide (a synthetic glucocorticoid). Nonhuman primate models have also played a contributory role in defining the genital malformations associated with diethylstilbestrol.

NEUROTOXICOLOGY One of the best known nonhuman primate models in neurotoxicity is the 1-methyl-4phenyl-1,2,3,6-tetrahydropyridine (MPTP)– treated monkey (Kaakkola and Teravainen, 1990; Bezard et al., 1997). The neurotoxic effects after exposure to MPTP in human and nonhuman primates resemble those associated with Parkinson’s disease: i.e., hypokinesia, rigidity, resting tremor, stooping posture, dysphagia, depletion of striatal dopamine, and loss of cells in the substantia nigra. Response to drug therapy (e.g., levodopa) is also similar in humans and nonhuman primates exhibiting these symptoms. Studies using neural grafts in MPTP-treated monkeys reported a reduction in parkinsonism. Studies using MPTP-treated monkeys will continue to provide important information regarding the pathophysiology and neurochemical effects associated with Parkinson’s disease. Studies of potential innovative or



long-term drug therapies will also continue to use this animal model. Another excellent example of an important nonhuman primate model is ocular toxicity caused by methanol exposure. Methanol (methyl alcohol or wood alcohol) poisonings have been reported since the turn of the 20th century and are characterized by severe metabolic acidosis; ocular toxicity; and, in the most serious cases, coma and death. Early studies using nonhuman primate models indicated that formate (formic acid), a metabolite of methanol, was responsible for the toxicity associated with methanol intake. Human and nonhuman primates display similar effects from high-dose methanol exposure, owing to their limited capacity, compared with rodents, to metabolize formate to carbon dioxide (Black et al., 1985). Elevated formate concentrations are believed to cause the optic disc edema and optic nerve lesions associated with methanol poisoning. As is the case with MPTP exposure and induction of a parkinsonian-like condition, treatment with formate in the monkey induces the optic nerve toxicity commonly associated with human methanol poisoning. Use of this animal model has aided the development of treatment strategies designed to mitigate the severe and frequently permanent consequences of acute high-dose methanol intake.

Neurobehavioral Toxicology


The highly evolved behavioral repertoire of nonhuman primates makes them excellent subjects for investigations of the functional effects of neurotoxicants. The nonhuman primate model is especially useful in studies of developmental exposures and effects, because monkeys, like humans, have relatively long periods of gestation, infancy, and adolescence. Studies can investigate possible critical periods in development for neurotoxicant effects. Special testing procedures are available for infant nonhuman primates that target milestones in cognitive and sensory development and physical growth. Macaque and human infants share certain limitations and abilities, particularly during the first months of life. The emergence of reflexes like sucking, rooting, grasping, clasping, and righting can be evaluated as early as postnatal day 1. Infant rhesus macaques (M. mulatta) developmentally exposed to lead exhibit lower muscle tonus and increased agitation on tests of neonatal reflexes and behavioral organization compared to controls (Levin et al., 1988).

Early cognition can be studied in monkeys during the first months of life using procedures identical to those used to evaluate human infants. Tests of object permanence are generally believed to measure coordinated reaching responses and spatial memory. Studies of in utero exposure to methylmercury in crab-eating macaques (M. fascicularis) indicated a delay in object permanence development. On average, infant crab-eating macaques exposed in utero to methylmercury exhibited object permanence a full month after controls (90 versus 60 days; Burbacher et al., 1990a). Visual recognition memory can be measured in both humans and monkey infants using a test in which novel visual stimuli are paired with familiar stimuli; looking times to each are recorded (Fagan, 1990). Visual preferences for novel stimuli are considered evidence for recognition memory because some aspects of the familiar stimuli must be retained in memory for the novelty response to occur. Deficits in visual recognition memory have been found in a number of monkey groups at high risk for poor developmental outcome, including those exposed to known human teratogens (methylmercury, ethanol; Burbacher et al., 1990a). Studies with human infants also reported reduced visual recognition scores in infants prenatally exposed to PCBs (Jacobson et al., 1985). The development of primate social behavior appears relatively sensitive to neurotoxicant exposure. Infants exposed in utero to methylmercury exhibited reduced levels of social play and spent more time engaged in passive, nonsocial behaviors (Burbacher et al., 1990b). Infant monkeys fed lead acetate daily from birth to 1 year of age demonstrated disrupted social development, resulting in decreased levels of social play and increased levels of fear and self-stimulation (Laughlin et al., 1991). These effects persisted after dosing was terminated. Infants exposed in utero to 2,3,7,8-tetrachlorodibenzo-p-dioxin (TCDD) initiated more play, retreated less frequently, and were displaced less often from preferred positions in the playroom (Schantz et al., 1992). TCDD-exposed monkeys also displayed increased levels of self-directed behaviors. Several learning and memory assessments have been developed for older infant, juvenile, and adult nonhuman primates. Tests have been designed to study both spatial and nonspatial memory, using simple and complex learning paradigms—e.g., discrimination, alternation, reversal, and concept learning (matching and nonmatching to sample). Computer-controlled


presentation of test stimuli allows the opportunity to test both monkeys and children on identical measures of cognition. Studies using these procedures have also been used to investigate the effects of in utero cocaine exposure on learning in rhesus monkeys (Morris et al., 1996b). Rice and associates have had one of the most productive neurobehavioral toxicology programs using nonhuman primate models to study caffeine, lead, PCBs, and methylmercury. Results from their research program described learning deficits on several test procedures in monkeys exposed to lead during development (Rice, 1996). The performance of the lead-exposed monkeys was characterized by an inability to attend to relevant cues and to keep pace with changing environmental contingencies. These effects are similar to those observed in children exposed to lead (e.g., attention deficits). The assessment of sensory functioning is a frequently overlooked area in neurobehavioral toxicology studies. Sensory tests not only are valuable tools in evaluating toxicant-related brain injury but also provide a measure of neurotoxicity that is relatively unencumbered by psychological variables, such as learning ability. Vision is probably the best studied of all sensory systems and is certainly the dominant sense in both human and nonhuman primates. Tests of this nature typically involve assessment of visual acuity and contrast sensitivity and are based on a signal-detection paradigm. Monkeys exposed to chronic low levels of methylmercury from birth exhibited impaired spatial vision relative to controls under conditions of both high and low luminance (Rice and Gilbert, 1982). Studies indicate that in utero exposure to methylmercury also impairs spatial vision in adulthood (Burbacher et al., 1999). Auditory and somatosensory functioning were also evaluated using the signal-detection paradigm. Auditory detection thresholds were studied in monkeys who were exposed to methylmercury during their first 7 years of life (Rice and Gilbert, 1992). Results showed a selective high-frequency hearing loss in treated animals. Somatosensory function was also evaluated in monkeys exposed to methylmercury or lead (Rice and Gilbert, 1995). Animals exposed to methylmercury demonstrated elevated vibration detection thresholds on this procedure, whereas results from the lead-treated monkeys were somewhat equivocal.

IMMUNOTOXICOLOGY Studies of the anatomy and function of the immune system of nonhuman primates re-

ported many similarities to that of humans (Bleavins and de la Iglesia, 1995). Using monoclonal antibodies raised against human antigens, researchers noted extensive cross-reactivities in several nonhuman primate species, including macaques (Tryphonas et al., 1996). For example, a study by Ozwara et al. (1997) examined the reactivity of 161 antihuman monoclonal antibodies in chimpanzees, rhesus macaques, and squirrel monkeys. Antibodies directed against T cell surface antigens and against cytokine receptors were examined for their reactivity with peripheral blood mononuclear cells. The results of the study indicated that 38 of 161 monoclonal antibodies reacted in all three nonhuman primate species; 112 monoclonal antibodies reacted in one or two of the species. Chimpanzees showed the highest cross-reactivity (65%), followed by rhesus macaques (45%) and squirrel monkeys (42%). Tryphomas et al. (1996) reported extensive cross-reactivities with antihuman monoclonal antibodies in M. fascicularis infants. An important finding in this study was the reported sex differences in the levels of CD4 monoclonal antibodies and for the CD4/CD8 ratio (females > males), a sex difference similar to that observed in humans. Bleavins and de la Iglesia (1995) reported the results of a study aimed at developing a delayed-type hypersensitivity procedure using crab-eating macaques (M. fascicularis). Delayed-type hypersensitivity was measured using the human multitest cell-mediated immunity (CMI) skin test, which includes seven antigens. Responses to the skin tests paralleled those observed in humans. The authors proposed the use of this delayed-type hypersensitivity procedure in M. fascicularis for preclinical safety testing. A number of sensitive methods are available for evaluating the effects of compounds on the immune systems of humans and animals. In a series of articles, Luster et al. (1993) described a screening battery for evaluating the potential immunotoxicity of compounds in mice. Five parameters were included in the battery (immunopathology, humoral-mediated immunity, CMI, nonspecific immunity, and host resistance challenge) in a two-tier approach. Studies using nonhuman primate models of immune system toxicity have included assessments of these parameters. For example, Tryphomas (1995) published a series of reports describing the effects of PCB exposure on the immune system of adult rhesus macaques. The results of the study indicated that low-level chronic (55 months) exposure to PCBs (Aroclor 1254) was



associated with changes in several immunological parameters in the rhesus macaque (Table 1.1.1). These changes were most likely the result of altered T cell and/or macrophage function. Pig-tailed macaques (M. nemestrina) have been used to study the immune effects associated with fetal alcohol exposure (Grossmann et al., 1993). Monkeys exposed to ethanol in utero were more susceptible to disease and exhibited reduced T lymphocyte proliferation and lower titers to tetanus toxoid than did nonexposed controls. The reduction in T cell proliferation was consistent with reports from studies of children with fetal alcohol syndrome (FAS) and rodent models of prenatal alcohol exposure. The authors noted that several of the effects seemed sex dependent and cautioned investigators about the need to control for sex in nonhuman primate studies of immune system effects.

RESPIRATORY TOXICOLOGY Reports of cross-species comparisons of the anatomy and physiology of the respiratory system and the rates of deposition, clearance, and

retention of inhaled particles have described many similarities between nonhuman primates and humans (Snipes, 1989, 1996; Nikula et al., 1997). For example, humans and nonhuman primates clear particles from the alveolar region more slowly, have larger alveoli and alveolar ducts, and have more complex acini than do rodents. Nonhuman primate models have been used extensively in studies of dust-induced pulmonary lesions (Snipes, 1996). In rats, chronic inhalation of poorly soluble dusts causes “lung overload,” which can result in altered pulmonary clearance and pulmonary fibrosis. Humans and nonhuman primates exhibit a different pattern of dust accumulation in the lungs after chronic exposure. Whereas rats show fast pulmonary clearance of dust and retain dust predominantly in macrophages within the alveoli, human and nonhuman primates exhibit a slower pulmonary clearance of dust and retain dust burdens in the pulmonary interstitium. The rat pattern of dust accumulation may be related to the increased susceptibility of this animal to alterations in pulmonary clearance after chronic dust exposure compared to nonhuman primates. The rodent animal model may not

Table 1.1.1 Immunological Parameters Assessed in PCB-Exposed Rhesus Monkeysa,b

Parameter

Resultc

Cell-mediated immunity Lymphocyte proliferation

D

Host-resistance challenge Pneumococcus titers

N

Nonspecific immunity Serum complement (CH50) Natural killer cells Serum thymosin Monocyte activation Total interferon Interleukin Tumor necrosis factor

I I I D Id D N

Humoral-mediated immunity CD2, CD4, CD8, CD20 IgM and IgG titers

Ne D

aSummarized bSerum


from Tryphonas (1995). hydrocortisone levels were normal in all exposure

groups. cI = increased, N = normal, D = decreased. dSignificant increase in low and high groups; significant decrease in moderate group. eSignificant decrease in percent of total T lymphocytes (CD2).


provide data relevant to the risk of pulmonary disease after chronic dust exposure in humans. In addition to dust particles, respiratory effects have been described in nonhuman primate models after exposure to diesel exhaust (Nikula et al., 1997), ozone (Dimitriadis, 1993), marijuana smoke (Flifeil et al., 1991), and different forms of beryllium (Haley et al., 1994).

CLINICAL CARCINOGENESIS In 1961, the National Cancer Institute began a program aimed at examining the susceptibility of nonhuman primates to chemicals that were known to cause tumors in rodents. Since then, the long-term carcinogenic activity of several therapeutic agents, food additives and compounds, environmental contaminants, Nnitroso compounds, and model rodent carcinogens have been evaluated. In addition, valuable data have been collected regarding the incidence of spontaneous tumors in several nonhuman primate species. Thorgeirsson et al. (1994) reported that the spontaneous tumor rate over a 32-year period for 181 rhesus monkeys was 2.8% for malignant tumors and 3.9% for benign tumors. For 130 crab-eating macaques and 62 African green monkeys, the corresponding rates were 1.5% and 0.8% and 8% and 0%, respectively. Continuous dosing studies with the artificial sweeteners (cyclamate or saccharin) over a 22-year period provided no evidence of carcinogenic effects. Fungal food contaminants such as aflatoxin B1 and sterigmatocystin, however, were found to be potent hepatocarcinogens. 2-Amino-3-methylimidazo[4,5f]quinoline (IQ), an imidazole heterocyclic amine (HCA) present in cooked meat, was also found to be a potent hepatocarcinogen, inducing malignant liver tumors in 65% of monkeys tested during a 7-year dosing period. Snyderwine et al. (1997) reported that IQ is activated in monkeys via N-hydroxylation carried out by cytochrome P-450 CYP3A4 and/or CYP2C9/10. Human hepatic microsomes have been shown to have a greater capacity to activate HCAs compared to rodents and nonhuman primates. Current estimates of the daily intake of HCAs are on the order of 1 to 20 mg/person. Based on animal data, the estimates of the cancer risk to humans associated with this intake of HCAs are 10−3 to 10−4. Nonhuman primate models have also been used to examine the uptake and metabolic characteristics of suspected carcinogens when different susceptibilities are observed in rodent models. For example, studies indicated that

mice are much more sensitive than rats to the carcinogenic effects of 1,3-butadiene and benzene (Henderson, 1996a,b). Studies using crabeating macaques reported a low uptake of 1,3butadiene after inhalation exposure. Concentrations of butadiene metabolites in the blood were 5 to 50 times lower in monkeys than in mice and 4 to 14 times lower than in rats. Studies of benzene also reported species differences in metabolism after inhalation exposure. Mice metabolize a greater fraction of a given dose of benzene than do rats and nonhuman primates. Mice also exhibit higher urinary concentrations of hydroquinone and its conjugates. Both rats and mice metabolize a higher fraction of benzene to ring-breakage metabolites than do nonhuman primates, as indicated by the levels of muconic acid in urine. Ring-breakage metabolites and hydroquinone have both been implicated in benzene carcinogenesis.

SUMMARY This unit describes several important uses of nonhuman primate models in toxicological research. The examples provided are by no means exhaustive. Nonhuman primates continue to be used in studies of drug metabolism and of the toxicokinetics of environmental pollutants. Monkeys are also likely to be used more as new biotechnology products are discovered. In all of these areas of research, monkeys represent a unique resource, given the close evolutionary history they share with humans. The decision to use nonhuman primate models should always be made after careful consideration of all other alternatives. When nonhuman primate models are deemed necessary, researchers bear a special responsibility to ensure that procedures to minimize pain and discomfort are used and that proper environmental enrichment programs are in place (Bloomsmith et al., 1991).

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Fagan, J.F. 1990. The paired-comparison paradigm and infant intelligence. Ann. N.Y. Acad. Sci. 608:337-364. Flifeil, S.E.G., Beals, T.F., Tashkin, D.P., Paule, M.G., Scallet, A.C., Ali, S.F., Bailey, J.R., and Slikker, W. 1991. Marijuana exposure and pulmonary alterations in primates. Pharmacol. Biochem. Behav. 40:637-642. Foster, W.G. 1992. Reproductive toxicity of chronic lead exposure in the female cynomolgus monkey. Reprod. Toxicol. 6:123-131. Grossman, A., Astley, S.J., Liggitt, H.D., Clarren, S.K., Shiota, F., Kennedy, B., Thouless, M.E., and Maggio-Price, L. 1993. Immune function in offspring of nonhuman primates (Macaca nemestrina) exposed weekly to 1.8 g/kg ethanol during pregnancy: Preliminary observations. Alcohol Clin. Exp. Res. 17:822-827. Nonhuman Primates as Animal Models for Toxicology Research

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Haley, P.J., Pavia, K.F., Swafford, D.S., Davila, D.R., Hoover, M.D., and Finch, G.L. 1994. The comparative pulmonary toxicity of beryllium metal and beryllium oxide in cynomolgus mon-

Levin, E., Schneider, M., Ferguson, S., Schantz, S., and Bowman, R. 1988. Behavioral effects of developmental lead exposure in rhesus monkeys. Dev. Psychobiol. 21:371-382. Luster, M.I., Portier, C., Pait, D.G., Rosenthal, G.J., Germolec, D.R., Corsini, E., Blaylock, B.L., Pollock, P., Kouchi, Y., Craig, W., White, K.L., Munson, A.E., and Comment, C.E. 1993. Risk assessment in immunotoxicology II. Relationships between immune and host resistance tests. Fund. Appl. Toxicol. 21:71-82. Mazue, G. and Richez, P. 1982. Problems in utilizing monkeys in toxicology. In Animals in Toxicological Research (I. Bartoseck, A. Guaitani, and E. Pacei, eds.) pp. 147-163. Raven Press, New York. Morris, P., Binienda, Z., Gillam, M.P., Harkey, M.R., Zhou, C., Henderson, G.L., and Paule, M.G. 1996a. The effect of chronic cocaine exposure during pregnancy on maternal and infant outcomes in the rhesus monkey. Neurotoxicol. Teratol. 18:147-154.


Morris, P., Gillam, M.P., Allen, R.R., and Paule, M.G. 1996b. The effect of chronic cocaine exposure during pregnancy on the acquisition of operant behaviors by rhesus monkey offspring. Neurotoxicol. Teratol. 18:155-166. Nikula, K.J., Avila, K.J., Griffith, W.C., and Mauderly, J.L. 1997. Sites of particle retention and lung tissue responses to chronically inhaled diesel exhaust and coal dust in rats and cynomolg u s m o n keys. Environ. Health Perspect. 105(Suppl. 5):1231-1234. Ozwara, H., Niphuis, H., Buijs, L., Jonker, M., Heeney, J.L., Bambra, C.S., Thomas, A.W., and Langermans, J.A.M. 1997. Flow cytometric analysis on reactivity of human T lymphocyte– specific and cytokine-receptor-specific antibodies with peripheral blood mononuclear cells of chimpanzee (Pan troglodytes), rhesus macaque (Macaca mulatta), and squirrel monkey (Saimiri sciureus). J. Med. Primatol. 26:164-171. Rice, D. and Gilbert, S. 1982. Early chronic lowlevel methylmercury poisoning in monkeys impairs spatial vision. Science 216:759-761. Rice, D. and Gilbert, S. 1992. Exposure to methylmercury from birth to adulthood impairs highfrequency hearing in monkeys. Toxicol. Appl. Pharmacol. 115:6-10. Rice, D. and Gilbert, S. 1995. Effects of developmental methylmercury exposure or lifetime lead exposure on vibration sensitivity function in monkeys. Toxicol. Appl. Pharmacol. 134:161169. Rice, D.C. 1996. Behavioral effects of lead: Commonalitie s betw ee n experimental an d epidemiologic data. Environ. Health Perspect. 104(Suppl. 2):337-351. Sakai, C.N. and Hodgen, G.D. 1988. Use of primate folliculagenesis models in understanding human reproductive biology and applicability to toxicology. Reprod. Toxicol. 1:207-221.

Schantz, S., Ferguson, S., and Bowman, R. 1992. Effects of 2,3,7,8-tetrachlorodibenzo-p-dioxin on behavior of monkeys in peer groups. Neurotoxicol. Teratol. 14:433-446. Schardein, J.L., Schwartz, B.A., and Kenel, M.F. 1985. Species sensitivities and prediction of teratogenic potential. Environ. Health Perspect. 61:55-67. Snipes, M.B. 1989. Long-term retention and clearance of particles inhaled by mammalian species. CRC Crit. Rev. Toxicol. 20:175-211. Snipes, M.B. 1996. Current information on lung overload in nonrodent mammals: Contrast with rats. Inhal. Toxicol. 8(Suppl.):91-109. Snyderwine, E.G., Turesky, R.J., Turteltaub, K.W., Davis, C.D., Sadrieh, N., Schut, H.A.J., Nagao, M., Sugimura, T., Thorgeirsson, U.P., Adamson, R.H., and Thorgeirsson, S.S. 1997. Metabolism of food-derived heterocyclic amines in nonhuman primates. Mutat. Res. 376:203-210. Thorgeirsson, U.P., Dalgard, D.W., Reeves, J., and Adamson, R.H. 1994. Tumor incidence in a chemical carcinogenesis study of nonhuman primates. Reg. Toxicol. Pharmacol. 19:130-151. Tryphomas, H. 1995. The use of nonhuman primates in the study of PCB immunomodulation. Hum. Exp. Toxicol. 14:107-110. Tryphomas, H., Lacroix, F., Hayward, S., Izaguirre, C., Parenteau, M., and Fournier, J. 1996. Cell surface marker evaluation of infant Macaca monkey leukocytes in peripheral whole blood using simultaneous dual-color immunophenotypic analysis. J. Med. Primatol. 25:89-105.

Contributed by Thomas M. Burbacher and Kimberly S. Grant University of Washington Seattle, Washington

The authors would like to thank Noelle Liberato for her dedicated assistance. The preparation of this manuscript was supported by NIH grants ES06673 and ES03745.



Statistical Approaches to the Design of Toxicology Studies Statistical methods provide an essential tool set for use across the field of toxicology. These methods may serve to perform any combination of three possible tasks. The most familiar is hypothesis testing—i.e., determining if two (or more) groups of data differ from each other at a predetermined level of confidence. The second function involves the construction and use of models, which is most commonly linear regression or the derivation of some form of correlation coefficient. Model fitting allows researchers to relate one variable (typically a treatment, or independent, variable) with other variables (usually one or more effects of dependent variables). The third function, reduction of dimensionality, is less commonly used than the first two and includes methods for reducing the number of variables in a system while only minimally reducing the amount of information, therefore making a problem easier to visualize and understand. Examples of such techniques are factor analysis and cluster analysis. A subset of the third function, discussed under Descriptive Statistics, is the reduction of raw data to single expressions of central tendency and variability (such as the mean and standard deviation). There is also a special subset (data transformation, which includes such things as the conversion of numbers to log

UNIT 1.2

or probit values) that is part of both the second and third functions of statistics. Figures 1.2.1, 1.2.2, 1.2.3, and 1.2.4 present a series of decision trees for selecting individual statistical techniques within the framework of the classification of methodologies. This unit presents an overview of statistics. Gad (1998) presents a much more extensive discussion. Salsburg (1986) and Krewski and Franklin (1991) have also published works devoted to the field of statistical analysis in toxicology; and although these are more narrow in scope, they provide useful insights.

DESCRIPTIVE STATISTICS Descriptive statistics is a fundamental starting place, used to convey the general nature of any set of collected data. The statistics describing any single group of data have two components. One of these describes the location of the data, and the other gives a measure of the dispersion of the data in and about this location. A fact that is often overlooked is that the choice of what parameters are used to convey these pieces of information implies a particular nature for the distribution of the data. Most commonly, for example, location is described by giving the (arithmetic) mean, and dispersion is described by giving the standard

What is objective of analysis?

To be able to predict effects/actions of agents?

To sort out which variables are important?

YES Modeling function; go to Figure 1.2.3

YES Reduction of dimensionality function; go to Figure 1.2.4

To determine if there are differences between groups of data?

YES Hypothesis-testing function; go to Figure 1.2.2

Figure 1.2.1 Overall decision tree for selecting statistical procedures. See Gad (1998) for an explanation of the statistical methods. Contributed by Shayne C. Gad Current Protocols in Toxicology (1999) 1.2.1-1.2.18 Copyright © 1999 by John Wiley & Sons, Inc.


1.2.1

1.2.2


F-test

Data not significant; no more tests necessary

Dunnett's

Cochran t -test

N1 = N2 N1

Student's t -test df = N1 – 1

Student's t -test df = (N2 + N2) – 2

Categorical (quantal) data

Distribution free multiple comparisons

Significant (+)

Fisher’s exact test

Data not significant; no more tests necessary

Not significant (–)

Krustal-Wallis nonparametric ANOVA

Significant (+)

Wilcoxon rank sum test (2 groups)

Not significant (–)

R ×C χ2

Frequency data, Comparison of 2 Comparison of ≥3 such as mortalities, groups or if the groups all with pathology findings variance in some variation within group ≥1 group = 0 Three or more groups (no variation within group) NO YES

Nonparametric data, such as percentage values, ranks

No

Figure 1.2.2 Decision tree for selecting hypothesis-testing procedure. *, if plot does not clearly demonstrate lack of normality, exact test may be employed; +, if continuous data, Kolmongorox-Smirnov test may be used; −, if discontinuous data, χ2 goodness-of-fit test may be used. See Gad (1998) for an explanation of the statistical methods.

Duncan's multiple range test

If answer to If answer to either is NO both is YES

Intracomparison only of groups vs. controls? Group sizes approximately equal?

N2

Not normal (+)

Normal (–)

Scatter* gram

Yes

Significant (+) Not significant (–) Not significant (–) Significant (+) Homogeneous Heterogeneous

ANOVA

Homogeneous Heterogeneous

Not significant (–) Significant (+)

F-max test

Continuous data, such as body weights, blood cell counts Comparison ≥3 Comparison of 2 groups groups

Assumed parametric data

Visually examine data: do they appear normally distributed?

Are data continuous or discrete in nature?

≥3

Continuous

Discrete

How many variables? 2

What is desired?

Relationships between variables Nonlinear components

Linear

Discriminant analysis

Multiple regression

Relationships between variables

Modeling

Measure of relationships

Nonlinear regression

Kendall's rank correlations

Nonlinear What is desired? Modeling

Nonlinear regression

Linear



Kendall's rank correlations

What is desired?

Correlation coefficient

Data type Percentage

Continuous

Probit/logit plots or moving averages

Linear regression or trend analysis

Figure 1.2.3 Decision tree for selecting modeling procedures. See Gad (1998) for an explanation of the statistical methods.

Establish objective of desired data reduction

Data to be linearized

Cyclic patterns Multito be identified dimensional or non-numeric and measured data to be assigned or removed relative values

Transformations Multidimensional scaling or nonmetric scaling

Multidimensional and/or nonmetric data to be grouped

Fourier analysis

Analysis to be Quantitative corrected for relationships single of multilinear/time dimensional data variable to be established Life table analysis

Cluster analysis

Principle components or canonical correlations

Figure 1.2.4 Decision tree for selecting reduction of dimensionality procedures. See Gad (1998) for an explanation of the statistical methods.


deviation (SD) or the standard error of the mean (SEM). For continuous variables, the character of a sample (x) may be described using the measures of central tendency and dispersion with which researchers are generally most familiar: the mean, denoted by the symbol X and also called the arithmetic average, is calculated by adding up all the values in a group then dividing by N (or n), the number of data points in a group. The SD is denoted by the symbol s for samples and σ for populations; in the former case it is calculated as

s=

∑x

2

−

d∑ X i

2

N

N −1

where X is the individual datum. If the total number of data in the group is N, then the SEM is calculated as SEM =

SD N

The use of these parameters (the mean with either the SD or SEM) to describe a group implies, however, that there is reason to believe that the data being summarized are from a population that is at least approximately normally distributed. If this is not the case, then the researcher should instead use a set of terms that do not have such a rigid underpinning: commonly, the median for location and the semiquartile distance for dispersion. When all the numbers in a group are arranged in a ranked order (i.e., from smallest to largest), the median is the middle value. If there is an odd number of values in a group, then the middle value is obvious (e.g., in the case of 13 values, the seventh largest is the median). When the number of values in the sample is even, the median is calculated as the midpoint between the (N/2)th and the ([N/2] + 1)th number. For example, the series of numbers 7, 12, 13, 19, the median value would be the midpoint between 12 and 13, which is 12.5.

EXPERIMENTAL DESIGN

Statistical Approaches to the Design of Toxicology Studies

A priori selection of statistical methodology (as opposed to the post hoc, or after the fact, approach) is as significant a portion of the process of protocol development and experimental design as any other and can measurably enhance the value of the study. The prior selection of statistical methodology is essential for the effective detailing of other parts of the protocol, such as the number of animals per

group and the sampling intervals for body weight. To make such a selection, the toxicologist must have both an in-depth knowledge of the area of investigation and an understanding of the general principles of experimental design, because the analysis of any set of data is dictated to a large extent by the manner in which the data are obtained. The four statistical principles of experimental design are replication, randomization, concurrent (“local”) control, and balance. Replication. Any treatment must be applied to more than one experimental unit (animal, plate of cells, etc.) to allow an estimate of the variability of the unit. This provides more accuracy in the measurement of a response than can be obtained from a single observation, because underlying experimental errors tend to cancel each other out. It also supplies an estimate of the experimental error derived from the variability among all the measurements taken (replicates). Randomization. This is practiced to ensure that every treatment will have its fair share of extreme high and extreme low values and allows the toxicologist to proceed as if the assumption of “independence” were valid. That is, there is no avoidable/known systematic bias in how one obtains data. Concurrent control. Comparisons between treatments should be made, to the maximum extent possible, between experimental units from the same group. That is, animals used as a control group should come from the same source and lot as the test-group animals. And, except for the treatment being evaluated, test and control samples should be maintained and handled in exactly the same manner. Balance. If the effect of several different factors is being evaluated simultaneously, the experiment should be laid out in such a way that the contributions of the different factors can be separately distinguished and estimated. The goal of all these principles is good statistical efficiency and the economizing of resources. Montgomery (1997) provides a comprehensive review of the area of experimental design. The first precise or calculable aspect of experimental design encountered is determining sufficient test- and control-group sizes to allow one to have an adequate level of confidence in the results of a study (i.e., in the ability of the study design with the statistical tests used to detect a true difference, or effect, when it is present). The statistical test contributes a level of power to such a detection. The power of a


statistical test is the probability that a test results in rejection of the null hypothesis, H0, when some other hypothesis, H, is valid. This is termed the power of the test “with respect to the (alternative) hypothesis H.” If there is a set of possible alternative hypotheses, the power, regarded as a function of H, is termed the power function of the test. When the alternatives are indexed by a single parameter Π, simple graphical presentation is possible. If the parameter is a vector Π, one can visualize a power surface. If the power function is denoted by ϑ(Π) and H0 specifies Π = Π0, then the value of ϑ(Π)— the probability of rejecting H0 when it is in fact valid—is the significance level. A test’s power is greatest when the probability of a type II error, or the probability of having a false-negative result (whereas a type I error refers to a false-positive result), is the least. Specified powers can be calculated for tests in any specific or general situation. Some general rules to keep in mind are 1. The more stringent the significance level, the greater the necessary sample size. More subjects are needed for a 1% level test than for a 5% level test. 2. Two-tailed tests require larger sample sizes than do one-tailed tests. Assessing two directions at the same time requires a greater investment in resources. 3. The smaller the critical effect size, the larger the necessary sample size. Subtle effects require greater efforts (i.e., larger sample sizes). 4. Any difference can be significant if the sample size is large enough. 5. The larger the power required, the larger the necessary sample size. Greater protection from failure requires greater effort (more precision or larger sample size). The smaller the sample size, the smaller the power—i.e., the greater the chance of failure. 6. The requirements and means of calculating necessary sample size depends on the desired (or practical) comparative sizes of test and control groups. This number (N), or the necessary sample size, can be calculated by using the following formula: N=

at

1

+ t2 d

2

f

2

s2

where t1 is the one-tailed t-value with N − 1 degrees of freedom (df) corresponding to the desired confidence level; t2 is the one-tailed t-value with N − 1 df corresponding to the

probability that the sample size will be adequate to achieve the desired precision; s is the sample standard deviation, typically derived from historical data and calculated as follows (with υ being the variable of interest): s=

1

2 υ1 − υ 2 f a ∑ N −1

and d is the acceptable range of variation in the variable of interest. This calculation is illustrated in Example 1.

Example 1 In a subchronic dermal study in rabbits, the principal point of concern is the extent to which the compound causes oxidative damage to erythrocytes. To quantitate this, the laboratory will measure the numbers of reticulocytes in the blood. What then would be an adequate sample size to allow the question at hand to be addressed with reasonable certitude? To determine this, use the one-tailed t-value for an infinite number of degrees of freedom at the 95% confidence level (i.e., P ≤0.05). A set of t-tables shows this number to be 1.645. From prior experience, it is known that the usual values for reticulocytes in rabbit blood are from 0.5 to 1.9 ×106/ml. The acceptable range of variation, zero, is therefore equal to the span of this range, or 1.4 (d). Likewise, an examination of the control data from previous rabbit studies shows that the sample standard deviation is 0.825 (s). When all these numbers are inserted into the equation for sample size (presented above), the required sample size (N) is calculated to be 2

a1.645 + 1.645f a0.825f2 = a1.4f 10.824 2 0.825f = a 1.96 2

= 4.556 In other words, in this case, in which there is little natural variability, measuring the reticulocyte counts of groups of only five animals each should be sufficient. A good approximation can be generated by substituting t-values for an infinite number of degrees of freedom.

OUTLIERS AND ROUNDING OF NUMBERS Outliers are extreme (high or low) values that are widely divergent from the main body




of a group of data and from what is the common experience. They may arise from an instrument (such as a balance) being faulty, they may be caused by the seemingly natural urge of some animals to frustrate research, or they may be indicative of a “real” value. Outlying values can be detected by visual inspection of the data; by use of a scattergram (discussed later); or (if the data set is small enough, which is usually the case in toxicology) by a large increase in the parameter estimating the dispersion of data, such as the standard deviation. When it is possible to solidly tie one of the above error-producing processes (such as a balance being faulty) to an outlier, it can be safely deleted from consideration. But if it is not possible to positively attribute such a cause to an outlier (even if the researcher has strong suspicions), the problem is much more complicated, for then such a value may be one of several other things. It could be a result of a particular cause that is the grounds for the entire study—i.e., the very “effect” that is being sought—or it could be because of the collection of legitimate effects that constitute sample error. As will be discussed later (see Characteristics of Screens) and as is now becoming more widely appreciated, in animal studies outliers can be an indication of a biologically significant effect that is not yet statistically significant. Variance inflation can result from such outliers and can be used to detect them. Outliers, in fact, by increasing the variability within a sample, decrease the sensitivity of the statistical tests and may actually preclude a statistically significant result. Alternatively, the outlier may be the result of an unobserved technician error. In this case the data point should be rejected—excluded from consideration with the rest of the data. Identifying legitimate cases (based on technical—i.e., actual technique-based measurement—error) for rejection from statistical analysis is the most complicated aspect of outlier issues. When the number of digits in a number is to be reduced (because of limitations of space or to reflect the extent of significance of a number), it is necessary to round it off. Failure to have a rule for performing this operation can lead to both confusion and embarrassment for a facility (during such times as study audits). One common rule follows. A digit to be rounded is not changed if it is followed by a digit 5 or by a 5 followed

by other nonzero digits, it is increased to the next highest number. When the digit to be rounded is followed by 5 alone or 5 followed by zeros, it is unchanged if it is even but increased by one if it is odd. Examples of this rule for a case in which the numbers must be reduced to 3 digits follows: 1374 becomes 137 1376 becomes 138 13852 becomes 139. The rationale behind this procedure is that over a period of time the results should even out, because as many digits will be increased as are decreased.

METHODS The brevity of this unit enforces a focus on applications at the expense of methodology. In addition to Gad (1998), the reader is urged to consult Sheskin (1997) for detailed presentations and considerations of actual statistical methodologies.

Randomization Randomization is the act of assigning a number of items (e.g., plates of bacteria or test animals) to groups in such a manner that there is an equal chance for any one item to end up in any one group. This is a control against any possible unconscious bias in assignment of subjects to test groups. One variation on this is what is called a censored randomization, which ensures that the groups are equivalent in some aspect after the assignment process is complete. The most common example of a censored randomization is one in which it is ensured that the body weights of the test animals in each group are not significantly different from those in the other groups. This is done by examining weights by analysis of variance after group assignment, then rerandomizing if there is a significant difference at some nominal level, such as P ≤ 0.10. The process is repeated until there is no difference. A second valuation is stratified (or blocked) randomization, by which members of subclasses (such as litter mates) are assigned to different groups. There are several alternatives for actually performing the randomization process. The three most commonly employed are card assignment, use of a random number table, and use of a computerized algorithm. In the card-based method, individual identification numbers for items (e.g., plates or animals) are placed on separate index cards. These cards are then shuffled and placed one at a time in succession into piles corresponding to


the required number of test groups. The results are the random group assignment. The random number table method requires that one assign unique numbers to the test subjects and have access to a random number table. For example, one may simply set up a table with a column for each group to which subjects are to be assigned. Start from the head of any one column of numbers in the random number table (each time the table is used, a new starting point should be used). If there are fewer than 100 test subjects, use only the last two digits of each random number in the table. If there are more than 99 but less than 1000, use only the last three digits. To generate group assignments, read down a column of the table, one number at a time. As you come across digits that correspond to a subject number, assign that subject to a group (enter its identifying number in a column), proceeding from left to right and filling one row at a time. After an animal is assigned to a group, any duplication of its unique number is ignored. Use as many successive columns of random numbers as needed to complete the process. The third (and now most common) method is to use a random-number generator that is built into a calculator or computer program. Instructions for the use of such programs are provided by the manufacturer or developer. One is also occasionally required to evaluate whether a series of numbers (such as an assignment of animals to test groups) is random. This requires the use of a randomization test, of which there are a large variety. The χ2 test (described later), can be used to evaluate the goodness of fit to a random assignment. If the result is not critical, a simple sign test will work. For the sign test, first determine the middle value in the numbers being checked for randomness. Then go through a list of the numbers assigned to each group, scoring each as a “+” (greater than the middle number) or “−” (less than the middle number). The number of pluses and minuses in each group should be approximately equal.

APPLICATIONS Median Lethalities The data required to calculate the median lethality (LD50) or the median effective dose (ED50) include several dosage (or exposure) levels, the number of animals dosed, and the number that died. If the purpose is only to establish the median effective dose in a rangefinding test, then testing four or five animals

per dose level, using Thompson’s method of moving averages, is the most efficient methodology and will give a sufficiently accurate solution. With two dose levels, if the ratio between the high and low dose is ≤2, even total and no mortality at those two dose levels will yield an acceptable, accurate median lethal dose, although a partial mortality is desirable. If, however, the purpose is to estimate a number of toxicity levels (LD10, LD90) and to establish the slope of the dose/lethality curve more precisely, it is common to use at least 10 animals per dosage level and to employ the log/probit regression technique. Note that in the equation Yi = a + bxi, b is the slope of the regression line and that the method already allows one to calculate 95% confidence intervals about any point in this line. Tests of significance between two or more such sets of data (i.e., slopes of mortality curves) may readily be done by the t-type test (discussed earlier) at any one set of points, such as the LD50 values of two curves. The confidence interval at any one point will be different from the interval at other points and must be calculated separately. In addition, the nature of the probit transform is such that toward the extremes—LD10 and LD90, for example—the confidence intervals will “balloon,” i.e., become very wide. Because the slope of the fitted line in these assays has a large uncertainty in relation to the uncertainty of the LD50 itself (the midpoint of the distribution), much caution must be used with calculated LDx values other than LD50 values.

Body and Organ Weights Body weight and the weights of selected organs are among the sets of data normally collected in studies for which animals are repeatedly dosed with (or exposed to) a chemical. Body weight is frequently the most sensitive parameter for indicating an adverse effect. How to best analyze this and in what form to analyze the organ weight data (as absolute weights, weight changes, or percentages of body weight) have been the subjects of a number of articles (Weil and Gad, 1980). The author’s experience has been that the following procedures are appropriate if the sample sizes are sufficient (≥10). With smaller sample sizes, the normality of the data becomes increasingly uncertain and nonparametric methods, such as that of KruskalWallis, may be more appropriate (Zar, 1974). 1. Organ weights as percentages of total body weights are calculated. 2. Body weights are analyzed either as weights or as changes in body weight. Even if



Association of Changes in Biochemical Parameters with Actions at Particular Target Organs

Table 1.2.1

Parameter 1

Organ system Blood Heart Lung Kidney

Liver

Bone Intestine Pancreas

Albumin

—

—

—

Dec.

Dec.

—

—

—

ALP

—

—

—

—

Inc.

Inc.

Inc.

—

Total bilirubin

Inc.

—

—

—

Inc.

—

—

—

BUN

—

—

—

Inc.

Dec.

—

—

—

Calcium

—

—

—

Inc.

—

—

—

—

Cholinesterase —

—

—

Inc.

Dec.

—

—

—

CPK

—

Inc.

—

—

—

—

—

—

Creatinine

—

—

—

Inc.

—

—

—

—

Comments Produced by liver; very significant reductions require extensive liver damage Elevations usually associated with cholestasis; bone alkaline phosphatase tends to be higher in young animals Elevations usually associated with cholestasis owing to obstruction or hepatopathy Estimates blood filtering capacity of the kidneys; does not become significantly elevated until kidney function is reduced 60% to 75% Can be life-threatening and result in sudden death Found in plasma, brain, and RBC Most often elevated as a result of skeletal muscle damage but can also be produced by cardiac muscle damage; can be more sensitive than histopathology Also estimates blood-filtering capacity of kidney, as does BUN, but is more specific continued

the groups were randomized properly at the beginning of a study (no group significantly different in mean body weight from any other group, and all animals in all groups within two standard deviations of the overall mean body weight), there is an advantage to using the computationally slightly more cumbersome changes in body weight. 3. Bartlett’s test is performed on each set of data to ensure that the variances of the sets are homogeneous. 4. As appropriate, the sequence of analysis outlined in the decision trees (see Figs. 1.2.1, 1.2.2, 1.2.3, and 1.2.4) is followed.

Clinical Pathology


A number of clinical chemistry and hematology parameters are now assessed in the blood and urine collected from animals in chronic toxicity studies. In the past (and still, in some places), the accepted practice has been to evaluate these data using univariant-parametric methods (primarily t-tests and/or analysis of

variance, ANOVA); however, it can be shown that this is not the best approach on a number of grounds. First, such biochemical and blood cell parameters are rarely independent of each other, and interest is not often focused on just one parameter. Rather, there are batteries of parameters associated with toxic actions at particular target organs. For example, increases in creatine phosphokinase, α-hydroxybutyrate dehydrogenase, and lactate dehydrogenase occurring together strongly indicate myocardial damage. In such cases, the interest is not in a significant increase in just one of these, but in all three. Table 1.2.1 gives a short summary of the association of various clinical chemistry parameters with known target organ toxicities. Similarly, changes in serum electrolytes (sodium, potassium, and calcium) interact with each other; a decrease in one is frequently tied to an increase in one of the others. Loeb and Quimby (1989) provide an excellent source of detailed understanding of the meaning and de-


Table 1.2.1

Parameter 1

Association of Changes in Biochemical Parameters with Actions at Particular Target Organs, continued

Organ system Blood Heart Lung Kidney

Liver

Comments

Bone Intestine Pancreas

Glucose

—

—

—

—

—

—

—

Inc.

GGT

—

—

—

—

Inc.

—

—

—

HBDH LDH

— —

Inc. Inc.

— Inc.

— Inc.

Inc. Inc.

— —

— —

— —

Total protein

—

—

—

Dec.

Dec.

—

—

—

SGOT

—

Inc.

—

Inc.

Inc.

—

—

Inc.

SGPT

—

—

—

—

Inc.

—

—

—

SDH

—

—

—

—

Inc. or Dec.

—

—

—

Alterations other than those associated with stress are uncommon and reflect an effect on the pancreatic islets or anorexia Elevated in cholestasis; a microsomal enzyme, the levels of which often increase in response to microsomal enzyme induction — Increases usually the result of skeletal muscle, cardiac muscle, and liver damage; not very specific Absolute alterations are usually associated with decreased production (liver) or increased loss (kidney); can see increases in cases of muscle “wasting” (catabolism) Present in skeletal muscle and heart; most commonly associated with damage to these structures Elevations usually associated with hepatic damage or disease Liver enzyme; can be quite sensitive but is fairly unstable; samples should be processed as soon as possible

Abbreviations: Inc., increase of chemistry values; Dec., decrease in chemistry values; ALP, alkaline phosphatase; BUN, blood urea nitrogen; RBC, red blood cells; CPK, creatinine phosphokinase; GGT, gamma glutamyl transferase; HBDH, hydroxybutyric dehydrogenase; LDH, lactic dehydrogenase; SGOT, serum glutamic oxaloacetic transaminase (also called aspartate aminotransferase; AST); SGPT, serum glutamic pyruvic transaminase (also called alanine amino transferase; ALT); SDH, sorbitol dehydrogenase.

tails of laboratory animal clinical chemistries. Furthermore, for some parameters—owing to their biological background or the measurement method—the data are frequently either not normally distributed (particularly because of being markedly skewed) or not continuous. This can be seen in some of the reference data for experimental animals in Weil (1982). Both normality (a normal distribution) and continuous data are underlying assumptions in the parametric statistical techniques most commonly used and described in this unit. Finally, it should always be kept in mind that it is rare for a change in any single parameter to be biologically significant. Rather, because parameters are so interrelated, patterns of changes in parameters should be expected and analyzed.

Incidence of Histopathologic Lesion In recent years, there has been as increasing emphasis on histopathologic examination of many tissues collected from animals in subchronic and chronic toxicity studies. Although it is not true that only those lesions that occur at a statistically significant increased rate in treated/exposed animals are of concern (because there are cases in which a lesion may be of such a rare type that the occurrence of only one or a few such treated animals raises a flag), it is true that, in most cases, a statistical evaluation is the only way to determine if what is seen in treated animals is significantly worse than what is seen in control animals. And although cancer is not the only concern, among the possible classes of lesion it is the one that is of greatest interest.




Typically, comparison of incidences of any one type of lesion between controls and treated animals are made using χ2 or Fisher’s exact test with a modification of the denominators. Too often, experimenters exclude from consideration all those animals (in both groups) that died before the first animals were found with a tumor at that site. Two major controversial questions are involved in such comparisons: should they be based on one-tailed or two-tailed distribution, and what are the effects and implications of multiple comparisons? The one- versus two-tailed distribution controversy revolves around the question of which hypothesis is being properly tested in a study such as a chronic carcinogenicity study. Is the tumor incidence different between the control and treated groups? In such cases, the hypothesis is bidirectional and therefore a two-tailed distribution is being tested against. Or is the question whether the tumor incidence is greater in the treated group than in the control group? In the latter case, the hypothesis is unidirectional, and only the right-hand tail of the distribution is under consideration. The implications of the answer to this question are more than theoretical; significance is much greater (exactly double, in fact) in the one-tailed case than in the two-tailed. For example, a set of data analyzed by Fisher’s exact test, which would have a two-tailed P level of 0.098 and a onetailed level of 0.049, would be flagged as significantly different. The multiple comparisons problem is a much more lively one. In chronic studies, the lesion/tumor incidence on each of a number of tissues, for each sex and species, is tested, and each result is flagged if it exceeds the fiducial limit of P ≥ 0.05. The point to ponder here is the meaning of “P ≥ 0.05.” This is the level of the probability of making a type I error (incorrectly concluding that there is an effect when, in fact, there is not). It is necessary to accept the fact that there is a 5% chance of producing a false positive from this study. The tradeoff is a much lower chance (typically 1%) of a type II error (i.e., of passing as safe a compound that is not safe). These two error levels are connected; to achieve a lower type II level inflates the type I level. The problem in this case is that making a large number of such comparisons involves repeatedly taking the chance of “finding” a false-positive result. The set of lesions and/or tumor comparisons described above may number >70 tests for significance in a single study, which

will result in a large inflation of false-positive findings. The extent of this inflated false-positive rate (and what can best be done to reduce its effects) has been discussed and estimated with a great degree of variability. Salsburg (1977) estimated that the typical National Cancer Institute (NCI)–type cancer bioassay has a probability of type I error of between 20% and 50%. Fears et al. (1977), however, estimated it as being between 6% and 24%. Without some form of correction factor, the false-positive rate of a series of multiple tests can be calculated as being equal to 1 − 0.95N, where N is the number of tests and the selected alpha level (type I error rate) is 0.05. What, then, is a proper use of such results? Or, conversely, how can the researcher control for such an inflated error rate? There are statistical methods available for dealing with this multiple comparisons problem. One is the use of Bonferroni inequalities to correct for successive multiple comparisons (Wilks, 1962). These methods have the drawback that there is some accompanying loss of power, expressed as an inability to identify true positives properly. A second approach is to use the information in a more mature decision-making process. First, the historical control incidence rates (such as are given for the B6C3FI mouse and Fischer-344 rats in Fears et al., 1977) should be considered; some background incidences are so high that these tissues are null and void for making decisions. Second, one should not look for just a single significant incidence in a tissue but rather for a trend. For example, the following percentages of a liver tumor incidence were found for the female rats of a study: (a) control, 3%; (b) 10 mg/kg, 6%; (c) 50 mg/kg, 17%; and (d) 250 mg/kg, 54%. In this study only the incidence at the 250-mg/kg level might be statistically significant; however, the trend through the levels suggests a dose response. Looking for such a trend is an essential step in a scientific assessment of the results; and one of the available trend analysis techniques should be used.

Reproductive Toxicology Reproductive implications of the toxic effects of chemicals have become increasingly important. Along with other types of studies that are closely related (such as teratogenesis and dominant lethal mutagenesis), reproduction studies are now common companions to chronic toxicity studies.


One point that must be keep in mind with all the reproduction-related studies is the nature of the appropriate sampling unit. Put another way: What is the appropriate N in such a study—the number of individual pups or the number of litters (or pregnant females)? Fortunately, it is now fairly well accepted that the first case (using the number of offspring as the N) is inappropriate (Weil, 1970). The real effects in such studies are actually occurring in the female that receives the dosage or exposure to the chemical or that is mated to a male that received a dosage or exposure. What happens to her, and to the development of the litter she is carrying, is biologically independent of what happens to every other female/litter in the study. This cannot be said for each offspring in each litter; the death of, or other change in, one member of a litter can and will be related to what happens to every other member in numerous fashions. Or the effect on all the offspring might be similar for all those from one female and different or lacking in another. As defined by Oser and Oser (1956), there are four primary variables of interest in a reproduction study. First, there is the fertility index, which may be defined as the percentage of attempted matings (i.e., each female housed with a male) that resulted in pregnancy; pregnancy is determined by a method such as the presence of implantation sites in the female. Second, there is the gestation index, which is defined as the percentage of mated females, as evidenced by a dropped vaginal plug or a positive vaginal smear, that deliver viable litters (i.e., litters with at least one live pup). Two related variables that may also be studied are the mean number of pups born per litter and the percentage of total pups per litter that are stillborn. Third, there is the viability index, which is defined as the percentage of offspring born that survive at least 4 days after birth. Finally, there is the lactation index, which is the percentage of those animals per litter alive at 4 days that survive to weaning. In rats and mice, this is classically taken to be 21 days after birth. An additional variable that may reasonably be included in such a study is the mean weight gain per pup per litter. Given that N is at least 10 (proper sample size is discussed in Developmental Toxicology), it is possible to test each of these variables for significance using a method such as the Wilcoxon-Mann-Whitney U test, or the Kruskal-Wallis nonparametric ANOVA. If N 6-in. from the front of the vertical laminar flow hood, as the sterile zone begins there. Prepare fish-derived cell cultures 1. Turn on the laminar flow hood, and wipe all surfaces with 70% ethanol solution. 2. Examine the confluent culture of RTgill-W1 cell flask under an inverted phase-contrast microscope. Note the general appearance of the culture, checking for inadvertent microbial contamination or unexpected rounding and detachment of the fish-derived cells. Cultures to be passaged should appear normal and confluent (i.e., covering the bottom of the flask completely) or close to confluence. The Use of Fish-Derived Cell Lines

3. Under the laminar flow hood, aspirate the old medium and add 1.5 ml of 0.53 mM versene to the flask. Swirl it around gently, leave for 1 min, and aspirate it off.

1.5.2 Supplement 15


4. Add 1 ml versene and 1 ml trypsin solution to the flask, replace the cap, and observe under the inverted phase-contrast microscope. Do not leave the cells in trypsin for >5 min as cellular digestion and cell death may ensue. The cells will begin to detach from the culture surface in ∼1 to 3 min. Older cultures that have not been passaged for a long period of time are difficult to detach. Ideally, the cells will detach individually, and form a single-cell suspension.

5. Add 3 ml Leibovitz’s complete medium containing FBS to the flask. Pipet the medium up and down, directing the stream towards the bottom of the flask, to make sure that all cells are dislodged and resuspended in the medium. Trypsin inhibitors in FBS stop the action of the trypsin.

6. Transfer the cell suspension to a sterile 15-ml centrifuge tube and centrifuge 5 min at 200 × g, room temperature, in a tabletop centrifuge. 7. Aspirate the supernatant from the 15-ml centrifuge tube, being careful not to aspirate the cell pellet. Leave a small amount of supernatant (0.25 ml) over the cell pellet. Flick the centrifuge tube to resuspend the cells in the small volume of medium. Plate cells to maintain cell line 8a. Add 10 ml fresh medium to the centrifuge tube and transfer 5 ml to each of two 75 cm2 tissue culture flasks. Add 5 ml medium to each flask. 9a. Examine flasks using a phase-contrast microscope. Note whether cells have detached as single cells or as clumps and whether the suspension has been distributed equally between the two flasks. 10a. Allow the cells to grow at 18° to 22°C. When the cultures are confluent (7 to 10 days), split 1:3 or harvest to use for an experiment. Plate cells in microwell plates 8b. Add 4 ml Leibovitz’s complete medium containing FBS. Determine the cell density using a hemacytometer (APPENDIX 3B). Using fresh medium, adjust cell density to 1 × 105 cells/ml if using 48-well plates and to 1.5 × 105 cells/ml if using 96-well plates. 9b. If using a 48-well tissue-culture microwell plate, add 5 × 104 RTgill-W1 cells in 500 µl of Leibovitz’s complete medium to 40 wells of a 48-well tissue-culture microwell plate. Use the remaining eight wells as blanks by adding Leibovitz’s complete medium alone. If using a 96-well tissue-culture microwell plate, add 3 × 104 RTgill-W1 cells in 200 µl of Leibovitz’s complete medium to 84 wells of a 96-well tissue culture plate. Use the remaining twelve wells as blanks by adding Leibovitz’s complete medium alone. 10b. Allow the cells to grow for 3 to 4 days in the dark at 18° to 22°C to form a confluent cell monolayer for the 48-well plates and for 2 to 3 days for the 96-well plates. Growing cells over a 2- to 4-day period allows for a more consistent cell density and better adherence of cells than newly initiated, dense cultures.

Expose fish-derived cell lines to putative toxicants 11. Turn on the laminar flow hood, and wipe all surfaces with 70% ethanol solution. 12. Examine the plated microwell plate under the inverted phase-contrast microscope. Note the general appearance of the cell culture, making sure that the bottom of the wells contain a confluent monolayer and inadvertent microbial contamination is absent. Toxicological Models


Supplement 15

13. Make serial dilutions (working solutions) from test compound stock solutions in carrier solution (e.g., DMSO). If the carrier solution is an organic solvent, prepare working solutions such that the concentrations are at least 200 times the final concentrations desired in the culture wells. Serial dilutions are necessary to ensure that the solvent is diluted sufficiently in the culture medium in the wells in order minimize interference due to the solvent with cell viability and/or toxicant uptake. For more water-soluble compounds, a dilution series can be prepared directly in culture medium. However, many organic toxicants, including PAHs, will require the use of an organic carrier solvent.

14. Remove growth medium from plates by inverting over a catch basin. Drain plates further for a few seconds on a small stack of paper towels. 15. Rinse each culture well with 500 µl L-15/ex solution if using 48-well plates or 200 µl L-15/ex solution if using 96-well plates. Remove this rinsing medium by inverting over a catch basin. Drain plates further for a few seconds on a small stack of paper towels. Then add 500 µl L-15/ex solution to each well if using 48-well plates or 200 µl L-15/ex solution if using 96-well plates. 16. Expose cells to the test compound in a vertical laminar flow hood. For example, if working solutions were prepared in DMSO at 200 times the final concentration required, pipet 2.5 µl test compound to each well containing 500 µl/well L-15/ex solution in 48-well plates using a positive displacement digital micropipet. If using a 96-well plate, pipet 1.0 µl test compound dissolved in DMSO to each well containing 200 µl/well L-15/ex solution. Because cells in L-15/ex solution are particularly sensitive to DMSO, tilt the plates before adding the test compound to increase the volume of medium above the cells as a protective layer. Dispense the 2.5 ìl or 1 ìl test compound above the level of the medium, and then touch the droplet to the surface of the medium to aid in dispersion. In addition, reduce the light level in the flow hood to avoid irradiation of cells in the presence of the toxicant.

17. In a 48-well plate, dose six wells (five wells with confluent cell monolayers and one well without cells to serve as a blank) for each of seven concentrations of compounds and the DMSO control. In a 96-well plate, dose eight wells (seven wells with confluent cell monolayers and one without cells to serve as a blank) for each of twelve concentrations of compounds and the DMSO control. Wrap plates in Parafilm to prevent evaporation during the exposure period. The use of plate sealing foils, which are commercially available, e.g., Nunc or Greiner, is recommended in order to minimize well-to-well transfer and losses due to evaporation of more volatile toxicants.

For exposure of fish-derived cells to toxicants in the dark 18a. For exposures in the dark, incubate plates up to 48 hr at 18°to 22°C. 19a. If not specifically desired (see below), avoid exposure of the plates to light. Although L-15/ex is unlikely to be affected by irradiation, many toxicants may be altered chemically.

The Use of Fish-Derived Cell Lines

20a. At the end of the dark exposure, remove plates from the incubator and continue with the cytotoxicity assay for assessing metabolic activity with the fluorescent dye alamar blue, proceed to step 23 (for membrane integrity assessment, use CDFA-AM, see Alternate Protocol 1; or for lysosomal activity assessment, use neutral red, see Alternate Protocol 2).

1.5.4 Supplement 15


For exposure of fish-derived cells to toxicants in the presence of irradiation 18b. Turn on the lamps of the radiation exposure chamber at least 15 min prior to use in order to allow lamps to warm up and emit a stable radiation. The radiation chamber should contain at least two fixtures to hold lamps, e.g., one UV-A and one UV-B fluorescent lamp, and it should be shielded from any radiation outside the chamber. Ideally, lamps should be linked to a transformer in order to allow for easy manipulation of the radiation output.

19b. Place the appropriate unit of the spectroradiometer into the radiation exposure chamber in order to perform UV radiation measurements as described in the user manual of the spectroradiometer. Ensure that the appropriate unit of the spectroradiometer is positioned the same distance from the UV lamps as the culture plate would be. Also, place the lid of the culture plate on top of the unit to measure the amount of UV irradiation passing through the lid of the plate.

20b. Using the transformer, adjust UV intensities as desired. For example, a 10:1 ratio of UV-A to UV-B may be desirable in order to mimic the ratio of these two UV components in nature. If no transformer is available, radiation intensity may be adjusted by varying the distance between the exposure tray and the lamps but this method is somewhat cumbersome.

21b. Once the radiation exposure chamber has been set up, place tissue culture plates into the chamber in an atmosphere of air for the required exposure period. Place a second tissue culture plate into the chamber at the same time but protected from radiation. This second plate serves as the dark control.

22b. At the end of the UV exposure, remove plates from the chamber and proceed with the cytotoxicity assay for assessing metabolic activity using alamar blue, proceed to step 23 (for membrane integrity using CDFA-AM, see Alternate Protocol 1; or for lysosomal activity using neutral red, see Alternate Protocol 2). Assess metabolic activity using alamar blue 23. Prepare a 5% (v/v) working solution of alamar blue in L-15/ex solution. Alamar blue is purchased as a ready-to-use solution in quantities of 25 ml and 100 ml. When stored in the dark at 2° to 8°C and kept aseptically, alamar blue can be used for at least 1 year.

24. Remove exposure medium from plates by inverting over a catch basin. Drain plates further for a few seconds on a small stack of paper towels. 25. Add 100 to 150 µl of 5% alamar blue working solution to each well of a 48-well plate, or 50 to 100 µl per well to a 96-well plate. The general rule is to at least cover the growth surface of the wells although some plate readers may require slightly larger volumes for accurate readings.

26. Incubate the plates in the dark at 18° to 22°C for 30 min. Although longer incubation times are also possible, the yield of fluorescent units from alamar blue can decline if the incubation period is too long (O’Brien et al., 2000).

27. Measure alamar blue on the fluorometric plate reader at excitation and emission wavelengths of 530 and 590 nm, respectively. Toxicological Models


Supplement 15

ALTERNATE PROTOCOL 1

EVALUATION OF TOXICITY IN FISH-DERIVED CELL LINES USING CFDA-AM TO ASSESS MEMBRANE INTEGRITY Although esterase substrates have been used as a measure of cell membrane integrity since the 1960s (Rotman and Papermaster, 1966), 5-carboxyfluorescein diacetate acetoxymethyl ester (CFDA-AM) is an example of one developed to improve this application (Haugland, 1996). CFDA-AM diffuses into cells rapidly and is converted by non-specific esterases of living cells from a nonpolar, nonfluorescent dye into a polar, fluorescent dye, 5-carboxyfluorescein (CF), which diffuses out of cells slowly. In this protocol, fish cells in microwell cultures are exposed for a period of time to a putative toxicant, and after removal of the putative toxicant, CFDA-AM is added and the capacity of the cells to produce CF is measured. Additional Materials (also see Basic Protocol) 4 mM 5-carboxyfluorescein diacetate acetoxymethyl ester (CFDA-AM, see recipe) RTgill-W1 cells in a 48-well or 96-well plate exposed to toxicants (see Basic Protocol, steps 1 to 22) 1. Prepare a working solution of 4 µM CFDA-AM by diluting the 4 mM CFDA-AM stock solution 1:1000 in L-15/ex solution. 2. Remove exposure medium from RTgill-W1 cells in a 48-well or 96-well plate exposed to toxicants by inverting over a catch basin. Drain plates further for a few seconds on a small stack of paper towels. 3. Add 100 to 150 µl of 4 µM CFDA-AM working solution to each well of a 48-well plate or 50 to 100 µl of 4 µM CFDA-AM working solution to each well of a 96-well plate and incubate the plate in the dark for 30 to 120 min at 18°to 22°C. 4. Measure CF fluorescence on the fluorometric plate reader at respective excitation and emission wavelengths of 485 and 530 nm. Alamar blue (see Basic Protocol, steps 23 to 27) and CFDA-AM can be added together to perform the two assays in a single step (Schirmer et al., 1997) because the fluorescent products of the two indicator dyes can be detected at different emission wavelengths without interfering with each other. The advantage of doing so is the conservation of material and time as fluorescent readings are taken on the same culture wells. Thus, in order to perform the two assays together, add the appropriate amount of alamar blue to make a 5% (v/v) working solution in L-15/ex solution and then dilute the CFDA-AM stock solution (4 mM in DMSO) 1:1000 in that same volume of L-15/ex solution. Proceed with the incubation period as described above in step 3. Inasmuch as alamar blue and CFDA-AM do not affect the viability of cells, fluorescent dyes can be removed after fluorescent measurement and replaced by culture medium to allow the cells to recover for a period of time after which the indicator dyes can be re-applied.



EVALUATION OF TOXICITY IN FISH-DERIVED CELL LINES USING NEUTRAL RED TO ASSESS LYSOSOMAL ACTIVITY Although first used to evaluate cell viability in virology, neutral red (NR) has been utilized most intensively in in vitro toxicology. The principle is that viable cells accumulate NR (3-amino-7-dimethylamino-2-methylphenazine hydrochloride) in lysosomes (Borenfreund and Puerner, 1984). NR can be applied before or after the exposure of cell cultures to toxicants, so that the measured endpoint represents either the release or uptake of the dye (Borenfreund and Puerner, 1984; Reader et al., 1990). Measurements can be done either spectrophotometrically (Borenfreund and Puerner, 1984) or fluorometrically (Essig-Marcello and van Buskirk, 1990). In this protocol, the use of NR after the exposure of fish cells to toxicants and the fluorometric measurement of any subsequent changes in NR uptake are described.

1.5.6 Supplement 15


Additional Materials (also see Basic Protocol) Neutral red stock solution (see recipe) RTgill-W1 cells in a 48-well or 96-well plate exposed to toxicants (see Basic Protocol, steps 1 to 22) Neutral red fixative solution (see recipe) Neutral red extraction solution (see recipe) Orbital shaker 1. Prepare a working solution of 33 µg/ml neutral red by diluting the neutral red stock solution 1:100 in L-15/ex solution. 2. Remove exposure medium from RTgill-W1 cells in a 48-well or 96-well plate exposed to toxicants by inverting over a catch basin. Drain plates further for a few seconds on a small stack of paper towels. 3. Add 100 to 150 µl of 33 µg/ml neutral red working solution to each well of a 48-well plate or 50 to 100 µl of 33 µg/ml working solution to each well of a 96-well plate and incubate the plate in the dark for 60 min at 18°to 22°C. 4. Remove the neutral red working solution by inverting over a catch basin and drain a few seconds on a small stack of paper towels. It is critical to remove all the neutral red working solution from each well, especially in a 96-well plate.

5. Rinse wells once with 100 µl/well of neutral red fixative solution. The rinsing step removes any excess neutral red that has not been localized in lysosomes.

6. Add 100 µl/well of neutral red extraction solution to solubilize the lysosomal neutral red. Place plates on an orbital shaker and shake at ∼40 rpm for 10 min. 7. Measure neutral red fluorescence on the fluorometric plate reader at excitation and emission wavelengths of 530 and 645 nm, respectively. In the interest of conserving material and for performing cell viability assays on the same set of cells, the neutral red assay can be performed on the same plate previously used to assess alamar blue (see Basic Protocol) and/or CFDA-AM fluorescence (see Alternate Protocol 1). After termination of the alamar blue and CFDA-AM exposures, remove the dye solution(s) and start by adding neutral red as described above. However, inasmuch as the neutral red assay will terminate the cell culture because cells will be fixed, a separate plate needs to be used if the alamar blue/CFDA-AM plates are to be used to study recovery.

REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2E; for suppliers, see SUPPLIERS APPENDIX.

CFDA-AM stock solution, 4 mM Dissolve the 5-mg vial of CFDA-AM (Molecular Probes) in 2.35 ml of sterile anhydrous DMSO (final concentration 4 mM) in a laminar flow hood. Dispense into sterile 1.5-ml microcentrifuge tubes in 50-µl aliquots to prevent degradation from thawing and refreezing. Wrap each microcentrifuge tube in aluminum foil to prevent light degradation. Store dessicated up to 1 year at −20°C to avoid ester hydrolysis due to moisture. Toxicological Models


Supplement 15

L-15/ex solution Development of this modified medium is outlined in Schirmer et al. (1997) and is based on the constituents of basal medium, L-15 (Leibovitz, 1963). Make up all components from cell-culture grade reagents (Sigma) and prepare in cell-culture grade, distilled water. Salt solution A: In 600 ml water, dissolve: 80 g NaCl, 4.0 g KCl, 2.0 g MgSO4, 2.0 g MgCl2 Salt solution B: In 100 ml water, dissolve: 1.4 g CaCl2 Salt solution C: In 300 ml water, dissolve: 1.9 g Na2HPO4, 0.6 g KH2PO4 Autoclave each solution separately and store up to 1 year at room temperature. Sodium pyruvate solution: In 100 ml of water, dissolve 5.5 g sodium pyruvate. Filter sterilize through a 0.2-µm filter, dispense in 5.7-ml aliquots, and store up to 1 year at −20°C. Galactose solution: In 100 ml of water, dissolve 9.0 g galactose. Filter sterilize through a 0.2-µm filter, dispense in 5.7-ml aliquots, and store up to 1 year at − 20°C. To prepare L-15/ex solution: To 500 ml of sterile cell culture-grade, distilled water, add aseptically: 34.0 ml salt solution A 5.7 ml salt solution B 17.0 ml salt solution C 5.7 ml sodium pyruvate solution 5.7 ml galactose solution Store L-15/ex solution up to 1 year at room temperature Leibovitz’s L-15 complete medium containing FBS To 500 ml of Leibovitz’s L-15 medium (Sigma), aseptically add 50 ml fetal bovine serum (FBS; Sigma). Final FBS concentration in culture medium is 8.9%, which is usually referred to as 10%. Aseptically add 10 ml of penicillin/streptomycin (100 IU/ml penicillin, 100 µg/ml streptomycin; Sigma). Store up to 1 year at 4°C. Neutral red extraction solution Prepare 1% (v/v) acetic acid and 50% (v/v) ethanol in deionized, distilled water. Store for up to 1 year at room temperature in the dark. Neutral red fixative solution Prepare 0.5% (v/v) formaldehyde and 1% (w/v) CaCl2 in deionized, distilled water. Store for up to 1 year at room temperature in the dark. Neutral red stock solution Dissolve 3.3 mg of neutral red powder (Sigma) per milliliter of Dulbecco’s PBS (D-PBS; Sigma or Life Technologies) in an amber vial. Store for up to 1 year at 4°C. Alternatively, neutral red stock solution can be purchased dissolved in D-PBS at a concentration of 3.3 mg/ml (Sigma).

Trypsin solution Dissolve 100 mg trypsin (Sigma) in 10 ml of Ca2+- and Mg2+-free Hank’s balanced salt solution (Sigma) to make a trypsin stock solution. Dispense 0.5 ml of this solution into 9.5 ml of Ca2+- and Mg2+-free Hank’s balanced salt solution. Store for up to 1 year at −20°C. The Use of Fish-Derived Cell Lines

1.5.8 Supplement 15


COMMENTARY Background Information The many in vitro toxicology tests can be divided into two types: general (basal) cytotoxicity tests and tests of differentiated cell function (Flint, 1990). The general type consists of tests that measure cytotoxic phenomena only, e.g., the inhibition of cell proliferation. One premise of this approach is that all toxic phenomena are fundamentally related to an impairment of some aspect of cellular activity in vivo. Therefore, toxicity in vivo should be expected if the test agent is bioavailable to a target tissue at concentrations that are observed to impair cell viability in vitro. A potential weakness of this approach is the observation that rather than being a general phenomenon of all tissues, toxicity in vivo is often limited to a small group of organs and cells within these organs. Thus, specific toxic effects might occur at concentrations well below those causing general cytotoxicity. The second class of in vitro tests attempts to overcome this weakness by monitoring a cellular function specific for the differentiated state of the cultured cell. From a risk assessment point of view, the tests for differentiated cell functions are usually more valuable (Flint, 1990). To date, general cytotoxicity tests mostly have been used in fish toxicology and are described in this unit. Many general cytotoxicity tests have been described. They measure impairment of cellular activities by potentially toxic treatments. However, since their first introduction, assays of cytotoxicity and/or cell viability have been criticized as to their meaning and have aroused debate as to which assay is most appropriate (Schrek, 1965; Shaw, 1994). Cytotoxicity assays that monitor reproductive capacity have been described as being the most comprehensive because they integrate the soundness of the entire cellular machinery (Shaw, 1994). However, proliferation assays are not ideal for all purposes. They usually reveal little about the specific cellular events that lead to impaired proliferation and can miss subtle, transitory effects. They give little insight into the potential short-term impact of an effect on cells to the integrity of an organ or tissue. Also, colony formation and proliferation rates are impractical endpoints with most fish cell lines because the cells grow slowly. As an alternative to proliferative endpoints, assays of cell viability and cell injury can be performed (Shaw, 1994), such as the ones described here.

Although numerous assays of cell viability have been developed, those that utilize fluorometric indicator dyes are perhaps best. First, more and more dyes are becoming commercially available to evaluate distinct cellular parameters. Second, the development of fluorometric multiwell plate readers has made the use of fluorometric dyes easy and rapid, such as for the assays in this unit. The microwells conserve material resources by reducing the number of cells needed and increasing the number of replicates. The plate readers have the potential for high interlaboratory reproducibility and can be coupled to computers in order to manage data quickly and easily.

Critical Parameters and Troubleshooting Temperature The choice of temperature for growth, exposure, and assay of fish cell cultures is flexible and can be dictated by practical considerations or by scientific objectives. This is because fish cells can be grown over a wide temperature range, e.g., from 5° to 25°C for salmonid cells (Bols et al., 1992). The medium used in this unit is based on Leibovitz’s L-15, which does not need a 5% CO2 atmosphere in order for the pH to be buffered (Leibovitz, 1963). As a result, the “incubator” for fish cell cultures in CO2-independent media, such as L-15, can be a desk drawer at room temperature, or a temperatureregulated chamber, such as a conventional incubator or a refrigerator. For all the protocols that are described in this unit with salmonid cells, temperatures from 18° to 22°C give consistent results, and fluctuations within this range have little or no effect on the outcome. Toxicant preparation The acts of dissolving compounds and of adding them to cell cultures can be the source of several problems. Unfortunately, these are difficulties that cannot really be solved, but are important to consider when interpreting results. The problems are most severe for hydrophobic environmental contaminants, such as the PAHs. Several alternative strategies for toxicant delivery have to be considered. One is whether or not to use carrier solutions to dissolve the compounds. The second is whether or not to completely remove the medium that was used to initiate the cell cultures from the microwells and to replace it with an equivalent volume of



Supplement 15

toxicant solution. For compounds that are going to be presented without the use of a carrier, there is usually no choice, the original medium has to be replaced with the toxicant solution. Except for extremely water-soluble compounds, the highest doses in dose-response curves cannot be achieved by adding small aliquots of a concentrated solution. By contrast, high concentrations of hydrophobic compounds can be achieved in carrier solutions, allowing them to be added to cultures in small aliquots. Potential problems with carriers, exposure media, and presentation strategies for toxicants are further discussed below. Carriers such as dimethylsulfoxide (DMSO) have nearly always been used to dissolve environmental contaminants for toxicity testing, but they potentially can influence the results. For example, fluoranthene that was dissolved in L-15/ex solution with DMSO was slightly more photocytotoxic to RTgill-W1 cells than fluoranthene dissolved in L-15/ex solution without DMSO (Schirmer et al., 1997). Differences can also be found between carriers. DDT in DMSO was more cytotoxic to tilapia brain cells than DDT in acetone (Parkinson and Agius, 1987). The induction of 7ethoxyresorufin o-deethylase (EROD) activity in RTL-W1 cells was better after exposure to 2,3,7,8-tetrachlorodibenzo-p-dioxin (TCDD) in DMSO than in isooctane (Clemons et al., 1994). Also, the carrier could influence the induction potency of some compounds (Yu et al., 1997).


Exposure medium The medium in which cells are incubated during exposure to environmental contaminants is another variable that can influence results. An extremely simple one is used in this unit, which contains salts, galactose, and pyruvate at specified concentrations in the basal medium, Leibovitz’s L-15 (Leibovitz, 1963), which is termed L-15 exposure or L-15/ex (Schirmer et al., 1997). L-15/ex has several advantages. For photocytotoxicity studies, the absence of vitamins and aromatic amino acids prevents the inadvertent generation of toxicants from these compounds during the UV treatment. L-15/ex is also advantageous for detecting toxicants that cause cytotoxicity through the generation of reactive oxygen species (ROS). This is because expression of their toxicity should be aided by the absence in L-15/ex of most antioxidants. The one exception might be the presence of pyruvate. For mammalian cells, pyruvate is part of an antioxidant defense

(O’Donnell-Tormey et al., 1987). A limitation of L-15/ex is that exposure times are restricted to several days, as ultimately nutrient deprivation will cause cell death. The short exposure times mean that toxicants acting by inducing particular cellular processes, such as xenobiotic metabolism, and causing accumulative damage might be missed. Yet, RTgill-W1 cells survive in L-15/ex for at least 100 hr (Schirmer et al., 1997). Although addition of fetal bovine serum (FBS) to the medium extends the life of cell cultures, the antioxidants of FBS prevents the detection of fluoranthene photocytotoxicity (Schirmer et al., 1997). Also, FBS can alter the bioavailability of toxicants (Schirmer et al., 1997; Hestermann et al., 2000). Dosing strategies One strategy for presenting toxicants to cells is to completely remove the medium that was used to initiate the cell cultures from the microwells and add an equivalent volume of solution with the toxicant under study. Both steps present difficulties. The medium removal step has two problems. First, removal must be done quickly without damaging cells, which can be done by aspirating off the medium. But, this sometimes causes cell death. Death usually appears within minutes of aspiration and occurs in large patches over the culture surface. Routinely examining cultures with an inverted phase-contrast microscope shortly after aspiration easily identifies this problem. Alternatively, the medium can be removed by inverting the plate over a catch basin and blotting with a paper towel. This is the recommended medium removal technique because the cells are not subjected to the same force as aspiration may involve. Second, complete removal of the initiating medium over the cell cultures has the potential to change the physiology of cell cultures. An example of this is the rapid induction of 7-ethoxyresorufin o-deethylase (EROD) activity upon the removal of medium from cultures of the rainbow trout liver cell line, RTLW1 (Segner et al., 2000). For short-term exposures (as described in this unit), this should not be a problem. Replacing the medium with toxicant solution is problematic for hydrophobic compounds. A concentrated solution of toxicant in a carrier such as DMSO must be serially diluted in an aqueous solution to prepare concentrations for dose-response curves. Hydrophobic compounds have a tendency to stick to the walls of containers and pipets used to prepare these solutions. Thus, the final toxicant concentrations that the cultures receive can be

1.5.10 Supplement 15


lower than the concentrations apparently added. The problem cannot be solved by simply measuring the concentrations through analytical chemistry methods, for this approach also requires the use of containers and pipets that can lead to an underestimation of final concentrations. The second presentation strategy is to use micropipets to add small volumes (≤10 µl) of the toxicant in carrier solvent to the medium over the cells in microwells. Dosing in this manner must be done very carefully with carriers, such as DMSO. The small volume of DMSO sometimes falls as a blob directly onto the cells, immediately killing them. Again, examining cultures with an inverted phase-contrast microscope easily identifies this problem. As mentioned, the difficulty can be avoided by keeping the pipet tip close to the medium surface and allowing the surface tension to disperse the DMSO rapidly and evenly through the culture. This second presentation strategy has at least two advantages. Changing the medium is avoided, which also avoids any possible changes in the cells caused by this act. Final culture concentrations are closer to the apparent toxicant concentration that is added. In extreme cases, a compound might appear to be toxic with this dosing method, but not by the method of preparing the toxicant in exposure medium and using this solution to replace the growth medium of cell cultures. If the results of the two different dosing strategies are profoundly different, they should be reported as such, as this will aid others in replicating the results in any future studies and will stress the subtlety of the cytotoxicant’s actions. UV irradiation For the concurrent exposure of cultures to environmental contaminants and UV light, the radiation exposures can be expressed in different ways, sometimes causing confusion. The most important distinction is between energy units, which refer to the wave-like character of radiation, and quantum units, which reflect the corpuscular character. The conversion from energy content to photon content is defined by Planck’s equation. Fluence rates are used to refer to the quantity of radiation per area per time. Thus, Wm−2 and Jm−2 sec−1 are expressions of energy fluence rates, whereas moles(photons)m−2sec−1 is an expression of photon fluence rates. In contrast to the fluence rates, the total amount of radiation received by the cells can be expressed as fluence, that is quantity per area. Thus, the energy fluence

(Jm−2= Wm−2sec) or the photon fluence (mol(photons)m−2) can be calculated by multiplying the fluence rate by the time of irradiation (in seconds). For example, a photon fluence rate of UV-B at 313 nm of 1.4 µmol(m−2)sec−1, which has been shown to be environmentally relevant (Oris and Giesy, 1987), is equivalent to an energy fluence rate at that wavelength of 53 µWcm−2(or 53 µJcm−2sec−1). If irradiation is performed for 2 hr, the energy fluence that the cells are exposed to is 0.4 Wcm−2sec (or 0.4 Jcm−2). Inasmuch as UV radiation exposure decreases with increasing distance from the UV source, it is important to measure UV radiation at the position at which culture plates will be placed. In order to account for absorption of UV radiation by culture plate lids, place a lid between the UV source and the radiation measurement device. The irradiation of cells in the presence of tissue culture plate lids is highly recommended for two reasons. First, the lids ensure sterility during the illumination process. Second, the plate lids absorb any radiation below a wavelength of 290 nm, a filtering process which, under natural conditions, is carried out by a stratospheric ozone (Schirmer et al., 1997). While measurements of UV irradiance should be done frequently to ensure that UV intensities of the fluorescent lamps are as required, initial measurements should confirm that the culture medium covering the cells does not detrimentally affect UV penetration. To study this, a lid-covered tissue culture plate with and without culture medium should be placed between the UV source and the spectroradiometer and the values compared. If measurements are the same, it can be concluded that the culture medium used has no discernible effect on UV penetration. In contrast, lower UV intensities measured in the presence of culture medium indicate that cells obtain less UV radiation than anticipated. For example, the authors found that UV intensities were not significantly affected upon passage through a 4.7 mm path of L-15/ex medium, which equals 500 µl L-15/ex medium in a 48-well tissue culture plate. In contrast, the same passage of UV in the presence of 10% FBS led to a reduction of UV readings of 27% for UV-B and 9% for UV-A (Schirmer et al., 1997). Another factor to consider is temperature. In the presence of a small fan and a distance of the tissue culture plates from the UV lamps of at least 15 cm, the authors did not find temperature to rise in the tissue culture medium within 2 hr of UV-irradiation at a photon fluence rate of 10 µmol m−2



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sec-1UV-A and 1 µmol m−2sec−1UV-B (Schirmer et al., 1998b). However, longer exposures or higher UV intensity may potentially lead to an increase in temperature. Toxicant removal Termination of toxicant exposures prior to the addition of the fluorescent indicator dyes to cell cultures must be done carefully and consistently. Inverting plates over a catch basin is recommend over aspiration. Removal must be done rapidly but without damaging cells. Viewing control cultures with an inverted phase-contrast microscope will identify any problems. Of course, at this stage, the loss of cells from toxicant-treated cultures can be expected. Fluorescent microwell plate readers A number of manufacturers (e.g., Molecular Devices, Applied Biosystems) make fluorescent plate readers, and likewise, microwell plates are produced by several companies (e.g., Falcon, Costar, Nunc). Although all fluorescent plate readers are likely suitable, the crucial issue is to make sure the microwell plate correctly matches the plate reader. Under most circumstances, only the appropriate settings need to be chosen, as the plate readers have been designed to accept most plates from various manufacturers. However, sometimes the dimensions for a particular microwell plate must be obtained from the manufacturer and entered into the plate reader. As mentioned earlier, the minimum volume of fluorescent indicator dyes needed for accurate measurement might vary for different microwell plate readers. Thus, initial experiments should determine the most suitable volume.

Anticipated Results


Data analysis: Calculation of EC50 The raw fluorescent units resulting from the cell viability assays are used to evaluate the toxicity of the chemical being tested. Cell viability is expressed as a percent of non-toxicant exposed cells (% of control). For each concentration of toxicant, there is one well that contains no cells (no cell control). Both the well with no cells and the wells with cells are exposed to the toxicant. Prior to calculating percent of control, subtract the fluorescent units (FU) for wells without cells from the experimental (ex) and control (con) values with cells. To calculate the cell viability (% of control) use the following formula:

% of control = (FUex cells −FUex no cells) ×100/(average [FUcon − FUcon no cells]) Data for each well of each concentration are expressed as a percent of control. Then, the average and standard deviation for each concentration are calculated. These values are used to calculate the EC50 for the toxicant. Dose-response data typically follow a sigmoidal relationship and can be analyzed by nonlinear regression in most graphing software such as SigmaPlot (Jandel Scientific). The data is fitted to the four-parameter logistic function for continuous response data. The logistic function is: y(d) = Ymin+ (Ymax−Ymin)\{1 + exp[− g(ln(d) −ln(EC50))]}−1 where y(d) is the % cell viability at the dose d, Ymin is the minimum percent cell viability, Ymaxis the maximum percent cell viability, gis a slope parameter, EC50is the dose that produces 50% of cell viability. Inasmuch as cell viability data are expressed on a 0% to 100% basis, the four-parameter equation is simplified to a two-parameter equation because Ymax and Ymin are constants of 100% and 0%, respectively: y(d) = 0% + (100% −0%) {1 + exp[− g(ln(d) −ln(EC50))]}−1 Interpretation of results With each fluorescent indicator dye (alamar blue, CFDA-AM, and NR), a reduction in fluorescent unit readings in experimentally treated cultures relative to the readings in control cultures indicates cytotoxicity or a loss of cell viability. The use of multiple dyes has the potential of revealing the mechanism(s) behind the cytotoxicity. However, the results and the interpretation can be straightforward or complicated, depending on the toxicant under study. Both a simple (Fig. 1.5.1) and a complex example (Fig. 1.5.2) are presented. When the dose-response curves for the three indicator dyes are identical or very similar for cell cultures after short exposures to toxicants, e.g., illustrated in Figure 1.5.1 for 1,4-dimethyl naphthalene, this indicates that the toxic mechanism is general membrane damage, which includes impairment of organelle membranes, such as those for mitochondria and lysosomes, as well as the plasma membrane. Because this loss of cell viability occurs quickly, it has come to be known as direct cytotoxicity to indicate that cellular metabolism of the toxicant is unlikely to be involved



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CFDA-AM (EC50 = 13.87 µM) neutral red (EC50 = 24.62 µM) 0 1

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Figure 1.5.1 Effect of 1,4-dimethyl naphthalene on viability of RTgill-W1 cultures. After the cultures had been exposed for 2 hr, cell viability was assessed with alamar blue (circles), CFDA-AM (squares), and neutral red (triangles). Results were expressed as a percentage of the readings in control wells exposed to L-15/ex solution with DMSO.

(Schirmer et al., 1998a). The terms ultra-fast cell death, or less preferably, necrosis, have been suggested for cell death that occurs quickly in mammalian cell cultures in response to strong stimuli (Blagosklonny, 2000). Ultrafast cell death appears before the activation of caspase, which is a characteristic of apoptosis (Blagosklonny, 2000). For mammalian cells, Blagosklonny (2000) has suggested that a time frame for the development of a decline in cell viability can be used to distinguish ultra-fast cell death (2 to 16 hr), apoptosis (16 to 36 hr), and slow cell death (>36 hr). As cellular phenomena take longer to develop in fish cells being grown at 18° to 22°C than in mammalian cells at 37°C, the time frame for these in fish cell lines might be increased considerably. Alternatively, the dose-response curves with the different indicator dyes can be unlike one another. Several examples already have appeared in the literature. One combination of outcomes is a decline in cell viability as meas-

ured with alamar blue being accompanied by little or no change in cell viability as measured with CFDA-AM. Such results have been seen with benzo[a]pyrene (BaP) and 6, 12-BaP quinone (Schirmer et al., 2000). As the reduction of alamar blue to a fluorescent product is now thought to indicate cellular metabolism rather than specifically mitochondrial activity (O’Brien et al., 2000), the results are interpreted as indicating that these compounds impair metabolism without impacting plasma membrane integrity. Another outcome combination is that of a greater decline in cell viability as measured with neutral red than in the cell viability monitored with alamar blue and CFDA-AM. This was seen in studies on the photocytoxicity of acenaphthylene, acenaphthene, phenanthrene, fluoranthene, pyrene, anthracene, and benzo[g,h,i]perylene (Schirmer et al., 1998b) . The interpretation of these results is that specific lysosomal damage has occurred immediately after concurrent exposure to these comToxicological Models


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pounds and UV radiation, with little or no impairment of plasma membranes and cellular metabolism. The CFDA-AM assay appears to monitor impairment to plasma membranes, but Figure 1.5.2A illustrates a perplexing outcome. A decline in fluorescent readings occurs at lower concentrations of 2-ethyl phenanthrene with CFDA-AM than with alamar blue, resulting in a lower EC50 for this toxicant. If the CFDA-AM measures membrane integrity, then cellular metabolism evaluated with alamar blue would not be expected to continue with little or no impairment while the integrity of the plasma membrane has been lost. An explanation can be advanced for this apparent anomaly. When carried out as described here, a decrease in fluorescent readings with CFDA-AM actually measures a decline in the total esterase activity within a microwell cell culture. The decrease in esterase activity with toxicant treatment could be achieved in two general ways: the loss of plasma membrane integrity and/or specific inhibitory actions on cellular esterases. In turn, the loss of plasma membrane integrity could decrease culture esterase activity in two slightly different ways. The first of these would be the complete or partial lysis of the cells upon toxicant exposure so that the esterases are released into the medium and lost when the medium is removed and replaced with the CFDA-AM solution. Another possible cause for the diminution of esterase activity is a change in plasma membrane integrity so that cytoplasmic constituents are lost to the medium but the esterases remain contained within the cells, which are still attached to the surface of the microwells. This change in the cytoplasmic milieu would be less able to support maximal esterase activity. Alternatively, the toxicant treatment could leave membrane integrity unimpaired but specifically interfere with cellular esterases, causing activity to decline. Examples of this would be a toxicant interfering with the uptake of the substrate, CFDA-AM, across the plasma membrane or inhibiting the catalytic activity of the esterases. The results in Figure 1.5.2A in which increasing 2-ethyl phenanthrene concentrations cause a more precipitous decline in CFDA-AM readings than in alamar blue read-


ings are likely an example of a toxicant impairing esterase activity rather than the plasma membrane. Additional information into mechanisms of toxicity can be obtained by applying the indicator dyes both immediately after terminating toxicant exposure and later after a period of potential recovery. The assays with alamar blue and CFDA-AM are non-toxic and can be applied on the same microwell cultures for both time points. For the NR assay, which requires cultures to be extracted, separate microwell cultures must be used. Differences between dose-response curves immediately and 24 hr after toxicant exposures have been found, suggesting impairments at specific cellular sites and the capacity of cells to repair them. For example, alamar blue readings decreased immediately after the end of exposures to BaP and 6, 12 BaP quinone, but recovered 24 hr later (Schirmer et al., 2000). As the CFDA-AM readings showed no changes, these results suggested that short exposures to these compounds transitorily disrupted cellular metabolism, although continuous exposure would ultimately lead to cell death. By contrast, a comparison of Figures 1.5.2A and 1.5.2B suggests that a 2-hr exposure to high concentrations of 2-ethyl phenanthrene initiates a process leading to irreparable cellular damage that is expressed as a loss of cell viability 24 hr later. As the development of this cytotoxicity occurs quickly, and for most of the time in the absence of significant amounts of test compound in the medium, the cytotoxicity is unlikely to be due to the generation of toxic metabolites by cell metabolism during the recovery period. Instead, during the 2-hr exposure period, 2-ethyl phenanthrene likely partitions into cell membranes, and in cultures with high concentrations, reaches levels that begins disrupting the plasma membrane. As a membrane-bound toxicant would not be easily removed upon termination of toxicant exposure, the process of plasma membrane impairment would continue during the recovery period, leading to cell death. The above discussion illustrates some of the complexities that might be anticipated. Likely, all the possible scenarios for cellular responses to toxicants as measured with these three indi-

Figure 1.5.2 (at right) Effect of 2-ethyl phenanthrene on viability of RTgill-W1 cultures. After the cultures had been exposed for 2 hr (A), cell viability was assessed with alamar blue (circles), and CFDA-AM (squares). The exposure medium was replaced with complete medium and the cells were incubated for an additional 24 hr (B), after which viability was reassessed with alamar blue (hexagons) and CFDA-AM (diamonds). Results were expressed as a percentage of the readings in control wells exposed to medium with DMSO.



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cator dyes have yet to be described. Several additional complicated scenarios will likely be revealed only by examining more compounds with these indicator dyes. For example, under some circumstances, neutral red readings might increase (Zhang et al., 1990). Overall, the methods described in this unit allow the rapid and inexpensive screening of toxicants for fish cells and at the same time give potential insight into their mechanism(s) of toxicity. In the future, understanding better the cellular function(s) being monitored with each indicator dye will improve their utility in identifying toxicity mechanisms.

Time Considerations One must consider the time to culture the fish-derived cell lines in preparation for exposure to the toxicants. The time between subculturing of the stock culture is between 7 and 10 days if a 1 to 3 split is used routinely. Once the flask is confluent, the cells are transferred to a microwell culture plate allowing the cells to attach and become a confluent monolayer on the bottom of each well, which will take ∼2 to 3 days depending on the use of 48-well or 96-well plates. The time for exposure to a toxicant can vary. Commonly, 2-hr exposures are done with toxicants that are believed to act through a necrotic process of ultra-fast cell death. Longer exposures, commonly 24- or 48-hr exposures, can be done to test toxicants that do not have a response within 2 hr. In terms of the cytotoxicity assays, all three assays can be done using one microplate. Thus, in 50% of all larval cells) form an extralarval cluster (defect III, first phase) after exposure to 20 µM cpf. (C) Exogastrulation (defect II, second phase) after exposure to 40 µM cpf. (D) Transformed cells accumulate in the blastocoel without extruding (defect I) after exposure to 5 µM cpf. (E) Exogastrulation (defect I, second phase) after exposure to 5 µM cpf. (F) At stage 19, larvae appear normal after exposure to 40 µM cpf with 40 µM dimethylaminoethyl arachidonate (AA-DMAE). Imaging was done 62 hr after fertilization (A,B,D,F) or 10 hr later (C,E). Abbreviations: dl, dwarf larva; ec, extralarval cell cluster; tr, transformed (pigmented) cells. Scale bar, 50 µm.

Sea Urchin Embryos and Larvae as Biosensors for Neurotoxicants



A

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Figure 1.6.7 Effects of chlorpyrifos (cpf) introduced at stage 1b (30 min after fertilization, 10°C) on the development of the sea urchin, S. droebachiensis. (A) Exposure to 160 µM cpf. (B) Control stage 8 (8 hr 45 min after fertilization) specimen. (C) Exposure to 40 µM cpf. (D) Control stage 10 (22 hr 25 min after fertilization) specimen. (E) Mushroom-like larvae after exposure to 40 µM cpf. (F) Control stage 20 (64 hr 30 min after fertilization) specimen. Abbreviations: dl, dwarf larva; ec, extralarval cell cluster; tr, transformed (pigmented) cells. Scale bar, 50 µm. Modified from Buznikov et al., 2001b.



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A

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ec ec

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bp bp bp

Figure 1.6.8 Effect of L-nicotine introduced at stage 12 (35 hr after fertilization, 8°C) on development of the sea urchin, S. droebachiensis. (A-C) Development of extralarval cell cluster near the vegetal pole of larvae after exposure to 200 µM L-nicotine. Imaging was done (A) 47, (B) 51, and (C) 60 hr after fertilization. (D) Normal larva at stage 15 (51 hr after fertilization) after exposure to 200 µM L-nicotine and 50 µM imechine. (E) Nearly normal larva at stage 15 after exposure to 200 µM L-nicotine and 40 µM dimethylaminoethyl docosahexaenoate (DHA-DMAE). (F) Extralarval cell cluster near the animal pole of a larva 60 hr after fertilization following exposure to 50 µM L-nicotine and 0.2 µM phorbol 12-myristate 13-acetate (PMA). Abbreviations: bp, blastopore; ec, extralarval cell cluster; hl, hyaline layer (enfoliated). Scale bar, 50 µm.


lysis; Fig. 1.6.9), but, like nicotine and cpf, the period of maximal sensitivity is the MBT. When these lipophilic choline derivatives are introduced during the MBT, they, like cpf, evoke extrusion of transformed cells and formation of extralarval cell clusters at the animal pole (Fig. 1.6.9). In keeping with the fact that it acts on a different neurotransmitter, ritanserin, an antagonist of 5-HT2 receptors, has very different pharmacological characteristics and does not evoke extrusion of transformed cells. Instead, it causes the formation of a multilayered em-

bryonic or larval wall (Fig. 1.6.10). The period of maximal sensitivity to ritanserin does, however, coincide with the MBT, reflecting the fact that different neuromodulators participate together in assembly of the embryo at a critical developmental period. Developmental malformations evoked by cholinergic ligands can be prevented by antagonists of nicotinic acetylcholine receptors (nAChRs) but not by atropine, an antagonist of muscarinic cholinergic receptors. The most effective antidotes for L-nicotine are Dtubocurarine, imechine, QX-222, and some



A

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Figure 1.6.9 Effects of 40 µM arachidonoylcholine (AA-choline) on development of the sea urchin, S. droebachiensis and protective action of 40 µM dimethylaminoethyl arachidonate (AA-DMAE). (A) AA-choline produces one-cell multinuclear embryos, which are beginning to undergo cell lysis. (B) AA-choline with AA-DMAE produces normal hatched larvae (stage 11). (C) AA-choline produces typical mushroom-like larvae at later stages. (D) AA-choline with AA-DMAE produces normal larvae (stages 12 to 13). Substances were introduced at stage 1b (15 min after fertilization; A, B) or at stage 10 (28 hr after fertilization; C,D). Specimens were imaged 3 hr (A) or 27 hr (B) later or 40 hr after fertilization (C,D). Abbreviations: ap, animal pole; ec, extralarval cell cluster; vp, vegetal pole. Scale bars, 50 µm.

other competitive and noncompetitive drugs, especially hydrophilic (quaternary and bisquaternary) compounds. Lipophilic nAChR antagonists such as dimethylaminoethyl arachidonate (AA-DMAE) and dimethylaminoethyl docosahexaenoate (DHA-DMAE) are less effective or ineffective (Buznikov et al., 2001c). Quite the opposite situation exists for cpf and AA-choline, for which AA-DMAE or DHADMAE provides full protection (Fig. 1.6.9), compared with only partial or even insignificant protection by hydrophilic nAChR antagonists. None of the cholinergic antagonists tested prevents the developmental malformations evoked by ritanserin, which acts on 5-HT2 receptors. All rescue compounds for ritanserin are 5-HT receptor agonists, especially very

lipophilic compounds such as serotonamide of arachidonic acid (AA-5-HT), serotonamide of docosahexaenoic acid (DHA-5-HT), and serotonamide of eicosapentaenoic acid (EPA-5-HT; Fig. 1.6.11), which completely rescue the ritanserin phenotype. 5-HT itself (which is hydrophilic) provides only partial protection, however, and the very hydrophilic compound N,N,N-trimethylserotonin iodide (5-HTQ) does not protect at all (Fig. 1.6.10). This suggests that putative intracellular 5-HT receptors (or more precisely, functional equivalents of 5-HT receptors) are the targets for all of the substances that rescue the ritanserin phenotype (and apparently for endogenous 5-HT also). Neurotransmitter receptors ultimately affect gene expression, and hence cell replication and



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Figure 1.6.10 Effect of ritanserin introduced at stage 11 (9 hr 15 min after fertilization, 21°C) on development of the sea urchin, L. variegatus. (A) Exposure to 10 µM ritanserin produces blastulae with multilayered cell walls. (B) Exposure to 10 µM ritanserin with 40 µM serotonamide of arachidonic acid (AA-5-HT) produces almost normal larvae at stage 15. (C) Exposure to 10 µM ritanserin with 40 µM N,N,N-trimethylserotonin iodide (5-HTQ) provides no protection. (D) Control larvae, stage 15. Imaging was done 15 hr after fertilization. Scale bar, 50 µm.


differentiation, by stimulating or inhibiting the activity of intracellular signaling pathways. Phorbol 12-myristate 13-acetate (PMA; 0.01 to 0.2 µM), an activator of protein kinase C, when introduced together with L-nicotine during the MBT, enhances and alters the reaction of sea urchin larvae to nicotine. The relative number of transformed cells increases sharply, and they are extruded at the animal pole (Fig. 1.6.8) rather than the vegetal pole. In the presence of PMA, cleaving embryos become sensitive to L-nicotine (as seen by the formation of multinuclear one-cell embryos and their quick lysis). The developmental effects of L-nicotine with PMA are very similar to the those of AA-choline, not just morphologically but also pharmacologically (as shown by the complete protective action of AA-DMAE and DHADMAE and the incomplete or insignificant protection by D-tubocurarine, imechine, or QX-

222). These malformations can be prevented or reduced by 1-(5-isoquinolinesulfonyl)-2methylpiperazine) (H-7), whereas similar effects of cpf are potentiated by this protein kinase C inhibitor. HA-1004, an inhibitor of protein kinase A, does not affect the sensitivity to any cholinergic neurotoxicants studied. In addition, cleaving sea urchin embryos completely or partly lose their sensitivity to both L-nicotine with PMA and AA-choline when incubated in calcium-free artificial sea water (CFASW). These results provide cogent examples of how the sea urchin model can be used to dissect the intracellular events underlying the mechanisms by which neurotoxicants perturb development. The obvious next step is to pursue the parallel signaling events in the mammalian brain so as to determine if they too reflect actions on these signaling intermediates.



HO HO O O NH NH

NH NH

EPA-5-HT

AA-5-HT HO

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NH NH

DHA-5-HT

Figure 1.6.11 Structure of serotonamides of polyenoic fatty acid. Abbreviations: AA-5-HT, serotonamide of arachidonic acid; DHA-5-HT, serotonamide of docosahexanoic acid; EPA-5-HT, serotonamide of eicosapentanoic acid.

As another example of how this model helps to explore mechanisms of developmental toxicity, the authors have evaluated the effects of chlorpyrifos-oxon (a metabolite of cpf known to be the active inhibitor of cholinesterase). Unlike cpf itself, cpf-oxon does not perturb sea urchin development (Buznikov et al., 2001b). Therefore, the developmental effects of cpf are not associated with inhibition of cholinesterase activity, and the underlying cholinergic component reflects the actions of cpf itself directed towards nAChRs and their associated signaling pathways. In contrast to the ease with which these conclusions were reached with the sea urchin model, comparable mechanistic conclusions in mammalian models would require extensive evaluations of placental transfer, maternal effects, and systemic toxicity of cpf-oxon, all of which are avoided with the invertebrate model. Based on these studies, the authors hypothesize that cpf, AA-choline, and DHA-choline have dual (cholinergic and noncholinergic) actions on sea urchin development. In other words, they can act as agonists of nAChRs coupled to L-type Ca2+ channels together with

signal transduction cascades involving PKC and intracellular Ca2+. Such actions are consistent with the dual effects of cpf on mammalian central nervous system (CNS) development (Slotkin, 1999). nAChRs located at the cell surface are already present at the one-cell stage of sea urchin development and during blastulation (Buznikov et al., 1997b, 1998; Ivonnet and Chambers, 1997); they can be available to some cholinergic ligands that evoke malformations during cleavage divisions (e.g., lipophilic cholines and L-nicotine with PMA). The typical changes in morphogenetic cell movements that occur during the MBT, however, appear to be coincident with rapid changes in expression or function of nAChRs that take place immediately prior to gastrulation. Judging by these morphological and pharmacological data, it appears that cholinergic neurotoxicants may disturb genomic regulatory signals mediated by endogenous acetylcholine during the MBT. Other regulatory signals may be mediated by endogenous 5-HT acting on 5-HT2 receptors, which are disturbed by ritanserin (Buznikov et al., 2001a). Future studies to test this hypothesis should be very informative.



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Invertebrate models such as sea urchin embryos and larvae may ultimately provide us with tools for rapid, high-throughput screening of neurotoxicants that affect mammalian brain development. They may also provide a framework for characterizing the cellular events underlying the neuroteratogenesis of a wide variety of compounds and environmental agents that converge on neurotrophic signals or their downstream signaling cascades. There are, of course, limitations to this model. The sea urchin, like culture-based models, cannot elucidate the importance of maternal-fetal pharmacokinetics and metabolism in determining the concentration of neuroteratogens that reach the fetus. Accordingly, the thresholds necessary to elicit effects in sea urchins cannot provide an absolute guide to the appropriate calculation of exposure limits for regulatory purposes, although they can be used for physiologically based pharmacokinetic models that provide a measure of exposure level of the fetal mammalian brain. On the other hand, the sensitivity of sea urchin embryos and larvae seen with the compounds used in these protocols is quite comparable to that of in vitro mammalian models, such as whole embryos, neuronal cell lines, or CNS cultures (Slotkin, 1999). The authors’ results indicate the potential utility of this model system (or of similar invertebrate model systems) to screen compounds for potential neuroteratogenic activity in a comparative manner and to assist in the identification of heretofore unsuspected cellular targets underlying neuroteratogenesis.

Critical Parameters and Troubleshooting


There are three main critical parameters of experiments with sea urchin embryos. (1) The specimens in the embryo suspensions must be maintained at an optimal density. (2) The optimal temperature for the particular species must be constantly maintained. (3) The eggs, embryos, and larvae must be protected from mechanical injuries, evaporation of water from wells of culture plates, and direct bright light. These are the main reasons that embryos or larvae are of an unsatisfactory quality. Sometimes specimens develop relatively normally up to the beginning of gastrulation, but then their subsequent development can be disturbed. If the relative number of abnormal larvae is 5% to 10% or more in controls, then the experiment needs to be aborted. It is then necessary to assess which conditions (e.g., transportation

and maintenance of adult sea urchins) contributed to the disturbances. Developmental dynamics are one of the main peculiarities of developing embryos as biosensors. Some neurotoxicants tested on sea urchin embryos inhibit, disturb, or block cleavage divisions, whereas others start to evoke developmental malformations at the midblastula stage or later. It is possible to determine the critical periods of vulnerability for different neurotoxicants and to use this information for subsequent analysis of results. Therefore, the timing of experiments is a critical parameter that is worth special attention.

Anticipated Results

Cpf (40 µM or greater) added at stages 10 to 12 should evoke the maximal effect (Buznikov et al., 2001b; defect IV), where ∼90% of embryonic or larval cells are transformed, extruded from the blastocoel at the animal pole, and are observed to cover the surface of the larvae (Fig. 1.6.6A). Lower cpf concentrations (20 to 30 µM) should produce an effect that is strong but less than maximal (defect III), where 50% to 75% of all extruded embryonic or larval cells form a mushroomlike cap (Fig. 1.6.6B). Defects III and IV are incompatible with further development. Cpf at 10 to 15 µM should produce a moderate effect (defect II), where the extralarval cap includes ∼20% to 40% of larval cells (Fig. 1.6.6C) and can be lost, resulting in dwarf larvae that are motile but incapable of further normal development. Defect II is often accompanied by exogastrulation. Cpf at 2.5 to 5 µM should evoke the accumulation of transformed cells in the vegetal half of the blastocoel followed by exogastrulation, but not the extrusion of transformed cells (Fig. 1.6.6D,E and Buznikov et al., 2001b; defect I). These exogastrulated larvae can develop as far as stages 20 to 22. Cpf at 0.5 to 1 µM should not disturb development up to at least stage 25 (Fig. 1.6.4). If cpf is introduced at the one-cell stage or during the first cleavage divisions, embryos should develop quite normally up to stage 10 to 11. In this case, typical developmental malformations may become noticeable when control larvae begin gastrulation (Fig. 1.6.7). AA-choline and DHA-choline (10 to 40 µM) or L-nicotine (25 to 100 µM) with PMA (0.05 to 0.2 µM) added at the one-cell stage evoke the formation of multinuclear embryos followed by rapid cell lysis (Fig. 1.6.9). If the embryos are incubated in CFASW, they are



more or less insensitive to these effects. The same substances added at stages 10 to 12 should evoke malformations (Fig. 1.6.8) very similar to those caused by cpf. In the case of choline derivatives, the dependence of developmental malformations on the concentration of ligand is clear: 20 to 40 µM AA-choline or DHAcholine produces maximal effects, whereas 2.5 to 5 µM are near-threshold concentrations. All developmental malformations described above can be fully prevented by lipophilic cholinergic compounds (e.g., 40 µM AADMAE or DHA-DMAE) introduced simultaneously with cpf or 1 to 2 hr earlier (Fig. 1.6.6F). Nonlipophilic cholinergic compounds such as D-tubocurarine, imechine, or QX-222, however, only partially protect against cpf. L-nicotine alone (50 to 100 µM and even 200 µM) should not disturb the cleavage divisions but evokes the extrusion of cells near the vegetal pole of larvae during gastrulation (Fig. 1.6.8). This effect is prevented by different nonlipophilic nAChR antagonists (D-tubocurarine, imechine, QX-222) but not by atropine; the protective action of lipophilic cholines is incomplete or even insignificant. Ritanserin (2.5 to 10 µM), added during the MBT, blocks gastrulation and evokes the formation of additional cell layers in the larval wall. AA-5-HT, DHA-5-HT, or EPA-5-HT used in equimolar or higher concentrations completely prevents this typical malformation, whereas the protective action of 5-HT is incomplete but observable. 5-HTQ provides no protection against ritanserin.

Time Considerations If cpf, lipophilic choline, or ritanserin are introduced at stage 10 (24 hr after fertilization for S. droebachiensis, 8°C or 7 hr after fertilization for L. variegatus, 21°C), anomalies become evident by 20 hr or 5 to 6 hr later, respectively, with maximal effects occurring after the next 8 to 10 hr (S. droebachiensis) or 2 to 3 hr (L. variegatus).

Literature Cited Bezuglov, V.V., Zinchenko, G.N., Nikitina, L.A., and Buznikov, G.A. 2001. Arachidonoylcholine and N,N-dimethylaminoethyl arachidonate are new cholinergic compounds. Bioorg. Khim. 27:227-230. Bottger, S.A. and McClintock, J.B. 2001. The effects of organic and inorganic phosphates on fertilization and early development in the sea urchin Lytechinus variegatus (Echinodermata: Echinoidea). Comp. Biochem. Physiol. C Toxicol. Pharmacol. 129:307-315.

Buznikov, G.A. 1990. Neurotransmitters in Embryogenesis. Harwood Academic Publishers, Chur, Switzerland. Buznikov, G.A. and Podmarev, V.I. 1990. The sea urchins Strongylocentrotus droebachiensis, S. nudus, and S. intermedius. In Animal Species for Developmental Studies, Vol. 1. Invertebrates (T.A. Dettlaff and S.G. Vassetzky, eds.) pp. 253285. Plenum, New York and London. Buznikov, G.A., Shmukler, Y.B., and Lauder, J.M. 1996. From oocyte to neuron: Do neurotransmitters function in the same way throughout development? Cell. Mol. Neurobiol. 16:533-559. Buznikov, G.A., Jokanovic, M., Kovacevic, N., and Rakic, L.J. 1997a. Sea urchin embryos and larvae as biosensors for screening and detailed study of pharmacologically active substances. Arch. Toxicol. Kinet. Xenobiot. Metab. 5:393400. Buznikov, G.A., Koikov, L.N., Shmukler, Y.B., and Whitaker, M.J. 1997b. Nicotine antagonists (piperidines and quinuclidines) reduce the susceptibility of early sea urchin embryos to agents evoking calcium shock. Gen. Pharmacol. 29:4953. Buznikov, G.A., Marshak, T.L., Malchenko, L.A., Nikitina, L.A., Shmukler, Y.B., Buznikov, A.G., Rakic, L.J., and Whitaker, M.J. 1998. Serotonin and acetylcholine modulate the sensitivity of early sea urchin embryos to protein kinase C activators. Comp. Biochem. Physiol. A 120:457462. Buznikov, G.A., Lambert H.W., and Lauder, J.M. 2001a. Serotonin and serotonin-like substances as regulators of early embryogenesis and morphogenesis. Cell Tissue Res. 305:177-186. Buznikov, G.A., Nikitina, L.A., Bezuglov, V.V., Lauder, J.M., Padilla, S., and Slotkin, T.A. 2001b. An invertebrate model of the developmental neurotoxicity of insecticides: Effects of chlorpyrifos and dieldrin in sea urchin embryos and larvae. Environ. Health Persp. 109:651-661. Buznikov, G.A., Bezuglov, V.V., Nikitina, L.A., Slotkin, T.A., and Lauder, J.M. 2001c. Cholinergic regulation of sea urchin embryonic and larval development. Ross. Fiziol. Zh. Im. I. M. Sechenova 87:1548-1556. Cameron, R.A., Mahairas, G., Rast, J.P., Martinez, P., Biondi, T.R., Swartzell, S., Wallace, J.C., Poutska, A.J., Livingston, B.T., Wray, G.A., Ettensohn, C.A., Lehrach, H., Britten, R.J., Davidson, E.H., and Hood, L. 2000. A sea urchin genome project: Sequence scan, virtual map, and additional resources. Proc. Natl. Acad. Sci. U.S.A. 97:9514-9518. Czihak, G. (ed.) 1975. The Sea Urchin Embryo: Biochemistry and Morphogenesis. Springer, New York. Dettlaff, T.A. and Dettlaff, A.A. 1961. On relative dimensionless characteristics of development duration in embryology. Arch. Biol. 72:1-16. Dinnel, P.A., Link, J.M., Stober, Q.J., Letourneau, M.W., and Roberts, W.F. 1989. Comparative sen-



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sitivity of sea urchin sperm bioassay to metals and pesticides. Arch. Environ. Contam. Toxicol. 18:748-755. Falugi, C. 1993. Localization and possible role of molecules associated with the cholinergic system during “non-nervous” developmental events. Eur. J. Histochem. 37:287-294. Harvey, E.B. 1956. The American Arbacia and Other Sea Urchins. Princeton University Press, Princeton, N.J. Ivonnet, P.I. and Chambers, P.L., 1997. Nicotinic acetylcholine receptors of neuronal type occur in plasma membrane of sea urchin eggs. Zygote 5:277-287. Slotkin, T.A. 1998. Fetal nicotine or cocaine exposure: Which one is worse? J. Pharmacol. Exp. Ther. 285:931-945. Slotkin, T.A. 1999. Developmental cholinotoxicants: Nicotine and chlorpyrifos. Environ. Health Persp. 10(Suppl. 1):71-80. Stephens, R.E. 1972. Studies on the development of the sea urchin Strongylocentrotus droebachien-

sis. I. Ecology and normal development. Biol. Bull. 142:132-144.

Internet Resources http://www.stanford.edu/group/Urchin Describes all procedures related to maintaining adult sea urchins and obtaining and handling sea urchin embryos and larvae.

Contributed by Gennady A. Buznikov N.K. Koltzov Institute of Developmental Biology Moscow, Russia Theodore A. Slotkin Duke University Medical Center Durham, North Carolina Jean M. Lauder University of North Carolina School of Medicine Chapel Hill, North Carolina




Zebrafish: An Animal Model for Toxicological Studies During the last century, teleosts (bony fish) have been widely used as sentinels for assessing environmental hazards including aquatic pollutants. Species used for assessing environmental toxicity include small fish such as fathead minnow, medaka, and zebrafish, as well as large fish such as trout, salmon, and catfish. The underlying biology of environmental toxicity in teleosts has been extensively studied and well described. Molecular biology and genetics have recently been used to elucidate the underlying mechanisms of toxicity in zebrafish and medaka and to predict effects in mammals. The versatile zebrafish is now incorporated in biopharmaceutical programs for elucidating human disease and for preclinical drug discovery and screening.

ZEBRAFISH AS AN ANIMAL MODEL Zebrafish (Danio rerio), a member of the Cyprinidae family, originate in South Asia. Zebrafish prefer warm water, but they thrive in many environments. Zebrafish have been used as a model organism for biological research since the 1930s. The zebrafish embryo has been extensively studied and described (Kimmel et al., 1995) since then and has become a popular model for developmental biology, toxicology, and recently for drug discovery (Peterson et al., 2000; Parng et al., 2002a). In addition, embryogenesis has been investigated using several large-scale genetic screens (Driever et al., 1996; Haffter et al., 1996). The model is now well established in many areas of biological research. Zebrafish are easy to breed and inexpensive to maintain. Adult fish grow to 4 to 5 cm long, and reach sexual maturity in 3 to 4 months. In addition, each female can lay 200 to 300 eggs per week. Zebrafish embryos develop rapidly; embryogenesis is complete five days after fertilization. Because the embryo is transparent, morphological structures and internal organs, including brain, eyes, heart, liver, and kidney can be easily visualized using light microscopy, without the need for surgery. Organ-specific and overall developmental toxicity can be assessed visually or quantified using dyes. Enzyme induction and gene expression can also be easily monitored. In addition, many intracellular biomarkers, such as glutathione, reac-

tive oxygen species, and protein and DNA-adduct formation can be studied in the whole animal. Because of its small size, a single embryo can be maintained in fluid volumes as small as 100 µl for the first six days of development and can be raised in individual wells of microtiter plates. Zebrafish embryos are remarkably permeable. Small molecules added directly to the fish water easily diffuse into the embryos, simplifying drug dispensing and assay processing (Peterson et al., 2000; Parng et al., 2002a). Large or lipophilic molecules can be injected into several hundred embryos in one hour. Because the embryo develops rapidly, compound screening can be completed in a few days. These features make the zebrafish a unique vertebrate model for high-throughput chemical screening, which is useful for preclinical drug discovery and toxicological evaluation (Fishman, 2001; Table 1.7.1).

ZEBRAFISH DEVELOPMENTAL AND GENETIC STUDIES Developmental and genetic studies using whole animals have typically been performed with invertebrates, including fruit fly and nematode. However, these animal models are not closely related to humans and lack many vertebrate organs, hence their use for predicting human toxicity is limited. Zebrafish are more closely related to humans and share many biological traits, genes, developmental processes, anatomy, physiology, and behavior. The size of the zebrafish genome is ∼50% of the mouse genome (Matthews, 2001). In addition, there is extensive synteny between zebrafish and mammalian genomes, and zebrafish genes are ∼75% homologous to human genes on average (Barbazuk et al., 2000). Zebrafish orthologs for some genes that are known to play key roles in human diseases have been identified, and results from large-scale mutant screens have demonstrated that mutations in some of these zebrafish orthologs display phenotypes similar to those present in human diseases (Dooley and Zon, 2000; Ward and Lieschke, 2002).

Developmental Stages Compared with other vertebrates, the embryonic development of zebrafish is rapid. Cell

Contributed by Chaojie Zhang, Catherine Willett, and Trisha Fremgen Current Protocols in Toxicology (2003) 1.7.1-1.7.18 Copyright © 2003 by John Wiley & Sons, Inc.

UNIT 1.7


1.7.1 Supplement 17

Table 1.7.1

Zebrafish as an Animal Model for Toxicity

Advantages

Disadvantages

Ease of maintenance

Use for toxicity testing not standardized by regulatory agencies in the U.S. Physiology less well characterized than mammals

Large number of offspring: ∼200 eggs per female every two weeks Rapid embryogenesis (6 days) Rapid generation cycle (2–3 months) Well characterized developmental stages Transparent embryo: internal organs can be visualized during embryogenesis Embryos are permeable to small molecules, and large or lipophilic molecules can be microinjected Embryos can be directly used to monitor water, soil, and waste water samples Small test sample size required per animal Genetics are well studied with many mutants available and easy to generate Genomic sequence is nearly complete with many genes cloned Transgenic reporter and disease lines available and easy to generate Methods and guidelines for toxicity studies established in Europe

Zebrafish: An Animal Model for Toxicological Studies

division begins shortly after fertilization, and by 6 hr post fertilization (hpf) gastrulation has begun (Fig. 1.7.1). The body plan is established within the first 20 hpf; by 24 hpf the heart is beating and there is a rudimentary circulatory loop (Stainier and Fishman, 1994). Hatching from the chorion occurs at ∼72 hpf, and the hatching rate is usually >80% (Creaser, 1934; Hisaoka and Firlit, 1960). By 96 hpf, the animal can swim and eat, although it still receives nourishment from the yolk. The majority of zebrafish morphogenesis is completed during this time; a marker for this stage is protrusion of the lower jaw and inflation of the swim bladder. By 120 hpf (5 days), all the internal organs, including heart, liver, kidney, and pancreas are fully developed and functioning. The stages of early development have been carefully described, and these stages are considered the standard for developmental biologists (Kimmel et al., 1990). The staging described by Kimmel was performed at 28°C, and this temperature is consistently used in developmental studies; however, much of the toxicity literature describes experiments performed at 26°C. Since development of the exothermic zebrafish embryo is affected by small changes

in temperature, it is important to consider experimental temperature when comparing results.

Mutagenic Screens In the late 1970s, George Streisinger at the University of Oregon recognized that zebrafish embryos have many advantages that make them superior to C. elegans and Drosophila for developmental and genetic studies (Streisinger et al., 1981). Dr. Streisinger used X rays to generate chromosomal abnormalities and developed a technique for producing homozygous diploid fish that made it possible to detect rare recessive mutations and to produce large clones of genetically identical fish. Streisinger’s colleagues at the University of Oregon continued to lay the foundations for establishing zebrafish as a model for developmental genetics. Normal development has been described in considerable detail, and a number of mutations that affect embryogenesis have been described (Kimmel et al., 1995). In the late 1980s, based on the work performed by Streisinger and his colleagues, several developmental biologists recognized that zebrafish would be an extremely useful verte-

1.7.2 Supplement 17


1 hpf

6 hpf

24 hpf

48 hpf

1.0 mm

120 hpf

Figure 1.7.1 Zebrafish development occurs rapidly. By 1 hpf (top left), two cell divisions have occurred. Gastrulation begins at 6 hpf (top center). By 24 hpf (top right), the organ primordia have been laid down and the brain has significantly developed. The heart is beating and a rudimentary circulatory loop through the main body axis has been established. By 48 hpf (middle panel), pigment has begun to form; the body axis has elongated. By 120 hpf (bottom panel), organ morphogenesis is complete and most organs are functioning. The swim bladder has inflated and the embryo can swim and eat on its own. At this point the embryo is considered a larva.

brate for extensive genetic studies. Beginning in 1992, large-scale screens for mutations affecting embryonic development were conducted by Nüsslein-Volhard’s and Driever’s groups in Germany and Boston, respectively (Driever et al., 1996; Haffter et al., 1996). These experiments resulted in the identification and characterization of a large number of mutants affecting many aspects of embryonic development, and the results were jointly published in a dedicated volume of Development (1996, vol. 123). Since then, cell lineage, embryonic axis formation, nervous system development, heart and muscle formation, and differential regulation of gene expression have been extensively studied at various stages of development in wild-type and mutant embryos (Stainier et al., 1993; Driever et al., 1994; Lee et al., 1994; Mullins et al., 1994; Strahle and Blader, 1994; Driever, 1995; Solnica-Krezel et al., 1995; Wilson et al., 1995; Lele and Krone, 1996). Many mutated genes have been cloned, which was an arduous process since chemical mutagenesis used in the screens typically results in point mutations (Talbot and Schier, 1999). Several cloned genes are orthologs of mammalian genes and these mutations result in similar phenotypes as those in humans. Thus, zebrafish is potentially a predictive model for human disease (Dooley and Zon, 2000; Ward and Lieschke, 2002).

In the mid-1990’s, a genetic screen was performed by insertional mutagenesis (Golling et al., 2002). Since this mutagenic process is laborious compared to chemical mutagenesis, this screen yielded fewer mutants, but the mutated genes are tagged by insertions so they can be easily cloned. Several mutants were found in known genes, and several novel genes were identified (Golling et al., 2002). At least one of the mutants resembles a mutation that causes kidney disease in humans (Sun and Hopkins, 2001). Ongoing research into the development and genetics of zebrafish will further strengthen the utility of zebrafish as a model organism for biological studies, including toxicity.

Gene Inactivation by RNA Interference Gene replacements and targeted deletions are not yet feasible in zebrafish; however, it is possible to remove a specific gene product by interfering with translation, a method called RNA interference (RNAi). In zebrafish, a type of RNAi using synthesized oligomers called morpholinos has been successfully used to knockdown expression of a number of genes (Nasevicius and Ekker, 2000; and references in Genesis, 2001, 30:89-200). RNAi knockdown can be used to: (1) identify the function of novel genes, (2) identify possible candidate genes when cloning a gene by position, or (3) inves-



Supplement 17

tigate whether a gene cloned by homology is a functional ortholog of a mammalian gene.

Transgenic Reporter Lines Transgenic fish are a powerful tool for studying gene regulation and expression, disease mechanisms and progression, and drug efficacy and toxicity. Transgenic lines can be created that express a visible marker, such as green fluorescent protein (GFP), under the regulation of specific promoters. These lines can serve as reporters to detect changes in expression of the marked gene and can serve as sentinels to detect the presence of bioactive compounds. The procedure for generating transgenic zebrafish is well established (Stuart et al., 1988, 1990; Westerfield et al., 1992; Lin et al., 1994). The use of live reporter gene markers such as GFP simplifies the screening for progeny that carry the transgene. Recently, high germline transmission rates (nearly 50%) were achieved by attaching DNA to nuclear localization signal peptides or by coinjecting I-SceI meganuclease (Liang et al., 2000; Thermes et al., 2002). Transgenic lines using organ-specific promoters expressing GFP have been created for vessels, kidney, blood, central nervous system, and lymphocytes (Jessen et al., 1999; Goldman et al., 2001; Huang et al., 2001). These lines have been used to study gene regulation, organ development, and the effect of mutations and drugs on development. Several transgenic lines have been developed to assess toxicity. A line expressing GFP from the heat shock protein 70 promoter is currently used to study organ toxicity of heavy metals (Blechinger et al., 2002). Transgenic lines have been created that combine several response elements, such as those for aromatic hydrocarbons, electrophiles, metals, estrogen, and retinoic acid, driving the expression of luciferase or GFP, for use as sentinels of aquatic pollution (Carvan et al., 2000; Kinoshita et al., 2000). Another transgenic line has been developed for screening estrogenic activity (Legler et al., 2002; Schreurs et al., 2002). Advantages of these lines include the ability to distinguish aquatic pollutants using living biosensors and to elucidate toxicity response to various toxins in whole animals.

TOXICOLOGICAL STUDIES Zebrafish: An Animal Model for Toxicological Studies

Starting in the 1950s, zebrafish have been used to test the toxicity of synthetic chemicals and natural products (Battle and Hisaoka, 1952; Hisaoka and Hopper, 1953, 1957;

Hisaoka, 1955). The toxic effects of some metals (e.g., zinc, selenium, mercury, and copper) and some organic solvents (e.g., phenol, aniline, and cyclohexane) were also assessed in zebrafish (Vitozzi and De Angelis, 1991). Because of the convenience and low cost of maintaining and handling zebrafish, subsequent studies in all areas of toxicology were performed in order to understand the adverse effects of chemicals and to predict results in humans. In experiments performed using zebrafish, lethality and malformation (during embryonic development) were used as general parameters for assessing toxicity. These studies demonstrated that zebrafish are amenable to chemical screens and exhibit good dose responsiveness to toxicity. In addition, the zebrafish embryo may have a predictive value similar to mammalian teratogenicity and toxicity assays (Van Leeuwen et al., 1990; Ensenbach and Nagel, 1995). However, since few compounds were tested, and effects on individual tissues and organs were not assessed, more studies are needed to further validate the correlation between the zebrafish and mammalian toxicity tests.

Ecotoxicological Studies Ecotoxicological studies are performed to detect, manage, and monitor the presence of environmental pollutants in water, soil, and waste. Assays using zebrafish have been developed to directly monitor water, soil, and wastewater quality (Ruoppa and Nakari, 1988). Zebrafish have also been used to assess risks, predict hazards associated with petroleum products and by-products, and evaluate working conditions in the petrochemical and mining industries (Vitozzi and De Angelis, 1991; Lele and Krone, 1996). In addition, pesticides, polychlorinated biphenyls (PCBs), dioxin, and some of their derivatives have been studied in zebrafish embryos (Burkhardt-Holm et al., 1999; Wiegand et al., 2000, 2001; Anderson, 2001; Andreasen et al., 2002). Because zebrafish develop rapidly, many toxicological endpoints can be evaluated a few days after fertilization. According to Schulte and Nagel (1994), toxicological endpoints at 72 hpf fall into two major groups. Type I endpoints are used for general morphological evaluation of development and include egg coagulation, embryo gastrulation, somite formation, tail extension, heart function, spontaneous movement, and hatching frequency. Type II endpoints are more detailed and include quantification of number of somites, blood circula-

1.7.4 Supplement 17


tion, eye development, spontaneous movement at 24 hpf, cardiac function measured by heart rate, otolith development, melanocyte development, skeletal malformation, and delayed hatching (Zhu and Shi, 2002). In Europe, zebrafish are widely accepted as an assay system for ecotoxicological studies. A number of recent guidelines from the Organisation of Economic Co-operation and Development (OECD; Guidelines 203, 210, and 212; OECD, 1992a,c, 1998) recommend zebrafish for aquatic toxicity testing. Different guidelines recommend using zebrafish at various stages, such as egg- and sac-fry-stage fish, early-lifestage fish, and juvenile fish. When testing a compound for acute toxicity (lethality), Guideline 203 (Fish Acute Toxicity Test; OECD, 1992a) recommends that medium lethal concentration (LC50) be obtained using zebrafish 2.0 ± 1.0 cm long (i.e., 2- to 3-month-old juvenile to adult fish), using an incubation time of 96 hr. Guideline 210 (Fish, early-life stage toxicity test; OECD, 1992c) suggests the use of early life stage fish, from 4 hpf to day 16, for chemical testing. Briefly, 4-hpf embryos are exposed to a chemical, usually under flowthrough conditions, until all the fish are free swimming and feeding exogenously. Observation continues until day 16. Lethal and sublethal effects on parameters such as growth are assessed and compared with controls. Since larval and early fry stages are the most sensitive stages for assessing developmental effects of xenobiotics, OECD Guideline 212 (Fish, short-term toxicity test on embryo and sac-fry stages; OECD, 1998) focuses on developmental toxicity in the early fish, from shortly after fertilization to 5 days post-hatching. Each guideline includes a detailed discussion of the principle of the test, a description of the method, procedures, and formats for data reporting and analysis. Zebrafish embryos have been used to assess the toxicity of heavy metals, organic pesticides, and organic reagents (Vitozzi and De Angelis, 1991; Zhu and Shi, 2002). The no-observed-effect level (NOEL) and lowest-observed-effect level (LOEL) were determined after treatment with copper, mercury, lead, nickel, chromium, zinc, and cobalt for 16 days, starting at 4 hpf. Treatment effects included delayed hatching and developmental toxicity (Zhu and Shi, 2002). Organic pesticides, such as astrazine, malathion, carbaryl, urea, and lindane have been tested and the LC50 values were reported (Dave, 1987; Gorge and Napel, 1990; Dave and Xiu, 1991; Groth et al., 1993; Schulte and Nagel, 1994; Zhu and Shi, 2002). The LC50 of

40 organic-solvent reagents were assessed after incubating for 48 or 96 hr (Zhu and Shi, 2002). Zebrafish-derived cell lines are also used for these purposes. Toxicity studies with polychlorinated dioxin compounds, biphenyl, and antifouling compounds have also been reported (Henry, 2000; Okamura, 2002). Results from these experiments showed that the zebrafish embryo is sensitive to environmental toxins and is a reliable and convenient model for toxicity testing. Furthermore, zebrafish can be used to monitor and elucidate the toxicological mechanisms underlying embryo development and to identify organs targeted by specific compounds.

Endocrine Disruption There is increasing awareness that both man-made and naturally occurring chemicals may mimic or interfere with the endocrine processes in humans. Such substances have been termed endocrine disruptors (EDCs). A range of synthetic chemicals and some natural compounds have been reported to mimic estrogen effects in aquatic organisms (Pelissero and Sumpter, 1992; Sumpter and Jobling, 1995; Harries et al., 1996). Currently, there is a great demand for new assay methods, predictive endpoints, and reliable animal models for endocrine disruption testing and assessment (EDTA). EDCs interfere with homeostasis by interacting with receptors for endogenous hormones, or they can exert effects in a receptorindependent manner. Estrogens and their analogs, such as 17-β-estrodiol (E2) and 17-αethinylestradiol (EE2), are the most potent EDCs identified so far. OECD recommends the use of several fish species for monitoring estrogens and estrogen analogs (OECD, 1992b,c, 2000). According to these Guidelines, teleosts such as fathead minnow, zebrafish, and medaka are the most promising models. Although fathead minnow, zebrafish, and medaka are all small fish and have similar generation cycles (3 to 4 months), zebrafish has several inherent advantages for EDTA studies, including a unique process of sexual development that differs from either the fathead minnow or medaka. Sex determination in zebrafish is not genetically determined: sex ratios are dependent on population density and food availability during development. In addition, gonadal development in zebrafish passes through a stage of juvenile hermaphroditism (Takahashi, 1977; Chan and Yenung, 1983). This means that short-term exposure may affect the genetic regulation of both female and male



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Comparison of Toxicity in Zebrafish and Mammalian Modelsa

Table 1.7.2

Zebrafish LC50/log LC50 (mg/liter)

Compound

Target organs

Mammalian models LC50/log LC50 (mg/kg)

Model (dose route)

Target organs

Acetaminophen

252/2.4

Liver

500/2.69

Mouse (i.p.)

Aspirin

100.9/2.0

Gastrointestinal, teratogenic, kidney, muscle

167/2.22

Mouse (i.p.)

Caffeine

108/2.03

127/2.1

Mouse (i.p.)

Col-3

1.0/0

Behavioral, neuronal Teratogenic, liver, Gastrointestinal

75/1.87

Rat (oral)

Developmental, liver Teratogenic, liver, cardiovascular, kidney Liver, gastrointestinal, kidney Teratogenic, liver, kidney Developmental Teratogenic, liver

315/2.5

Mouse (oral)

170/2.23

Mouse (i.v.)

Kidney, ureter, bladder

410/2.61

Mouse (i.p.)

1500/3.17

Rat (oral)

4.0/0.6 268/2.4

Mouse (i.v.) Rat (oral)

21/1.35

Mouse (i.v.)

Liver, heart, kidney, gastrointestinal Liver, kidney, cardiovascular Kidney Teratogenic, carcinogenic, liver, reproductive Liver, cardiovascular

Cyclophosphamide 650/2.8 Cyclosporin A

69/1.83

Dexamethasone

324/2.51

DCA

72/1.85

Didemnin B DNT

6.2/0.79 23/1.4

Doxorubicin

30.3/1.51

Ecteinascidin

0.42

Ethanol

11,180/4.0

Epirubicin

16.3/1.2

Flavopiridol

2.2/0.34

5-FU

3.3/0.52

Fujisawa peptide

30/1.48

Teratogenic, liver, cardiovascular, kidney Teratogenic, kidney Teratogenic, neuronal, craniofacial Cardiac, hemorrhage Teratogenic, liver, gastrointestinal

—

—

7,000/3.8

Rat (oral)

16/1.2

Mouse (i.v.)

3/0.48

Rat (oralc)

Liver, kidney

18.9/1.2

Rabbit (oral)

Teratogenic, heart, kidney

5.1/0.7

Rat (i.v.)

Liver, kidney, gastrointestinal Gastrointestinal, kidney, ureter, cardiovascular, muscle Behavioral, neuronal Gastrointestinal, kidney, liver, bone marrowb Teratogenic, liver

— Liver, teratogenic, neuronal, craniofacial Carcinogenic, mutagenic, cardiac Teratogenic, liver, gastrointestinal, spleen, bone marrow (Arneson et al., 1995) Reproductive, liver, kidney Lung, heart continued


1.7.6 Supplement 17


Table 1.7.2

Comparison of Toxicity in Zebrafish and Mammalian Modelsa, continued

Zebrafish LC50/log LC50 (mg/liter)

Compound Geldanamycin Ibuprofen Methotrexate

3.13/0.49 5.56/0.74 454/2.6

Naproxen

13.2/1.12

Phenylurea dithiocarbamate PCBs

1.5 10/1

Staurosporine SU5416

0.012/−1.92 1.0/0

Tacrine TCDD

11.13/1.04 0.98/−0.015

Trithiophene Vinblastine sulfate

2.7/0.43 90.9/1.96

Mammalian models

Target organs

LC50/log LC50 (mg/kg)

Model (dose route)

Target organs

Liver Liver, kidney Gastrointestinal, liver, kidney, teratogenic Liver, gastrointestinal Teratogenic, liver

1.0/0 495/2.69 180/2.3

Mouse (i.p.) Mouse (i.p.) Rat (oral)

Liver Kidney Gastrointestinal, liver, teratogenic

435/2.63

Mouse (i.v.)

Gastrointestinal

Unknown

—

Liver, gastrointestinal Liver, kidney Teratogenic, liver, cardiovascular Teratogenic, liver Cardiovascular, liver, kidney, teratogenic, growth inhibition, neuronal Liver, muscle Liver, gastrointestinal, developmental

8000/3.9 Unknown Unknown

Rat (i.v.) — —

Liver Liver Unknown Unknown

20/1.3 0.114/−0.94

Mouse (i.p.) Rat (oral)

Liver Cardiovascular, liver, kidney, teratogenic, neuronal

110/2.04 3.05/2.48

Rat (i.p.) Rat (oral)

Liver Mutagenic, bone marrow, depression, liver, developmental

aAbbreviations: DCA, dichloroacetic acid; DNT, 2,4-dinitrotoluene; 5-FU, 5-fluorouracil; PCBs, polychlorinated biphenyls. bSee Page et al. (1998). cMaximum tolerance dose.

development, resulting in a change of sex or sexual characteristics. However, sexual differentiation in medaka begins before hatching in females and at ∼13 days post hatching in males (Kime and Nash, 1999). Gonad development in the fathead minnow is less well characterized. Moreover, the sex ratio has been well characterized for zebrafish under specific conditions (Fenske et al., 1999), but it is not known for either fathead minnow or medaka. Therefore, use of zebrafish for EDTA studies may produce more accurate and predictive results than other species. Furthermore, due to the small size of zebrafish compared to larger species such as rainbow trout, zebrafish assays can substantially reduce the cost, time, and space required for testing.

It has been reported that treatment with estrogen or its analogs changed the sex ratio of the siblings of exposed female zebrafish, caused necrosis in testes, and induced over-expression of vitellogenin protein (VTG; Kime, 1998; Olsson et al., 1999). VTG is an egg yolk protein synthesized in the liver and excreted into blood in the female fish. Estrogen is believed to regulate the expression of VTG. Due to the low concentration of circulating estrogen in male and juvenile fish, their blood levels of VTG are usually low. An increase in VTG in nonfemale fish indicates the presence of EDCs. Therefore, monitoring the blood levels of VTG in male fish has become a reliable endpoint for EDTA studies (Sumpter and Jobling, 1995). Toxicological Models


Supplement 17

Whole-fish homogenate is used in EDTA tests because it is difficult to obtain enough blood from zebrafish. Using a zebrafish-specific anti-VTG antibody, VTG at concentrations as low as 40 ng/g fish can be detected by ELISA. Other approaches include immunohistochemical studies, RT-PCR to detect VTG mRNA, histological analysis of gonad development, reproduction rates, and reproductive strength of the following generation. For histological examination after chemical exposure, the gonad is removed and sectioned. Sections are fixed on a slide and examined under a light microscope for any morphological change or necrosis. Studies related to reproduction include the ability to lay eggs, the fertilization rate, and the fertility of the subsequent generation. It has been reported that EDCs caused developmental arrest of eggs from exposed females (Kime and Nash, 1999; Olsson et al., 1999). PCBs and their metabolites, di-2-ethylhexyl phthalate, 2,3,7,8-tetrachlorodibenzo-pdioxin (TCDD), and toxaphene have been tested on zebrafish (Holm et al., 1995; Orn et al., 1998; Olsson et al., 1999). It has been proposed that TCDD exerts its antiestrogenic effect by binding to the Ah-receptor, rather than by interfering with the estrogen receptor. Although the mechanism of action is not fully known, TCDD has been shown to reduce egg number and cause abnormality in offspring of mature females that ingested TCDD (Wannemacher et al., 1992; Fahraus-Van Ree and Payne, 1997; Gillesby and Zacharewski, 1998). However, which signal transduction pathway is involved in toxicity of PCBs is controversial. PCBs, like TCDD, have been reported to bind to the Ah-receptor (Cook et al., 1997). In contrast, PCB 104 was capable of mimicking the effect of E2 by inducing ER mRNA (Billsson et al., 1998). Since the mechanisms of endocrine disruption are quite complicated, additional research is needed in this area.

TOXICITY TESTING OF THERAPEUTICS: A COMPREHENSIVE APPROACH USING ZEBRAFISH


The authors have recently developed a family of assays in which chemical toxicity can be rapidly assessed in zebrafish. These methods include acute toxicity testing, target organ toxicity, developmental toxicity and teratogenicity, and an assay for apoptosis. Using 30 chemicals which were commercially available or provided by the National

Cancer Institute (NCI), the authors demonstrated the feasibility of using zebrafish embryos to study toxicity response by assessing LC50, developmental teratogenicity, and specific organ toxicity (Table 1.7.2). The data show good correlation with LC50 values in other mammalian models (Fig. 1.7.2). In addition, as expected, >90% of the compounds showed specific tissue, organ, and behavioral toxicity. These results suggest that zebrafish is an excellent model organism for drug toxicity testing. LC50 results showed that a few of the compounds (trithiophene, naproxen, ibuprofen, Col-3, DCA, PCBs, and 2,4-DNT) were more toxic in zebrafish than in mammals, perhaps because testing in zebrafish was performed on embryos, a developmental system, while the comparative mammalian toxicity was obtained using adult animals. The finding that a subset of compounds is more toxic in zebrafish than in mice or rats suggests that zebrafish may be a more sensitive model for chemical toxicity. Zebrafish embryos exhibit responses to xenobiotic chemicals similar to those in mammals, including induction of xenobiotic enzymes and generation of oxidative stress (Wiegand et al., 2000; Dong et al., 2001). In addition to developmental defects (see below), changes in gene expression can be used for toxicogenomic studies. Although zebrafish orthologs of many genes involved in human toxicity have been identified (Table 1.7.3), expression of the cytochrome gene CYP 1A1, the first cytochrome gene cloned and characterized, is examined in most zebrafish toxicity studies. CYP 1A1 has been shown to be up-regulated in a zebrafish liver cell line treated with TCDD and β-naphthoflavone (BNF; Miranda et al., 1993). Additionally, CYP 1A1 has been shown to be functionally active in embryos in late gastrula stage, and its expression can be induced by TCDD or hexachlorobenzene (HCB; Mizell and Roming, 1997). Recent studies designed to assess effects of several drugs on toxicity response demonstrate that CYP 1A1 expression was up-regulated in zebrafish embryos treated with 10 µM tacrine and 50 µM phenytoin, and down-regulated with 10 µM ibuprofen and 100 µM doxorubicin (Fig. 1.7.3). These results suggest that, similar to mammalian models (Sinz and Woolf, 1997; Nakamura et al., 2000), CYP P450 in zebrafish is responsive to drug treatment (Parng et al., 2002b; Semino et al., 2002). Expression of several genes has been examined in response to a number of drugs (Parng et al., 2002b; Semino et al., 2002), and currently DNA arrays are

1.7.8 Supplement 17


mammals, log LC50 (mg/kg)

5 4 3 2 1

y = 1.1283x

0 −1 −1

0

1 2 3 4 Zebrafish, log LC50 (mg/liter)

5

Figure 1.7.2 Comparative LC50 values. LC50 values obtained with zebrafish (expressed as mg/liter in Table 1.7.2) are compared with the corresponding LD50 values (mg/lg) obtained with mammals, in logarithmic scale. A graphic representation of the data was developed; the line represents the best fit between the two sets of values (slope = 1.13). LD50 values for mammals were obtained from the NIH TOXNET database, NCI, and others. Table 1.7.3

Examples of Zebrafish Homologs of Mammalian Toxicity Related Genes

Gene Apoptosis Bad Bax Bcl-L Caspase 3 Casp8 Mcl 1b Mcl-1A Survivin 2 TRAIL-like protein TNF Cell cycle CDK5 Cyclin B1 Cyclin D1 Cyclin E Proliferation CBP/p300 IGF-1 IGF-2 STAT3 Neurotrophin-7 Ngr1 NurD Pax-6 PDGF WT-1

GenBank no.a AF231017 AF231015 AAK81706 AB047003 AF273220 AF441284 AF302805 AY057058 AF250041 AF250042 AF203736 NM_131513 NM_131025 NM_130995 AF359242 AF268051 AF194333 AJ005693 AF055906 AF036149 AF036148 X63183 AF200951 AY028627

Gene Stress response Egr1 Heat shock factor 1a Heat shock factor 1b Max MDM2 Mix1 c-Myc Nueronal nitric oxide synthase P53 Xenobiotic AhR1 AhR2 Catalase COX-1 COX-2 CYP 1A1 CYP 2E1 CYP 2A6 CYP 3A4 ERalpha ERβ1 ERβ2 Glutathione-S-transferase Pyruvate carboxylase

aSee http://www.psc.edu/general/software/packages/genbank/genbank.html.

GenBank no.a U12895 AF159134 AF159135 L11711 AF010255 AAA19324 L11710 AF219519 NM_131327 NM_131028 NM_131264 AAF89686 AY028584 AY028585 AF210727 AA605954 AI545376 BF717373 AB037185 AJ414566 AJ414567 NM_131734 NM_131550



Supplement 17

− +

doxorubicin

− +

ibuprofen

phenytoin

tacrine

− +

− +

CYP1A1

GAPDH

Figure 1.7.3 CYP 1A1 expression in zebrafish embryos. Zebrafish embryos (96 hpf) were treated with 0.1% DMSO (−) or 10 µM tacrine, 50 µM phenytoin, 10 µM ibuprofen, or 10 µM doxorubicin (+) for 4 hr. RNAs were isolated, and RT-PCR was used to analyze expression of both CYP 1A1 and GAPDH, a housekeeping gene. CYP 1A1 was up-regulated by tacrine and phenytoin treatment and down-regulated by ibuprofen and doxorubicin treatment.

being generated to assess patterns of gene expression and toxicity (by Phylonix, MWG Biotech, and Oakridge National Labs). Although these results suggest that P450 metabolism in zebrafish may be similar to that in mammals, the function of P450 in zebrafish has not been directly studied.

ORGAN TOXICITY


Organ toxicity studies include examining liver, gastrointestinal (GI), and kidney morphologies. Liver and GI toxicity is assessed using a staining method that relies on the presence of biotinylated proteins in the liver and gut: biotin is visualized by the application of streptavidin-peroxidase followed by a chromogenic substrate for peroxidase to specifically visualize the liver and intestine after drug treatment (Fig. 1.7.4A). The data showed that treating a 24-hpf zebrafish embryo with 100 µM dexamethasone for 5 days caused a dramatic reduction in liver size. Kidney toxicity is assessed by staining with an antibody against the mouse sodium/potassium ATPase (Fig. 1.7.4B). Antibody staining outlines the pronephric duct (black arrows) and the newly developed nephron (white arrow) at 48 hpf. Treating embryos for 24 hr with 5 µM brefeldin A had little effect on the pronephric duct, but completely inhibited nephron development. Using the same methods described above, organ toxicity was quantified by staining with soluble substrate. The intensity of the resulting colorimetric product is proportional to the size of the organ. In addition to quantifying the toxic response, this method can be automated to

process a large number of animals using a microplate reader. The results show that dexamethasone (from 1 to 100 µM) reduced liver and GI size in 144-hpf zebrafish embryos in a dose dependent manner (Fig. 1.7.5). A functional assay for fatty acid metabolism in the gut has also been developed (Farber et al., 2001). This assay relies on the change in the fluorescence of phospholipase A2 after cleavage in the gut and can be used to identify chemicals that alter lipid metabolism. Developmental toxicity in zebrafish embryos has been assessed after treatment with valproic acid (VPA), an anticonvulsant drug, and twelve other related substances (Herrmann, 1993). It was shown that VPA and the twelve other compounds cause retardation, developmental arrest, and malformation, including edema, brain deformation, shortened and bent tail, and bipartite axiation of the posterior trunk. These findings show that results with zebrafish embryos are similar to those observed in hydroids and in mammalian embryo culture systems. Furthermore, zebrafish development is highly synchronous and the developmental stages can be accurately determined, making zebrafish embryos a suitable model for drug toxicity testing. The authors have developed a detailed system to assess developmental toxicity in response to chemicals and therapeutics treatment (see below).

Developmental Toxicity and Teratogenicity Developmental toxicity is a major aspect of environmental pollutants and a significant



A

B E

T G E T G

Figure 1.7.4 Visual assessment of liver and kidney organ toxicity. (A) Zebrafish embryos (24 hpf) treated for 5 days were used to determine the effects of dexamethasone on liver toxicity. The embryos were fixed with paraformaldehyde, incubated with streptavidin-peroxidase, and stained with a chromogenic dye to specifically label biotinylated enzymes in the liver and gut. Arrows indicate the position of the liver. Top, untreated embryo; bottom, embryo treated with 100 µM dexamethasone for 5 days. E, eye; G, gastrointestine; T, tail. Scale bar = 1 mm. (B) Zebrafish embryos (48 hpf) were fixed with methanol and formalin and stained with a mouse antibody against sodium/potassium ATPase. Black arrow indicates the position of the pronephric duct; white arrow indicates the nephron. Top, untreated embryo; bottom, embryo treated with 5 µM brefeldin A for 24 hr.

120 % of Control

100 80 60 40 20 0

0

1 10 Dexamethasone µ/M)

100

Figure 1.7.5 Quantification of liver toxicity after dexamethasone treatment: generation of dose response. Larva 144 hpf show a dose response to 1 to 100 µM dexamethasone on liver size detected and quantified by peroxidase and stained with a soluble dye. Ten zebrafish were analyzed for each concentration. The absorbance was detected at 405 nm. Values are expressed as a percentage of control (% Control) ± standard deviation.

problem with numerous pharmaceutical agents. Several mammalian models are typically used for developmental toxicity studies, including rat and mouse whole-embryo, organ, and tissue cultures (Chatot et al., 1980; Cicurel and Schmid, 1988). However, mammalian whole-embryo cultures are complicated: embryos must be explanted with the visceral yolk sac and ectoplacental cone intact for culture,

and the embryo culture time is limited to 48 hr (Bechter et al., 1991). Partly due to the embryonic attributes described above, zebrafish is a much simpler model system for assessing toxicity. The frog embryo teratogenesis assayXenopus (FETAX) is well established for evaluating developmental toxicity (Van Leeuwen et al., 1990; Dawson, 1991; Fort et al.,



Supplement 17

A

B

Figure 1.7.6 Morphological characteristics for developmental index assessment. Comparison of zebrafish (120 hpf) untreated (A) and treated (B) with 5 µM brefeldin A. Treated zebrafish shows heart edema (black arrow), necrotic liver (white arrow), undeveloped intestine (large arrow), underdeveloped jaw (gray arrow), and lack of swim bladder (arrowhead).

B 125 100 75 50

LC50 = 4 µM EC50 = 0.48 µM

25 0 0.0001 0.001 0.01 0.1 Log [conc.] µM

1

10

TI = 10.4

% Malformation/mortality

% Malformation/mortality

A

125 100 75 50 25 0 1E- 1E- 1E- 0.001 0.01 0.1 1 06 05 04 Log [conc.] µM

LC50 = 0.1 µM EC50 = 0.001 µM 10

TI = 10.4

Figure 1.7.7 Dose-response curves for developmental toxicity of flavopiridol (A) and TCDD (B). Developmental malformation (diamonds) and mortality (squares) are shown.


1991; Stringer and Blankememyer, 1993). However, it appears that the frog is not responsive to halogenated aromatic hydrocarbons (HAHs), such as 2,3,7,8-tetrachlorodibenzo-pdioxin (TCDD; Beck and Powell, 2002), a major class of environmental toxins. Another complication of FETAX is that many compounds require activation by metabolic liver enzymes; since toxicity is evaluated before the embryonic frog liver develops, FETAX assessment of some compounds must be done in the presence of liver extract (Fort et al., 1991, 1998). Furthermore, since frog embryos are not transparent, it is difficult to inspect effects on internal organs. The authors have developed a scoring system for evaluating developmental toxicity using zebrafish embryos. The scoring system consists of 20 toxicological endpoints, including heart shape, swim bladder inflation, jaw development, body edema, hemorrhage, several aspects of organ malformation, and necrosis (Fig.

1.7.6). The authors determine the mortality (LC50), malformation (EC50), no-observed-effect level for malformation (NOEL), and teratogenic index (TI) by generating dose-response curves for mortality and developmental malformation. The authors have assessed the effects of eight chemicals, including environmental contaminants and common drugs, and more studies are ongoing. The results are shown in Fig. 1.7.7 and Table 1.7.4. Treatment by semistatic emersion (compounds replenished daily) was initiated at 24 hpf (day 1) and terminated at 120 hpf (day 5). Mortality was monitored each day and used to obtain the dose-response curve to establish the LC50 value—i.e, the percentage lethality was plotted against the concentration to obtain the concentration that caused 50% lethality (LC50). To obtain the dose-response curve for malformation, the developmental score was plotted against the concentrations for each compound. The concentration that caused



Table 1.7.4

Comparison of Chemical Teratogenicitya,b

Chemicals

Zebrafish TI

Zebrafish LC50 (mg/liter)

FETAX (TI)

LC50 or MTD in other species

Teratogenicity in mammals

Col-3 2,4-DNT Epirubicin Flavopiridol Methotrexate Staurosporine SU 5416 TCDD

3 1.7 0.75 10.4 18.2 1.0 4.3 100

1.0 23 16.3 2.2 454b 0.012 1.0 0.98

75 mg/kg 24.3 mg/liter 16 mg/kg 3 mg/kg (MTD) 504 mg/liter Unknown Unknown 0.81 mg/kg

Rat (oral) 96-hpf fathead minnow Mouse (i.v.) Rat (oral) FETAX — — Streptopelia rioria (oral)

Mouse, rat Mouse, rat, rabbit — Mouse, rat, hamster Mouse, rat, hamster — Unknown Mouse, rat, rabbit

aAbbreviations: DNT, 2,4-dinitrotoluene; FETAX, frog embryo teratogenesis assay-Xenopus; MTD, maximum tolerated dose; TCDD, 2,3,7,8,-

tetrachlorodibenzo-p-dioxin. bFETAX TI value 19.3. See Bantle (1995).

50% malformation is defined as the medium effective concentration (EC50). The teratogenic index (TI) is defined as the ratio of LC50/ EC50. The test compound is considered teratogenic if the TI value is >1.5 (Bantle et al., 1989, 1990; Bantle, 1995). Dose-response curves for EC50 and LC50 for flavopiridol (a cyclin-dependent kinase inhibitor used as an experimental anticancer compound), and TCDD, are shown in Figure 1.7.7. Developmental toxicity of the eight compounds was studied using the scoring system and compared with data for other species (Table 1.7.4). Although the authors have only assessed eight compounds to date, the data strongly suggest that zebrafish embryo is a suitable model for studying developmental toxicity and teratogenicity.

TOXICITY-RELATED CELL DEATH Cell death often occurs in response to exposure to toxic compounds. General cell death, or necrosis, is easily scored in the developing embryo by localized opacity within an organ (Fig. 1.7.6). Apoptosis, or programmed cell death, is a mechanism of ridding the organism of unwanted or damaged cells and is involved in organ toxicity of many classes of compounds. The pathways and mechanisms of apoptosis in zebrafish appear to be similar to those in mammals. For example, specific caspase inhibitors have been shown to inhibit DNA fragmentation in zebrafish embryos, similar to mammals (Inohara and Nunez, 2000). Prevention of apoptosis in zebrafish embryos using caspase inhibitors has also been reported (Chan and Yager, 1998). Homologs of many genes in the apoptotic pathway in ze-

brafish have been identified and cloned, including nine Bcl-2 family genes (Mcl-1a, Mcl-1b, BLP1, Bcl-xl, Bax, Bad, Nip3, Nip3L, NR13), and seven caspase member genes (caspase 2, 3, 4, 6, 8, 9, 13; Parng et al., 2002a). Other apoptosis-related genes identified in zebrafish include four IAPs (inhibitor of apoptosis), four death receptor genes, ten apoptosis related kinases, and other related transcriptional genes and molecules (Ikegami et al., 1997). Therefore, zebrafish is a promising animal model for studying apoptosis mechanisms and gene regulation in response to environmental toxins and for evaluating the potential toxicity of therapeutics. To detect apoptotic cells in live embryos, the authors developed apoptosis assays that rely on fluorescent reagents, including acridine orange and caspase substrates. Apoptotic cells are stained with fluorescent dyes that are easily detected in vivo with an epifluorescent microscope. In developing zebrafish embryos, naturally occurring apoptosis is consistently detected in the dorsal neural tube, hatching glands, retina, lens, cornea, inner ear, and olfactory organs (Bader et al., 1982; Yabu et al., 2001; Parng et al., 2002a); therefore, reagents that alter this developmental apoptosis pattern can be effectively screened in transparent embryos and the target organs or cells of pro-apoptotic reagents can be identified. As shown in Figure 1.7.8, increased apoptosis can be visualized in the liver of neomycin-treated embryos. In addition, these assays can be adapted for quantitative microtiter plate analysis using the same approach described for organ toxicity screens. Toxicological Models


Supplement 17

A

A

E

SB L GI

B

B E

L SB GI

Figure 1.7.8 Neomycin-induced liver apoptosis in zebrafish. Zebrafish embryos (96 hpf) were stained with acridine orange. (A) Untreated zebrafish. (B) Zebrafish treated with 2.5 µg/ml neomycin for 24 hr. Left panels show light micrographs and right panels, fluorescent micrographs. Neomycin induced apoptosis in the liver (arrow in lower right panel). Intestine (GI) is fluorescent in both treated and untreated zebrafish due to ingestion of dye. Abbreviations: L, liver; E, ear; SB, swim bladder.

SUMMARY Zebrafish is a unique animal model for assessing toxicity. It is a vertebrate that completes morphogenesis in 120 hpf, and embryos are transparent and develop ex utero. Since zebrafish has become a popular model for geneticists and developmental biologists, information about all aspects of zebrafish biology is rapidly accumulating. In addition, for decades zebrafish has been used to evaluate environmental toxicity, and formal protocols have been established by the OECD. Exploratory research for evaluating toxicity using many classes of compounds, from environmental toxins to therapeutics, shows that, in general, responses in zebrafish are similar to those in mammals. All of these factors indicate that this model organism will become a valuable tool for assessing environmental toxicity and for predicting drug toxicity.

LITERATURE CITED


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Dawson, D. 1991. Additive incidence of development malformation for Xenopus embryos exposed to a mixture of ten aliphatic carboxylic acids. Teratology 44:531-546.

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Blechinger, S.R., Warren, J.T., Kuwada, J.Y., and Krone, P.H. 2002. Developmental toxicology of cadmium in living embryos of a stable transgenic zebrafish line. Environ. Health Perspect. 110:1041-1046.

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Burkhardt-Holm, P., Oulmi, Y., Schroeder, A., Storch, V., and Braunbeck, T. 1999. Toxicity of 4-chloroaniline in early stages of zebrafish (Danio rerio): II. Cytopathology and regeneration of liver and gills after prolonged exposure to waterborne 4-chloroaniline. Arch. Environ. Contam. Toxicol. 37:85-102. Carvan, M.J., Dalton, T.P., Stuart, G.W., and Nebert, D.W. 2000. Trangenic zebrafish as sentinels for aquatic pollution. Ann. N.Y. Acad. Sci. 919:133147. Chan, S.T.H. and Yenung, W.S.B. (eds.) 1983. Sex Control and Sex Reversal in Fish Under Natural Conditions. Academic Press, New York. Chan, D.W. and Yager, T.D. 1998. Preparation and imaging of nuclear spreads from cells of the zebrafish embryos: Evidence for large degradation intermediates in apoptosis. Chromosoma 107:39-60. Chatot, C., Klein, N.W, Piatek, J., and Pierro, L.J. 1980. Successful culture of rat embryos on human serum: Use in the detection of teratogens. Science 207:1471-1473. Cicurel, L. and Schmid, B. 1988. Postimplantation embryo culture for the assessment of the teratogenic potential and potency of compounds. Experientia 44:833-840. Cook, P., Zabel, E., and Peterson, R. 1997. The TCDD toxicity equivalence approach for characterizing risks for early life-stage mortality in trout. In Chemically Induced Alternations in Functional Development and Reproduction of Fishes: Proceeding from a Session at the 1995 Wingspread Conference (R. Rolland, M. Gilbertson, and R. Peterson, eds.) pp 9-27. SETAC Press, Pensacola, Fl. Creaser, C.W. 1934. The technic of handling the zebrafish (Brychyanio rerio) for the production

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Fort, D., Stover, E.L., Propst, T., Faulkner, B.C, Vollmuth, T., and Murray, F. 1998. Evaluation of the developmental toxicity of caffeine and caffeine metabolites using the frog embryo teratogenesis assay -Xenopus (FETAX). Food Chem. Toxicol. 36:591-600.

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Orn, S., Andersson, P., Forlin, L., Tysklind, M., and Norrgren, L. 1998. The impact on reproduction of an orally administrated mixture of selected PCBs in zebrafish (Danio rerio). Arch. Environ. Contam. Toxicol. 35:52-57. Page, J.G., Rogers, T.S., Fulton, R.B., Thompson, C.R., Hill, D.L., Smith, A.C., and Tomaszewski, J.E. 1998. Toxicity of COL-3 to rats and monkeys. Proc. Am. Assoc. Cancer Res. 39:596. Parng, C., Seng, W.L., Semino, C., and McGrath, P. 2002a. Zebrafish: A preclinical model for drug screening. Assay Drug Dev. Technol. 1:41-48. Parng, C., Semino, C., Zhang, C., and McGrath, P. 2002b. Gene expression profiling in zebrafish after drug treatment. Toxicologist 66:LB75. Pelissero, C., and Sumpter, J.P. 1992. Steroids and “steroids-like” substances in fish diets. Aquaculture 107:283-301. Peterson, R.T., Link, B.A., Dowling, J.E., and Schreiber, S.L. 2000. Small molecule developmental screens reveal the logic and timing of vertebrate development. Proc. Natl. Acad. Sci. U.S.A. 97:12965-12969. Ruoppa, M. and Nakari, T. 1988. The effects of pulp and paper industry waste water on the fertilized eggs and alevins of zebrafish and on the physiology of rainbow trout. Water Sci. Technol. 20:201-202. Schreurs, R., Lanser, P., Seinen, W., and Van der Burg, B. 2002. Estrogenic activity of UV filters determined by an in vitro reporter gene assay and an in vivo transgenic zebrafish assay. Arch. Toxicol. 76:257-261. Schulte, C. and Nagel, R. 1994. Test acute toxicity in the embryo of zebrafish, Brachydanio rerio, as an alternative to the acute fish test: Preliminary results. Altern. Lab. Anim. 22:12-19. Semino, C., Zhang, C., Parng, C., and McGrath, P. 2002. Zebrafish as an animal model to assess drug toxicity. Toxicologist 66:LB59.

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Stainier, D.Y.R. and Fishman, M.C. 1994. The zebrafish as a model system of cardiovascular development. Trends Cardiovasc. Med. 4:117-121. Stainier, D.Y.R., Lee, R.K., and Fishman, M.C. 1993. Cardiovascular development in the zebrafish. Development 119:31-40. Strahle, U. and Blader, P. 1994. Early neurogenesis in the zebrafish embryos. FASEB J. 8:692-698. Streisinger, G., Walker, C., Dower, N., Knauber, D., and Singer, F. 1981. Production of clones of homozygous diploid zebra fish (Brachydanio rerio). Nature 291:293-296. Stringer, B. and Blankememyer, J. 1993. Rapid measurement of toxicity using electrochromic



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dyes and frog embryos. Bull. Environ. Contam. Toxicol. 51:557-563.

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Stuart, G.W., Vielkind, J.R., McMurray, J.V., and Westerfield, M. 1990. Stable lines of transgenic zebrafish exhibit reproducible patterns of transgene expression. Development 109:577-584. Sumpter, J.P. and Jobling, S. 1995. Vitellogenesis as a biomarker for estrogenic contamination of the aquatic environment. Environ. Health Perspect. 103:173-178. Sun, Z. and Hopkins, N. 2001. vhnf1, the MODY5 and familial GCKD-associated gene, regulates regional specification of the zebrafish gut, pronephros, and hindbrain. Genes Dev. 15:32173229. Takahashi, H. 1977. Juvenile hermaphroditism in the zebrafish (Brachydanio rerio). Bull. Faculty of Fisheries Hokkaido University 28:57-65. Talbot, W.S. and Schier, A.F. 1999. Positional cloning of mutated zebrafish genes. In The Zebrafish: Genetics and Genomics (H.W.III Detrich, M. Westerfield, and L. Zon, eds.) pp 260-286, Academic Press, San Diego. Thermes, V., Grabher, C., Ristoratore, F., Bourrat, F., Choulika, A., Wittbrodt, J., and Joly, J. 2002. I-Scel meganuclease mediates highly efficient transgenesis in fish. Mech. Dev. 118:91. Van Leeuwen, C.J., Grootellar, E.M., and Niebeek, G. 1990. Fish embryos as teratogenicity screens: A comparison of embryotoxicity between fish and birds. Ecotoxicol. Environ. Saf. 20:42-52. Vitozzi, L. and De Angelis, G. 1991. A critical review of comparative acute toxicity data on freshwater fish. Aquatic Toxicol. 19:167-204. Wannemacher, R., Rebstock, A., Kulzer, E., Schrenk, D., and Bock, K. 1992. Effects of 2,3,7,8-tetrachlorodibenzo-p-dioxin on repro-

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Contributed by Chaojie Zhang and Catherine Willett Phylonix Pharmaceuticals Cambridge, Massachusetts Trisha Fremgen Northeastern University Boston, Massachusetts




Preclinical Models of Parkinson’s Disease

UNIT 1.8

Parkinson’s disease (PD) is a neurodegenerative disorder in which pigmented midbrain neurons progressively die producing a dopamine (DA) deficit in the striatum, which manifests as an akinetic movement disorder. Experimentally induced striatal DA depletion in animals is a valid model of parkinsonism (Schultz, 1982). The capacity of certain substances to damage catecholaminergic neurons has been used extensively to produce DA deficiency in animals (Ungerstedt, 1971a,b; Burns et al., 1983; Annett et al., 1992). This unit describes methods for inducing parkinsonism in nonhuman primates and rodents using the neurotoxins 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine (MPTP; Basic Protocols 1 and 4 and Alternate Protocol 1) and 6-hydroxydopamine (6-OHDA; Basic Protocols 2 and 3). Additionally, procedures for evaluating the animals are presented (Support Protocols 1 to 4). Other models are briefly reviewed in the Commentary. STRATEGIC PLANNING Model Selection: Species Rodent models of PD are good anatomical and biochemical models of the disease. However, they are not good functional models because the behavioral syndrome only superficially resembles some aspects of idiopathic PD, and the progressive nature of the disease is not reproduced. The complex behavioral repertoire of monkeys provides greater scope for determining the extent of functional recovery. In nonhuman primates, administration of MPTP can induce a stable parkinsonian syndrome that is remarkably similar to the idiopathic disease (Burns et al., 1983). All primates are susceptible to MPTP toxicity. However, marmosets show considerable motor recovery despite biochemical and histological evidence for extensive damage, so that in the chronic stage only mild akinesia and incoordination of movement remain. The choice of species for specific experiments depends on the cost, availability of the animals, the type of study proposed (some species, like squirrel monkeys, are difficult to train), and the size of the brain required (for certain neuroimaging studies, such as positron emission tomography or PET scanning, the large brain of baboons is preferable). All results and specifications concerning doses and behavior in this unit refer to macaques (Rhesus and Cynomolgus) in the MPTP models and marmosets for the 6-OHDA lesions. As in primates, MPTP and 6-OHDA are the most common toxicants used to induce DA depletion in rodents. Rats are the most commonly used experimental rodents, but they are rather insensitive to MPTP, requiring high doses and, with most strains, intracerebral administration of 1-methyl-4-phenyl-pyridinium (MPP+; the toxic metabolite of MPTP). So far, 6-OHDA lesioning continues to be the most popular parkinsonian model in rats. Lesion Different lesions yield appropriate animal models to evaluate different therapeutic strategies, such as DA replacement (cell-mediated or pharmacological), neuroprotection, and neuroregeneration, as well as physiopathological mechanisms of neurodegeneration. Complete DA lesions in experimental animals are considered, at least in a neuropathological sense, analogous to end-stage PD. Data obtained from studies using this type of model are useful for the evaluation of therapeutic strategies aimed at DA replacement, as the DA-mediated positive effects will not be complicated by the potential response of remaining host DA neurons. On the other hand, experimental PD models sparing some DA terminals are analogous to less severe stages of the disease and are appropriate for investigating the potential benefit of regenerative approaches (e.g., the use of trophic


Contributed by Krys S. Bankiewicz, Rosario Sanchez-Pernaute, Yoshitsugu Oiwa, Malgorzata Kohutnicka, Alex Cummins, and Jamie Eberling

1.8.1

Current Protocols in Toxicology (2003) 1.8.1-1.8.31 Copyright © 2003 by John Wiley & Sons, Inc.

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factors) that require the presence of some residual DA neurons. In the context of tissue transplantation, the relative contributions of DA replacement or graft-induced trophic responses of a particular procedure are probably best addressed when efficacy is compared in both types of model. Toxin MPTP The capacity of MPTP to induce persistent parkinsonism in humans (Davis et al., 1979) and in nonhuman primates (Burns et al., 1983) provides the opportunity to investigate both therapeutic approaches and possible pathogenic mechanisms of PD in primate models. Three models can be induced in primates by MPTP administration: (1) the unilateral model, induced by internal carotid artery (ICA) administration; (2) the bilateral model, induced by systemic (i.v., i.m., or s.c.) administration; and (3) the combined overlesioned or bilateral asymmetric model, induced by unilateral ICA plus i.v. administration. Basic Protocol 1 describes procedures for both ICA and systemic administration. Mice exhibit characteristic neuropathological and biochemical signs of DA system damage following systemic administration of MPTP (Basic Protocol 4). Loss of tyrosine hydroxylase (TH)–positive neurons is seen in the substantia nigra pars compacta (SNc) and decreased levels of DA and its metabolites are observed in the striatum. These changes are accompanied by akinesia-depressed spontaneous motor activity (Heikkila et al., 1985; Hallman et al., 1985; Donnan et al., 1987; Sundstrom et al., 1990). Rats are very resistant to MPTP; to produce significant DA damage, the toxic metabolite MPP+ has to be directly administered into the CNS. 6-OHDA 6-Hydroxydopamine has been used extensively to produce parkinsonism in rodents; however, few studies in primates have used this toxin (Schultz, 1982; Annett et al., 1992), most likely because it requires intracerebral administration. Basic Protocols 2 and 3 describe the respective procedures for stereotactic administration of this toxin in primates and rats. NOTE: All protocols using live animals must first be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) and must follow officially approved procedures for the care and use of laboratory animals. BASIC PROTOCOL 1


COMBINED ICA AND INTRAVENOUS ADMINISTRATION OF MPTP: THE OVERLESIONED (BILATERAL ASYMMETRIC) PRIMATE MODEL In both humans and nonhuman primates, administration of MPTP produces a stable parkinsonian syndrome (Burns et al., 1983). MPP+ is the toxic metabolite of MPTP (Nicklas et al., 1985), and conversion of MPTP to MPP+ is mediated by monoamine oxidase B (MAO-B). MPP+ accumulates selectively in neurons possessing a DA uptake system (Javitch et al., 1985) and is further concentrated in mitochondria. The basis of the toxic effects of MPP+ is only partially understood; retrograde axonal transport, melanin binding, and damage to the mitochondrial respiratory system (Nicklas et al., 1985) have been implicated in the pathogenesis of injury to DA neurons. Replacement of DA with L-3,4-dihydroxyphenylalanine (L-DOPA) is the most common therapeutic approach in PD, and thus far the most efficacious. L-DOPA response is a diagnostic criterion and staging tool in PD and, as such, it is included in this unit with other outcome measures for parkinsonian primates. Evaluation of the response of MPTP-lesioned nonhuman primates to L-DOPA is also useful for validating the sensitivity of the specific neurobehavioral measures used to assess the effect of novel therapies and to compare the relative efficacy of such therapies, taking into account both positive and adverse effects. Moreover, the direct effect of different therapies on the response to L-DOPA can be investigated.

1.8.2 Supplement 18


Materials Sterile saline: 0.9% (w/v) NaCl 10-mg vial of 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine HCl (MPTP⋅HCl; Sigma) Adult macaques (Rhesus and Cynomolgus; Sierra Biomedical or Charles River) Ketamine/xylazine Isoflurane Betadine 70% (v/v) ethanol 0.1 M HCl Peroxide 10- and 60-ml sterile syringes 10- or 30-ml sterile vial Animal balance (accurate to 0.1 g) Intravenous (i.v.) extension set Electric shaver Alcohol pads 22-G i.v. catheter Tracheal tube Surgical table Absorbant blue benchpads Stretch gauze and cotton swabs Sterile surgical tools: scalpel, large forceps, delicate curved forceps, scissors, needle holders, retractors, mosquito hemostats 27-G sterile needle Infusion pump (fitted for a 60-ml syringe) 3-0 Vicryl with needle Plastic, transparent millimeter-scale ruler 22-G angiocatheter Heated water pad Drapes 2/0 silk suture Towel clamps CAUTION: The most hazardous operations in conducting MPTP animal experiments are the preparation, handling, and injection of concentrated solutions of MPTP. Always use a chemical hood and skin protection (gloves, Tyvex laboratory coat) and mask. Any spilled MPTP should be degraded by spraying with 0.1 N HCl, while excess solution should be mixed 1:1 with 6 M H2SO4 and degraded by adding 4.7 mg potassium permanganate per 100 ml. The concentration of MPTP in the solution used for ICA administration is very low (0.03 to 0.07 mg/ml) and therefore quite safe. However, solutions for i.v. administration (even small volumes) are much more concentrated, and additional caution must be taken. NOTE: Animals should be adults and should interact well with the investigators so clinical evaluation can be performed. Prepare MPTP solution 1. Fit a 10-ml sterile syringe with a 20- or 23-G needle and fill with sterile saline. 2. Infuse 1 ml sterile saline into a sterile vial containing 10 mg MPTP⋅HCl. Aspirate the 1-ml solution back into the syringe to complete the 10-ml volume (final 1 mg/ml MPTP). 3. Transfer the entire 10-ml MPTP solution into a sterile 10- or 30-ml vial.



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4. Weigh the animal (within 0.1 g) and determine the total dose, adjusting for the HCl group using a 1.2× conversion factor. Particular caution should be exercised when calculating the dose for intracarotid administration. MPTP is toxic during the infusion (i.e., first pass effect) only because it undergoes peripheral conversion to MPP+, which cannot cross the blood brain barrier. For this route of administration, MPTP should be calculated according to brain size, which tends to be constant over wide body weight ranges, instead of per body weight. The latter can give the false impression that older (heavier) animals are more susceptible to MPTP, when in fact they are receiving larger doses for similar brain sizes. As a general guideline, 2 to 2.5 mg can be used for small (3 to 5 kg) animals, 3 to 3.5 mg for a 6- to 10-kg monkey, and 4 mg for large (>10 kg) animals.

5. Fill a sterile 60-ml syringe with 2 to 4 ml MPTP solution (2 to 4 mg MPTP). Fill the rest of the syringe with sterile saline and attach an i.v. extension set. The final concentration in the syringe is 2 to 4 mg/60 ml (0.033 to 0.067 mg/ml MPTP).

Prepare the animal 6. Anesthetize a macaque monkey with ketamine (10 mg/kg) and xylazine (1 mg/kg) in the home cage. Transfer to the procedure room. 7. Shave the calves and neck region of the animal using an electric shaver. Palpate the calf muscle and identify the saphenous vein. 8. Clean the skin with an alcohol pad. Using a 22-G i.v. catheter, cannulate the vessel and flush with saline. 9. Intubate the animal with a 3.5- to 5-mm (inside diameter) tracheal tube, according to the size of the animal. 10. Transfer the animal to a surgery room, place it on a surgical table with absorbant blue benchpads, and a heated water pad at 37°C. Maintain isoflurane anesthesia at 1.5% with an oxygen flow of ∼0.3 liters/min. 11. Hyperextend the head on the surgical table by placing stretch gauze through the canines and applying gentle retraction. Secure the head in position by tying the gauze to the surgery table. 12. Cover the animal completely with a series of sterile drapes. Cut out an opening in the drape for the neck incision and secure drapes with towel clamps. Scrub the surgery site with Betadine followed by 70% ethanol. 13. Make a midline incision through the skin of the neck with a sterile scalpel. 14. Using blunt dissection technique, locate and open the carotid sheath, exposing the common carotid artery, internal jugular vein, and vagus nerve (Fig. 1.8.1). Isolate the common carotid artery below the carotid bifurcation. 15. Locate the superior thyroid artery and the external carotid artery and temporarily clamp the vessels using mosquito hemostats.


Typically, both superior thyroid and external carotid arteries form a very short branch medially to the carotid bifurcation. Hemostats should be clamped on this branch, if possible. Another alternative is to clamp both the thyroid superior and external carotid (below the external maxillary branch) arteries using single hemostats distal to the carotid bifurcation. Use only mosquito hemostats with an adjusted clipping mechanism formed by bending handles inside.

1.8.4 Supplement 18


internal maxillary artery external maxillary artery

XII (hypoglossal nerve) X (vagus nerve)

superior thyroid artery

clamp location

ICA

*

ECA

CCA JV

Figure 1.8.1 ICA dissection. The common carotid artery (CCA) at the level of the bifurcation showing the internal carotid artery (ICA), the X and XII cranial nerves, the external carotid artery (ECA), and its first branches: the superior thyroid artery and the external maxillary artery. An asterisk marks the recommended location of the vascular clamp, in the ECA proximal to the superior thyroid artery origin. JV, jugular vein.

16. Attach a 27-G needle to the i.v. extension tubing and drain all the air from the needle by activating an infusion pump for several seconds. Drain all excess MPTP onto a sponge soaked with 0.1 M HCl solution. Perform ICA infusion 17. Insert the 27-G needle (with attached extension set and 60-ml MPTP syringe) into the internal carotid artery in a direction retrograde to the direction of blood flow. 18. Program the infusion pump to deliver 4 ml/min and infuse the entire volume from the syringe (2 to 4 mg MPTP). Toxicological Models


Supplement 18

19. Remove the vascular clamps (or mosquito hemostat) from the superior thyroid and external carotid arteries. Withdraw the needle from the common carotid artery and apply pressure for 5 min (until bleeding has stopped). 20. Clean the wound with peroxide, close the incision site at the neck with 3-0 Vicryl, and clean the sutured area. 21. Monitor the recovery of the animal and return it to the animal room. Record ipsilateral pupil mydriasis (dilation) by measuring the diameter of both pupils with a plastic transparent millimeter-scale ruler. In most cases, ipsilateral pupil mydriasis of 3 to 5 mm follows ICA infusion of MPTP solution. Absence of mydriasis does not indicate that lesion has not been successful, but its presence is a good indicator of unilateral damage. CAUTION: Whenever possible, keep the animals in a quarantine room for 48 to 72 hr. Animal excreta may contain considerable amounts of unmetabolized MPTP; thus, precautions should be taken to minimize skin contact with animal fur and excreta and to avoid inhalation of bedding dust from MPTP-treated animals. As a general precaution, 0.1 N HCl solution should be used to spray the bedding in monkey cages during the first 48 hr after MPTP administration, and the cages should then be washed with an acidic solution. This ensures that any free MPTP is converted to its nontoxic form (for safety concerns see Yang et al., 1988).

22. House animals for 2 weeks before i.v. administration. Administer systemic (i.v.) MPTP 23. Weigh the animal (within 0.1 g) and determine the total dose of MPTP required, adjusting for the HCl group using a 1.2× conversion factor. For i.v. administration, the recommended dose is 0.3 mg/kg body weight.

24. Anesthetize the animal with ketamine (10 mg/kg) and xylazine (1 mg/kg) in the home cage. Transfer the monkey to the procedure room. 25. Shave the calves using an electric shaver. Palpate the calf muscle and identify the saphenous vein. Clean the vein with an alcohol pad. 26. Using a 22-G angiocatheter, cannulate the vessel and flush with sterile saline. 27. Fill a sterile 3- to 6-ml syringe with the appropriate volume of 1 mg/ml MPTP solution (0.3 ml/kg or 0.3 mg/kg) and infuse the MPTP into the vein over 3 min. 28. Flush the line with 3 ml sterile saline. 29. Remove the needle and apply pressure to the saphenous vein until bleeding stops. All safety guidelines described for ICA administration should be followed (see annotation to step 21).

Evaluate motor behavior 30. One week after i.v. administration of MPTP, assess the syndrome using a motor rating scale for primates (Table 1.8.1) based on the unified Parkinson’s disease rating scale (UPDRS). Monitor response to L-DOPA (see Support Protocol 1) and activity (see Support Protocol 2).


1.8.6 Supplement 18


Table 1.8.1

Rating Scale for Parkinsonian Primatesa

Parameter

Score

Degree

Tremor (right arm/left arm)b

0 1 2 3 0 1 2 3 0 1 2 3 0

3 0 1 2 3 0 1 2 3 0 1 2 3 0

Absent Occasional or barely detectable (normal for aged), occurring while active Frequent or easily detectable, occurring while active or at rest Continuous or intense, occurring while active and at rest Absent Occasional episodes of short duration (10 sec) Uses all four limbs smoothly and symmetrically Walks slowly (normal for aged), noticeable limp Walks very slowly and with effort, may drag limb or refuse to bear weight Unable to ambulate Normal function: able to grasp/retrieve small objects, aims accurately, independent use Reduced ability in grasping/retrieving small objects, independent use, may have reduced aim Able to grasp small objects rarely, only with assistance, or with great difficulty Unable to grasp/retrieve small objects Quick, precise movements Mild slowing of movements (normal for aged) Slow deliberate movements with marked impairment initiating movements No movements Moves freely, alert, responsive Reduced activity (normal for aged), moves less frequently (without provocation) Minimal activity, moves with provocation, may have reduced facial expression Akinetic (essentially no movements) Requires no assistance for maintaining posture Requires assistance for standing Requires assistance for walking, or falls Face down, or unable to maintain posture Normal posture, stands erect Reduced posture (normal for aged), stands with feet apart, knees flexed Stooped posture, hunched, legs bent Unable to maintain posture, recumbent Immediate, robust response to provocation Slightly diminished or delayed response, open mouth threat Minimal response or longer delay, without open mouth threat No response to provocation Normal function, able to grasp/retrieve large objects accurately

1 2 3

Reduced ability/frequency of grasping/retrieving large objects Great difficulty in grasping/retrieving large objects, rarely used Unable to grasp/retrieve large objects

Freezing

Locomotion

Fine motor skills (right arm/left arm)b

1 2 3 Bradykinesia (right arm/left arm)b 0 1 2

Hypokinesia

3 0 1 2

Balance

Posture

Startle response

Gross motor skills (right arm/left arm)b

aAccording to mean scores in the scale, animals are classified into five stages. Stage 0: A maximum of 5 points is considered normal according to the

authors’ results in healthy animals. Stage 1: Total of 5-12 points; hemiparkinsonian monkeys do not show axial impairment. Stage 2: Total 12-20 points; mild to moderate bilateral symptoms. Stage 3: Total 21-30 points; moderate to severe bilateral symptoms, but without major systemic consequence. Stage 4: Total >30 points; severely damaged and can be difficult to manage due to feeding difficulties and complications of akinesia; may require DA replacement. bScore each arm separately and add the total.


Supplement 18


SYSTEMIC MPTP LESION IN PRIMATES The principal advantage of this approach is that the behavioral syndrome closely resembles that of Parkinson’s disease in humans. The main drawbacks are that it takes a long time to produce animals with stable and uniform lesions and, if the lesion is too extensive, it produces a considerable mortality. Total dose is not predetermined, but will depend on the highly variable response of each animal to the toxin. Therefore, careful clinical evaluation is mandatory throughout the lesion progression. The toxic effect is cumulative. Additional Materials (also see Basic Protocol 1) 3-ml syringes equipped with 26-G needles Cage with back-squeezing mechanism 1. Prepare MPTP as described (see Basic Protocol 1, steps 1 to 3). 2. Weigh the animal and determine the appropriate dose (0.33 to 1 mg/kg), adjusting for the HCl group with a 1.2× conversion factor. 3. Fill a sterile syringe with the appropriate volume of MPTP solution. 4. With the monkey in a cage with a back-squeezing mechanism, squeeze the back of the cage to hold the animal and inject the MPTP intramuscularly or subcutaneously into the thigh or arm. For better results, inject twice a week. Follow safety guidelines described for ICA administration (see Basic Protocol 1, step 21). If the animal shows general symptoms such as feeding difficulty or profound akinesia, stop the injections and reevaluate the motor syndrome (see Table 1.8.1) 2 to 4 weeks later.

5. Assess the syndrome as described (see Basic Protocol 1, step 30). BASIC PROTOCOL 2

UNILATERAL 6-OHDA LESION IN PRIMATES 6-OHDA uses the catecholamine uptake system to enter catecholamine neurons and kill the cells through oxidative mechanisms. Unilateral injection of 6-OHDA into the medial forebrain bundle of marmosets produces a severe loss of tyrosine hydroxylase-immunoreactive neurons in the ipsilateral SNc and DA depletion of >90% in the dorsal striatum, accumbens, and frontal cortex ipsilateral to the lesion (Annett et al., 1992). Levels of 5-hydroxytryptamine and noradrenaline (NA) are also decreased, but previous administration of NA uptake blockers (e.g., desipramine) limits the non-DA damage. The animals show ipsilateral spontaneous rotation, contralateral apomorphine-induced rotation, reduced spontaneous activity, contralateral sensorimotor neglect, and ipsilateral hand preference with variable impairment of hand skill (Annett et al., 1992). Persistent deficits are observed in animals with >95% DA depletion. Materials Sterile saline: 0.9% (w/v) NaCl Adult marmosets (Sierra Biomedical or Charles River) Ketamine/xylazine Isoflurane Betadine 70% (v/v) ethanol 4 mg/ml 6-hydroxydopamine (6-OHDA), HBr (Sigma) in 0.01% (w/v) ascorbate/0.9% (w/v) NaCl (protect from light)


Stereotaxic frame/tower (David Kopf Instruments) Manipulator arm Spinal needle

1.8.8 Supplement 18


5-ml syringes Intravenous (i.v.) line with 3-way stopcock 22-G angiocatheter Animal balance Electric razor Alcohol pads Surgical tape Tracheal tube Isoflurane inhalation chamber Sterile drapes, gauze, and rubber bands Towel clamps Tissue forceps Scissors Electric cauterizer Water heating pad Dremel drill with carbide bur excavating tip 10-µl Hamilton syringes and needle 3-0 Vicryl 2-0 silk sutures NOTE: Animals should be adults and should interact well with the investigators so clinical evaluation can be performed. Calibrate stereotactic frame 1. Fix the settings on the ear bars of a stereotactic frame so the bars are ∼1 mm apart. 2. Fit the manipulator arm with a 20-G spinal needle and position it on the stereotactic frame. Ensure that the mark readings on both ear bars are identical. 3. Advance the spinal needle into the space between the ear bars. Adjust the position of the needle along the anterior/posterior (A/P) scale to correlate with the plane of the ear bars. Prepare animal 4. Flush an i.v. line with sterile saline using a 5-ml syringe, then prime a 22-G angiocatheter and 3-way stopcock with saline. 5. In the animal room, weigh a marmoset and anesthetize with 10 mg/kg ketamine/1 mg/kg xylazine. Transfer animal to the surgery room. 6. Shave the calves, arms, and head for surgical and intravenous access using an electric razor. 7. Palpate the calf muscle or flex and extend the ankle to visualize the saphenous vein. Clean the shaved area with an alcohol pad. 8. Insert the 22-G angiocatheter into the saphenous vein and flush with saline to ensure patency. 9. Attach the 3-way stopcock with i.v. line to the catheter and start a slow saline drip (1 ml/min) to maintain patency of the vessel. Secure the i.v. line onto the calf muscle using surgical tape. 10. Intubate the animal with an appropriate size tracheal tube and maintain on isoflurane anesthesia by inhalation. Toxicological Models


Supplement 18

Position animal in stereotactic frame 11. Place the animal in the stereotactic frame and tighten the right ear bar to the frame. Elevate the animal’s head and insert the right ear bar into the ear canal. 12. Position the left ear bar in the animal’s left ear. Ensure that the animal’s eyes are parallel to the front plane of the frame. 13. Center the animal’s head such that the same setting for both ear bars is achieved. Position the eye bars in the inferior border of the orbits. Set the incisor bar and readjust the eyebars to the inferior orbital rim. Fix the incisor bar at an adequate height. 14. Clean the entire surface of the cranium by scrubbing in an outward circular motion with Betadine scrub followed by 70% ethanol. Spray the stereotactic frame with 70% ethanol. Use sterile gloves for cranium preparation. 15. Cover the animal completely with a heating pad and then cover it with a series of sterile drapes. Cut out an opening in the drape for the craniotomy and secure drapes with towel clamps. Administer 6-OHDA 16. Determine the target skin incision site using the baseline A/P coordinates as a guideline. Make a small sagittal incision through the skin and fascia using an electric cauterizer. 17. Using sterile gauze, retract the skin and fascia to expose the cranial surface. Secure the skin in place with towel clamps. 18. Cover a Dremel drill with sterile drapes and fix in place with sterile rubber bands. Use a carbide bur excavating tip and make a burr hole that exposes the dura. 19. Position the spinal needle over the first target site and manually advance the needle to the surface of the cranium. For marmosets, use the following coordinates with reference to stereotactic zero (Annett et al., 1992), with all five injection sites at A/P +6.5: one lateral (3 µl): lateral (L) ±3.2, ventral (V) 7.5 two central (2 µl each): L ±2.2, V 6.5 and V 7 two medial (2 µl each): L ±1.2, V 6 and V 7. 20. Insert the needle at the first target site, then touch the electric cauterizer to the needle and withdraw manually. 21. Place a 10-µl Hamilton syringe on the arm holder and infuse the selected volume of 4 mg/ml 6-OHDA at a rate of 0.5 µl/min. Leave the syringe in place for 5 min to avoid overflow. 22. Repeat steps 19 to 21 at the remaining four injection sites. 23. Suture the fascia with sterile 3-0 Vicryl. Close the incision site by subcutaneous suturing of the skin with 2-0 silk sutures. Clean and dry the incision site with sterile gauze and spray with Betadine.


Evaluate motor behavior 24. Evaluate nigral damage in vivo and post mortem using the methods described for the MPTP model (see Support Protocol 1 and Support Protocol 2). Also evaluate rotational behavior (see Support Protocol 3).



EVALUATION OF CHANGES IN MOTOR BEHAVIOR IN RESPONSE TO L-DOPA

SUPPORT PROTOCOL 1

Once the animals show a stable deficit (typically ∼6 weeks), it is possible to evaluate the changes in motor behavior following an intramuscular injection of L-DOPA methyl ester (M-L-DOPA) with a peripheral inhibitor of L-aromatic amino acid decarboxylase. Dosing with Sinemet preparation does not provide predictable results due to inadequate control of intake and stomach content. M-L-DOPA injection should be used to ensure adequate brain delivery, as estimation of the effective dose is highly inaccurate with oral administration. Local complications are not observed using intramuscular (i.m.) injection. The effect of L-DOPA can be evaluated using the activity or motor tasks shown in Table 1.8.1. For each animal, the response to saline injection should be used as control.

A M:8.4 S:515

BASELINE CRS:15 M:14.4 S:877

M:13.5 S:826

M:20.8 S:1267 M:18.3 S:567 CRS: 15

M:20.5 S:1251

M:16.8 S:1026

M:20.7 S:1285

M:11.2 S:684

M:13.7 S:837

M:6.2 S:379

M:31.6 S:1930

M:11.6 S:706

M:16.3 S:505 CRS: 15.5

max:103

B

M:44.3 S:2705

M:221 S:1346

BASELINE CRS:13.5 M:21.4 S:1303

max:192

M:14.6 S:888

BASELINE CRS:17 M:18.1 S:1103

max:186

M:21.6 S:1315

BASELINE CRS:14.8 M:26.2 S:1599

C

D

–120

–60

M:8.1 S:493 M:9.9 S:308 CRS: 14.3

M:15.4 S:476 CRS: 13.3

M:71.0 S:4334

M:19.3 S:1176

M:52.8 CRS: S:1638 4.5

M:68.1 S:4155

M:39.1 S:2388

M:40.4 S:1253 CRS: 9.8

0

M:14.7 S:456 CRS: 12.5

60

M:38.2 S:1183 CRS: 9.3

120

180

240

Time (min)

Figure 1.8.2 Activity and clinical rating scale (CRS) outcome of administration of increasing doses of M-L-DOPA in an overlesioned monkey. The drug was administered at time zero and activity is shown from 120 min before to 240 min following administration of (A) 5 mg, (B) 10 mg, (C) 20 mg, and (D) 40 mg M-L-DOPA. Mean and raw activity data for each segment is shown along with clinical score. During this experiment a shorter version of the scale was used; therefore, clinical score values are smaller than would be recorded using the scale as shown in Table 1.8.1. Increase of drug-induced activity is associated with clinical improvement in a dose-responsive manner. In panels, M, mean counts; S, sum of counts.



Supplement 18

Materials Lesioned animal (see Basic Protocol 1, Alternate Protocol 1, and Basic Protocol 2) L-3,4-Dihydroxyphenylalanine methyl ester (M-L-DOPA; Sigma) Benserazide (Sigma) Sterile saline: 0.9% (w/v) NaCl Cage with back-squeezing mechanism 3-ml syringes 26- to 30-G needles 1. Prepare the total dose of M-L-DOPA and add benserazide at a 1:10 benserazide/ML-DOPA ratio. Mix and dilute in 1 to 2 ml sterile saline. Appropriate doses might vary for different stages and between species, but typical doses are 5 to 50 mg/kg.

2. With a lesioned monkey in a cage with a back-squeezing mechanism, squeeze the cage and inject i.m. into the thigh of the animal using a 3-ml syringe with a 26- to 30-G needle. It is advisable to keep the monkeys in cages with back-squeezing mechanisms so they can easily be held for i.m. injections, removal of monitors, or other quick procedures.

3. Rate the response according to the parameters described in Table 1.8.1. Evaluate the response 45 to 60 min after injection and at later time points when duration of the response is relevant. It is useful to videotape the animals, particularly when they display abnormal movements that might be difficult to categorize. Figure 1.8.2 shows typical results of the procedure. SUPPORT PROTOCOL 2

MONITORING ACTIVITY TO ASSESS MPTP-TREATED MONKEYS Whole-body activity measurement is a useful indicator of DA lesion in MPTP-treated monkeys and correlates with the global motor score (Fig. 1.8.3). This objective measure approximates akinesia, which is most pronounced in animals with moderate to severe lesions. The baseline motor activity is reduced and the normal pattern disappears after the MPTP lesion. An example of the striking MPTP effect on activity is shown in Figure 1.8.4. The personal activity monitors (PAM) contain a biaxial piezoelectric sensor that is calibrated to detect threshold activities >0.024 × g acceleration. Sensitivity and epoch length is programmable. The acceleration signal is sampled and digitally integrated to quantify all activity under the signal curve. The information is converted to a reference scale of data counts or acceleration units (G). The PAM monitor can be inserted into a vest that the animal wears during testing or into a collar around the neck. Activity data are normally acquired over four to five days that should include a weekend. Materials Lesioned animal (see Basic Protocol 1, Alternate Protocol 1, and Basic Protocol 2) Ketamine/xylazine


Personal activity monitors (PAM; ActiTrac, Individual Monitoring Systems) PAM connector cable Computer Nylon cable ties Vetwrap Nylon collars/vest



Whole-body activity (mean counts)

70 r = – 0.68

60 50 40 30 20 10 0 10

12

14

16

18

20

22

24

CRS score (mean)

Figure 1.8.3 Correlation between clinical rating scale and whole-body activity in overlesioned hemiparkinsonian monkeys. Increased parkinsonian disability correlates well with a decrease in whole-body activity. Data from 28 overlesioned monkeys in stages 1, 2, and 3 are shown.

A

Baseline MEAN ACTIVITY (6AM-6PM) 28.4

Sunday

B

Monday

Tuesday

Wednesday

Sunday

Monday

14 WEEKS POST MPTP MEAN ACTIVITY (6AM-6PM) 1.7

Friday

Saturday

Figure 1.8.4 Whole-body activity measured by activity monitors (A) before and (B) 14 weeks after MPTP administration. Activity was recorded over 3 days.



Supplement 18

1. Connect a PAM to a computer. Check that the battery level for the PAM is >5.25 volts, and program the sensitivity, epoch length, and start time and date. Enter the animal and test data. ActiTrac has a battery life of 4 years and memory allows up to 62 days of continuous recording. An epoch time of 1 min provides adequate information for time periods >60 min; to detect short-lived changes, use 30 sec.

2. Place the PAM in a horizontal position such that the PAM label faces outward. Secure the PAM to the collar using nylon cable ties. Wrap the monitor and cable ties with Vetwrap. 3. Anesthetize a lesioned animal in its home cage with 7 mg/kg ketamine and 1 mg/kg xylazine. 4. Place a nylon collar around the animal’s neck and then wrap the buckle with Vetwrap. 5. Record the animal’s identification number, the date and time, and the PAM identification number. 6. Begin activity data acquisition 60 min after administration of anesthesia. As long as data analysis is done comparing similar time periods (excluding setup and removal), the exact delay between anesthesia and activity monitoring is not critical.

7. Anesthetize the animal as in step 3 and remove the PAM. 8. Connect the PAM to the interface and download information to a data file. 9. Extract 12-hr segments (e.g., 6:00 am to 6:00 pm) and summarize them in a table (Fig. 1.8.4). Depending upon the study, activity is normally evaluated at the following time points: prelesion baseline, postlesion baseline, and postexperimental treatment. Mean and total counts can be used to compare animals but, as there is considerable variability between subjects, it is better to compare relative values (e.g., percentage with respect to baseline). SUPPORT PROTOCOL 3

ROTATIONAL BEHAVIOR AS A MEASURE OF UNILATERAL NIGROSTRIATAL LESIONS In animals with a unilateral DA lesion (see Basic Protocol 2), there is an imbalance of motor activity such that they usually display spontaneous turning toward the side of the lesion. Administration of indirect DA agonists (e.g., DA-releasing drugs such as D-amphetamine) increases the imbalance and the ipsilateral rotation. Administration of direct DA agonists (e.g., apomorphine) evokes contralateral turning, which is considered to be the result of denervation hypersensitivity of DA receptors in the lesioned side. Rotational behavior in response to DA agonists grossly correlates with the severity of the lesion (it is, in fact, better correlated with asymmetry indices). Quantification of the rotational response can be accomplished in monkeys by videotaping the animals and counting the turns, and in rats by using specific devices called rotometers (Ungerstedt, 1971a,b). Suggested compounds and doses include 0.5 to 5 mg/kg D-amphetamine sulfate (Sigma) in saline and 0.025 to 0.25 mg/kg apomorphine⋅HCl (Sigma) in saline. Avoid amphetamine whenever possible, particularly if tests have to be repeated, as it might increase the mortality rate. For either compound, use the lower end of the range for primates (primates are susceptible to compound effects), the higher end for rats.




6-OHDA LESIONS IN RATS In rats, several models can be induced by unilateral intracerebral stereotactic injection of 6-OHDA into different brain structures. Bilateral application of 6-OHDA is rarely used because of high mortality due to diencephalic damage (adipsia and aphagia); hence, only unilateral models are discussed here.

BASIC PROTOCOL 3

Complete DA lesion can be induced by unilateral injection of 6-OHDA in the medial forebrain bundle or in the SNc (Ungerstedt and Arbuthnott, 1970; Ungerstedt, 1971a). These animals demonstrate a characteristic asymmetric motor behavior in response to antiparkinsonian drugs (Ungerstedt, 1971b) that enables distinction between drugs with predominantly DA receptor agonist activity from those with predominantly DA-releasing activity. Systemic administration of L-DOPA or direct DA receptor agonists leads to contralateral rotation (towards the undamaged side), while administration of DA-releasing substances (amphetamine, amantadine) leads to ipsilateral rotation (in the direction of the damaged side). This model is also useful in studies of DA replacement therapy and neuroprotection factors. Partial lesion models are induced by injection of 6-OHDA in the medial forebrain bundle (MFB) or SNc in smaller doses that leave a number of DA neurons intact (Zigmond and Strickler, 1989). These are useful models for the study of pathophysiological mechanisms and neuroregeneration. The selective A-9 lesion model can be induced by injection of 6-OHDA in the SNc, leaving the ventral tegmental area (VTA or A-10 region) neurons intact (Perese et al., 1989; Thomas et al., 1994), thus reproducing the selective vulnerability of DA neurons in the SNc observed in idiopathic PD. This model is useful to study the contribution by other DA areas to the restoration of the nigrostriatal pathway and the differential effects of protective and repair mechanisms on different DA nuclei. The striatal lesion model is induced by injections of 6-OHDA into the striatum, causing progressive retrograde degenerative changes of the corresponding DA neurons in the SNc (Sauer and Oertel, 1994; Lee et al., 1996). This is a useful model for pathophysiological, neuroregenerative, and neuroprotective studies. Materials Sprague-Dawley rats, 200 to 250 g Isoflurane Betadine 70% (w/v) ethanol 4 mg/ml 6-hydroxydopamine (6-OHDA, HBr; Sigma) in 0.01% (w/v) ascorbate/saline (protect from light) Sterile saline: 0.9% (w/v) NaCl Animal balance (accurate to 0.1 g) Isoflurane inhalation chamber Electric razor Stereotactic frame Scalpel Tissue forceps Scissors 10-µl Hamilton syringes and needles Dental drill Sutures or staples Toxicological Models


Supplement 18

Prepare animal 1. Weigh a Sprague-Dawley rat (within 0.1 g) and place in an isoflurane chamber until deeply anesthetized. 2. Position the animal in a stereotactic frame and fix the plastic tube connected to the anesthesia machine to the nose of the animal using surgical tape (make sure that the snout bar does not collapse the tube). Maintain isoflurane at ~1.5% with an oxygen flow of 2 to 3 liters/min. Alternative anesthetics such as ketamine/xylazine or chloral hydrate may also be used.

3. Shave the head with an electric razor. Clean the skin with Betadine and 70% ethanol. 4. Perform a midline incision with a scalpel and identify the bregma at the intersection of the coronal and the sagittal sutures. Administer 6-OHDA 5. Fill a 10-µl Hamilton syringe with the appropriate 6-OHDA solution. Attach syringe to the holder on the stereotactic frame. Adjust 6-OHDA solutions according to the region of injection: For medial forebrain bundle lesion: 2 µg/µl 6-OHDA in saline containing 0.1% (w/v) ascorbic acid For SNc lesion: 4 µg/µl 6-OHDA in saline containing 0.02% (w/v) ascorbic acid For striatal lesion: 0.4 µg/µl 6-OHDA in saline containing 0.1% (w/v) ascorbic acid For A-9 selective lesion: 2 µg/µl 6-OHDA in saline containing 0.02% (w/v) ascorbic acid. 6. Calculate the stereotactic coordinates for injection (Paxinos and Watson, 1986). Some examples of possible injection sites include: For medial forebrain bundle lesion: Anteroposterior (A/P) −2.2 mm, mediolateral (M/L) 1.5 mm with reference to bregma; ventrodorsal (V/D) −8.0 mm with reference to dura For SNc: A/P −5.4 mm, M/L 2.2 mm with reference to bregma, V/D −7.5 mm with reference to dura For striatal lesion: A/P +0.5 mm, M/L 2.8 mm with reference to bregma, V/D −4.5 mm with reference to dura For A-9 selective lesion: 1st lesion: A/P +3.5 mm, M/L 1.9 mm, V/D −7.1 mm with reference to lambda and dura with needle bevel directed rostrally 2nd lesion: A/P +3.5 mm, M/L 2.3 mm, V/D −6.8 mm with reference to lambda and dura. 7. Adjust the incisor bar in the animal until the heights of lambda and bregma skull points are equal. 8. Drill a burr hole at the target site using a dental drill. 9. Lower the needle of the Hamilton syringe through the dura down to the selected depth and start injection at a rate of 0.5 to 1 µl/min. Adjust volumes (and doses) of 6-OHDA according to the region of injection: Preclinical Models of Parkinson’s Disease

For medial forebrain bundle lesion: 4 µl solution (8 µg 6-OHDA) For SNc lesion: 2 µl solution (8 µg 6-OHDA) For striatal lesion: 20 µl solution (8 µg 6-OHDA) For A-9 selective lesion: 2 µl solution (4 µg 6-OHDA).



10. Leave the needle in place for 5 min and withdraw slowly (1 mm/min). 11. Close scalp margins with sutures or staples. Remove the animal from the stereotactic frame and place it in its home cage. Put food on the floor of the cage and monitor the animal’s weight for 3 days after surgery. Supplement the diet (e.g., with fruit) if there is >10% weight loss. Evaluate behavior 12. Evaluate rotational behavior (see Support Protocol 3). MPTP LESION IN MICE This model is induced by systemic administration of MPTP, which causes transient impairment of DA system function in these animals. It is a simple and useful model for studies on the pathophysiology of neurodegenerative processes as well as the effects of neurotrophic and neuroprotective agents.

BASIC PROTOCOL 4

Many different protocols have been used with various doses, routes of administration, and species. An example of the model induced by intraperitoneal administration of MPTP to C57Black mice, using four injections of 10 mg/kg (total 40 mg/kg), is described (Ricaurte et al., 1987). Materials C57Black mouse, age 8 to 12 months 1-Methyl-4-phenyl-1,2,3,6-tetrahydropyridine HCl (MPTP⋅HCl; Sigma) Isotonic saline Animal balance (accurate to 0.1 g) 1-ml syringe equipped with 26-G needle 1. Weigh a C57Black mouse (within 0.1 g). 2. Prepare a 1 mg/ml MPTP solution in isotonic saline. 3. Fill a 1-ml syringe, equipped with a 26-G needle, with MPTP solution to provide a dose of 10 mg/kg (e.g., 0.4 ml for 40-g mouse). 4. Hold mouse in dorsal recumbency with left leg immobilized. Insert the needle in the lateral aspect of the lower left abdominal quadrant through the skin and musculature, and immediately lift the needle against the abdominal wall and inject the solution. 5. Repeat the injection three times at 1-hr intervals. 6. Observe animals for any changes in general locomotion, stooped posture and piloerection (see Support Protocol 4). MONITORING ACTIVITY IN MPTP-TREATED MICE Open-field locomotor activity can be measured to evaluate the toxic effect of MPTP in mice and correlates with the content of dopamine in the striatum and nucleus accumbens (Leroux-Nicollet and Costentin, 1986). However, MPTP induces an acute increase in activity in some mouse strains, as the predominant effect in the early stage is inhibition of DA reuptake by MPTP. This effect can be blocked with neuroleptics. In C57 Bl/6 mice, MPTP produces a decrease in baseline activity (∼60% in albino and 40% in black mice) that is prevented by administration of monoamine oxidase inhibitors. Amphetamine induces an increase in locomotor activity in normal animals that is absent in MPTP-lesioned animals. Apomorphine induces a decrease in locomotor activity that is not significantly different between normal and MPTP-injected animals.

SUPPORT PROTOCOL 4



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Measurement of locomotor activity can range from simple observation to sophisticated automated procedures. In general, these instruments use an array of infrared photobeams and reveal the activity of the animal by the number and pattern of beam interruptions. The Digiscan Animal Activity Monitor (Omnitech Electronics) consists basically of a Plexiglas cage with infrared monitoring sensors at determined distances. A Digiscan analyzer collects data and transmits them to a computer for storage and further use. Several variables can be recorded, such as horizontal activity, movement time, total distance, average speed, average distance per horizontal movement, and number of movements. Materials MPTP-lesioned mouse (see Basic Protocol 4) Dexamphetamine sulfate Saline: 0.9% (w/v) NaCl Automated open-field instrument (e.g., Digiscan Animal Activity Monitor; Omnitech Electronics) 1-ml syringe with 26- to 30-G needle 1. Place an MPTP-lesioned mouse in a Digiscan cage and record activity for the selected period of time. It is mandatory to perform pre-exposure habituation to get a reliable baseline.

2. Prepare dexamphetamine sulfate at 0.5 mg/ml in saline and place in a 1-ml syringe with a 26- to 30-G needle. 3. Weigh mouse and inject intraperitoneally at 1.5 mg/kg. 4. Place the animal in the Digiscan cage and record data for ≥30 min. It is important to perform all experiments at the same time of day as activity varies with circadian cycles.

COMMENTARY Background Information


Evaluation of DA deficit in primates Many different methods have been used to rate motor impairment induced in primates by MPTP and 6-OHDA. Rotational behavior and reach-and-grasp timed tasks are usually employed to quantify motor deficits in unilateral models. Marked delay in initiating movements on the side contralateral to ICA MPTP administration manifests clinically as “neglect” but, unlike cortical neglect, the animals respond when stimuli are maintained long enough, showing that the apparent inattention is really a profound hypokinesia. In these animals, this characteristic parkinsonian sign can be measured independently. In most studies in which the functional capabilities of unilaterally lesioned monkeys have been examined, there are obvious deficits in gross motor behavior evident upon direct observation or detected by the use of automated monitoring devices or measurement of volun-

tary reaching movements. However, these measures provide little information about the cognitive abilities of the animal. Delayed responses and delayed alternation tests, which are measures sensitive to disruption of the frontostriatal system, are altered in monkeys treated with MPTP. These changes occur in monkeys with measurable motor deficits and also in animals treated chronically with low doses of MPTP that retain normal motor function by clinical assessment (Schneider and Kovelowski, 1990). Neuroimaging techniques such as positron emission tomography (PET), single photon emission tomography (SPECT), and functional magnetic resonance imaging (MRI) are powerful methods that allow in vivo evaluation of DA function in parkinsonian monkeys. However, selecting the most appropriate approach depends largely on available resources. PET and the DA metabolism tracer [18F]-6-fluoro-LDOPA (FDOPA) have been used extensively to evaluate DA function in the striatum of normal



SPECT β-CIT

PET FMT

stage 1

stage 2

stage 3-4

Figure 1.8.5 Neuroimaging representation of mild (stage 1), moderate (stage 2), and severe (stage 3-4) MPTP-induced DA lesion in overlesioned monkeys. Note that the side ipsilateral to the intracarotid infusion of MPTP is severely depleted at all stages. The contralateral side shows a progressive DA depletion on both (top row) SPECT β-CIT [2β-carbomethoxy-3β-(4-iodophenyl)tropane, a cocaine analogue with a high affinity for the dopamine transporter] and (bottom row) PET FMT images as the animals display more severe signs of PD. This black-and-white facsimile of the figure is intended only as a placeholder; for full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm.

and MPTP-treated monkeys (Doudet et al., 1989; Pate et al., 1993) but not in midbrain structures due to limitations in both tomograph resolution and quantification of FDOPA metabolism. The use of the DA tracer [18F]-6-fluoroL-m-tyrosine (FMT; Melega et al., 1989) provides a high level of image contrast and a high signal-to-noise ratio, reflecting its more restricted metabolism in both peripheral and central tissues, and permitting the evaluation of DA activity in the SNc and striatum (Eberling et al., 1997, 1998; Jordan et al., 1997). For examples of SPECT and PET images, see Figure 1.8.5, Figure 1.8.6, and Figure 1.8.7. 123Iodine-2β-carbomethoxy-3β-(4-iodoph enyl) tropane ([123I]β-CIT) has been used to measure DA transporter activity in both human and nonhuman primates using SPECT (Laruelle et al., 1993). Although [123I]β-CIT has high affinity for both DA and serotonin transporters, the high density of DA transporters in the

striatum makes striatal tracer uptake a relatively specific indicator of striatal DA transporter density (Laruelle et al., 1993). Postmortem studies have shown a reduction in DA transporter density in the striatum of PD patients that parallels the loss of DA, and SPECT [123I]βCIT studies have shown a reduction in striatal DA transporters in PD and an association between clinical signs and striatal uptake of [123I]β-CIT. Discussing detailed neuroimaging methods is beyond the scope of this unit, but as shown in Figure 1.8.5, both PET and SPECT can be used successfully to determine the severity of a lesion in monkey striatum following MPTP administration. ICA MPTP administration as a model of active degeneration of the nigrostriatal DA pathway An interesting approach to study neuroprotection in MPTP-treated monkeys is based on



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2.00 putamen

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Figure 1.8.6 Following ICA administration of MPTP, DA terminals degenerate first, followed by degeneration of DA neurons in the SNc 5 weeks later. During the time period between MPTP administration and DA cell death in the SNc there is a transient upregulation of activity in the SNc that can be measured by PET. This upregulation represents the time course of cell death and can be used as an indicator of DA cell degeneration. Asymmetry ratios were constructed to evaluate differences in FMT accumulation between ipsilateral (the side of ICA MPTP infusion) and contralateral sides at eight different time points. Asymmetry ratios for the putamen and SNc were calculated as the Ki value for the ipsilateral side minus the Ki value for the contralateral side divided by the mean of the two Ki. A negative asymmetry ratio indicates that Ki was lower on the ipsilateral side, while positive ratios indicate higher Ki values on the ipsilateral side. *p < 0.0001 significantly less than baseline; †p = 0.001, ‡p = 0.002, and §p = 0.04 significantly greater than baseline; **p < 0.05) significantly greater than 1 week, 2 weeks, 3 weeks.


the finding that following ICA MPTP administration, MPP+ is sequestered in both DA terminals and cell bodies as a result of its selective uptake by the DA transporter system. This initiates molecular changes that ultimately result in neuronal degeneration, most likely by compromising mitochondrial function. Within the first few weeks following MPTP administration, DA levels are profoundly decreased in the striatum with little or no morphological changes in the nigral cell bodies. MPTP-induced nigrostriatal degeneration begins almost immediately in the striatal terminals and results in cell body degeneration within 4 to 6 weeks (Fig. 1.8.7 and Fig. 1.8.8). The difference in the time course of degeneration in the striatum and SNc suggests that the primary mechanism of toxicity in the terminals is MPP+, while cell body degeneration may result from both retrograde degeneration and retrograde transport of MPP+ from the striatum (Javitch et al., 1985; Herkenham et al., 1991). Importantly, although the time course of degeneration is clearly ac-

celerated, the MPTP model includes the key elements of idiopathic PD, including selective and progressive degeneration of the nigrostriatal DA pathway (Fig. 1.8.7). The use of FMT, along with higher resolution PET tomographs, allows direct longitudinal evaluation of DA activity in the SNc as well as in the striatum (Eberling et al., 1997; Fig. 1.8.6). Following ICA infusion of MPTP, activity of DA neurons is upregulated during degeneration of the DA pathway, which is completed within 5 weeks. This pattern of MPTP-induced degeneration can be used to study neuroprotection. Following ICA administration of MPTP, >95% of TH-immunoreactive cells in the SNc will degenerate within 4 to 5 weeks. There is very little, if any, between-animal variation in TH-positive cell degeneration in the SNc following the successful ICA infusion of MPTP. Active neuroprotective agents should be able to protect DA neurons following ICA administration of MPTP. Such neuroprotection might be demon-



MPTP

3 days

5 days

4 weeks

Figure 1.8.7 Low-power photomicrography showing tyrosine hydroxylase immunoreactivity in the striatum and SNc in monkeys at 3 days, 5 days, and 4 weeks following unilateral intracarotid infusion of MPTP. Terminals in the striatum degenerate first, shortly after MPTP administration, while DA cell bodies are not affected (note lack of TH staining in the striatum but not in SNc at 3 and 5 days). It takes 4 to 6 weeks for DA cells in the SNc to degenerate.

strated by preventing upregulation and/or extending survival of DA cells in the SNc on the side of MPTP infusion during the first 3 to 4 weeks following MPTP infusion. Other models of PD in primates Isoquinolines. For some time now there has been an active search for endogenous compounds that act like MPTP. Isoquinoline (IQ) derivatives such as tetrahydroisoquinoline (TIQ), 1-benzyl-TIQ, and (R)-1,2-dimethyl5,6-dihydroxy-TIQ[(R)-N-methyl-salsolinol)] are DA-derived alkaloids that have shown potential neurotoxicity (Nagatsu and Yoshida, 1988). TIQs, like MPTP, may be activated via N-methylation by N-methyltransferase and via oxidation by MAO. TIQs as well as MPP+ inhibit complex I of the electron transport system in mitochondria, thereby reducing ATP formation and producing oxygen radicals. Although the properties of TIQs are similar to those of MPTP, the neurotoxicity is weaker. Chronic administration produces parkinsonism in primates with decreased DA, biopterin, and tyrosine hydroxylase activity (Nagatsu and Yoshida, 1988). Long-term neurotoxic effects of IQs remain to be further examined in primates.

Aged monkeys. DA cells are lost during the normal course of aging at a rate of ∼5% per decade in monkeys (Fearnley and Lees, 1991). Rhesus monkeys that are over 22 years of age display many Parkinson-like changes that appear to be age-related. Some, but not all, hypokinetic aged monkeys respond to L-DOPA administration as shown in Figure 1.8.9. In a recent study, motor deficits in aged monkeys were shown to correlate with decreased DA markers (Emborg et al., 1998). However, because decreased locomotion of aged monkeys can also be attributed to factors that are not DA-mediated, the response to L-DOPA might be used as a selection marker for animals that show DA-mediated hypokinesia. More work needs to be done to fully characterize aged monkeys as a potential model of PD. Other models of PD in mice Weaver mutant. This is a genetically induced model of cerebellum atrophy accompanied by brain DA deficiency due to abnormal development and additional degeneration of the striatal DA system. The adult weaver mutant develops striatal DA deficiency related to a reduction in the number of DA neurons in the SNc (A-9). The neuropathological and biochemical



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H&E

H&E

TH-I R

TH-I R

MPTP Side

Figure 1.8.8 Three weeks following ICA administration of MPTP, this animal showed clear signs of unilateral deficit. Upon PET examination, activity in the SNC was still elevated while the signal from the putamen was absent. The animal was sacrificed one day after PET, and degenerating but still TH-positive neurons were present in the SNc. No TH staining was detected in the putamen. Abbreviations: H&E, hematoxylin and eosin staining; TH-IR, tyrosine hydroxylase immunoreactivity.


changes manifest as ataxia and fine tremor (Schmidt et al., 1982). This is a convenient model for studying potential DA trophic effects. Homozygous weaver animals of the C57Black strain are obtained by breeding heterozygote pairs. At ∼2 weeks of age, homozygous weaver animals are recognized behaviorally by poor righting response, ataxia, hypotonia, and tremor, and anatomopathologically by visible atrophy of the cerebellum. DA-deficient mouse model. This model is created by inactivation of the TH gene resulting in an inability to synthesize DA. These mice show no TH immunoreactivity in midbrain DA neurons and their terminals have extremely low DA concentrations. Behaviorally they manifest hypoactivity, adipsia, and aphagia leading to negative growth rate and death by 4 weeks of age. L-DOPA reverses the behavioral changes, and continuous administration can lead to a nearly normal growth rate. These animals have normal development of DA midbrain neurons and nigrostriatal projections. The model is use-

ful for DA replacement and pharmacological studies.

Critical Parameters and Troubleshooting MPTP primate models Handling MPTP. The experimenter should always use a chemical hood and skin protection (gloves, laboratory coat) and mask. Any spilled MPTP or excess solution should be degraded by potassium permanganate (see cautionary note in Basic Protocol 1). In case of exposure, MAO-B inhibitors block the conversion to MPP+, thus preventing the toxic effects. ICA administration. Selective injection in the ICA minimizes contralateral involvement. For this route, the dose should be calculated according to brain size, which is fairly stable for a wide range of body weights. In macaques there appears to be no direct correlation between the severity of the parkinsonian lesion and the dose of MPTP or the weight and age of the animal. Some monkeys are more suscepti-



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Figure 1.8.9 Activity tracing in a 24-year-old female rhesus monkey over 3 days. This animal displayed an increase of body activity 5 to 10 min following intramuscular administration of 30 mg/kg M-L-DOPA with benserazide. Only some of the hypokinetic monkeys examined showed an increase in activity in response to M-L-DOPA, suggesting that hypokinetic features in aged animals are not always due to decreased DA levels in the striatum. M, mean counts; S, sum of counts.

ble to MPTP than others and, in general, old females are more prone to MPTP toxicity than young males. The choice of anesthetics is also relevant, as the widely used ketamine inhibits DA uptake and can thus decrease MPTP toxicity by inhibiting MPP+ uptake into the DA cells. Acute stage. Administration of MPTP produces mydriasis, piloerection, tachycardia, and tachypnea lasting 5 to 15 min. Repeated injections produce involuntary facial (perioral) movements, facial grimacing, retrocollis, and dystonic flexion of the extremities lasting 10 to 30 min. Intake of food and water should be carefully monitored in the acute phase; some animals might require tube feeding and LDOPA support. In the first 48 hr following ICA administration of MPTP, the animal might present partial motor seizures with occasional secondary generalization; this should be adequately treated with diazepam at 0.5 mg/kg. The degree of motor impairment is quite variable at acute stages. The severity of the motor signs increases over the first 2 weeks. This phase, which can be life threatening in severely affected animals, is followed by a period of

partial improvement during the following 4 to 6 weeks, probably related to functional recovery of non-DA neurons and/or compensatory mechanisms within the nigrostriatal system. In these first weeks, dystonic posturing is frequently observed in severely lesioned animals in addition to akinesia, dyskinesia, and occasionally oculogyric crises. L-DOPA response. Dyskinesias occur in several primate species after MPTP and chronic L-DOPA therapy (Schneider, 1989), as in the majority of patients with chronic PD who are treated with L-DOPA. In general, dyskinesias develop more rapidly in MPTP-induced parkinsonism—both in humans and nonhuman primates—than in patients with idiopathic PD, but it is exceptional when this complication appears with an acute challenge. It is nevertheless advisable to avoid the use of high doses producing significant dyskinesias as this might invalidate the test. Model. The overlesioned bilateral model has several advantages over other models. It is a practical model in which animals are very seldom affected by MPTP to the extent that they



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Figure 1.8.10 Relationship between Ki values for the (A) caudate and (B) putamen, and clinical rating scores for stage 2 and 3 monkeys. Monkeys with more severe clinical signs showed lower Ki values in both regions.


cannot take care of themselves. The success rate of the model is very high. Typically 80% to 90% of animals that are enrolled into the project are considered good study subjects, and the mortality rate is very low (e.g., 97% DA depletion). In general the probability of spontaneous behavioral recovery depends on the extent of DA depletion. However, animals with a similar degree of DA denervation show different recovery profiles in behavioral tests, probably resulting from regional anatomical variation in DA content. Thus, it is necessary to obtain data from a group of animals to control for spontaneous recovery. Rotation in response to apomorphine is fairly stable over time and correlates with other motor scores. Amphetamine induces rotation, but does not increase motor activity and does not correlate with other measures. 6-OHDA rat model The 6-OHDA rat model is rather simple and reliable, although the rotational response must be tested to select rats with severe lesions (>99% DA depletion) for use in long-term experiments. This means that it is necessary to lesion a large number of animals to produce enough well-lesioned animals. High doses (>40 µg) have a considerable mortality rate (e.g., 3% to 5%), as do bilateral lesions. The volume of the infusion is relevant, as diffusion depends directly on volume. For medial forebrain bundle (MFB) lesions, small volumes are recommended to keep the lesion limited; for striatal infusion, large volumes are necessary to cover the whole nucleus.



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Figure 1.8.11 Tissue concentration of DA in the caudate nucleus and putamen in parkinsonian monkeys at stages 1 (n = 3), 2 (n = 7), and 3 (n = 5) shown as a percentage of (A) ipsilateral striatum and (B) contralateral striatum tissue concentration in normal (n = 4) monkeys.

MPTP administration in mice Systemic MPTP administration in mice causes damage to the nigrostriatal rather than mesolimbic terminal areas (Donnan et al., 1987). The severity and duration of this specific damage depends on total dose, administration regimen (multiple injections are more effective than single), route of administration (subcutaneous is the most effective), strain (C57Black strain is the most sensitive), and age (older animals are more sensitive; Heikkila and Sonsalla, 1987; Date et al., 1990). The neurotoxic effect of MPTP can be augmented by coadministration of diethyldithiocarbamate, acetylaldehyde, or ethanol. Recovery of behavioral, neuropathological, and biochemical changes is observed after a variable period of time, most frequently over a period of months depending on the dose of MPTP and cotreatment regimen (Hallman et al., 1985; Donnan et al., 1987).

Anticipated Results Motor behavior Primates. Unilateral administration of MPTP and 6-OHDA is attended by persistent contralateral motor impairment; both upper and lower limbs show the characteristic parkinsonian signs of bradykinesia, rigidity, cogwheeling, and tremor. Spontaneous locomotor activity usually consists of continuous circling towards the lesioned side. Treatment with LDOPA/carbidopa or the directly acting mixed D1 and D2 receptor agonist apomorphine alleviates motor deficits and reverses the direction of turning (Bankiewicz et al., 1986). The apomorphine-induced turning response in a direction away from the lesioned side is very stable over time (for ≥5 years) and can also be quantified. Assessment of drug-induced turning response and function of the contralateral arm can be used for evaluation of new experimental therapeutic strategies.



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Figure 1.8.12 Correlation between levels of DA as measured by PET and HPLC in stage 2 and 3 overlesioned hemiparkinsonian monkeys in (A) caudate and (B) putamen.


Systemic and overlesioned models using MPTP produce a bilateral parkinsonian syndrome displaying axial signs, such as stooped posture and impaired balance. In these animals evaluation can be made using a specific scale similar to the UPDRS, such as the one shown in Table 1.8.1. Figure 1.8.2 shows the effect of increasing doses of L-DOPA on motor activity and scores in a stable parkinsonian primate. Rodents. In rats, turning behavior is the easiest way to assess the degree of DA denervation. Apomorphine induces contralateral rotation in animals with >90% striatal DA depletion. In animals with partial lesions, apomorphine does not induce significant turning but

amphetamine induces ipsilateral rotation demonstrating an asymmetry in DA content. In mice, a decrease of spontaneous locomotor activity and an increase in response to amphetamine can be observed. In vivo neurochemical changes CSF. After unilateral administration of MPTP in monkeys, the level of monoamine metabolites in cerebrospinal fluid (CSF) initially drops. However, after 3 months, only the levels of homovanillic acid remain low (Burns et al., 1983). In hemiparkinsonian monkeys, the initial decrement in CSF monoamine metabolites and DA levels is >60%; three weeks after




1.8.27


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Substantia nigra pars lateralis

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Figure 1.8.13 Anatomical representation of midbrain DA structures in normal and stage 1, 2, and 3 overlesioned hemiparkinsonian monkeys. In stage 1, DA cells in the SNc (A-9 region) are severely depleted while the contralateral side shows no significant cell death in this region. Note that TH-positive cells in the SNc are further depleted in stage 2 and 3 monkeys, which corresponds to progressive parkinsonian signs that are seen in these monkeys. Regions of the ventral tegmental area are partially spared at all stages.

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Figure 1.8.14 Results of in vivo microdialysis of three hemiparkinsonian (stage 1) monkeys. Samples were collected over 20 min from the caudate and putamen on both sides of the brain and were analyzed for levels of DA. Extracellular levels of DA were detected only on the control side (A), with putamen showing higher levels of DA than caudate nucleus. KCl and amphetamine induced DA release only on the control side, suggesting complete DA lesion on MPTP-treated side (B).

the lesion they are decreased by only 40% to 50% (Bankiewicz et al., 1990). Microdialysis. Extracellular fluid (ECF) levels of catecholamines in brain tissue can be examined in vivo by microdialysis (Fig. 1.8.14). Using a small probe (27-G), it is possible to measure the regional distribution and concentration of DA and its metabolites in brain ECF (Wang et al., 1990). With this technique it is also possible to examine the relationship between changes in levels of different neurotransmitters in brain ECF. Methodology for performing microdialysis experiments in MPTP-treated monkeys is described elsewhere (Wang et al., 1990).


Postmortem analyses Parkinson’s disease is characterized by loss of DA neurons in the SNc leading to DA depletion in the striatum. Consequently, evaluation of nigral cell loss and striatal DA deficiency are the main post mortem outcome measures for all of these models. Immunohistochemistry. The distribution of neuronal damage within the SNc in the MPTPtreated animal model (see Basic Protocol 1) is also of interest because histological changes in this region are exhibited in a specific pattern in PD. MPTP is selectively toxic, affecting mostly the nigrostriatal DA system while leaving the mesolimbic system relatively intact. The midline population of DA cells in the midbrain

(ventral tegmental area, paranigralis nucleus, and paraventricular area), the parabrachial nucleus, and parts of nucleus lateralis are affected by MPTP (German et al., 1988). Furthermore, the damage closely resembles the histological findings in PD. TH immunoreactivity in DA midbrain nuclei and anatomic schemes of each level are shown in Figure 1.8.12 for different stages. MPTP-treated monkeys in stage 1 (hemiparkinsonian) show a dramatic loss of DA cells (>95%) in the SNc on the MPTP-lesioned side, whereas animals at stage 2 and 3 display further loss of DA neurons on the contralateral side as they become more parkinsonian (Fig. 1.8.13). This pattern follows changes observed by neuroimaging (Figs. 1.8.5 and 1.8.6) and biochemical techniques (Fig. 1.8.11). The decrease of TH immunoreactivity in the SNc (cell bodies) and striatum (projections) is observed in all models described here. However, it is transient in mice and, in partial lesions, a significant recovery can be observed over time. Unilateral injection of 6-OHDA in marmosets (see Basic Protocol 2) produces a decrease in TH immunoreactivity in the SNc of 90% to 97% on the contralateral side (Annett et al., 1992). In rats treated with 6-OHDA (see Basic Protocol 3), loss of TH-positive nigral cells depends on the type of lesion. Lesions of the



MFB and SNc at high doses should achieve a complete nigral depletion. Partial lesions should respect VTA and the medial ventral tier. Striatal lesions cause a cell loss in the ipsilateral SNc that is correlated with the dose and volume of the striatal injection. Neurochemical changes. MPTP administration (see Basic Protocol 1) causes a profound reduction in striatal DA with no clear differences between caudate and putamen levels. After ICA MPTP administration, DA is almost completely absent in the caudate and putamen on the lesioned side. The difference in DA concentration between the normal and the MPTP-treated side is >96% (Fig. 1.8.11). This represents almost total DA denervation of the caudate and putamen on the MPTP-lesioned side. However, greater concentrations of DA are found in areas that receive innervation from remaining DA cells in the ventral tegmental area (e.g., nucleus accumbens septi, area olfactoria, septum pellucidum). In the model that combines ICA and i.v. MPTP administration, DA depletion on the side contralateral to ICA administration is much less severe than on the side of ICA administration and correlates with the severity of parkinsonian signs. DA concentration, as measured by postmortem striatal punches (Fig. 1.8.11), correlates well with in vivo PET measurements of DA activity (Fig. 1.8.12). In marmosets lesioned with 6-OHDA (see Basic Protocol 2), the reduction of DA in the ipsilateral striatum is usually >97% (Annett et al., 1992). In rats lesioned with 6-OHDA (see Basic Protocol 3) and showing a robust rotational response to apomorphine, DA levels are decreased by >99% in the contralateral striatum.

Time Considerations For Basic Protocol 1, ICA injection takes 30 to 60 min and i.v. injection takes 15 min. To obtain overlesioned animals, a 2-week waiting period is required between ICA and i.v. administration. It is then necessary to wait 6 to 8 weeks (acute stage and spontaneous recovery) to establish a stable parkinsonian syndrome. Clinical rating takes 99% DA depletion). These animals show a stable deficit with consistent druginduced rotational behavior over time appropriate for such studies. Basic Protocol 4 is a very simple and quick procedure as animals can be used right away but only for a short period of time. Animals show minimal or no behavioral symptoms after weeks to months.

Literature Cited Annett, L.E., Rogers, D.C., Hernandez, T.D., and Dunnett, T.B. 1992. Behavioral analysis of unilateral monamine depletions in the marmoset. Brain 115:825-856. Bankiewicz, K.S., Oldfield, E.H., Chiueh, C.C., Doppman, J.L., Jacobowitz, D.M., and Kopin, I.J. 1986. Hemiparkinsonism in monkeys after unilateral internal carotid artery infusion of 1methyl-4-phenyl-1,2,3,6-tetrahydropyridine (MPTP). Life Sci. 39:7-16. Burns, R.S., Chiueh, C.C., Markey, S.P., Ebert, M.H., Jacobowitz, D.M., and Kopin, I.J. 1983. A primate model of parkinsonism: Selective destruction of dopaminergic neurons in the pars compacta of the substantia nigra by N-methyl-4phenyl-1,2,3,6-tetrahydropyridine. Proc. Natl. Acad. Sci. U.S.A. 890:4546-4550. Date, I., Felten, D.L., and Felten, S.Y. 1990. Long term effect of MPTP in the mouse brain in relation to aging: Neurochemical and immunocytochemical analysis. Brain Res. 519:266-272. Davis, G.C., Williams, A.C., Markey, S.P., Ebert, M.N., Caine, E.D., Reichert, C.M., and Kopin, I.J. 1979. Chronic parkinsonism secondary to intravenous injection of meperidine analogs. Psychiatry Res. 1:249-254. Donnan, G.A., Kaczmarczyk, S.J., Mckenzie, J.S., Rowe, P.J., Kalnins, R.M., and Mendelsohn, F.A. 1987. Regional and temporal effects of 1methyl-4-phenyl-1,2,3,6-tetrahydropyridine on dopamine uptake sites in mouse brain. J. Neurol. Sci. 81:261-271. Doudet, D.J., Miyake, H., Finn, R.T., Mclellan, C.A., Aigner, T.G., Wan, R.Q., Adams, H.R., and Cohen, R.M. 1989. 6-18F-L-Dopa imaging of the dopamine neostriatal system in normal and clinically normal MPTP-treated rhesus monkeys. Exp. Brain Res. 78:69-80. Eberling, J.L., Bankiewicz, K.S., Jordan, S., Vanbrocklin, H.F., and Jagust, W.J. 1997. Pet studies of functional compensation in a primate model



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of Parkinson’s disease. NeuroReport 8:27272733. Eberling, J.L., Jagust, W.J., Taylor, S., Bringas, J., Pivirotto, P., Vanbrocklin, H.F., and Bankiewicz, K.S. 1998. A novel MPTP primate model of Parkinson’s disease: Neurochemical and clinical changes. Brain Res. 805:259-262. Emborg, M.E., Ma, S.Y., Mufson, E.J., Levey, A.I., Taylor, M.D., Brown, W.D., Holden, J.E., and Kordower, J.H. 1998. Age-related declines in nigral neuronal function correlate with motor impairments in rhesus monkeys. J. Comp. Neurol. 401:253-265. Fearnley, J.M. and Lees, A.J. 1991. Aging and Parkinson’s disease: Substantia nigra regional selectivity. Brain 114:2283-2301. German, D.C., Dubach, M., Askari, S., Speciale, S.G., and Bowden, D.M. 1988. 1-Methyl-4phenyl-1,2,3,6-tetrahydropyridine-induced parkinsonian syndrome in Macaca fascicularis: Which midbrain dopaminergic neurons are lost? Neuroscience 24:161-174. Hallman, H., Lange, J., Olson, L., Strömberg, I., and Johnsson, G. 1985. Neurochemical and histochemical characterization of neurotoxic effects of 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine on brain catecholamine neurons in the mouse. J. Neurochem. 44:117-127. Heikkila, R.E. and Sonsalla, P.K. 1987. The use of the MPTP-treated mouse as an animal model of Parkinsonism. Can. J. Neurol. Sci. 14:436-440. Heikkila, R.E., Hess, A., and Duvoisin, R.C. 1985. Dopaminergic neurotoxicity of 1-methyl-4phenyl-1,2,3,6-tetrahydropyridine (MPTP) in the mouse: Relationships between monoamine oxidase, MPTP metabolism and neurotoxicity. Life Sci. 36:231-236. Herkenham, M., Little, M.D., Bankiewicz, K., Yang, S.-C., Markey, S.P., and Johannessen, J.N. 1991. Selective retention of MPP+ w ithin the monoaminergic systems of the primate brain following MPTP administration: An in vivo autoradiographic study. Neuroscience 40:133-158. Javitch, J.A., D’amato, R.J., Strittmatter, S.M., and Snyder, S.H. 1985. Parkinsonism-inducing neurotoxin N-methyl-4-phenyl-1,2,3,6-tetrahydropyridine: Uptake of the metabolite N-methyl4-phenylpyridine by dopamine neurons explains selective toxicity. Proc. Natl. Acad. Sci. U.S.A. 82:2173-2177. Jordan, S., Eberling, J.L., Bankiewicz, K.S., Rosenberg, D., Coxson, P.G., Vanbrocklin, H.F., O’Neil, J.P., and Emborg, M.E. 1997. 6(18F)Fluoro-L-m-tyrosine: Metabolism, PET kinetics and MPTP lesion in primates. Brain Res. 750:264-276.


Laruelle, M., Baldwin, R.M., Malison, R.T., ZeaPonce, Y., Zoghbi, S.S., al-Tikriti, M.S., Sybirska, E.H., Zimmermann, R.C., Wisniewski, G., Neumeyer, J.L., et al. 1993. SPECT imaging of dopamine and serotonin transporters with [123I]β-CIT: Pharmacological characterization of brain uptake in nonhuman primates. Synapse 13:295-309.

Lee, C.S., Sauer, H., and Bjorklund, A. 1996. Dopaminergic neuronal degeneration and motor impairments following axon terminal lesion by intrastriatal 6-hydroxydopamine in the rat. Neuroscience 72:641-653. Leroux-Nicollet, I. and Costentin, J. 1986. Acute locomotor effects of MPTP in mice and relationships with dopaminergic systems. In MPTP: A Neurotoxin Producing a Parkinsonian Syndrome (S.P. Markey, N. Castagnoli, A.J. Trevor, and I.J. Kopin, eds.) pp. 419-424. Academic Press. Melega, W.P., Perlmutter, M.M., Luxen, A., Nissenson, C.H., Grafton, S.T., Huang, S.C., Phelps, M.E., and Barrio, J.R. 1989. 4-(18F)Fluoro-L-mtyrosine: An L-3,4-dihydroxyphenylalanine analog for probing presynaptic dopaminergic function with positron emission tomography. J. Neurochem. 53:311-314. Nagatsu, T. and Yoshida, M. 1988. An endogenous substance of the brain, tetrahydroisoquinoline, produces parkinsonism in primates with decreased dopamine, tyrosine hydroxylase and diopterin in the nigrostriatal regions. Neurosci. Lett. 87:178-182. Nicklas, W.J., Vyas, R.E., and Heikkila, R.E. 1985. Inhibition of NADH-linked oxidation in brain mitochondria by 1-methyl-4-phenyl-pyridine, a metabolite of the neurotoxin 1-methyl-4-phenyl1,2,5,6-tetrahydropyridine. Life Sci. 36:25032508. Oiwa, Y., Eberling, J.L., Nagy, D., Pivirotto, P., Emborg, M.E., and Bankiewicz, K.S. 2003. Overlesioned hemiparkinsonian nonhuman primate model: Correlation between clinical, neurochemical and histochemical changes. Front Biosci. 8:A155-A166. Pate, B.D., Kawamata, T., Yamada, T., Mcgeer, E.G., Hewitt, K.A., Snow, B.J., Ruth, T.J., and Calne, D.B. 1993. Correlation of striatal fluorodopa uptake in the MPTP monkey with dopaminergic indices. Ann. Neurol. 34:331-338. Paxinos, G. and Watson, C. 1986. The Rat Brain in Stereotaxic Coordinates. Academic Press, New York. Perese, D.A., Ulman, J., Viola, J., Ewing, S.E., and Bankiewicz, K.S. 1989. A 6-hydroxydopamineinduced selective parkinsonian rat model. Brain Res. 494:285-293. Ricaurte, G.A., Irwin, I., Forno, L.S., DeLanney, L.E., Langston, E., and Langston, J.W. 1987. Aging and 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine-induced degeneration of dopaminergic neurons in the substantia nigra. Brain Res. 403:43-51. Sauer, H.W. and Oertel, H. 1994. Progressive degeneration of nigrostriatal dopamine neurons following intrastriatal terminal lesions with 6-hydroxydopamine—A combined retrograde tracing and immunocytochemical study in the rat. Neuroscience 59:401-415. Schmidt, M.J., Sawyer, B.D., Perry, K.W., Fuller, R.W., Foreman, M.M., and Ghetti, B. 1982. Dopamine deficiency in the weaver mutant mouse. J. Neurosci. 2:376-380.



Schneider, J.S. 1989. Levodopa-induced dyskinesias in parkinsonian monkeys: Relationship to extent of nigrostriatal damage. Pharmacol. Biochem. Behav. 34:193-196. Schneider, J.S. and Kovelowski, C.J. 1990. Chronic exposure to low doses of MPTP. I. Cognitive deficits in motor asymptomatic monkeys. Brain Res. 519:122-128. Schultz, W. 1982. Depletion of dopamine in the striatum as an experimental model of parkinsonism: Direct effects and adaptive mechanisms. Prog. Microbiol. 18:121-166.

insonian nonhuman primates. Exp. Neur. 110:181-186. Yang, S.C., Markey, S.P., Bankiewicz, K.S., London, W.T., and Lunn, G. 1988. Recommended safe practices for using the neurotoxin MPTP in animal experiments. Lab. Anim. Sci. 38:563-567. Zigmond, M.J. and Strickler, E.M. 1989. Animal models of parkinsonism using selective neurotoxins: Clinical and basic implications. Int. Rev. Neurosci. 31:1-79.

Sundstrom, E., Fredriksson, A., and Archer, T. 1990. Chronic neurochemical and behavioral changes in MPTP-lesioned C56BL/6 mice: A model for Parkinson’s disease. Brain Res. 528:181-188.

Contributed by Krys S. Bankiewicz University of California San Francisco San Francisco, California

Thomas, J., Wang, J., Takubo, H., Sheng, J.G., Dejesus, S., and Bankiewicz, K.S. 1994. A 6-hydroxydopamine-induced selective parkinsonian rat model: Further biochemical and behavioral characterization. Exp. Neurol. 126:159-167.

Rosario Sanchez-Pernaute McLean Hospital and Harvard Medical School Boston, Massachusetts

Ungerstedt, U. 1971a. Striatal dopamine release after amphetamine or nerve degeneration revealed by rotational behaviour. Acta Physiol. Scand. Suppl. 367:51-68.

Yoshitsugu Oiwa Wakayama Medical University Wakayama, Japan

Ungerstedt, U. 1971b. Postsynaptic supersensitivity after 6-hydroxydopamine induced degeneration of the nigrostriatal dopamine system. Acta Physiol. Scand. Suppl. 367:69-93.

Malgorzata Kohutnicka Institute of Psychiatry and Neurology Warsaw, Poland

Ungerstedt, U. and Arbuthnott, G.W. 1970. Quantitative recording of rotational behavior in rats after 6-hydroxydopamine lesions of the nigrostriatal dopamine system. Brain Res. 24:485-493.

Alex Cummins National Institute of Mental Health Bethesda, Maryland

Wang, J., Skirboll, S., Aigner, T.G., Saunders, R.C., Hsiao, J. and Bankiewicz, K.S. 1990. Methodology of microdialysis of neostriatum in hemipark-

Jamie Eberling University of California Davis Davis, California



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CHAPTER 2 Assessment of Cell Toxicity INTRODUCTION

E

nvironmental agents that are injurious to cells trigger a spectrum of responses that can range from cell death to adaptation, repair, and proliferation. Cell death, in turn, can take the form of apoptosis—a programmed form of cell death that requires gene expression—or necrosis. A host of environmental agents have been implicated in toxic cell injury, among them free radicals (nitrogen, oxygen, and carbon derivatives), which are known to interact with cellular constituents including membrane lipids and chromagen materials.

There are various ways of evaluating cellular toxicity and cell death, and a number of different indices for assessing cellular response to toxic stimulus or injury. UNIT 2.1 outlines the current understanding of what constitutes cell toxicity and how it should be defined. UNIT 2.2 presents a number of methods that can be used to discriminate between the apoptotic and necrotic modes of cell death. Because many of the distinguishing criteria are not specific to one mode, more than one method should be used to make the determination. It is now known that the mitochondrial function is changed in apoptosis. UNIT 2.10 provides protocols for assessment of various aspects of mitochondrial function. UNIT 2.3 describes immunochemical methods for detecting the covalent binding of chemicals to proteins as a means to investigate the interactions of potentially cytotoxic chemicals with cellular constituents. UNIT 2.4 describes methods for detecting lipid peroxides, free radicals formed by the interaction of oxygen derivatives with membrane proteins. Oxygen free radicals have been implicated in both apoptosis and necrosis. provides protocols for measuring intracellular free Ca2+ using fluorescence spectroscopy or digital imaging microscopy. Changes in cellular Ca2+ homeostasis are intimately linked to cell death, with the cascade of events that they set in motion culminating in cellular necrosis and/or apoptosis. In addition, intracellular calcium is used in signaling and as a second messenger. UNIT 2.5

UNIT 2.6 provides protocols that can be used to detect cytotoxicity, which is operationally defined as the effect toxicants have on cell growth, metabolic functions, or viability. The protocols permit detection of physiological and/or morphological changes that may have occurred as a consequence of toxicant exposure. Sixteen protocols are provided to detect cytotoxicity in vitro. These protocols can be used to screen chemicals efficiently and economically before testing in animals.

describes in situ hybridization for analyzing specific cellular mRNA in frozen and cryostat-sectioned tissues. The method combines cell and molecular biology techniques for detection and localization of specific nucleic acid sequences within individual cell or tissue preparations. Many protocols have been developed for different types of in situ hybridization. This unit includes methods using either radiolabeled or digoxigenin-labeled probes.

UNIT 2.7

Assessment of Cell Toxicity Contributed by Mahin D. Maines Current Protocols in Toxicology (2004) 2.0.1-2.0.2 Copyright © 2004 by John Wiley & Sons, Inc.

2.0.1 Supplement 19

UNIT 2.8 presents a basic insight into the theoretical and practical possibilities and limitations of confocal microscopy. It also provides practical guidelines for the novice user of the technique. The method makes it possible to view a very thin optical plane in a specimen by excluding light from planes above and below the focal plane. Although there are several different types of confocal microscopes developed for specialized purposes, this unit concentrates on confocal scanning laser microscopy (CSLM).

The heat shock family of proteins (HSP) rapidly responds to chemical and environmental insults. The HSP70 family of proteins comprises a major family of stress-responsive proteins that have in common various functions in cellular defense mechanisms. UNIT 2.9 provides procedures that allow measurement of response of different isotypes of the HSP70 family at the transcription and translation levels. Mahin D. Maines

Introduction

2.0.2 Supplement 19


Current Concepts in Cell Toxicity Toxicology is sometimes defined as the study of the adverse or harsh effects of toxic agents on living cells, tissues, or whole organisms. The term “toxic agent” is generally used to refer to both naturally occurring toxic substances (toxins) and to harmful products or byproducts (toxicants) generated from human activities, which may have diverse effects on living cells. These agents can be subclassified in various ways—for example, according to their target organ (e.g., liver, kidney, or reproductive organs), industrial use (e.g., food additives or pesticides), source (e.g., animal or plant toxins), and effects (e.g., cancer or liver injury). When a cell is exposed to toxic agents, it will undergo certain molecular or biochemical transformations. The biotransformations or alterations caused by the cell’s exposure to harmful products is generally referred to as cell toxication or cell toxicity. On the other hand, the biotransformations or cellular alterations that lead to the elimination of toxicants or the prevention of toxic metabolite formation from these compounds are called “cell detoxification.” Cell toxicity can have reversible (short-lived or repairable) effects, or affect the cell irreversibly, leading to mutations, malignant transformation, or cell death. The final outcome of being in contact with toxic agents depends mainly on the nature of the toxic substance, the dose applied, and the exposure time. At low doses or short exposure times, small quantities of the toxic agent will come in contact with the cell and be delivered into its cytosol. This may lead to toxicity resulting from the mere presence of the agent in the cell, or the agent may interact with an array of target molecules. Such toxicity initiates a series of events leading to minor and potentially reversible injuries. These events can cause a temporary halt in some of the cellular functions. Similarly, the presence of toxic agents or cellular injuries caused by them may activate the cell’s defenses—detoxification or repair mechanisms—to enable it to withstand such harsh environments and to repair the injuries that have been inflicted upon the cell. The cell’s defense mechanisms involve rapid production or activation of specific molecules—examples of which are heat-shock proteins (HSPs), metallothioneins (MTs), superoxide dismutases (SODs), and catalase— which enable the cell to withstand or tolerate

selective toxic agents. A number of xenobiotics (e.g., carbon monoxide, heavy-metal ions, and strong acids and bases) are directly toxic, whereas the toxicity of the others is largely due to production of metabolites. The mechanism of detoxification includes glutathione, cytochrome P-450, Cu, Zn-SOD, and Mn-SOD. However, at high doses or prolonged exposure times, the toxic agents cause irreversible cellular damage, to the extent that the rate of toxication exceeds that of detoxification, leading to a general cellular dysfunction and cell death. This unit attempts to define cell toxicity that leads to cellular demise, with a strong emphasis on cell death via both apoptosis and necrosis, by summarizing some of the more recent developments in cellular, molecular, and biochemical studies of the events that govern the induction and execution of cell death.

CELL DEATH Toxic cell death can occur via two processes, which are fundamentally different in their nature and biological significance (Kerr et al., 1972). These are termed necrosis and apoptosis and their features are summarized in Table 2.1.1. Necrosis, or “accidental” cell death, was the classic model and was thought to be the universal mode of cell death until apoptosis was identified in 1972 as a separate mode of cell death. In general, necrosis is considered to be a passive process (Trump and Ginn, 1969) that is usually caused by extreme trauma or injury to the cell (Kerr et al., 1972). In tissues, necrosis is often seen as a lesion or patch consisting of many disintegrated cells. Typically, necrosis involves irreversible changes within the nuclei (such as karyolysis), loss of cytoplasmic structure, dysfunction in various organelles (especially mitochondria), and, finally, cytolysis as a result of high-amplitude swelling. The release of the dying cell’s contents into the extracellular space can cause further injury or even death of neighboring cells, and may result in inflammation or infiltration of proinflammatory cells into the lesion, leading to further tissue damage (Haslett, 1992). The first references to a different type of cell death came from developmental studies. The concept of a programmed physiological cell death in developmental biology or embryology refers to the type of cell death that occurs at

Contributed by Afshin Samali, Boris Zhivotovsky, and Sten Orrenius Current Protocols in Toxicology (1999) 2.1.1-2.1.16 Copyright © 1999 by John Wiley & Sons, Inc.

UNIT 2.1

Assessment of Cell Toxicity

2.1.1

Table 2.1.1

Features of Apoptosis and Necrosis

Characteristics

Apoptosis

Necrosis

Stimuli Occurrence Reversibility

Physiological or pathological Single cells Limited

Pathological (injury) Groups of cells Limited

Shrinkage and formation of apoptotic bodies Lost (early) Present Absent

Swelling and later disintegration Lost (late) Absent Present

Blebbing Late-stage swelling Present

Blebbing proir to lysis Very early swelling Present

Convolution of nuclear outline and breakdown (karyorrexis)

Disappearance (karyolysis)

Presenta Presenta

Absenta Absenta

Absent Present

Present Present

Present Present Present High Present Compaction in uniformly dense masses HMW and internucleosomal Present Present

Absent —a Present Low Present Clumping not sharply defined

Cellular level Cell shape Adhesion between cells Phagocytosis by other cells Exudative inflammation Cellular organelles Membranes Cytoplasm Mitochondrial permeability transition Nucleus

Biochemical level Gene activation Requirement for protein synthesis Lysosomal enzyme release Activation of non-lysosomal enzymes Activation of caspases Cleavage of specific proteins Changes in cytoskeleton Level of ATP required Bcl-2 protection Nuclear chromatin DNA breakdown RNA degradation Phosphatidylserine exposure aThis

Current Concepts in Cell Toxicity

Randomized —a Absent

feature is not a universal event or there are conflicting reports.

specific times during the development of the organism. Glucksmann (1951) clearly emphasized the existence of this phenomenon for the first time. This was followed by a surge of interest in 1960s (for a review see Lockshin, 1981). In the 1970s, histochemical studies of lysosomal changes in hepatic ischemia provided direct evidence for the existence of two distinct types of cell death (Kerr, 1971; Kerr et al., 1972). It was observed that rounded masses that developed in dead hepatic tissue contained cells that were morphologically different from those found in necrosis. These differences initially caused some confusion: the process was mistaken for a variation of necrosis and was

called “shrinkage necrosis” (Kerr, 1971). The following year, Kerr, Wyllie, and Currie proposed the term “apoptosis” to describe the distinct morphological changes associated with this form of cell death (Kerr et al., 1972). Nowadays, apoptosis commonly refers to the process of cell death leading to the appearance of the morphological changes, rather than to the morphological characteristics themselves, which set this form of cell death apart from necrosis. Apoptosis has been described as a form of cellular suicide, since death appears to result from induction of active processes within the cell itself. Typically, apoptosis involves margi-


nation and condensation of nuclear chromatin (pyknosis), cytoplasmic shrinkage, membrane blebbing, nuclear fragmentation, and finally, formation of apoptotic bodies. A cell that is dying, whether by apoptosis or necrosis, undergoes rapid changes which are reflected in its structure, morphology, and biochemistry. These are generally a result of various enzymes activated through elaborate signaling pathways. Since the first description of apoptosis in 1972, there has been a surge of interest in the study of apoptosis, and very little attention has been given to necrosis. This lack of parallel study of the two processes is also reflected in the degree of detail provided here describing apoptosis and necrosis.

STRUCTURAL CHANGES DURING CELL DEATH Considerable biochemical changes occur within the apoptotic cell, which facilitate neat packaging and removal of apoptotic bodies by neighboring cells. However, as described above, necrosis is accompanied by high-amplitude cell swelling. Modifications in the cytoskeleton and cytoplasmic membrane are required for both shrinkage and swelling to occur. During apoptosis, this results in the loss of cell-cell contact, untethering of the plasma membrane, and rapid blebbing or zeiosis (Saunderson, 1982). Similarly, necrotic cells undergo membrane blebbing, but in contrast to apoptosis, the tracts of contiguous dying necrotic cells collapse without loss of contact with neighboring cells. This results in clumps of necrotic cells rather than single cells. Tissue transglutaminase, a Ca2+-dependent protein-glutamine γ-glutamyltransferase, is induced and activated in liver hyperplasia and in glucocorticoid-treated thymocytes (Fesus et al., 1987). This enzyme cross-links cytokeratin, a component of the cytoskeleton, through ε-(γglutamyl) lysine bonds. During apoptosis there is a significant increase in transglutaminase mRNA, as well as the the protein itself, its enzyme activity, and protein-bound (γ-glutamyl) lysine (Fesus et al., 1987). It is thought that transglutaminase activity may stabilize apoptotic cells and inhibit membrane leakage during the early stages of the process by forming a shell around the cell. Transglutaminase activity, however, does not appear to change during necrosis. For example, it was recently reported that ethanol-induced cytotoxicity in astroglial cells was due to necrotic cell death, and that it occurred in the absence of any change in transglutaminase activity (Holownia et al., 1997).

A critical part of apoptosis, in contrast to necrosis, is the efficient recognition and removal of the apoptotic cells by phagocytes (Fadok et al., 1992). This involves the rearrangement and biochemical alteration of the plasma membrane in the dying cell. There are a number of different changes that occur in the plasma membrane of apoptotic cells. One such change results in the alteration of carbohydrates on the plasma membrane, which helps preferential binding of macrophages to apoptotic cells (Duvall et al., 1985). Recognition of apoptotic cells by macrophages can also be mediated via the vitronectin receptor (CD36; Savill et al., 1990). The loss of membrane phospholipid asymmetry in the membrane of cells undergoing apoptosis is yet another significant change leading to the externalization of phosphatidylserine at the surface of the cell, thereby enabling cell recognition by macrophages (Fadok et al., 1992). Development of fluorescently labeled annexin V, which binds specifically to phosphatidylserine residues, enables detection of this externalization in apoptotic cells (Koopman et al., 1994). However, under in vitro culture conditions, where phagocytic cells are absent, apoptotic cells and their fragments lyse in a process very similar to that observed in necrosis. This phenomenon is termed secondary necrosis or post-apoptotic necrosis. Despite such systematic changes in the plasma membranes of apoptotic cells, there are no reports of any changes in the composition of the plasma membranes of necrotic cells. Necrotic cells do not undergo phagocytosis by other cells, and the spillage of their cellular contents through plasma membrane disruption into the extracellular milieu induces cellular inflammatory responses. Cell-volume changes are another feature of cell death. Interestingly, the cell volume changes during apoptosis and necrosis go in different directions. Apoptotic cell shrinkage, which leads to a loss of 30% to 50% of the cell volume (Saunderson, 1982), is due to budding of the endoplasmic reticulum. Vesicles thus generated migrate and fuse to the plasma membrane, releasing their contents into the extracellular region. This process requires energy (ATP), since water is moved against the osmotic gradient (Saunderson, 1982). Mitochondria, which are postulated to remain structurally and functionally intact during apoptosis, provide the necessary energy. In contrast, during necrosis the cell loses control of ion flux, resulting in a reduction in concentration gradients of Na+, K+, Ca2+, and Mg2+, as well as changes in



osmotic pressure. This, in turn, leads to uptake of water, giving rise to high-amplitude swelling of the cell and its organelles. This process is energy-independent and does not require a large amount of ATP.

MACROMOLECULAR DEGRADATION DURING CELL DEATH Cell death is associated with activation or increase in the activity of lipases, nucleases, and different classes of proteases. In apoptosis there is an efficient and neat packaging of the cell contents into apoptotic bodies. This requires the rearrangement and breakdown of lipids, proteins, and nucleic acids. Another possible reason for efficient degradation of macromolecules in apoptotic cells prior to their phagocytosis is that contents from dead cells are thus prevented from entering the phagocyte or the neighboring cells and causing the same injuries that may have cost the dying cell its life. Which enzymes and substrates are activated depends on the induction stimuli and/or the cell type. In necrosis, the types of enzymes activated appear to be different, with very little overlap, if any, with the apoptotic enzymes. These enzymes are largely lysosomal acidic enzymes with less specific substrates than the apoptotic enzymes. However, these necrotic enzymes are thought to be activated in late apoptosis and in the absence of phagocytosis, when the cells undergo secondary necrosis. Thus, there is a similarity between the features of necrotic cells and those of apoptotic cells undergoing secondary necrosis. The major macromolecules and the mode of their degradation during apoptosis and necrosis are discussed below.

DNA Degradation


Three patterns of DNA degradation are already known to occur during apoptosis. One or more of these may occur during the progression of apoptosis in a single cell (for review see Zhivotovsky et al., 1994). These are: singlestrand nicks (Gorczyca et al., 1992), chromatin cleavage with the formation of large (50- to 200-kbp) fragments (Brown et al., 1993; Zhivotovsky et al., 1994), and finally formation of nucleosome-size fragments of 180 to 200 bp, which produce the ladder pattern that has long been accepted as a biochemical hallmark of apoptosis (Wyllie, 1980). One or more nuclear endonucleases have been suggested as being responsible for these patterns of DNA fragmentation, since isolated

nuclei can be induced to produce the same pattern. A number of endonucleases have been identified in different cell systems, with different ion requirements for their activity (for review see Zhivotovsky et al., 1994). For example, the endonuclease in thymocytes is Ca2+/Mg2+-dependent, whereas the endonuclease operating in HL-60 cells appears to function independently of these ions (Fernandes and Cotter, 1993). Endonuclease activity in cells undergoing apoptosis can be regulated by proto-oncogenes and tumor-suppressor genes, such as c-myc, Ha-ras, bcl-2, and p53, as shown by analysis of genetic control of susceptibility to apoptosis (Arends et al., 1993). The apoptotic endonuclease or endonucleases cleave DNA at the exposed linker regions, and it has been suggested that these nucleases are topologically constrained rather than sequence constrained. Results obtained from cloning and sequencing DNA fragments from apoptotic cells show that the nuclease has no preference for specific DNA sequences or for the type of DNA to cleave. This randomness may be functionally important, and reflects the known properties of the endonucleases. The finding that apoptotic endonucleases are constitutive enzymes and that their activity can be modulated by different signals in cells undergoing apoptosis underlines the important role of nucleases in both physiological and pathophysiological processes. Although the role of intracellular endonucleases is not yet understood, one may speculate that a major function is to maintain genomic stability. Unfortunately, both purpose and mechanisms of DNA fragmentation in apoptotic cells are still unclear. Such fragmentation could serve to destroy the genetic information of unwanted cells and thus act as an irreversible step in the process. On the other hand, it may be simply a mid-to-late event that reflects ion redistribution and subsequent chromatin hypersensitivity to endonuclease(s) present in apoptotic cells. Alternatively, it may facilitate cleavage of DNA before uptake of apoptotic bodies during phagocytosis. Whatever the reason, it is still important to understand the mechanisms of this intriguing step of apoptosis. As compared to apoptosis, at least two patterns of DNA degradation are recognized as occurring during necrosis. The first is the formation of both single- and double-strand DNA breaks. The second is randomized chromatin fragmentation. It is unclear if there is a link between these two steps of chromatin disintegration, although the probability for that is


rather high. DNA fragments isolated from necrotic cells contain 5′-OH and 3′-PO4 end groups. Among different endonucleases isolated and characterized up to now, only one is able to produce such DNA fragments—i.e., DNase II, which is localized in lysosomes. This makes sense, since the activation of lysosomal enzymes has been observed in necrotic cells. Several publications have implicated the involvement of DNase II in apoptosis as well; however a critical role for this enzyme in apoptosis appears improbable in view of at least two considerations. First, DNase II is a lysosomal enzyme and lysosomes are intact until very late stages of apoptosis. Second, DNA fragments isolated from apoptotic cells contain 3′-OH end groups, not 3′-PO4 groups as in necrotic cells. The search for enzymes responsible for cleaving of chromatin during necrosis is still in progress in many laboratories.

RNA Degradation Changes in ribosomal counts occurring in response to cell injury have been known for a number of years. This phenomenon may be a result of the release of ribosomes from the endoplasmic reticulum at particular stages leading to necrosis (for review see Bowen, 1981). A detailed analysis of the role of ribosomes in the pathogenesis of liver-cell necrosis was conducted by Bernelli-Zazzera (1975). He showed that ischemia resulted in a decrease in the number of ribosomes, and that stripped rough endoplasmic reticulum from ischemic cells bound fewer added ribosomes than did the reticulum from normal cells. We have concluded that there is a “loosened relationship” between endoplasmic reticulum membranes and ribosomes in ischemic livers. However, it is unknown how specific these changes are for necrosis and what relationship they bear to this type of cell death. There are also a number of reports that RNA, as well as DNA, is susceptible to cleavage during apoptosis. The changes in ribosomal RNA (rRNA) during apoptosis are best studied. rRNA consists of conserved and divergent (or variable) domains numbered from D1 to D12 (for review see Houge and Doskeland, 1996). The cleavage of rRNA during apoptosis selectively affects the two largest divergent domains, D2 and D8, in the large 28S rRNA molecule of the 60S ribosomal subunit, while the 18S rRNA molecule in the 40S ribosomal subunit remains unaffected (Houge et al., 1993). However, the pattern of rRNA cleavage occurring during apoptosis cannot be reproduced in necrotic

cells or when exposing cell lysates to random RNase activity, suggesting that it is specific to apoptotic cells. Recently it was demonstrated that rRNA and DNA cleavages can occur independently in apoptotic cells (Samali et al., 1997). In other words, the previously observed correlation is likely to be coincidental. The absence of apoptotic rRNA cleavage in some cell types suggests that this phenomenon is tightly regulated and unrelated to DNA fragmentation. Therefore it appears that the pattern of fragmentation seen during apoptosis is not part of a scheme for general macromolecular disintegration and degradation, but rather a trait present in only some cell types.

Protein Degradation The prerequisite for proteolysis in apoptosis is well documented. Several proteolytic activities were implicated in both the induction and execution steps of cell killing (for review see Zhivotovsky et al., 1997). Early evidence of a role for proteases in apoptosis came from studies on granule proteases (granzymes/fragmentins). These investigations identified granzymes involved in the exocytosis pathway of lymphocyte-mediated cytotoxicity, which are responsible for the lethal damage inflicted by these cells upon target cells (Shi et al., 1992). More recently, a family of aspartic acid–specific cysteine proteases have been discovered in mammalian cells, which share homology with the Caenorhabditis elegans death gene, ced-3 (Yuan et al., 1993). The newly adopted nomenclature for these enzymes, caspases, refers to the aspartic acid–specific cysteine protease activity specific to this family (Alnemri et al., 1996). At least thirteen of these proteases have now been purified and their genes cloned (for review see Zhivotovsky et al., 1997). The activation of the caspase family of proteases has been detected in numerous tissues and cell types and may function as a common pathway through which apoptotic mechanisms operate. Some procaspases, following translation, become autocatalytically cleaved at specific P1 aspartic acid residues, thereby generating a tetrameric cysteine protease holoenzyme that recognizes a characteristic cleavage motif (WXXD, DEXD, or XEXD) in target proteins (Nicholson and Thornberry, 1997). The effects of caspases in apoptosis appear to be accomplished by the cleavage of numerous proteins located in the cytoskeleton, cytoplasm, and nucleus, although the significance of these cleavages to the cell-death process is still unclear.




There have been a large number of reports on caspase involvement in apoptosis; however, there is yet very little evidence for the involvement of this family of proteases in necrosis. Recent work from our own group has shown that menadione-induced necrosis of hepatoma cells is independent of caspase activation (A. Samali et al, 1999). Furthermore, Tomaselli and co-workers (Armstrong et al., 1997) observed the activation of caspase 3 (CPP32/apopain) in cerebellar granule neurons undergoing apoptosis but not necrosis. These results appear to be in line with the work carried out in our laboratory which have shown that after the acute, necrotic death of cerebellar granule neurons exposed to glutamate, the remaining neurons undergo a delayed, apoptotic death (Ankarcrona et al., 1995). However, we did not detect any caspase activation during the “necrotic” step of our experimental model (M. Ankarcrona, B. Zhivotovsky, S. Orrenius, and P. Nicotera, unpub. observ.). Apart from the caspase family of proteases, a number of other proteolytic enzymes are also implicated in cell death. Studies from our laboratory have suggested the involvement of Ca2+dependent proteolytic activity in oxidant injury in the liver (Nicotera et al., 1986). Although the substrates for this protease activity during cell injury remain largely unidentified, it appears that cytoskeletal and membrane-integral proteins may be a major target for this proteolytic event during chemical toxicity (Mirabelli et al., 1989). Calpain, another Ca2+-dependent neutral protease and member of the papain family of cysteine proteases, degrades a number of key cellular proteins including proto-oncogenes, steroid-hormone receptors, protein kinases, and cytoskeletal proteins (Croal and DeMartino, 1991, Vanags et al., 1996). Calpain, one of the degradative nonlysosomal proteases, is also implicated in necrosis (Arai et al., 1990). Enhanced calpain activity has been observed in anoxic hepatocytes and neurons. Moreover, inhibition of calpain by acidosis, calpain protease inhibitors, and glycine delays anoxic injury. Precise mechanisms responsible for the stimulation of calpain activity during either necrosis or apoptosis are still unclear. More recently, it has been shown that hepatocellular carcinoma cells resist necrosis during anoxia by preventing phospholipase-mediated calpain activation (Arora et al., 1996). However, it is unknown whether sustained calpain activity promotes necrosis as a “deathase” by degrading key cytoskeletal proteins such as spectrin or by acting as a “signaling protease.” Therefore, a better

understanding of the protein substrates cleaved in necrosis will be required before the role of calpains in necrosis can be elucidated. Another example of noncaspase proteases that may be involved in apoptosis is seen in the plasminogen activator system, which is a major proteolytic complex responsible for the breakdown of the extracellular matrix. Expression of the plasminogen activator inhibitor type 2 (PAI2) in HeLa cells, which do not synthesize PAI-2, protects them from tumor-necrosis factor– (TNF-) induced apoptosis (Dickinson et al., 1995). Interestingly PAI-2 has a high degree of structural similarity to crmA, which inhibits interleukin 1β–converting enzyme– (ICE)–induced apoptosis (Dickinson et al., 1995). PAI-2 itself is not spared during apoptosis, and it loses its activity after cleavage by other proteases (Jensen et al., 1994). There are no reports about the involvement of this protease in necrosis. An alternative protease system, which is thought to play a role in cell death, is that of proteasomes. The contribution of proteasomal activity during apoptosis has primarily been evaluated by using inhibitors. There are several observations that show inhibition of proteasomal activity by “specific” peptide inhibitors either fails to prevent receptor-mediated apoptosis (B. Zhivotovsky, D.H. Burgess, I. Ares-Pörn, and S. Orrenius, unpub. observ.) or drives cells into apoptosis (Shinohara et al., 1996). Currently there is no evidence that these enzymes activate caspases. Thus, a conclusive role for this enzymatic system in apoptosis remains to be elucidated. The interaction between other enzyme activities, either proximal or distal to the cleavage and activation of the procaspases, and their role in apoptosis, is the subject of rapidly evolving research. The lytic nature of lysosomal enzymes and their release by various experimental procedures suggested long ago that they might be responsible for cell necrosis in damaged cells and tissues (for review see Bowen, 1981). DeDuve had already in the 1960s developed the concept of the lysosome as a “suicide bag” (deDuve, 1963). This hypothesis states that particle-bound acid hydrolase can be released into the surrounding cytoplasm under appropriate conditions and bring about the cell’s ultimate destruction. Such a response might occur during cellular injury, but lysosomes in fact are relatively stable organelles, which fragment and release their contents after cell death rather than before. Much attention was paid to acid hydrolases in the earlier publications. Hydro-


lases, however, were later shown to occur in association with the Golgi and/or Golgi–endoplasmic reticulum lysosomes, endoplasmic reticulum extracisternal spaces, and plasma membranes (for review see Bowen, 1981). In reference to inflammatory cells, it has been shown that acid hydrolases are directly transferred from the endoplasmic reticulum into foci of cytoplasmic degeneration (Van Lancker, 1975). It was later concluded that when cells die, areas of the cytoplasm become segregated for autolysis, and that these areas are supplied with hydrolases, either from primary lysosomes or directly from the endoplasmic reticulum. Furthermore, an accumulation of free acid phosphatases around ribosomes in the extracisternal space of the endoplasmic reticulum has been linked to this type of cell death (for review see Bowen, 1981). This early extracisternal buildup appears as a prelude to cell autolysis. On the other hand, histochemical studies showed that the liberation of acid phosphatases and esterases was not an early change (Kerr et al., 1972), and the authors of that study suggested that the release of lysosomal enzymes is not responsible for the initiation of necrotic cell death, but rather accompanies this type of death. There is no evidence for preferable degradation of any proteins by lysosomal proteases during necrosis. Thus, in contrast to apoptosis, the involvement of both lysosomal and nonlysosomal protease activities in necrosis was documented. However, the precise role of these protease activities in either induction or execution of necrosis, as well as the specific cleavage of cellular proteins, remain to be elucidated.

CELLULAR SIGNALING DURING CELL DEATH Signal transduction is thought to play a key role in the onset of both apoptosis and necrosis, and this may be mediated by an increase in intracellular Ca2+ levels, protein kinase C (PKC), cyclic adenosine monophosphate/protein kinase A (cAMP/PKA), or phosphatases. It has been demonstrated that glucocorticoidinduced cell death involves Ca2+ influx in lymphocytes and that this type of cell death can be mimicked using calcium ionophores (Kaiser and Edelman, 1978). Based on these observations it appears that intracellular Ca2+ levels may initiate a death signal, probably through the activation of Ca2+-dependent enzymes (proteases, phospholipases, and endonucleases) in this system. The evidence for a role of Ca2+ in necrotic cell killing is particularly strong in the

central nervous system. Ca2+ appears to mediate the neurotoxicity of cyanide, such heavy metals as lead and mercury, and organotin compounds (for review see Nicotera and Orrenius, 1996). Furthermore, intracellular Ca2+ overload, resulting from excessive stimulation of excitatory amino acid receptors and enhanced Ca2+ influx through membrane channels, appears to play an important role in ischemic brain damage. Much recent research has focused on glutamate-induced excitotoxicity and its contribution to brain damage in various diseases (Choi, 1992; Ankarcrona et al., 1995). The calcium ion plays a critical role in this process, and intracellular Ca2+ overload appears to mediate the lethal effect of N-methyl-D-aspartate (NMDA) receptor overactivation. This mechanism is responsible not only for the brain damage induced by certain neurotoxins, but also for excitotoxicity, which is strongly implicated in neuronal death following insults such as ischemia and trauma. Thus, NMDA receptor antagonist not only blocks Ca2+ influx and neuronal death elicited by glutamate or NMDA in vitro, but can also reduce the volume of infarction produced by focal ischemia in vivo. As discussed above, glutamate can trigger the onset of both apoptosis and necrosis in cerebellar granule cells (Ankarcrona et al., 1995). Radical scavengers and agents that inhibit the generation of nitric oxide by nitric oxide synthase were ineffective in preventing cell death in this system, whereas NMDA receptor/channel blockers prevented both necrosis and apoptosis in glutamate-treated cerebellar granule cells. These data suggest that Ca2+ overload–mediated NMDA-receptor-operated channels are sufficient to induce either necrosis or apoptosis in cerebellar granule neurons in vivo. It is clear that intracellular Ca2+ overload is an important factor in various in vitro models, and results from clinical studies appear to support this hypothesis. The Ca2+ overload seems to result from increased Ca2+ influx through both receptor-operated and L-type Ca2+ channels, as indicated by the neuroprotective effects of glutamate-receptor antagonists and L-type channel blockers. Various Ca2+-dependent degradative processes have been found to contribute to Ca2+-mediated necrotic cell killing in in vitro studies, and, although their relative contribution is unclear, it appears that perturbation of cytoskeletal organization, impairment of mitochondrial function, and/or activation of certain proteases may be of particular importance. For example, it has been proposed that activation of phospholipase A2 by a sustained increase in




cytosolic calcium plays an important role in necrotic cell killing. Support for this hypothesis comes from several observations that necrosis in the liver and heart caused by ischemia, as well as in hepatocellular carcinoma cells in anoxic condition, is prevented by inhibitors of this enzyme or by the inhibition of calpain activation (for review see Nicotera and Orrenius, 1996). Thus, it is likely that the calcium ion may play a determinant role in the necrotic killing process. One of the classic signaling pathways is that of PKC, which is a multifunctional serine/threonine kinase that utilizes diacylglycerol as a second messenger (Nishizuka, 1984). Twelve different isoforms of PKC have been identified to date and are classified according to their calcium dependence and phorbol ester binding activity (for review see Lavin et al., 1996). There are some reports as to the involvement of PKC in apoptosis. For example, it has been reported that treatment of mouse thymocytes with phorbol esters, which activate PKC, induces apoptosis (Kizaki et al., 1989). Similarly, apoptosis induced by the calcium ionophore A23187 can be inhibited by the PKC inhibitor H7. However, most of the evidence to date indicates that PKC activation inhibits apoptosis (McConkey et al., 1996). This paradox may be explained by the large number of isoforms, which may be differentially regulated during apoptosis (Forbes et al., 1992). Similarly, the role of protein phosphatases during apoptosis is not clear. There are some reports that a number of cell lines react to okadaic acid by undergoing apoptosis (Boe et al., 1991), while there are other reports that the inhibition of protein phosphatase activity blocks apoptosis in some other cell lines (Baxter and Lavin, 1992). From these reports it appears that protein activation/modification, through reversible protein phosphorylation by kinases and protein phosphatases rather than by de novo protein synthesis, may play a more central role in the regulation of apoptosis in some cells. Receptor-mediated killing involves activation of a cascade of signaling events leading to activation of caspases. For example, apoptosis induced by ligation of the CD95 (Fas/Apo-1) receptor results in association of several cytotoxicity-dependent Apo-1-associated proteins (CAPs; Trauth et al., 1989). The aggregated receptor, together with the CAP proteins (1 to 4) form the death-inducing signaling complex or DISC. DISC formation upon engagement of the receptor is essential for CD95-induced

apoptosis. CAP1/2 were identified as the serine phosphorylated Fas-associated death domain (FADD) and CAP3/4 as Fas-like (FLICE) or caspase-8, pro-enzyme and activated enzyme (for review see Wallach, 1997).

ROLE OF MITOCHONDRIA DURING CELL DEATH Work from several laboratories has indicated that mitochondrial damage may be a common event in the development of cell injury caused by various toxic agents (Nicotera and Orrenius, 1996). It was postulated that mitochondrial damage is initially manifested by a decrease in the mitochondrial membrane potential followed by ATP depletion (Zamzami et al., 1995). During necrosis, these changes occur irreversibly and usually lead to disruption of the mitochondrial structure. These changes set necrosis aside from apoptosis, during which mitochondrial structure remains morphologically intact. Mitochondria have nevertheless been implicated in apoptosis ever since the discovery that the Bcl-2 protein localizes to the outer mitochondrial membrane (Hockenbery et al., 1990). Moreover, by using different cellfree systems, it has been shown that nuclear apoptosis depends on the presence of ATP (Lazebnik et al., 1993; Newmeyer et al., 1994; Kass et al., 1996). It has been demonstrated that cytochrome c can induce cleavage and activation of procaspase 3 in cytochrome c–minus cytoplasmic extracts (Liu et al., 1996). It has also been shown that cytochrome c is released from the mitochondrial intermembrane space into the cytosol by cells undergoing apoptosis. In this case cytochrome c release is not preceded by changes in mitochondrial membrane potential (Kluck et al., 1997). Two groups have presented evidence that a high ATP level is required during the apoptotic process and suggested that the level of intracellular ATP determines whether a cell will die by apoptosis or necrosis (Nicotera and Leist, 1997; Tsugimoto, 1997). During exposure to glutamate, both mitochondrial membrane potential and ATP levels decline in many neurons (Ankarcrona et al., 1995). In neurons with irreversibly dissipated mitochondrial potentials, necrosis rapidly ensued. The surviving population recovered both mitochondrial membrane potential and energy levels and subsequently underwent delayed apoptosis. It was postulated that the loss of energy and the onset of rapid necrosis simply prevents the activation of the default apoptotic program. This postulate was supported by the observation that treatment of


cerebellar granule cells with a combination of glutamate and the irreversible mitochondrial uncoupler carbonyl cyanide m-chlorophenylhydrazone (CCCP) resulted in necrosis for most of the neuronal population, rather than in the delayed apoptosis observed with glutamate alone. There is abundant evidence for an ATP requirement in several apoptosis models, although the critical ATP-dependent steps in the process have not been characterized in detail. They are likely to be linked to both the signaling and the execution phases of apoptosis. It was suggested (Tsugimoto, 1997) that the caspase protease cascade is regulated by ATP-dependent reactions and that the protease cascade might not proceed in an autonomous fashion. In addition, it has been shown (Yasuhara et al., 1997) that the active nuclear transport mechanism, which requires ATP hydrolysis, is involved in the apoptotic changes of the nuclei. It is likely that an another ATP-dependent step or steps downstream from caspase activation is important for perpetuation of the apoptotic process. This fact could explain the previously published data on the ATP requirement for induction of nuclear apoptosis, induced by different cytoplasmic extracts (Lazebnik et al., 1993; Newmeyer et al., 1994; Kass et al., 1996). Thus, it is not surprising that the depletion of cellular ATP blocks various events in apoptosis. The question that remains to be answered is whether, and how, ATP depletion redirects the death process towards necrosis when cells have been triggered with apoptotic stimuli, such as anti-CD95 antibodies. In addition to the possible requirement of ATP for caspase activation, redirection could also be explained by an unknown mechanism, which shortens the survival time of cells that would eventually die by necrosis resulting from energy deprivation. The next important question that remains to be resolved is an apparent contradiction between, on the one hand, the obligatory induction of the mitochondrial permeability transition associated with a cessation of mitochondrial ATP synthesis, and, on the other hand, a need for maintenance of the intracellular ATP level during development of apoptosis, which may take many hours or days. In addition, hyperproduction of reactive oxygen species (ROS) results from the loss of mitochondrial membrane potential, yet the cell needs to maintain a reducing environment for optimal caspase activity. These controversies could be resolved if the proposed apoptosis-induced changes were restricted to a small subset of

mitochondria in the cell. Maintenance of the energy level required by cells to undergo apoptosis could occur solely by glycolysis. This would be consistent with the observation by Jacobson et al. (1993) that cells lacking mitochondrial DNA and a functional respiratory chain can still undergo apoptosis. It is very likely that the mitochondrial permeability transition associated with a cessation of mitochondrial ATP plays an important role in necrotic cell death, whereas leakage of cytochrome c from morphologically intact mitochondria via a yet unknown and more selective pathway than outer-membrane rupture following permeability transition–induced mitochondrial swelling plays an important role in apoptotic cell death.

GENETIC MODULATION OF APOPTOTIC CELL DEATH In recent years, enough evidence has been gathered to suggest a role for the protein products of a number of oncogenes and tumor-suppressor genes, as well as stress proteins and acute phase proteins, in the modulation cell death. These can be grouped into genes whose products are positive or negative regulators of cell death. The genes with a key role in the modulation of cell death and their effects on both apoptosis and necrosis are reviewed below.

c-myc The c-myc proto-oncogene is a short-lived nuclear protein and a member of a family closely related genes that also include v-myc, n-myc, and b-myc. The Myc family plays a key role in the regulation of cell proliferation (for review see Ryan and Brinie, 1996). It has been demonstrated that c-myc is capable of driving apoptosis (Evan et al., 1992). For instance, when T cell hybridomas were stimulated through their CD3/T cell receptor, they rapidly underwent apoptosis that could be blocked using an antisense oligonucleotide to c-myc mRNA. Furthermore, in fibroblasts, c-myc overexpression can induce apoptosis (Evan et al., 1992). The decision to proliferate or die in cells overexpressing c-myc seems to depend on the presence or absence of extracellular survival factors.

p53 The p53 protein is the product of a tumorsuppressor gene whose loss of function occurs in over half of all human tumors, thus implicating p53 inactivation in tumorigenesis. Nevertheless, the main functions of p53 appear to be in mediating the cellular response to DNA dam-



age and maintaining genomic stability (Kastan et al., 1991). Many forms of genotoxic stress induce a rapid increase in p53 protein levels, through both stabilization of the protein (Kastan et al., 1991) and an increase in p53 mRNA levels (Sun et al., 1995). p53 induces growth suppression by regulating cell-cycle arrest and/or inducing apoptosis. Reintroduction and overexpression of wild-type p53 in cells that have lost endogenous p53 function may induce apoptosis (Yonish-Rouach et al., 1991). In this regard, p53 is shown to be required for both irradiation-induced G1 arrest and apoptosis in several cell systems (Lowe et al., 1993). Induction of growth arrest by p53 has been shown to be transcription activation–dependent and appears to involve induction of p21 (Waf1/Cip/sdi1) and growth arrest and DNA damage-inducible gene 45 (Gadd45). For review see YonishRouach (1996). Although transcriptional activation appears to be involved in the induction of growth arrest, little is known about the possible downstream elements in this apoptotic pathway. p53 has been shown to induce apoptosis in the presence of transcriptional activation, in which case p53 may induce expression of Bax (Miyashita and Reed, 1995) and/or CD95 (Owen-Schaub et al., 1995). However, recent reports suggest that transcriptional activation is not essential for the induction and progression of p53-induced apoptosis (for review see Yonish-Rouach, 1996).

Bcl-2 family


This is a family of genes in which many members hold key positions in the apoptotic machinery. The proteins in this family share at least two highly conserved regions, which have been referred to as Bcl-2 homology 1 and 2 (BH1 and BH2) domains (Oltvai et al., 1993). Bcl-2 was the first member of this family to be identified in follicular B cell lymphomas (Bakhshi et al., 1985). Several other family members, which were discovered later, may function to inhibit apoptosis, including Bcl-2, Bcl-xL, Bcl-W, Bfl-1, A1, and MCL-1, while others function to accelerate the rate of apoptosis once it has been induced, including Bax, Bcl-xs, Bag, Bik, Bim, Bak, Bar, and Bad. Bcl-2 protein is localized to the mitochondria, endoplasmic reticulum, and nuclear membrane (Hockenbery et al., 1990). Bcl-2 appears to have antioxidant properties, as it can inhibit apoptosis induced by reactive oxygen species (Hockenbery et al., 1993). However, it has also been demonstrated that Bcl-2 can inhibit cell death under conditions that preclude the forma-

tion of reactive oxygen species (Jackobson et al., 1993). Therefore, the ability of Bcl-2 to protect against the damaging effects of free radicals does not appear to completely account for the suppression of cell death by Bcl-2. Bax is more ubiquitously expressed than Bcl-2 (Krajewski et al., 1994). The bax gene promoter contains four p53-binding sites, and it has been shown that the levels of its protein can be up-regulated by p53 (Miyashita and Reed, 1995). It is suggested that the ability of Bcl-2 to suppress apoptosis depends on the relative proportion of Bcl-2 homodimers and Bcl-2/Bax. In addition to Bcl-2, Bax can also heterodimerize with other Bcl-2-related proteins, including Bcl-xL, Bcl-xS, MCL-1, and A1, as shown by the yeast two-hybrid assay (for review see Reed, 1997). These reports suggest that Bcl-2 family members function in part through protein-protein interactions, and that susceptibility to apoptosis is determined by multiple competing dimerizations in which Bax is a common partner. However, the ultimate mechanism by which members of this family may induce or suppress apoptosis is not understood. It has been shown recently that overexpression of Bcl-2 can also inhibit necrosis at least in neuronal cells when induced by such diverse stimuli as viruses, hypoxia, oxidative stress, or exposure to toxicants (Lezoulac’h et al., 1996; Shimizu et al., 1996). Whether this effect is cell-specific remains unclear. The mechanism of action of Bcl-2 in cells undergoing necrosis may simply involve antioxidant activity. The second possibility is that Bcl-2 prevents the consequences of mitochondrial dysfunction (collapse of the mitochondrial transmembrane potential, uncoupling of the respiratory chain, hyperproduction of superoxide anions, disruption of mitochondrial biogenesis, outflow of matrix calcium and glutathione, and release of soluble intermembrane proteins), all of which can lead to a bioenergetic catastrophe culminating in the disruption of plasma membrane integrity which is specific for necrosis (Kroemer, 1997). However, more work should be done to justify this hypothesis and to understand the mechanism of Bcl-2-induced protection from necrosis.

Ras Members of the Ras family of proteins play an essential role in the control of normal cell growth and may induce transformation. Certain evidence has led a number of groups to advocate that Ras, in addition to inducing transformation, can also prevent cell death. Overex-


pression of the ras gene is shown to inhibit apoptosis in a number of cases (Wyllie et al., 1987; Arends et al., 1993). Ras appears to be a focal point for the convergence of many signaling pathways, especially during receptor/ligand interaction, such as epidermal growth factor (EGF), fibroblast growth factor (FGF) and nerve growth factor (NGF), T cell receptor–CD3 (TCR-CD3), and CD95. The question as to which protein downstream of the pathway is the key effector of the anti-apoptotic signal through Ras is unclear, but it may involve the activation of MAP kinase or phosphotidylinositol 3 kinase or both.

Stress Proteins The stress proteins are a set of evolutionarily conserved proteins that are synthesized or activated in response to stress (for review see Lindquist and Craig, 1988). The main function of these proteins is to afford protection to cells in times of stress. When the stress element is removed, these cells continue to function normally and the levels drop back to the normal. Heat-shock proteins (HSPs), a subgroup of stress proteins, were first demonstrated to be induced as a response to elevated temperatures. There is strong evidence suggesting that the induction of HSPs coincides with the acquisition of tolerance to higher doses of stress, which may otherwise be lethal to the cell. In this regard, it was demonstrated that a mild heat shock induces the rapid synthesis of a number of HSPs (Jaattela et al., 1992; Mosser and Martin, 1992) and that heat-shocked or thermotolerant cells demonstrated a greater degree of resistance to environmental stress (Jaattela et al., 1992), probably by resisting apoptosis (Samali and Cotter, 1996). Furthermore, it was demonstrated that thermotolerant or HSP-overexpressing cells are more resistant to cell death induced by hyperthermia or growth-factor withdrawal (Mailhos et al., 1993). This may suggest a possible role for HSPs in the resistance mechanism. Furthermore, HSPs are also implicated in drug resistance (McClean and Hill, 1992), and heat-shocked or HSP-overexpressing cells become more resistant to the cytotoxic effects of some anticancer drugs (Oesterreich et al., 1993) that are otherwise capable of inducing both apoptosis and necrosis (Martin et al., 1990).

CONCLUSIONS Cell toxicity leading to cell death may occur via apoptosis or necrosis. The mode of cell death depends on the nature of the toxic agent,

but in extreme situations the final outcome is cell disruption and disintegration. In other words, necrosis is the acute type of cell death or the cellular response to extreme toxic injury. During necrosis, chromatin adopts a highly flocculated form, and the DNA from these cells is digested randomly to give a smear when analyzed by agarose gel electrophoresis. The main feature of necrosis is an increase in cell volume. The rapid increase in cell volume results in membrane rupture and cell lysis (for review see Trump and Berezesky, 1995). After two decades of research and development in the field of cell death, it is now universally accepted that apoptosis is an essential strategy for maintaining dynamic balance and equilibrium of living systems, and is observed to occur as a normal mechanism in development and homeostasis, as well as an altruistic mode of toxic cell death. Classification of cell death is now based on morphological and biochemical criteria, their circumstantial occurrence, or a combination of both. Although morphological characterization of apoptosis and the features distinguishing it from necrosis have been well documented, progress in our understanding of the mechanisms underlying the process has been quite slow. There is now much evidence that classic apoptosis and necrosis represent only definitive endpoints for these two processes. However, both types of cell death can occur either simultaneously or sequentially in the same cell suspension or tissues depending on type and/or dose of toxic insult. This suggests that some early biochemical and morphological events might be similar or even identical in both types of cell death. If this is the case, different downstream events might be important in determining whether the cell will undergo apoptosis or necrosis. It could also be that after intensive toxic treatment, the cellular apoptotic machinery becomes suppressed, after which necrosis would predominate. The events summarized in Table 2.1.1 can be observed at different levels of cellular organization. Thus, both physiological or pathological stimuli can induce apoptosis, but only pathological injury leads to necrosis. Apoptosis, in contrast to necrosis, occurs in individual cells in an asynchronous fashion without damaging neighboring cells. Both processes can be protected by certain inhibitors before a point of no return is reached. Once the cascade of caspase proteolytic activities becomes active, apoptosis cannot be completely reversed. The point of no return during necrosis remains to be elucidated. Neverthe-




less, cells can be rescued from necrosis as well as from apoptosis. Cellular shrinkage is a hallmark of apoptosis and is seen in many if not all cell types that undergo this type of death. Cell-volume regulation is a complex phenomenon, involving the homeostasis of intracellular ions such as chloride, sodium, and potassium. As the most abundant intracellular ion, K+ has been implicated in several of the regulatory mechanisms governing cell volume, including regulatory volume decrease. It has been shown that intracellular K+ content decreases in cells undergoing apoptosis induced either with etoposide or dexamethasone (Barbiero et al., 1995). Work in the authors’ laboratory has shown a potential link between glutathione (GSH) efflux and apoptotic cell shrinkage (Nobel, 1997). Inhibitors of apoptotic GSH efflux were able to inhibit cell shrinkage in thymocytes induced to undergo apoptosis by etoposide without affecting DNA fragmentation. Moreover, the GSH efflux was accompanied by K+ efflux in Jurkat cells treated with anti-CD95 antibody. The K+ efflux was not blocked either by the inhibitor of GSH efflux or by any known inhibitors of K+ channels and cotransporters tested. Concomitant with apoptotic K+ efflux, significant inhibition of Na+/K+-ATPase activity was observed. Inhibition of this enzyme by ouabain caused a K+ efflux of similar magnitude to that induced by anti-CD95 antibody, and it potentiated CD95mediated cell shrinkage. This suggested that with Na+/K+-ATPase activation, K+ and GSH effluxes facilitated apoptotic cell shrinkage, which was followed by the formation of apoptotic bodies. In contrast, necrotic cells showed extensive swelling, which resulted in subsequent disintegration of cells. Apoptotic cells lose adhesion at the very early stages, while necrotic cells stay adherent together until very late stages. As mentioned before, individual apoptotic cells can be efficiently recognized either by neighboring cells or professional macrophages and can be removed by phagocytosis. This step of the apoptotic process involves rearrangements of and dramatic changes in the plasma membrane. Removal of the dead-cell fragments before their plasma membranes are lysed has important implications for other cells within the tissues. It is assumed that apoptosis occurs without inflammation. Lysis of plasma membranes in necrotic cells seems to precede the removal of these remnants, and exudative inflammation is a common event during necrosis.

The significance of apoptosis is based on the fact that apoptotic cells tend to be “environmentally friendly” and package their contents into membrane-bound vesicles ready for ingestion by phagocytic cells, without releasing their contents into the intercellular matrix. Hence there is no inflammatory response. Apoptosis is also an altruistic cell death, in that damaged or injured cells commit suicide to allow the neighboring cells to continue to proliferate without being affected by the death of the neighbor. In addition, sacrifice of individual abnormal cells benefits the whole organism. Most if not all mechanisms leading to cell death involve molecular damage that is potentially reversible by cellular repair mechanisms. Therefore, if the repair mechanisms operate effectively, they may prevent cell death. It could be that in cells undergoing apoptosis, the cellrepair machinery is repressed by specific signals, which program cells to die. On the other hand, in cells undergoing necrosis this machinery might be directly suppressed by extremely strong insult (high doses of toxicants or irradiation). Since apoptosis is a gene-directed process and many genes are involved in the regulation of this multistage action, the repression of repair processes can also be regulated on this level. There is no evidence for expression of any specific genes during necrosis, which is why it is difficult to discuss the possible requirement of gene regulation for necrotic cell death. Thus, it seems that classic necrosis is a passive process. An inhibition of the activity of a number of gene products involved in apoptosis can switch the death pathway from apoptosis to necrosis. Also rapid changes in the cellular ATP levels, discussed above, and an imbalance in the ratio of nitric oxide and superoxide, can switch the form of cell death. The search on both molecular and biochemical levels for mechanisms of apoptosis and necrosis, along with attempts to understand the differences/similarities between these two types of cellular response to toxicants, are currently the hottest topics in biological research.

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Contributed by Afshin Samali, Boris Zhivotovsky, and Sten Orrenius Karolinska Institutet Stockholm, Sweden



Determination of Apoptosis and Necrosis

UNIT 2.2

Cell death may occur by two mechanisms: apoptosis, or programmed cell death, and necrosis, or cell death due to injury or trauma (see UNIT 2.1). Both types of cell death have their own specific and distinct morphological and biochemical hallmarks. Apoptotic cells share a number of common features, such as phosphatidylserine (PS) exposure, cell shrinkage, chromatin cleavage, nuclear condensation, and formation of pyknotic bodies of condensed chromatin. Necrotic cells exhibit nuclear swelling, chromatin flocculation, loss of nuclear basophilia, breakdown of cytoplasmic structure and organelle function, and cytolysis by swelling. Cell death can be induced by a wide variety of stimuli, such as growth factor withdrawal, heat shock, cold shock, radiation, heavy metals, genotoxic drugs, and a number of biological ligands such as Fas-L and tumor necrosis factor (TNF). Most if not all of these can induce both apoptosis and necrosis in a time- and dose-dependent manner. A variety of techniques have been developed to assess cytotoxicity in untreated and/or toxicant-treated cells, most of which are based on the loss of plasma membrane integrity. Membrane disruption can be detected based on the uptake of vital dyes (see Basic Protocol 1) or the release of cellular contents into the extracellular milieu (see Basic Protocol 5). However, during apoptosis, the cell membrane remains intact for a relatively long time, and therefore some of these assays cannot detect the early stages of apoptosis. Alternatively, assays that detect all types of cell death do not discriminate between apoptosis and necrosis. Therefore, more specific techniques have been developed to determine cell death, and the combination of several methods is required to distinguish between the two separate types. These techniques rely on specific morphological and molecular or biochemical changes associated with these two processes. This unit describes some of the techniques most commonly used to detect cell death. Morphological assays include trypan blue exclusion (see Basic Protocol 1), differential staining (see Basic Protocol 2), Hoechst staining (see Basic Protocol 3), flow cytometry to measure cell size and DNA content (see Basic Protocol 4), and annexin V binding to detect externalized phosphatidylserine (see BAsic Protocol 5). Methods to detect chromatin cleavage include TUNEL assays for whole cells (see Basic Protocol 6) and paraffin sections (see Alternate Protocol 1), DNA fragmentation assays using whole cells (see Basic Protocol 7), assays of total genomic DNA (see Alternate Protocol 2), analysis of DNA fragmentation by agarose gel electrophoresis (see Alternate Protocol 3), phenol extraction of DNA for analysis of fragmentation (see Alternate Protocol 4), a quantitative assay for DNA fragmentation (see Basic Protocol 8), and detection of DNA fragmentation by pulsed-field gel electrophoresis (see Basic Protocol 9). Caspase activity is assessed by detecting proteolytic cleavage of fluorescent substrates (see Basic Protocol 10) or by immunodetection (see Basic Protocol 11). Protocols are also provided for Cytospin preparations from cell suspensions (see Support Protocol 1) and preparation of SDS-polycrylamide minigels (see Support Protocol 2). Table 2.2.1 describes how these assays are used to distinguish apoptotic cells from necrotic cells. MORPHOLOGY ASSAYS Loss of membrane integrity, which occurs late in apoptosis and relatively early in necrosis, can be detected by cellular uptake of the vital dye trypan blue (see Basic Protocol 1). Apoptosis and necrosis also can be identified morphologically based on the criteria described elsewhere (Bowen, 1980). These include cell shrinkage, nuclear condensation, and cleavage for apoptotic cells; and nuclear swelling, chromatin flocculation, and loss Contributed by Boris Zhivotovsky, Afshin Samali, and Sten Orrenius Current Protocols in Toxicology (1999) 2.2.1-2.2.34 Copyright © 1999 by John Wiley & Sons, Inc.


2.2.1

Table 2.2.1

Methods for Determination of Apoptosis and Necrosis

Method

Protocol

Morphological/cytological analysis Trypan blue staining Basic Protocol 1 Differential staining

Basic Protocol 2

Hoechst staining

Basic Protocol 3

Flow cytometry

Basic Protocol 4

Annexin V binding and PI staining

Basic Protocol 5

Assays for chromatin cleavage TUNEL assay Basic Protocol 6 and Alternate Protocol 1 Whole-cell DNA Basic Protocol 7 fragmentation Total genomic DNA Alternate Protocol 2 fragmentation Simple protocol for DNA Alternate Protocol 3 fragmentation Phenol extraction for DNA Alternate Protocol 4 fragmentation Quantitative assay of DNA Basic Protocol 8 fragmentation Basic Protocol 9 Pulsed-field detection of high-molecular-weight DNA Assays for caspase activation Specific substrate cleavage Basic Protocol 10 assay Immunodetection of Basic Protocol 11 caspase activation

Result for Apoptosis No staining (early apoptosis) Blue staining (late apoptosis) Membrane blebbing, chromosome condensation and nuclear shrinkage, cytoplasmic constriction and loss of cell volume Increased blue fluorescence, fragmented or condensed nuclei Reduced forward and side scatter Positive fluorescence response for annexin V binding; exclusion of PI

Necrosis Blue cells Nuclear swelling, chromatin flocculation, membrane blebbing and disruption, appearance of cell “ghosts” Blue nucleus with diffuse staining Increased forward and side scatter Positive fluorescence; uptake of PI

Detection of strand breaks

Detection of strand breaks

DNA ladder

DNA smear

DNA ladder

DNA smear

DNA ladder

DNA smear

DNA ladder

DNA smear

Increase in the percentage of fragmented DNA High-molecular-weight bands

Increase in the percentage of fragmented DNA DNA smear

Positive detection of cleaved fluorescent substrate Positive detection of cleavage products of procaspase

No changes in fluorescence detected No cleavage products detected

of nuclear basophilia for necrotic cells. The levels of both apoptosis and necrosis in a particular cell population can be estimated from Cytospin preparations (see Support Protocol 1) stained with RAPI-DIFF (see Basic Protocol 2 Figure 2.2.1), Hoechst stain (see Basic Protocol 3), or propidium iodide (see Basic Protocol 4). BASIC PROTOCOL 1


Measurement of Cell Death by Trypan Blue Exclusion This common cell viability assay is based on the ability of a cell with an intact membrane to exclude the dye trypan blue. Therefore, this assay allows one to distinguish between cells with intact and disrupted membranes. Since this method does not give an indication of the mode of cell death, it should be used only in conjuction with a more informative morphological method.


A

control

B

apoptosis

C

necrosis

Figure 2.2.1 RAPI-DIFF staining of Cytospin preparation of HL-60 cells. Cells were treated with different concentrations of a cytotoxic drug to induce apoptosis or necrosis. (A) Untreated cells, (B) apoptotic cells, and (C) necrotic cells.

Materials Cell suspension to be assessed 2× PBS tablets, pH 7.2 to 7.4 (Sigma) or 2× PBS (APPENDIX 2A) 0.2% (w/v) trypan blue (Sigma) in 2× PBS (store up to 1 to 2 months at 4°C) Hemacytometer: improved Neubauer type (Karl Hecht; Baxter) or equivalent Coverslips (e.g., Chance Propper) Light microscope 1. Remove from the cell suspension a sample containing ∼5 × 104 cells. 2. Add an equal volume of 0.2% trypan blue, mix, and incubate 1 to 2 min at room temperature to permit dye uptake. 3. Load samples onto the hemacytometer. 4. Count the total number of cells and the number of unstained cells in five of the major sections of the hemacytometer. Calculate the average number of cells per section. 5. Calculate the number of cells/ml culture medium by multiplying the average number of cells per section by the dilution factor (2 in this case) and hemacytometer index (104). Determine the percentage viability according to the following formula: % viability =

number of unstained cells × 100 total number of cells

Control cells and cells in early stages of apoptosis exclude trypan blue; cells in the late stages of apoptosis and necrotic cells take up the dye and appear as blue cells.

Differential Staining of Cells This staining procedure involves three steps: first cells are fixed in 100% methanol, then the nuclei are stained with an acid dye, and finally the cytoplasm is stained using a basic dye. Such three-step procedure allows differential staining and contrast between the cytoplasm and the nucleus. Apoptotic cells exhibit membrane blebbing, chromatin condensation and nuclear shrinkage, cytoplasmic constriction and loss of cell volume, and formation of apoptotic bodies. Necrotic cells undergo nuclear swelling, chromatin flocculation, cell membrane blebbing and disruption, and finally cell lysis resulting in the appearance of “ghost cells” (see Figure 2.2.1). This method can also be applied to cells growing attached to coverslips or chambered culture slides.

BASIC PROTOCOL 2



Materials Cytospin preparations of cells (see Support Protocol 1) 100% methanol Acid dye: 0.1% (w/v) eosin Y/0.1% (w/v) formaldehyde/0.4% (w/v) sodium phosphate dibasic/0.5% potassium phosphate monobasic Basic dye: 0.4% (w/v) methylene blue−polychromed/0.4% (w/v) azure/0.4% (w/v) sodium phosphate dibasic/0.5% (w/v) monobasic potassium phosphate DPX mountant (in solution; BDH) Coverslips (e.g., Chance Propper) Microscope slides (e.g., Menzel-Glaser) Light microscope 1. Fix cells after Cytospin preparation (see Support Protocol 1) by immersing the slide ten times in 100% methanol. The three solutions required for this procedure are available as a kit, called RAPI-DIFF Stain kit, from Diagnostic Developments. The solutions can be stored 1 to 2 months at room temperature.

2. Stain the cell nuclei by dipping the slide in the acid dye ten times. 3. Stain the cell cytoplasm by dipping the slide in basic dye ten times. 4. Rinse the slide by dipping in distilled water, let it air dry, and mount it using DPX mountant. 5. Score the cells as normal, apoptotic, condensed and/or fragmented nuclei, or necrotic swollen cells, based on their morphological appearance under a light microscope (see UNIT 2.1 and Fig. 2.2.1). BASIC PROTOCOL 3

Hoechst Staining of Cells Hoechst 33342, a bisbenzimide dye, is a cell-permeant, minor group−binding DNA stain that fluoresces bright blue upon binding to DNA. It is water soluble and relatively nontoxic. Hoechst 33342 can be excited with the UV lines of the argon-ion laser and most conventional fluorescence excitation sources, and it exhibits relatively large Stokes shifts (excitation and emission maxima ∼350 and 460 nm), making it suitable for multicolor labeling experiments (Pollak and Ciancio, 1990). Cells are scored as apoptotic if they have fragmented nuclei. This method can also be applied to cells growing attached to coverslips or chambered culture slides. Materials Cell suspension 1× and 2× PBS, pH 7.2 to 7.4 (from Sigma 2× PBS tablets, or see APPENDIX 2A) 4% (w/v) paraformaldehyde (see recipe) 10 µg/ml Hoechst 33342 dye (Molecular Probes) in PBS (APPENDIX 2A) 50/50 (v/v) PBS/glycerol Microscope slides (e.g., Menzel-Glaser) Coverslips (e.g., Chance Propper) Fluorescent microscope Additional reagents and equipment for Cytospin preparations (see Support Protocol 1)


1. Place an aliquot of cell suspension containing 0.3−0.5 × 106 cells in a microcentrifuge tube, and centrifuge 5 min at 1000 × g, 4°C.


2. Remove and discard supernatant. Resuspend the dry pellet in 30 to 50 µl of 4% paraformaldehyde. 3. Spin the fixed cells onto a glass slide using a Shandon/Lipshaw Cytospin centrifuge (see Support Protocol 1). Alternatively, spread the cells onto a slide with a pipet tip, and air dry slide. 4. Stain slide with 10 µg/ml Hoechst dye. Hoechst 33342 dye can be diluted from 100× stock kept in the dark at 4°C.

5. Leave slide in the dark for 10 min at room temperature. 6. Rinse by dipping slide five times in distilled water, and let it air dry in the dark. 7. Mount slide with coverslip using PBS/glycerol. Monitor fluorescence with a blue filter in the fluorescence microscope, and score cells with fragmented nuclei as apoptotic. This method is useful for discrimination of apoptotic from nonapoptotic (untreated or necrotic) cells.

Cytospin Preparation of Cells for Analysis Cytospin preparations can be used in many protocols for cell biology. Once slides are prepared they can be stained with variety of dyes and analyzed using light, fluorescent, and confocal microscopy, depending on the dye type. Before staining, slides can be stored up to 1 year at −20°C covered with aluminum foil. Fixed and stained slides can be stored indefinitely once mounted permanently.

SUPPORT PROTOCOL 1

Materials Cell suspension PBS (APPENDIX 2A) Cytospin centrifuge and cups (Shandon/Lipshaw) 1. Place a sample of cell suspension containing ∼0.5−1 × 105 cells in a microcentrifuge tube, and centrifuge 5 min at 1000 × g, 4°C. 2. Remove and discard supernatant. Resuspend pelleted cells in 100 µl PBS. 3. Add suspension to the Cytospin cup and slide setup, and centrifuge 2 min at 500 × g, 4°C. 4. Air dry the slide. Flow Cytometric Analysis to Detect Apoptotic Cells Many studies report that a reduction in cell size and buoyant density accompany apoptotic cell death (Yamada and Ohyama, 1980); thus, apoptotic cells appear smaller and more dense than their normal counterparts. These changes are accurately detected in a majority of cells by their light-scattering properties as measured by a flow cytometer (Darzynkiewicz et al., 1994; Robinson et al., 1999). The forward scatter (FSC), which indicates cell size, and the side scatter (SSC), which reveals the degree of granularity of the cell, are monitored in cells undergoing apoptosis. The DNA content profile, which is regularly used in studies of the cell cycle, indicates cells’ positions in the cycle depending on their DNA content as measured by propidium iodide (PI) staining (fluorescence). Dead cells have a higher amount of subdiploid DNA which accumulates in the pre-G1 position of cell cycle profiles. The protocol described here uses flow cytometry of fixed cells to simultaneously assess changes in cell size and granularity as well as differing DNA content.

BASIC PROTOCOL 4



Materials Cell suspension 70% ethanol, ice cold 1× PBS, pH 7.2 to 7.4 (APPENDIX 2A) 50 µg/ml propidium iodide (Sigma) in 1× PBS (APPENDIX 2A for PBS; store up to 1 month at 4°C) 25 mg/ml RNase A (e.g., Sigma) in water Flow cytometer (e.g., Becton Dickinson) and tubes 1. Place a sample of cell suspension containing ∼1 × 105 cells in a microcentrifuge tube, and centrifuge 5 min at 1000 × g, 4°C. Remove and discard supernatant. 2. Add 1 ml ice-cold 70% ethanol dropwise while vortexing (to avoid clumping) to resuspend cells. Incubate 30 min on ice to allow cell fixation. 3. Centrifuge 5 min at 1000 × g, 4°C, to pellet cells. 4. Decant ethanol. Add 1 ml of 50 µg/ml propidium iodide containing 20 µl of 25 mg/ml RNase A and incubate 15 min at room temperature. 5. Set up the flow cytometer for forward scatter and side scatter (FSC and SSC) to determine cell size and granularity, together with FL-2 content for DNA (560 nm excitation, 640 nm emission; see manufacturer’s instructions). The reduction in FSC and SSC seen in apoptotic cells is due to the decrease in cell volume and increase in cellular granularity. In necrotic cells there is an increase in both FSC and SSC, reflecting an increase cell volume and loss of cell granularity. BASIC PROTOCOL 5

Assessment of Annexin V Binding to Detect Externalized Phosphatidylserine and Propidium Iodide Exclusion A typical feature of apoptosis is the rearrangement and loss of plasma membrane phospholipid asymmetry as a result of externalization of phosphatidylserine (PS) from the inner leaflet of the plasma membrane to the outer leaflet. It has been demonstrated that annexin V can bind to PS in a calcium-dependent manner. This property of annexin V has made it a very useful tool for detection of apoptosis at the early stages of the process (Koopman et al., 1994), especially in conjuction with propidium iodide. Materials Cell suspension PBS, pH 7.2 to 7.4 (APPENDIX 2A) 1 to 3 µg/ml annexin V–FITC (Bender Biosystems, Nexin Research, Roche, or Pharmingen) HEPES buffer, pH 7.4 (see recipe) HEPES buffer (see recipe) containing 50 µg/ml propidium iodide (e.g., Sigma) Flow cytometer (e.g., Becton Dickinson) and tubes 1. Place a sample of cell suspension containing ∼2–4 × 105 cells in a microcentrifuge tube, and centrifuge 5 min at 1000 × g, 4°C. 2. Remove and discard supernatant. Resuspend cells in 200 µl annexin V–FITC diluted 4:1 in HEPES buffer/propidium iodide. Incubate 5 min at room temperature in the dark. 3. Analyze cells by flow cytometer for both green (annexin V–FITC; FL-1, 515 nm excitation, 545 nm emission) and red (PI; FL-2, 560 nm excitation, 640 nm emission) fluorescence according to the manufacturer’s instructions (see Figure 2.2.2).


Cells that are apoptotic bind annexin V, display green fluorescence, and exclude propidium iodide. In contrast, necrotic cells with a disrupted plasma membrane display uptake of both annexin V (green fluorescence) and propidium iodide (red fluorescence; see fig 2.2.2).


Propidium iodide uptake

control

104

apoptosis

necrosis

103 102 101 100 100

101

102

103

104

PS exposure

Figure 2.2.2 Flow cytometric analysis of phosphatidylserine exposure of Jurkat cells treated with different concentrations of a cytotoxic drug. Cells were double stained for annexin V and propidium iodide. Apoptotic cells are positive for annexin V only, whereas necrotic cells are positive for both annexin V and propidium iodide. (A) Untreated cells, (B) apoptotic cells, (C) necrotic cells. (This figure is reproduced by kind permission from Dr. M. Hampton, Institute of Environmental Medicine, Karolinska Institute, Stockholm.)

ASSAYS FOR CHROMATIN CLEAVAGE Chromatin cleavage is a hallmark of apoptosis and involves the formation of high-molecular-weight (>50-kbp) and nucleosome-sized (200-bp) DNA fragments. The high-molecular-weight fragments can be separated by pulsed-field gel electrophoresis, and the nucleosome-sized fragments, when separated on a conventional agarose gel, demonstrate a ladder pattern. The DNA from necrotic cells, on the other hand, has a random and general cleavage pattern and produces a smear when electrophoresed on either a pulsed-field or conventional gel (see Figure 2.2.3). Prior to the formation of the cleavage products, endogenous nucleases must generate a large number of DNA strand breaks. These strand breaks in DNA can be detected by attaching biotin-conjugated nucleotides to the 3′ hydroxyl termini in a reaction catalyzed by exogenous terminal deoxynucleotidyl transferase (TdT). A number of methods can be used to identify apoptotic or necrotic DNA. All of these methods are equally successful and require the same number of cells. TUNEL Assay for DNA Fragmentation in Cells Induction of apoptosis results in the generation of single-strand DNA breaks. These can be detected using the TdT-mediated dUTP-biotin nick end-labeling (TUNEL) method (Gavrieli et al., 1992). This method requires cell fixation with cross-linking agents such as formaldehyde, which, unlike some of the alcohols (such as ethanol), prevents the extraction of degraded DNA. In other words, this fixation step prevents the loss or reduction of the cellular DNA content after the extensive washings and staining involved in this protocol. Once cells are fixed with formaldehyde, treatment with alcohols does not affect the DNA content of cells.

BASIC PROTOCOL 6

Materials Cell suspension 100% methanol, −20°C PBS, pH 7.2 to 7.4 (APPENDIX 2A) 1% (v/v) formaldehyde 70% ethanol, ice cold continued



M

A

1

2

3

MW (bp)

MM 1 2 3

B

MW (kbp)

680 1033 242.5 653

394

48.5

Figure 2.2.3 Gel electrophoresis of DNA from Jurkat cells treated with different concentrations of a cytotoxic drug. (A) Conventional gel: M, marker; lanes 1 to 3, DNA from apoptotic, untreated, and necrotic cells, respectively. The DNA ladder, when present, is a strong indicator of apoptosis. (B) Pulsed-field gel: MM, markers; lanes 1 to 3, DNA from untreated, apoptotic, and necrotic cells, respectively.

Terminal deoxyribonucleotidyltransferase (TdT; 25 U/µl) and 10× buffer (0.3 M Tris base/1.4 M sodium cacodylate, pH 7.2/1 mM DTT; e.g., Boehringer Mannheim) 25 mM CoCl2 1 mM Bio-16-dUTP (e.g., Boehringer Mannheim) Termination buffer: 300 mM NaCl/30 mM sodium citrate (e.g., Sigma or equivalent) Staining buffer (e.g., Sigma; see recipe) FACS fluid (e.g., Becton Dickinson; optional; for flow cytometry) DPX mountant (BDH) 5 µg/ml propidium iodide in PBS (APPENDIX 2A) Flow cytometer (e.g., Becton Dickinson) and tubes; or fluorescent microscope Coverslips (e.g., Chance Propper) Microscope slide (e.g., Objektträger, Menzel-Glaser) 1. Wash a sample of cell suspension containing ∼106 cells by centrifuging 5 min at 1000 × g, 4°C, discarding supernatant, and resuspending in PBS. Transfer to microcentrifuge tube and centrifuge 5 min at 1000 × g, 4°C. 2. Remove and discard supernatant. Resuspend cells in 1% formaldehyde and fix for 15 min on ice. 3. Centrifuge 5 min at 1000 × g, 4°C. Remove and discard supernatant. Resuspend cell pellet in PBS, and repeat centrifugation. Remove and discard supernatant. 4. Resuspend cell pellet in 0.5 ml PBS and add to 5 ml ice-cold 70% ethanol. Cells can be stored in ethanol for several weeks at −20°C. Determination of Apoptosis and Necrosis

5. Centrifuge 5 min at 1000 × g, 4°C. Remove and discard ethanol. Resuspend cells in PBS, and repeat centrifugation.


6. Prepare elongation buffer (50 µl per sample): 41.5 µl ultrapure water 5 µl 10× TdT buffer 2 µl 25 mM CoCl2 1 µl 1 mM Bio-16-dUTP 0.5 µl 25 U/µl TdT enzyme. Add 50 µl elongation buffer to each cell pellet and resuspend. Incubate 30 min at 37°C. It is recommended that more than the required amount of elongation buffer be prepared; that is, multiply the total number of samples + 1 by 50 ìl (the volume of the elongation buffer per sample) when calculating how much buffer to prepare. This buffer needs to be made fresh.

7. Add 5 ml PBS and centrifuge 5 min at 1000 × g, 4°C. Remove and discard supernatant. The reaction may first be stopped by incubating cells with termination buffer for 15 min at room temperature, though omission of this step does not appear to affect the results.

8. Resuspend cell pellet in 100 µl staining buffer, and incubate 30 min at room temperature. Again, it is recommended that extra buffer be prepared.

9. Add 2 ml PBS, resuspend cells, and centrifuge 5 min at 1000 × g, 4°C. Remove and discard supernatant. Repeat. 10. Add 1 ml of 5 µg/ml propidium iodide and incubate 30 min at room temperature in the dark. 11. Centrifuge 5 min at 1000 × g, 4°C, and remove supernatant. 12a. For flow cytometry: Resuspend cells in sufficient amount of PBS or FACS fluid and measure fluorescence (for FITC, 488 nm excitation, 520 ± 20 nm emission, FL-1 channel; for PI, 560 nm excitation, 640 nm emission, FL-2 or FL-3 channels). 12b. For fluorescence microscopy: Resuspend cells in 50 to 100 µl of PBS and spin them onto a slide as described under Cytospin preparations (see Support Protocol 1). Mount slides with DPX mountant and observe fluorescence with green and red filters under microscope. Cells for fluorescence microscopy can be counterstained with hematoxylin for ≤30 sec, and then rinsed with water, before microscopy. The number of single-strand breaks is increased during the early stages of apoptosis (increase in green fluorescence) without any change in the DNA content (measured by PI staining). With time the DNA content is decreased (red fluorescence) due to formation of apoptotic bodies, which also results in the reduction of TUNEL signal.

TUNEL Assay in Paraffin-Embedded Sections The TUNEL assay can also be performed on paraffin-embedded tissue sections. Additional Materials (also see Basic Protocol 6) Paraffin-embedded tissue sections on slides 4% (w/v) paraformaldehyde (see recipe) or 4% formaldehyde, in PBS (APPENDIX 2A) 96%, 90%, and 80% ethanol Xylene BSA (e.g., Sigma) continued




10 mM Tris⋅Cl, pH 8 (APPENDIX 2A) 20 µg/ml proteinase K in 10 mM Tris⋅Cl (APPENDIX 2A for Tris⋅Cl) 3% (v/v) methanol 2% (w/v) BSA in PBS (APPENDIX 2A for PBS) ExtrAvidin-peroxidase (Sigma) diluted 1/50 in PBS/1% BSA/0.5% Tween 20 3-Amino-9-ethylcarbazole (AEC) Fluorescent microscope Coverslips (e.g., Chance Propper) Microscope slide (e.g., Objektträger, Menzel-Glaser) 1. Fix section in 4% paraformaldehyde or formaldehyde, then immerse in PBS. 2. Remove paraffin from sections by incubating slide 10 min at 70°C or 30 min at 58° to 60°C. 3. Rehydrate sections by sequential incubation in xylene (twice, 5 min each), 96% ethanol (twice, 3 min each), 90% ethanol (3 min), and 80% ethanol (3 min), followed by 3 min in water. Because paraffin traces might interfere with the enzymatic reaction in tissue sections, use of fresh solvents is recommended.

4. Treat sections with 10 mM Tris⋅Cl, pH 8, for 5 min, then incubate 15 min in 20 µg/ml proteinase K at room temperature. Wash slide four times in ultrapure water, 2 min each time. 5. Incubate sections in 3% methanol for 30 min at room temperature, and rinse by dipping five times in ultrapure water. 6. Prepare 100 µl per slide of elongation buffer (see Basic Protocol 6, step 6; the buffer needs to be fresh). Incubate the sections 60 min at 37°C in humid atmosphere. 7. Rinse the sections by dipping three times in ultrapure water, and incubate in termination buffer for 15 min at room temperature. 8. Place sections sequentially in PBS (5 min), 2% BSA (10 min), and PBS (5 min), all at room temperature. Addition of BSA prevents nonspecific binding of ExtrAvidin-peroxide.

9. Incubate the sections in ExtrAvidin-peroxidase for 15 min at room temperature. 10. Incubate the sections in 2% BSA, and wash four times in PBS. 11. Stain the sections with AEC for 30 min at 37°C, then rinse by dipping three times in ultrapure water. 12. Mount slides and observe for fluorescence. After mounting the covered slides can stored for many months at 4°C. Tissue sections, like cells, can be counterstained with hematoxylin for ≤30 sec, followed by rinsing in water. BASIC PROTOCOL 7


Detection of DNA Fragmentation in Whole Cells In 1976, Skalka et al. reported that the DNA in chromatin of irradiated lymphoid tissues degrades in vivo into oligonucleosome-length fragments. This observation was first linked to endonuclease activation in 1980 (Wyllie, 1980). Since then several methods have been developed to measure internucleosomal DNA fragmentation. This method was adopted from the protocol first described by Sorenson et al. (1990) and does not require DNA purification. Appearance of a DNA ladder will correspond to apoptosis, whereas a DNA smear will indicate necrosis. This also applies to all the alternate protocols in this section.


Materials 1× and 5× TBE buffer (see recipe), pH ∼8.0 at room temperature (do not adjust pH) SeaKem GTG agarose (FMC Bioproducts) Cell suspension 50 mg/ml RNase A (see recipe) 4× DNA loading buffer: 4× TBE buffer (see recipe) containing 40% (w/v) sucrose and 0.25% (w/v) bromphenol blue (e.g., Sigma; store up to 2 to 3 weeks at 4°C) Ultrapure agarose (Life Technologies) 10% (w/v) SDS (see recipe) 20 mg/ml proteinase K in water (store in aliquots up to 1 year at −20°C) 1× DNA loading buffer: 1× TBE buffer (see recipe) containing 10% (w/v) sucrose and 0.25% (w/v) bromphenol blue (e.g., Sigma) DNA marker VI (pBR328 DNA cleaved with BglI and HinfI; Boehringer Mannheim), 1 µl in 20 µl of 1× DNA loading buffer TE buffer, pH 8.0: 10 mM Tris⋅Cl/1 mM EDTA 10 mg/ml ethidium bromide (see recipe) Boiling water bath or microwave oven Gel electrophoresis apparatus: GNA-100 (Amersham Pharmacia Biotech), Buffer Puffer (Owl Scientific), or equivalent Power supply (Power-Pac 300, Bio-Rad, or equivalent) Shaker at 4°C MacroVue UV Transilluminator (Hoefer Scientific Instruments) or equivalent Photoman Polaroid gel documentation system (Hoefer Scientific Instruments) or equivalent Prepare agarose gel for electrophoresis 1. Prepare sufficient volume of 1× TBE to cast the gel and to fill the electrophoresis tank. It is important to use the same batch of electrophoresis buffer in both the electrophoresis tank and in the gel. Differences in pH or ionic strength can greatly affect the DNA mobility.

2. Weigh 0.9 g SeaKem agarose in a flask, and add 50 ml of 1× TBE (for GNA-100 gel electrophoresis apparatus). Heat in a boiling water bath or microwave oven until dissolved, then allow the agarose to cool to 60°C for 10 to 15 min. This is a standard 1.8% agarose gel. The volume should not occupy >40% of the volume of the flask. For the Buffer Puffer gel electrophoresis apparatus, use 2.7 g agarose and 150 ml TBE.

3. Pour gel into mold. Remove unwanted air bubbles. Place comb in the gel and allow gel to set 1 hr. Prepare sample 4. Place a sample of cell suspension containing 4−5 × 105 cells in a microcentrifuge tube, and centrifuge 3 min at 1000 × g, 4°C. If necessary, the pellet may be stored up to 2 to 3 weeks in microcentrifuge tubes at −20°C before proceeding.

5. Carefully remove supernatant and resuspend pellet in 16 µl ultrapure water. 6. Add 4 µl of 50 mg/ml RNase A (final concentration 10 mg/ml), mix, and leave 20 min at room temperature. 7. Add 5 µl of 4× DNA loading buffer.



8. Using a scalpel, cut out a piece of gel between the comb and the upper edge of the gel (do not remove the comb), leaving the gel above the first one or two wells intact. Prepare digestion gel 9. Prepare 5 ml of digestion gel by weighing 40 mg Ultrapure agarose, adding 2.75 ml ultrapure water, and stirring on a hot plate to dissolve. Do not use the SeaKem agarose in this step because it causes SDS to come out of solution. For the Buffer Puffer gel electrophoresis apparatus, prepare 10 ml of digestion gel.

10. When steam starts to rise from the flask, add 1 ml of 5× TBE and 1 ml of 10% SDS. When agarose has boiled and dissolved, take flask off the heat and allow gel to cool to 50°C. 11. Add 250 µl of 20 mg/ml proteinase K, mix gently, and pour into the gap above the main gel. Allow to cool, and remove the comb. Set up for electrophoresis 12. Mount the gel in the electrophoresis tank, and add just enough 1× TBE buffer to cover it to a depth of 1 mm. 13. Add 1 µl DNA marker VI to 30 µl of 1× DNA loading buffer and load the marker into the well that is separated from the digestion gel. Load all other samples into the wells connected with the digestion gel. For the Buffer Puffer apparatus, use 2 ìl DNA marker VI dissolved in 30 ìl of 1× DNA loading buffer.

14. Run the gel overnight (∼14 hr) at 20 V to facilitate digestion of the sample. 15. Turn the voltage up to 90 V and run for an additional 1.5 hr. For the Buffer Puffer apparatus, run gel 2 hr at 100 V after the initial 20-V overnight run.

Develop gel 16. Remove the gel from the tank and rinse with water. 17. Add 100 ml TE buffer and 40 µl of 50 mg/ml RNase A. Place gel on shaker for 3 to 4 hr. 18. Rinse the gel with distilled water. Add 100 ml TE buffer and 5 µl of 10 mg/ml ethidium bromide. Place gel on the shaker for 40 min. 19. Wash several times with fresh TE to remove ethidium bromide, and photograph under UV light. ALTERNATE PROTOCOL 2

Detection of DNA Fragmentation in Total Genomic DNA This method is a modified version of the protocol described by McGahon et al. (1995). The presence of ethidium bromide in the gel avoids later staining and destaining of the gel. Although Basic Protocol 7 and this alternate protocol are equally sensitive, the latter is less time consuming. In addition, the dry loading prevents the loss of material. Additional Materials (also see Basic Protocol 7) Lysis buffer: 2 mM EDTA/100 mM Tris⋅Cl, pH 8.0/0.8% (w/v) SDS (store at room temperature)



1. Place sample of cell suspension containing 4−6 × 105 cells in a 1.5-ml microcentrifuge tube. Centrifuge 5 min at 2000 × g, 4°C. 2. Remove and discard supernatant. Add 20 µl lysis buffer. 3. Add 2 µl of 50 mg/ml RNase A. Mix well by flicking the tip of the tube. Do not apply vigorous vortexing. Incubate at least 30 min at 37°C. 4. Add 10 µl of 20 mg/ml proteinase K, and incubate at least 1.5 hr at 50°C. 5. Add 8 µl of 4× DNA loading buffer. If necessary, after this step the samples can be stored for at least 1 week at 4°C.

6. Place 0.9 g SeaKem agarose (1.8%) in 50 ml of 1× TBE buffer in a flask (for GNA-100 gel electrophoresis apparatus). Heat in a boiling water bath or microwave oven until dissolved. Allow the agarose to cool to 60°C for 10 to 15 min. Add 3 µl of 10 mg/ml ethidium bromide. 7. Dry load the samples. Dissolve 1 µl DNA marker VI in 30 µl of 1 × DNA loading buffer and load into the gel. Apply low current (∼35 mA) for 7 hr or higher current (∼60 mA) for 4 hr. Samples are dry loaded and enough TBE buffer is added to the apparatus so it touches both sides of the gel but does not cover it. This is to ensure that the sample is not lost. After 10 min, when the samples have entered the gel, the current is stopped and more TBE is added so it covers the whole gel, and the current is then reapplied.

8. Photograph the gel under UV light. Simple Protocol for Detection of DNA Fragments This simple protocol was developed in our laboratory. It is particularly useful for detection of chromatin cleavage in lymphoid cells (Zhivotovsky et al., 1995). It is also very useful for different lymphoid cells and less so for hepatocytes and epithelial cells. For these types of cells, see Alternate Protocol 4.


Additional Materials (also see Basic Protocol 7) Lysis buffer, 4°C: 5 mM Tris⋅Cl, pH 8.0/20 mM EDTA/0.5% (v/v) Triton X-100 100% ethanol, −20°C 5 M NaCl (APPENDIX 2A) RNase T1/A stock (see recipe) Vacuum lyophilizer (e.g., Hetovac, Heto-Holten) Prepare DNA sample 1. Place sample containing 1−2 × 106 cells in a microcentrifuge tube, and centrifuge 5 min at 2000 × g, 4°C. Resuspend pellet in 1 ml PBS and repeat centrifugation. 2. Resuspend pellet in 250 µl TE buffer, add 250 µl of 4°C lysis buffer, and vortex. Refrigerate 30 min. 3. Centrifuge sample 15 min at 15,000 × g, 4°C. 4. Transfer supernatant to a fresh microcentrifuge tube. Add 1 ml −20°C ethanol and 30 µl of 5 M NaCl. Mix and place in −20°C freezer overnight. 5. Centrifuge sample for 15 min at 15,000 × g, 4°C. Remove and discard supernatant. 6. Place precipitate in vacuum lyophilizer for 20 to 25 min.



7. Add 20 to 30 µl TE buffer and 1 µl RNase T1/A stock. Incubate 1 hr at 37°C. 8. Add 1 µl of 20 mg/ml proteinase K, and incubate an additional 1 hr at 37°C. 9. Add 8 µl of 4× DNA loading buffer. Cast and run the gel 10. Dissolve 0.9 g SeaKem agarose in 50 ml TBE buffer (for GNA-100 gel electrophoresis apparatus) or 2.7 g SeaKem agarose in 150 ml TBE buffer (for Buffer Puffer gel electrophoresis apparatus). Heat in a boiling water bath or microwave oven until dissolved, then cool gel and pour into mold. 11. Mount the gel in the electrophoresis tank, and add just enough 1× TBE buffer to cover the gel to a depth of 1 mm. 12. Load DNA marker VI in the first well and samples in the rest of wells (see Basic Protocol 7, step 13). 13. Run the gel at 60 mA (for GNA-100) or 70 mA (for Buffer Puffer) until the bromphenol blue front is ∼1 to 2 cm from the end of the gel. 14. Stain gel with ethidium bromide and photograph under UV light (see Basic Protocol 7, steps 18 to 19). ALTERNATE PROTOCOL 4

Phenol Extraction of DNA Fragments for Agarose Gel Electrophoresis This protocol is a modification of the classical method described by Wyllie (1980). This method requires DNA purification. Although it takes much more time tha other techniques and needs many more cells for preparation, it yields very clean, protein-free DNA fragments. Additional Materials (also see Basic Protocol 7) Lysis buffer (see Alternate Protocol 3), 4°C 100% ethanol, −20°C 5 M NaCl (APPENDIX 2A) RNase T1/A stock (see recipe) TE-saturated phenol (see recipe) 24:1 (v/v) chloroform/isoamyl alcohol (store mixture at room temperature in a fume hood) 0.5% (w/v) SDS Prepare DNA sample 1. Place sample containing 2−5 × 106 cells in microcentrifuge tube, and centrifuge 5 min at 2000 × g, 4°C. 2. Resuspend pellet in 250 µl TE buffer, add 250 µl of 4°C lysis buffer, vortex, and refrigerate 30 min at 4°C. 3. Centrifuge sample 15 min at 15,000 × g, 4°C. 4. Transfer supernatant to fresh microcentrifuge tube. Add 1 ml −20°C 100% ethanol and 30 µl of 5 M NaCl. Mix and place in −20°C freezer overnight. 5. Centrifuge sample 15 min at 15,000 × g, 4°C. Remove and discard supernatant. 6. Add 500 µl TE buffer and 5 µl RNase T1/A stock to pellet. Incubate 30 min at 37°C.



7. Add 250 µl TE-saturated phenol and 250 µl of 24:1 chloroform/isoamyl alcohol. Vortex, and centrifuge for 2 to 3 min at 5000 × g, 4°C. Transfer upper layer to fresh 2-ml microcentrifuge tube. 8. Add 500 µl of TE buffer to lower layer; repeat step 7. Transfer upper layer to 2-ml microcentrifuge tube and mix with upper layer from step 7. 9. Extract the mixed samples from steps 7 and 8 with 1000 µl 24:1 chloroform/isoamyl alcohol and centrifuge again. 10. Transfer the upper layer to 1.5-ml microcentrifuge tubes, 500 µl per tube. Precipitate overnight with ethanol and NaCl as in step 4. 11. Repeat step 5. 12. Place precipitate in vacuum lyophilizer for 20 to 25 min. 13. Dissolve and mix precipitates from both microcentrifuge tubes in a total of 20 to 30 µl TE buffer/0.5% SDS. Add 5 µl of 4× DNA loading buffer. Cast and run the gel 14. Mix a 1.8% agarose gel by dissolving 0.9 g agarose in 50 ml TBE buffer (for GNA-100 gel electrophoresis apparatus) or 2.7 agarose in 150 ml TBE buffer (for Buffer Puffer). Cool gel and pour into mold. 15. Mount the gel in the electrophoresis tank, and add just enough 1× TBE buffer to cover the gel to a depth of 1 mm. 16. Load DNA marker VI in the first well (see Basic Protocol 7, step 13) and samples in the rest of the wells. 17. Run the gel at 60 mA (for GNA-100) or 70 mA (for Buffer Puffer) until the bromphenol blue front is ∼1 to 2 cm from the end of the gel. 18. Stain gel with ethidium bromide (see Basic Protocol 7, steps 18 to 19) and photograph under UV light. Quantitative Assay of DNA Fragmentation In addition to qualitative analysis by gel electrophoresis, DNA fragmentation can be quantitatively determined by using the diphenylamine reagent. This method, introduced by Burton (1956), cannot, however, discriminate between apoptotic and necrotic chromatin cleavage.

BASIC PROTOCOL 8

Materials Cell suspension to be assessed Lysis buffer (see Alternate Protocol 3), ice cold 10% (w/v) and 5% (w/v) trichloroacetic acid (TCA; keep at room temperature in dark flasks) Diphenylamine reagent (see recipe) 10-ml conical glass tubes Round-bottom glass tubes Water bath, 100°C 1. Prepare cell suspension containing 1−10 × 106 cells in a 1-ml volume. 2. Transfer 0.8 ml of cell suspension to microcentrifuge tube. Add 0.7 ml ice-cold lysis buffer. Vortex, and allow lysis to proceed 15 to 30 min at 4°C.



3. Centrifuge 15 min at 15,000 × g, 4°C. Transfer supernatant to labeled conical glass tube. 4. Add 0.65 ml of 5% TCA to the pellet in microcentrifuge tube, and add 1.5 ml of 10% TCA to the sample in labeled glass tube. 5. Precipitate both samples overnight (≥4 hr) at 4°C. 6. Centrifuge conical tube 10 min at 2500 × g, room temperature. 7. Remove supernatant after centrifugation and add 0.65 ml of 5% TCA to the pellet. Prepare two blank tubes with 0.65 ml of 5% TCA to be treated the same for the remaining steps. 8. Make hole on the top of each microcentrifuge tube from step 5, and cover each conical tube with a marble. 9. Boil both sets of tubes for 15 min in 100°C water bath. 10. Cool to room temperature. Centrifuge samples 5 min at 2500 × g, room temperature. 11. Transfer 0.5 ml of each supernatant (from both glass and microcentrifuge tubes) to labeled, round-bottom glass tubes. 12. Add 1 ml diphenylamine reagent to each tube. Incubate ≥4 hr at 37°C with a marble covering each tube. 13. Read absorbance at 600 nm in spectrophotometer. Set zero with blanks from step 7. Express results as the percentage of DNA fragmented:

% fragmented DNA =

BASIC PROTOCOL 9

absorbance of the supernatant × 100 absorbance of supernatant + pellet

Detection of High-Molecular-Weight Chromatin Fragments by Pulsed-Field Agarose Gel Electrophoresis Not all forms of apoptosis are accompanied by internucleosomal DNA fragmentation (Oberhammer et al., 1993a). However, the formation of high-molecular-weight apoptotic DNA fragments of 50 to 700 kbp (see Figure 2.2.3B), which was first observed by Walker et al. (1991), has been reported to occur in all the cell types studied to date (see Background Information in Commentary). One of the forms of pulse-field gel electrophoresis, field-inversion gel electrophoresis (FIGE), has been successfully used in several laboratories to identify high-molecularweight DNA fragments in apoptotic cells and a smear in the same range of DNA size in necrotic cells. The authors have used both vertical and horizontal gel chambers; resolution of DNA fragments is the same. This protocol is an adaptation of an earlier method described by Anand and Southern (1990).


Materials Cell suspension Agarose buffer (for molds; see recipe) SeaPlaque GTG low-melting-point agarose (FMC Bioproducts) 20 mg/ml proteinase K in water (store in aliquots up to 1 year at −20°C) Proteinase buffer (for plugs; see recipe) TE buffer, pH 8.0:10 mM Tris⋅Cl/1 mM EDTA


50 mM EDTA, pH 8.0 (APPENDIX 2A) SeaKem GTG agarose (FMC Bioproducts) 5× TBE buffer (see recipe) DNA size pulse markers: chromosomes from Saccharomyces cerevisiae (225 to 2200 kbp) and a mixture of λDNA HindIII fragments, λDNA, and λDNA concatemers (0.1 to 200 kbp; Sigma; supplied premade in syringe) Gel leveling table 100-µl insert molds (Amersham Pharmacia Biotech), stored in 0.1 M HCl 12- or 24-well tissue culture plates 50°C incubator 100°C water bath or microwave oven Pulsed-field gel electrophoresis system: vertical gel chamber with cooling elements (Protean II, Bio-Rad); horizontal gel chamber (HE 100B); power supply (PS 500 XT); and Switchback pulse controller (PC 500, Hoefer Scientific Instruments) Thermostatic circulator Multi Temp III (Amersham Pharmacia Biotech) Prepare sample plugs of agarose 1. Rinse insert molds extensively in distilled water and dry. Wrap Parafilm around the bottom of the molds and place them on ice to chill at least 10 min before pouring in the gel. Make sure to mark the sample wells with pencil or pen.

2. Place sample of cell suspension containing 1 × 106 cells in a microcentrifuge tube, and centrifuge 3 min at 2000 × g, 4°C. Remove and discard supernatant. Resuspend pellet in 100 µl of agarose buffer. 3. Prepare 1% low-melting-point agarose in agarose buffer, and place in a 60°C water bath until melted. 4. Add 100 µl of molten 1% agarose to the cell suspension, and mix with pipet. Immediately pipet mixture into two prechilled insert molds, 100 µl per mold. 5. Place the filled insert molds on ice for 10 to 15 min. 6. Remove Parafilm from the bottom of the insert molds, and dislodge each plug into separate well of a tissue culture plate. It is convenient to use 12- or 24-well tissue culture plates for this purpose.

7. Add 1 ml of proteinase buffer and 10 µl proteinase K (final concentration 0.2 mg/ml plug) to each well. Wrap Parafilm or clear tape around the plate, and incubate ≥24 hr on a shaker at 50°C. 8. Remove wrapping from plate, and remove buffer with pipet. If it is necessary to simultaneously analyze the low-molecular-weight DNA, it is possible to remove the DNA fragments that have leached out from the plugs during incubation with proteinase K by precipitating them from this buffer over 48 hr at −20°C. Add 0.2 volume of 10.5 M ammonium acetate and 2 volumes of cold absolute ethanol. This DNA can then be lyophilized, dissolved in 1× DNA loading buffer, and analyzed by conventional gel electrophoresis.

9. Wash each plug three times with 1 ml TE buffer, 2 hr per wash, on a shaker at 4°C. 10. Remove TE buffer, and add 1 ml of 50 mM EDTA. Plugs are now ready to put on a gel. To save samples for a long time, store the plugs in EDTA, in which DNA is stable for several months, at 4°C.



Prepare separating gel 11. Prepare 1% agarose (SeaKem GTG) gel in 0.5× TBE. For horizontal gels 12a. Dissolve 2.5 g agarose in 250 ml 0.5× TBE in a boiling water bath or microwave oven. Allow the agarose to cool to 60°C for 10 to 15 min. 13a. Seal the edges of the gel platform with tape, and place platform on leveling table. 14a. Pour the gel into the mold, and remove unwanted air bubbles. Place the comb into the gel, and allow to set for 1 hr. 15a. Remove the comb very carefully so as not to break the gel. Remove tape. For vertical gels 12b. Clamp two frosted glass plates together with 3-mm spacers in between. Seal the bottom with Parafilm to prevent leakage of warm agarose. 13b. Warm glass plates for 30 min in 50°C incubator. 14b. Melt agarose as in step 12a (1 g agarose in 100 ml 0.5× TBE), and pour into the prewarmed gel assembly to ∼5 mm below top of plates. Insert comb into the gel, and allow to set for 1 hr. 15b. Remove the comb very carefully so as not to break the gel, and wash the wells with 0.5× TBE to remove partially polymerized agarose. Load and run gel 16. Using a scalpel, cut out a 2-mm slab from one marker, and insert into first well of the gel with a spatula. 17. Repeat this procedure with the second marker. Make sure that the slab is flat on the bottom of the well. This applies to loading of samples as well.

18. To load sample, cut off 3 mm from sample plug, and insert it into individual well with a spatula. Repeat same procedure for all samples. Avoid introducing any air bubbles into the wells. Note and record the order of the samples!

19. Cement the slabs into wells with the remaining molten 1% agarose. 20. Transfer the loaded gel (either horizontal or vertical) into a precooled tank filled with 0.5× TBE. The buffer temperature should be 11° to 12°C at the beginning of the experiment.

21. Run the gel at a constant voltage with a suitable switcher program to achieve the desired resolution. The programs listed in Table 2.2.2 give good resolution of 50-, 300-, and 700-kbp fragments. 22. Remove gel carefully, and stain it in ethidium bromide for 1 hr (see Basic Protocol 7, step 18). Destain it in 0.5× TBE for 2 to 3 hr. 23. Visualize and photograph bands under UV light. Determination of Apoptosis and Necrosis


Table 2.2.2

Parameters for Running Pulsed-Field Agarose Gels

Running voltagea Procedure

Total running time

Horizontal gel

Vertical gel

170 V 10 min: run the DNA into gel with continuous forward pulse 6 hr: apply 20 sec forward time, 3:1 forward/back ratio, ramp factor 1.5b 6 hr: apply 10 sec forward time, 3:1 forward/back ratio, ramp factor 2 12 hr: apply 0.8 sec forward time, 3:1 forward/back ratio, ramp factor 12.5 24 hr, 10 min

200 V 10 min: run the DNA into gel with continuous forward pulse 2 hr: apply 12 sec forward time, 3:1 forward/back ratio, ramp factor 2c 3 hr: apply 2.4 sec forward time, 3:1 forward/back ratio, ramp factor 5.0 2.5 hr: apply 8.0 sec forward time, 3:1 forward/back ratio, ramp factor 3.0 7 hr, 40 min

a

Constant voltage This means that at the end of these 6 hr, the forward time should be 30 sec. c This means that at the end of these 2 hr, the forward time should be 24 sec. b

METHODS FOR ANALYSIS OF CASPASE PROTEOLYTIC ACTIVITY Several proteolytic activities are implicated in apoptosis and necrosis (see UNIT 2.1). The activation of caspases, a family of apoptotic proteases, plays a key role in apoptosis induced by diverse stimuli. Caspases, which have been detected in numerous tissues and cell types, are synthesized as a precursor form, or proenzyme, and an apoptotic signal converts the precursor into mature enzyme. Moreover, up to now there has been no evidence concerning the involvement of caspase activities in necrosis. Therefore the detection of caspase activity in cells can be used as a discriminating criterion to distinguish apoptosis from necrosis. All members of the caspase family of proteases share a number of amino acid residues crucial for substrate binding and catalysis. Despite their uniform requirement for an aspartate residue at the P1 position of the substrate site, caspases can be divided into three different groups according to their substrate preferences. The first group (caspases 1, 4, and 5) has the optimal substrate cleavage sequence WEXD; the second (caspases 2, 3, and 7) preferentially cleaves targets at the DEXD motif; and the third (caspases 6, 8, and 9) has the optimal substrate cleavage sequence (L/V)EXD (Thornberry et al., 1997). The predicted caspase specificities correspond to the cleavage sites in the different intracellular caspase targets. Analysis of caspase activation is also based on the substrate specificity of the enzymes. Described here are two methods recently developed and currently used for analysis of caspase activation. Detection of Caspase Activity by Specific Substrate Cleavage This assay was adopted from the protocol first described by Thornberry (1994). Detecting proteolytic cleavage of fluorescent substrates assesses caspase activity. To determine the specificity of the assayed samples for the substrate, appropriate aldehyde-conjugated inhibitors (see Table 2.2.3 in Reagents and Solutions, below) can be added at a final concentration of 50 nM to the complete buffer containing the substrate. The resulting inhibition is biphasic: initially V0 is equivalent to the substrate cleavage rate, and then a plateau in the rate of substrate cleavage occurs, usually within 5 to 8 min of the assay. As with all enzyme-substrate and -inhibitor interactions, the extent of inhibition is dependent on the concentration of the other components.

BASIC PROTOCOL 10



Materials Cell suspension 1× PBS, pH 7.2 to 7.4 (APPENDIX 2A) 1× substrate buffer (SB; see recipe) Caspase substrate and inhibitor stock solutions (see recipe) Liquid nitrogen 96-well tissue culture plates (e.g., LabSystems) Fluoroscan II (LabSystems) 1. Place duplicate samples containing 2−3 × 106 cells in microcentrifuge tubes, and centrifuge 5 min at 1000 × g, 4°C. 2. Remove and discard supernatant. Wash pellet with PBS, and centrifuge 5 min at 1000 × g, 4°C. 3. Remove and discard supernatant. Resuspend each pellet in 25 µl PBS (check pH, which should be neutral), and transfer into individual wells of a 96-well culture plate. The plate should be kept very cold (either by keeping it for some time at −20°C, or by having it “floating” on liquid nitrogen). If necessary, plates with cells can be stored for several days at −80°C.

4. Set up the computer control for the Fluoroscan II plate reader. Set temperature to 37°C. This takes about 3 to 4 min. Several computer programs are available for acquisition and data processing. The authors recommended Genesis II Windows-based microplate software (LabSystems and Life Sciences).

5. Immediately prior to use, prepare complete 1× SB by adding 1 µl of 200 mM stock solution of the substrate of interest (final concentration 50 µM) to 4 ml of 1× SB. 6. Dispense 50 µl of complete SB with substrate to each test well of cells at room temperature, and place plate immediately on the Fluoroscan II. The final concentration of substrate in the wells is 33.3 ìM.

7. Read samples. The maximum absorption for AMC (7-amino-4-methyl coumarin) is 354 nm. Fluorometric detection for AMC cleaved from peptide is at excitation 380 nm and emission 460 nm. BASIC PROTOCOL 11


Immunodetection of Active Caspases During Apoptosis All caspases are expressed as proenzymes (30 to 55 kDa) that contain three domains: an NH2-terminal domain, a large subunit (17 to 35 kDa), and a small subunit (10 to 12 kDa). Activation involves proteolytic processing between domains, followed by association of the large and small subunits to form a heterodimer. Thus, detection of large or small subunits by Western blotting can be used as a marker of caspase activation. All caspases can be described according to function as either activators or executioners of cell death. Their relative order appears to vary widely depending on the cellular background and apoptotic stimuli. Therefore, it is necessary to investigate the cleavage and activation of several caspases. Since the activation of the caspase 3–like proteases has been shown during most apoptotic process, it is reasonable to first investigate the activation of caspase 3 and then continue to look for the activation of caspases located up- and downstream (see UNIT 2.1). To compare the cleavage/activation of different caspases in the same samples. the membrane may be reprobed. Moreover, for densitometric analysis of results of protein degradation, the same membranes can be reprobed and stained with antibodies against marker proteins, such as glyceraldehyde-3-phosphate dehydrogenase or actin.


Materials Cell suspension PBS, pH 7.2 to 7.4 (APPENDIX 2A) Sample buffer (see recipe) Prestained SDS-PAGE markers, low range (e.g., Bio-Rad) 1× electrode buffer (see recipe) 4× high-salt Tris base buffer (HSB; see recipe) High-salt Tris base buffer with Tween 20 (HSBT; see recipe) Low-salt Tris buffer (LSB; see recipe) Transfer buffer (see recipe) Dry nonfat milk BSA fraction V (e.g., Sigma) Sodium azide Stripping buffer (see recipe) Primary antibodies: Anti–caspase 1 (Calbiochem) Anti–caspase 2 (Transduction Laboratories) Anti–caspase 3 (Pharmingen) Anti–caspase 6 (Research Diagnostics) Anti–caspase 7 (Transduction Laboratories) Anti–caspase 8 (Pharmingen) Anti–caspase 9 (Calbiochem) Anti–caspase 10 (Calbiochem) Appropriate secondary antibodies for the primary antibodies listed above: peroxidase-conjugated goat anti–rabbit IgG, goat anti–mouse IgG, and mouse anti–goat IgG (Pierce) Minigel (see Support Protocol 2) Anti-glyceraldehyde-3-phosphate dehydrogenase (Trevigen) Anti-actin (Sigma or Boehringer Mannheim) ECL Western blotting detection system (Amersham Pharmacia Biotech) or equivalent Boiling water bath Mini-Protean II Electrophoresis Cell (Bio-Rad) or equivalent Prot/Elec pipet tips for protein gel (Bio-Rad) Power supply (Power-Pac 300, Bio-Rad, or equivalent) 0.45-µM nitrocellulose membranes (Sartorius AG) Whatman 3MM filter paper Mini Trans-Blot Transfer Cell (Bio-Rad) or equivalent X-ray film (e.g., FujiFilm) Additional reagents and equipment for one-dimensional SDS-PAGE, minigels (see Support Protocol 2), and immunoblotting (UNIT 2.3) Prepare samples 1. Place 1 × 106 cells in a microcentrifuge tube and centrifuge 5 min at 2000 × g, 4°C. Resuspend pellet in 1 ml PBS, and centrifuge 2 min at 1000 × g, 4°C. Two methods may be used to extract proteins from cells for immunoblotting. Either the intact cells are resuspended directly in sample buffer, or an equal amount of proteins extracted from cells can be mixed with sample buffer. Direct resuspension of cells in sample buffer is described here.

2. Resuspend pellet in 100 µl sample buffer and incubate 4 min in boiling water bath. Let cool at room temperature. If samples become highly viscous as a consequence of the release of chromosomal DNA, shear the DNA either by sonication or by repeated passage through a 23-G hypodermic needle.



Set up and run gel electrophoresis 3. Release the gel holder from the casting stand (see Support Protocol 2). Assemble the inner cooling core with gels according to manufacturer’s instructions, and lower them into the buffer chamber. 4. Add 300 to 350 ml of 1× electrode buffer to the bottom of the chamber so that both the bottom 1 cm of the gel and electrode are covered. Add ∼120 ml of buffer to the upper buffer chamber, filling until the buffer reaches a level halfway between the short and long plates. Do not overfill the upper buffer chamber.

5. Remove the comb by pulling it straight up slowly and gently. 6. Load samples (20 to 30 µl) into wells under the electrode buffer with Prot/Elec Tips. Load one well with 5 µl of low-range prestained SDS-PAGE markers. 7. Attach the electrophoresis apparatus to an electric power supply. Run gel at constant 130 V in a 4° to 8°C cold room until the bromphenol blue reaches the bottom of the resolving gel (∼1.5 hr). 8. After electrophoresis is complete, turn off the power supply and disconnect the electrical leads. Immunoblot proteins 9. Remove cell lid and carefully pull inner cooling core out of lower chamber. Pour off upper buffer. 10. Remove glass plate sandwich from the inner cooling core. Push one of the spacers of the sandwich out to the side of the plates without removing it. Gently twist the spacer so the upper glass plate pulls away from the gel. 11. Mark gel orientation by cutting a right corner from bottom of gel, then immerse gel in transfer buffer. 12. Transfer the proteins from gel onto a nitrocellulose membrane by electrotransfer (see UNIT 2.3). CAUTION: It is important to wear gloves when handling gel, filter paper, and nitrocellulose membranes. The authors normally use a Mini Trans-Blot Module and Bio-Ice cooling unit, and transfer at a constant 100 V for 2 hr in a cold room. Control for transfer efficiency by checking transfer of prestained SDS-PAGE markers.

13. Transfer the nitrocellulose membrane into 1× HSB containing 5% dry nonfat milk, 1% BSA, and 0.05% sodium azide. Keep at least 1 hr (overnight is preferable) on a rocker in the 4° to 8°C cold room. 14. Wash membrane two times, 20 min each, in HSBT on a rocker at room temperature. 15. Put membrane into a plastic bag, and add desired primary antibody in appropriate dilution (usually 1:100 to 1:5000) in 1× HSB containing 1% BSA and 0.05% sodium azide. Incubate 1 to 2 hr on a rocker at room temperature. 16. Wash three to five times, 10 min each, with HSBT on a rocker at room temperature. Determination of Apoptosis and Necrosis

17. Add peroxidase-conjugated secondary antibody in appropriate dilution (usually 1:5,000 to 1:10,000) in 1× HSBT containing 1% BSA. Incubate for 1 hr on a rocker at room temperature.


18. Wash three times, 10 min each, with HSBT on a rocker at room temperature. 19. Wash two times, 30 min each, with LSB on a rocker at room temperature. No Tween 20 in this buffer!

20. Develop nitrocellulose membrane with ECL according to manufacturer’s instructions. 21. Cover membrane with plastic wrap, and immediately expose to X-ray film for 30 sec to 5 min in X-ray cassette. 22. Scan the film using a densitometer, and plot graph of optical density versus concentration. Following ECL detection, it is possible to reprobe the nitrocellulose membrane with other antibodies against different caspases or against marker proteins such as glyceraldehyde3-phosphate dehydrogenase or actin to either clarify the results or to obtain more information. To do this, proceed with the next steps. Figure 2.2.4 shows the results of a time-course study of caspase activation.

23. Submerge the membrane in stripping buffer, and incubate for 30 min at 50°C with agitation. 24. Wash the membrane with water, and repeat steps 14 to 22.

Caspase 3

0

15

30

45

60

75

90 min

p32

p20 p17

Caspase 7 p35

p20

Figure 2.2.4 Time course of caspase activation in Jurkat cells treated with anti-CD95 antibodies. Assessment of Cell Toxicity


SUPPORT PROTOCOL 2

Preparation of SDS-Polyacrylamide Minigels Minigels are useful because they require shorter run times and they use smaller quantities of reagents (the latter especially helpful in immunoassays). Materials 29% (w/v) acrylamide/1% (w/v) bisacrylamide stock (e.g., Bio-Rad) 10% (w/v) ammonium persulfate (APS; e.g., Bio-Rad): 100 mg in 1 ml water, prepared just before gel casting 1.5 M Tris⋅Cl, pH 8.8 (see recipe) 10% (w/v) sodium dodecyl sulfate (SDS; see recipe) TEMED (N,N,N′,N′-tetramethylethylenediamine; e.g., Bio-Rad) Isobutanol, water saturated 1.0 M Tris⋅Cl, pH 6.8 (see recipe) Mini-Protean II Electrophoresis Cell (Bio-Rad) Casting stand Additional reagents and equipment for one-dimensional SDS-PAGE (see APPENDIX 3) 1. Assemble the glass plates for Mini-Protean II system according to the manufacturer’s instructions, and check for leakage. 2. Prepare separating gel (12% gel/0.375 M Tris⋅Cl, pH 8.8). To make one set (two gels) for Mini-Protean II with 1.5-mm thick spacers, combine the following: 6.7 ml H2O 5 ml 1.5 M Tris⋅Cl, pH 8.8 200 µl 10% SDS 8 ml 29% acrylamide/1% bisacrylamide stock 100 µl 10% APS 10 µl TEMED. Because the molecular weights of caspases vary between 30 and 55 kDa, a 12% gel is recommended.

3. Pour separating gel into the gap between the glass plates. Leave sufficient space for stacking gel to be added later (the length of the comb teeth plus 4 to 5 mm). Using a pipet, carefully overlay the gel with water-saturated isobutanol. The advantage of using isobutanol is that the overlay solution can be applied rapidly with a pipet and bulb with very little mixing. If water is used, the overlay must be applied carefully with a needle and syringe, using a steady, even rate of delivery to prevent mixing.

4. Allow gel to polymerize 45 to 60 min. Rinse off the overlay solution completely with distilled water. This is especially important with isobutanol overlays. Do not allow alcohols to remain on the gels >1 hr, or the top of the gel will dehydrate. If necessary this gel can be stored 1 to 2 days at 4°C with distilled water on top.

5. Prepare the stacking gel (4% gel/0.125 M Tris⋅Cl, pH 6.8). For two gels (one set), combine the following:


7.3 ml H2O 1.25 ml 1.0 M Tris⋅Cl, pH 6.8 100 µl 10% SDS 1.3 ml 29% acrylamide/1% bisacrylamide stock 50 µl 10% APS 10 µl TEMED.


6. Pour stacking gel solution directly on top of the polymerized separating gel. Immediately insert a clean Teflon comb into the stacking gel solution, being careful to avoid trapping air bubbles. Top up with more stacking gel. Allow gel to polymerize 20 to 30 min. The gel cannot be stored at this stage and should be used within 1 to 2 hours.

REAGENTS AND SOLUTIONS Use Milli-Q-purified or other ultrapure water for all recipes and protocol steps. For common stock solutions, see APPENDIX 2; for suppliers, see SUPPLIERS APPENDIX.

Agarose buffer (for molds; used in Basic Protocol 9) Prepare 500 ml in H2O: 0.15 M NaCl 2 mM KH2PO4/KOH, pH 6.8 1 mM EGTA 5 mM MgCl2 Filter sterilize through 0.2-µm filter Store at 4°C Caspase substrate and inhibitor stock solutions Prepare 200 mM stock solutions of each substrate and inhibitor to be tested (see Table 2.2.3) in sterile ultrapure water. Dispense into 2-µl aliquots and store at −20°C. Diphenylamine reagent 100 ml glacial acetic acid 1.5 g diphenylamine 1.5 ml concentrated sulfuric acid 0.5 ml 16 mg/ml acetaldehyde stock Prepare just before use Prepare acetaldehyde stock in ultrapure water; store in a dark flask up to 1 month at 4°C.

DTT, 1 M Add 309 mg 1,4-dithiothreitol (DTT; Boehringer Mannheim) to 2 ml of 0.01 M sodium acetate, pH 5.2. Filter sterilize, dispense into 50-µl aliquots, and store at −20°C. Electrode (Tris-glycine) buffer, 10× Dissolve 90 g Tris base, 432 g glycine, and 30 g SDS in 3 liters water. Do not adjust pH with acid or base. Store up to 1 month at 4°C. If precipitation occurs, warm to 37°C before use. For one electrophoretic run, mix 50 ml of 10× stock with 450 ml water just before electrophoresis. Concentrations in 1× buffer: 25 mM Tris, 192 mM glycine, and 0.1% SDS, pH 8.3.

Table 2.2.3

Caspase Substrates and Inhibitors

Substrate

Inhibitor

Supplier (for substrate and inhibitor)

Ac-DEVD-AMC Ac-VEID-AMC Ac-IETD-AMC Ac-WEHD-AMC Z-YVAD-AMC Ac-LEHD-AMC

Ac-DEVD-CHO Ac-VEID-CHO Ac-IETD-CHO Ac-WEHD-CHO Z-YVAD-CHO Ac-LEHD-CHO

Peptide Institute Peptide Institute Peptide Institute Peptide Institute Enzyme Systems Products Enzyme Systems Products



Ethidium bromide, 10 mg/ml Stir 10 mg/ml ethidium bromide in water on magnetic stirrer for several hours. Store up to 2 months in a dark bottle at 4°C. HEPES buffer, pH 7.4 10 mM HEPES (N-hydroxyethylpiperazine-N-2-ethanesulfonic acid), pH 7.4 (adjusted with NaOH) 150 mM NaCl 5 mM KCl 1 mM MgCl2 1.8 mM CaCl2 Store up to several months at 4°C All components may be obtained from Sigma.

HSB (high-salt Tris buffer), 4× Dissolve 48.4 g Tris base and 233.76 g NaCl in 1.5 liters water. Adjust pH to 7.5 with 10 M HCl, and add water to 2 liters. Store up to several weeks at 4°C. Concentrations in 1× buffer: 50 mM Tris⋅Cl and 500 mM NaCl, pH 7.5.

HSBT (high-salt Tris buffer with Tween) Mix 500 ml of 4× HSB (see recipe) and 1500 ml water. Add 1 ml Tween 20 and gently mix. Store up to several weeks at 4°C. Concentrations in 1× buffer: 50 mM Tris⋅Cl, 500 mM NaCl, and 0.05% Tween 20, pH 7.5.

LSB (low-salt Tris buffer) Dissolve 12.1 g Tris base and 18 g NaCl in 1500 ml water. Adjust pH to 7.5 with 10 M HCl, and add water to 2000 ml. Store up to several weeks at 4°C. Concentrations: 50 mM Tris⋅Cl and 150 mM NaCl, pH 7.5.

NP-40, 0.1% (v/v) Add 1 µl of Nonidet P-40 (Boehringer Mannheim) to 1 ml water, vortex, and keep at 4°C. Paraformaldehyde, 4% (w/v) Add 8 g paraformaldehyde powder (e.g., Sigma) to 100 ml water and heat to 60°C in a fume hood. If necessary, add NaOH dropwise (one or two drops each time) to help dissolve the powder. When the solids have completely dissolved, let the solution cool to room temperature. Then add 100 ml of 2× PBS (APPENDIX 2A). Store at 4°C. Because this solution cannot be kept indefinitely, it is better to prepare small volumes every week.

Phenol, TE saturated Allow phenol to warm to room temperature, and then melt it at 68°C. Add hydroxyquinoline to a final concentration of 0.1%. Add a volume of TE buffer, pH 8.0 (10 mMTris⋅Cl/1 mM EDTA), equal to the volume of phenol, and vortex extensively. Allow the two phases to separate, and aspirate the upper aqueous phase. Repeat this saturation with TE buffer two to three times until volume of TE remains unchanged and the pH of the phenol phase has reached 8.0 (as measured with pH paper). Remove the aqueous phase, and add 0.1 vol fresh TE. Store up to 1 month in the dark at 4°C. Hydroxyquinoline, a yellow antioxidant, provides a convenient way to identify the organic phase during extraction. Determination of Apoptosis and Necrosis


Proteinase buffer (for treatment of plugs) Prepare 500 ml of buffer consisting of: 10 mM NaCl 10 mM Tris⋅Cl, pH 9.5 25 mM EDTA Add N-lauroylsarcosine (Sarkosyl) to 1% final concentration and dissolve using magnetic stirrer. Check pH, and if necessary adjust to 9.5 with NaOH. Filter through 0.2-µm filter and store at 4°C. RNase A, 50 mg/ml Dissolve RNase A (e.g., Sigma) at a concentration of 50 mg/ml in 10 mM Tris⋅Cl, pH 7.5/15 mM NaCl. Heat 15 min at 50°C. Allow to cool at room temperature. Divide into aliquots and store up to several months at −20°C. RNase T1/A stock Mix 50 µl of 500,000 U/ml RNase T1 and 50 mg of RNase A in 5 ml of 10 mM Tris⋅Cl, pH 7.5/15 mM NaCl. Heat for 15 min at 50°C. Allow to cool at room temperature. Divide into aliquots and store up to several months at −20°C. Sample buffer 625 µl 1.0 M Tris⋅Cl, pH 6.8 (62.5 mM final; see recipe) 2 ml glycerol (20% final) 2 ml 10% SDS (see recipe; 2% final) 0.5 ml 0.5% (w/v) bromphenol blue in H2O (0.025 %) 0.5 ml 2-mercaptoethanol (5% final) H2O to 10 ml Store up to several weeks at 4°C This buffer is derived from Laemmli (1970).

SDS, 10% (w/v) Dissolve 10 g of sodium dodecyl sulfate (SDS) in 90 ml of water by mixing on magnetic stirrer for several hours or by heating to 65°C to assist dissolution. Adjust the volume to 100 ml with water. Store at room temperature for up to 1 month. Staining buffer 54.2 µl water 25 µl 20× sodium cacodylate (SSC) stock solution 20 µl blotto (25% low-fat dried milk in PBS) 0.7 µl avidin-FITC (160×) 0.1 µl Triton X-100 Store up to several weeks at 4°C Stripping buffer 100 mM 2-mercaptoethanol 2% SDS 62.5 mM Tris⋅Cl, pH 6.8 (see recipe) Add 6.25 ml of 1 M Tris⋅Cl, pH 6.8; 0.77 ml β-mercaptoethanol; and 20 ml 10% SDS to 73 ml water and mix. This buffer should be prepared immediately before use.



Substrate buffer (SB) Concentrated stock: 100 mM HEPES (e.g., Boehringer Mannheim) 10% (w/v) sucrose 0.1% (w/v) CHAPS (3-[{3-cholamidopropyl}dimethylammonio]-1-propanesulfonate) (e.g., Sigma) Do not adjust pH. Divide into aliquots and store in 40-ml Falcon tubes or equivalent. Store up to 6 months at −20°C or 2 to 3 weeks at 4°C. 1× buffer: For each 96-well plate to be processed, add 25 µl of 1 M DTT (see recipe; 5 mM final) and 5 µl of 0.1% NP-40 (see recipe; 10−4% final) to 5 ml concentrated SB stock, and adjust pH to 7.25. This buffer can be kept for several hours at room temperature. TBE (Tris/borate/EDTA) buffer, 5× 445 mM Tris base 445 mM boric acid 10 mM trisodium EDTA Store at room temperature All components may be obtained from Sigma. Do not adjust pH, which should be about 8.0 at room temperature. When this solution is stored for a long time, a precipitate forms. To avoid problems, discard batches that develop a precipitate.

Transfer buffer Dissolve 22.13 g CAPS (3-cyclohexylamino-1-propanesulfonic acid) in 800 ml water. Adjust pH to 11.0 with 10 M NaOH (APPENDIX 2A). Add water to bring volume to 1 liter (0.1 M CAPS, pH 11.0), and store at 4°C. One hour prior to transfer, prepare the working solution. For one transfer, mix 100 ml of 0.1 M CAPS, pH 11.0; 700 ml water; and 200 ml methanol. Keep at 4°C to allow the exothermic reaction to cool down. Tris⋅Cl, 1.0 M, pH 6.8 Dissolve 36.3 g of Tris base in 150 ml of water; adjust pH to 6.8 with 10 M HCl. Add water to bring volume to 300 ml, and store at room temperature for up to 1 month. Tris⋅Cl, 1.5 M, pH 8.8 Dissolve 54.5 g of Tris base in 150 ml of water; adjust pH to 8.8 with 10 M HCl. Add water to bring volume to 300 ml, and store at room temperature for up to 1 month. COMMENTARY Background Information


A number of techniques have been developed to detect cytotoxicity and cell death in various cell types. The methods developed in the last two decades have been aimed at distinguishing between cell toxicity leading to apoptosis and that leading to necrosis. The choice of a particular method for analysis of cellular toxicity depends on the cell system; the toxin or toxicant; the expected mode of cell death; the type of information being sought; and, finally, technical limitations. There is no one single method that is satisfactory for obtaining all the information required. This is largely a

result of technical and practical limitations. Therefore, to make the right interpretation of data and to draw the correct conclusions regarding the mode of cell death, a combination of some of the methods mentioned above should be used. It is recommended that a combination of at least three criteria of cell death be evaluated: cell morphology (see Basic Protocol 4), DNA fragmentation (see Assays for Chromatin Cleavage), annexin V binding (see Basic Protocol 5), and/or caspase activation (see Basic Protocols 10 and 11). Although morphological changes are obviously secondary to biochemical alterations,


many of the published reports on cellular toxicity are based mainly on histological, cytological, and electron microscopy studies (Kerr et al., 1972; Bowen, 1980). The interpretation of histological data is based on the changes in membrane permeability. However, as mentioned in the introduction to this unit, analysis of plasma membrane integrity fails to identify cells in the early stages of apoptosis, although it can be used to identify necrotic cells in a population of interest. Alternative morphological methods draw on a mixture of optical and electron microscopy techniques. Combination of these methods allows one to distinguish between apoptotic changes—cell shrinkage, high cytoplasm density, membrane blebbing, compaction of chromatin, and formation of apoptotic bodies—and those indicative of necrosis—chromatin clumping, gross swelling of organelles, early membrane breakdown, and cell disintegration. In addition, confocal microscopy allows the combination of morphological analysis with subcellular localization of biochemical changes within damaged cells. Other changes that accompany cell death can be used as indicators of the mode of cell death. It has been shown that phagocytosis is a response to cell death and is particularly efficient in apoptosis. Apoptotic cells can be recognized by macrophages through a number of cell surface markers. The presence of PS on the outer plasma membrane is one such marker, and it can be detected efficiently by phagocytic cells, as blockage of PS renders phagocytosis less efficient. Therefore, PS exposure can be considered a biochemical feature of apoptosis with physiological importance (Koopman et al., 1994). Annexin V has recently been utilized as a probe to monitor changes in phospholipid distribution in the plasma membrane of cells undergoing apoptosis (see Basic Protocol 5). In combination with propidium iodine, annexin V staining can reliably discriminate apoptosis from necrosis. Flow cytometry is a useful technique to examine not only changes in the plasma membrane, but also changes in cell size and granularity (see Basic Protocol 4; Darzynkiewicz et al., 1994; Robinson et al., 1999). Apoptosis is accompanied by a dramatic loss of water and consequent cell shrinkage, which can be distinguished from necrosis by a difference in light-scatter pattern. This is a rapid technique to identify apoptotic versus necrotic cells and is very widely used nowadays. Several biochemical methods based on the detection of changes in chromatin structure have been developed. These vary from methods

for detecting single-strand DNA nicks or breaks, to those for demonstrating high-molecular-weight DNA fragments, to those for showing the subsequent internucleosomal cleavage. DNA nicks can be detected using TUNEL (see Basic Protocol 6; Gavrieli et al., 1992). However, this method apparently recognizes not only nicks formed during apoptosis, but also those formed during necrosis. Thus this method should always be used in combination with other more discriminating methods. Some other methods used for the analysis of DNA or chromatin damage appear to be better suited to discriminate between the two types of cell death. The different DNA gel electrophoresis techniques that are described here (see Basic Protocols 7 and 9; see Alternate Protocols 2, 3, and 4) can be utilized for a qualitative analysis (Wyllie, 1980; Sorenson, 1990). Although the quantitative analysis (see Basic Protocol 8) can provide limited information concerning the general cleavage of chromatin (Burton, 1956), the combination of different methods of gel electrophoresis provides more specific information concerning the type of chromatin cleavage (high-molecular-weight DNA fragments, DNA ladder, or smear). It has been shown that DNA laddering is not apparent in all apoptotic systems (Oberhammer et al., 1993a). In many types of cells, chromatin cleavage is restricted to the formation of highmolecular-weight DNA fragments (Walker et al., 1991; Oberhammer et al., 1993b). In such cases, again, the combination of two independent methods assists in drawing the correct conclusions in the study of cell death or toxicity. However, when DNA laddering occurs, it is highly indicative of apoptosis. Activation of the caspase family of proteases has been detected in numerous cell systems and appears to function as a pathway through which apoptotic mechanisms operate (for review see Zhivotovsky et al., 1997). Upon apoptotic triggering, a hierarchy of caspases is believed to become activated in a process where more proximal caspases cleave and activate downstream caspases, giving rise to a proteolytic cascade that serves to amplify the death signal. However, not all of these caspases normally function in cell death. A recent classification divided caspases into two groups, one involved in cytokine processing (caspase 1–like proteases) and the other playing a role in cell death (Alnemri et al., 1996; Cryns and Yuan, 1998). This latter group can be further subdivided according to function as activators or executioners of cell death. Up to now there



has been no evidence concerning the involvement of caspase activities in necrosis. Therefore the detection of caspase activity in cells can be used as a discriminating criterion to distinguish apoptosis from necrosis. Caspase activity is assessed by proteolytic cleavage of flurogenic substrates (see Basic Protocol 10) or by immunodetection of protein bands corresponding to active caspases (see Basic Protocol 11). In summary, a combination of morphological and biochemical methods is recommended to discriminate between the mode of cell death, especially if the manifestations of cell death or the stimuli used have not been previously characterized in the cell type of interest.



Morphological assays A characteristic feature of apoptosis is that membrane integrity of the dying cell is maintained long after the process has begun. However, a reduction in membrane integrity usually occurs during the late stages of apoptosis when the cells undergo a process commonly called “secondary necrosis” or “postapoptotic necrosis.” Therefore, a disadvantage of the trypan blue exclusion assay (see Basic Protocol 1) is underestimation of the level of apoptosis in a cell population. If the cytopsin preparations are understained after a full cycle of fixation and differential staining (see Basic Protocol 2), then the procedure can be repeated a second time until the desired staining is achieved. If overstaining is the problem, preparations can be destained by immersing them in 100% methanol for up to 5 sec, followed by rinsing with water. Because membrane blebbing is a feature common to both apoptosis and necrosis, one should take care not to use this as the sole criterion for determination of the mode of cell death. If assessing the nuclear changes in apoptotic cells is problematic, then use other conventional methods of inducing apoptosis in cells as a positive control, stain cells treated both ways with Hoechst dye (see Basic Protocol 3), and observe nuclear changes under the UV microscope. If no apoptotic changes are evident in cells, try to increase the time of incubation with apoptosis-inducing agents. If ethanol is added too quickly to the cells for flow cytometry (see Basic Protocol 4), they may clump together, which in turn may cause blockage of the flow cytometer. Fixation of the cells in ethanol may result in dehydration and

shrinkage of cells, but incubation in propidium iodide solution made in PBS rehydrates the cells. Also, if all cells dehydrate to the same extent, this should not interfere with obtaining accurate results. Binding of annexin V to PS is Ca2+ dependent, and therefore the correct Ca2+ concentration is required for optimal binding (see Basic Protocol 5). Care should be taken when analyzing necrotic cells or cells in the late stages of apoptosis, when the cells are undergoing secondary necrosis. At this stage the plasma membrane is damaged and annexin V can enter the cell and bind to the PS in the inner leaflet of the membrane. Thus, it is possible to get false-positive results. It is therefore important to run a dye exclusion assay to confirm the results. A suitable assay would be assessment of propidium iodide (PI) exclusion, since PI cannot enter cells with intact membranes and its presence can be assessed simultaneously with annexin V binding by flow cytometry. Therefore, if a cell is positive for annexin V and negative for PI, one can conclude that apoptosis is the mode of cell death. However, if doublepositive results are obtained, necrosis is implicated (see Figure 2.2.2). Chromatin cleavage assays The TUNEL technique (see Basic Protocol 6) is used to detect DNA strand breaks. There are rare situations when apoptosis proceeds without DNA degradation. Conversely, extensive DNA degradation may accompany necrosis. Thus, one should always use another independent assay, along with the TUNEL method, to confirm whether cells have undergone apoptosis. Costaining cells with propidium iodide or Hoechst is a very useful method to discriminate between apoptosis and necrosis in samples that have stained positively with the TUNEL technique. TUNEL-positive cells that express condensed, fragmented nuclei after staining with PI or Hoechst are apoptotic, whereas those with diffuse staining of the nuclei with PI or Hoechst are necrotic. If a high background is present in TUNELtreated samples, make sure that there is no DNase contamination in the test tubes or buffers. It is also possible that the number of DNA strand breaks is so large that one cannot quantify the degree of cell labeling. To eliminate this probelm, use fewer cells or decrease the incubation time. For exogenous enzymes, such as the TdT enzyme used in TUNEL, to enter the cell, the plasma membrane has to be permeable. In


many cases, the fixation procedure is sufficient to permeabilize cells; however, sometimes an additional short incubation on ice or short (1to 2-min) treatment with proteinase is required. To avoid loss of low-molecular-weight DNA, the permeabilized cells have to be fixed with formaldehyde before permeabilization. This fixation cross-links DNA fragments to other cellular constituents and prevents their leakage during the permeabilization step. The procedures for assessment of DNA fragmentation in whole cells (see Basic Protocol 7) may cause the samples to become viscous and difficult to load into gel sample wells. If this is the case, dilute samples with 1× DNA loading buffer. If no detectable DNA laddering is observed in a population of cells that are known to be apoptotic, increase the number of cells used for the assay. If it is difficult to observe distinctive bands because of high background, wash gel extensively with water. If an unusual band appears in the middle of the gel in all lanes, incubate gel longer with RNase (step 17). For analysis of fragmentation in total genomic DNA (see Alternate Protocol 2), the reagents are optimized for the stated number of cells. If larger numbers of cells are used, RNA and protein digestion will not be complete, which may interfere with electrophoresis. In addition, the procedures cause the samples to be viscous. Therefore, for minimum sheering of DNA, it is better to use pipet tips that have the ends (3 to 4 mm) cut off with a razor blade to make wide-bore tips. If no laddering is observed in DNA phenol extracted from a population of apoptotic cells (see Alternate Protocol 4), the DNA may have been lost in the organic phase during extraction. The aqueous phase should be removed carefully during extraction to prevent DNA loss. Another possible reason for failure to detect laddering may be degradation of DNA by exogenous nucleases. All solutions must be free of nuclease contamination. If, instead of a DNA ladder, a smear appears on the gel, this indicates random and general cleavage of chromatin, indicative of necrosis (see Figure 2.2.3A, lane 3). For quantitative assays of DNA fragmentation (see Basic Protocol 8), if absorbance of DNA in supernatant samples is below the sensitivity of detection, increase the number of cells. If all absorbance is present in the supernatant and absent from the pellet, then the pellet was mistakenly transferred together with supernatant to the labeled conical glass tubes (step 4). There could be several reasons for large fragments not to appear on the pulsed-field

electrophoresis gel (see Basic Protocol 9). If the material is still in the wells, the pulse generator power supply may not have been working properly. If some of the material is in the wells, but no sharp bands are visible and the material moved in the wrong direction, the current may not have been applied in the correct direction. If there is no material in the wells, the chromatin may have been digested by exogenous exonucleases during the preparation. If very little material is in the well, a bigger slab may have to be cut from the sample plugs. If too much material is in the lane, a smaller slab may have to be cut from the sample plugs. If thin stripes appear in any lanes in the gel, it means that either a bubble was in the well or the slab was not placed flat. Sometimes there is a leakage of buffer from the top reservoir, which may also influence the results. Caspase assays Release of a fluorogenic group from the peptide substrates is dependent on caspase activity and is proportional to the percentage of apoptotic cells in a cell population. In lysates of 106 cells, proportions of apoptotic cells of 5% to 10% can be detected by this method (see Basic Protocol 10). However, the lower limit of detection of caspase activity in cellular lysates of apoptotic cells varies with the kinetics of the apoptotic process, the percentage of apoptotic cells in a population, and the efficiency of cellular response to the apoptosis-inducing agent. If caspase activity is too low, first verify that the cells of interest are apoptotic using the morphological criteria described in this unit. Second, if cells are apoptotic, confirm that a sufficient number of cells and the right concentration of substrate are being used. If caspase activity is too high, use fewer cells. For immunoassays of caspase activation (see Basic Protocol 11), too many bands on the film may indicate too high a concentration of primary and/or secondary antibodies. If the background of nonspecific binding of immunological probes is unacceptably high, increase the length of time washing with HSBS and LSB. If only a band corresponding to native procaspase is on the film, verify that the cells are apoptotic using other methods described in this unit. If there is no band on the film, increase duration of incubation with primary antibodies, up to 18 hr at room temperature. If a white spot on the film makes it difficult to detect the protein band of interest, a bubble may have been introduced during blotting and the whole procedure must be repeated.



If a band on the film is too strong, decrease exposure time of X-ray film to ECL-stained membrane. If gel runs too fast or too slow and poor resolution of protein occurs, decrease or increase voltage, respectively, or check buffer recipe. If no prestained markers appear on the nitrocellulose filter, check transfer buffer recipe, or reposition the location of gel towards the cathode and anode.

Anticipated Results


Morphological assays In the trypan blue exclusion assay (see Basic Protocol 1), several thousand cells are sufficient to quantify the number of trypan blue−positive cells and to count the percentage of cells with an intact plasma membrane. Necrotic cells with rapidly disrupted membrane take up trypan blue as well. Differential staining (see Basic Protocol 2) is one of the fastest and easiest methods for identifying dead cells. Cells are viewed under a light microscope (40× magnification), and ∼300 cells are counted from separate fields of view and scored as normal, apoptotic, or necrotic based on the following morphological characteristics: membrane blebbing, chromatin condensation and nuclear shrinkage, cytoplasmic constriction and loss of cell volume, and formation of apoptotic bodies for apoptotic cells; nuclear swelling, chromatin flocculation, cell membrane blebbing and disruption, and finally cell lysis resulting in the appearance of “ghost cells” for necrotic cells (see Figure 2.2.1). Because chromatin condensation is one of the early events of apoptosis, apoptotic cells stained with Hoechst 33342 have increased blue fluorescence compared with live cells similarly stained (see Basic Protocol 3). The intensitivity of the blue fluorescence changes with the incubation time, so that an optimal incubation time should be found for each cell system for discrimination of apoptotic cells (usually 5 to 10 min). For quantification of apoptosis, a minimum of 150 cells, from 10 to 15 fields, should be evaluated for the changes described above. During flow cytometry analysis (see Basic Protocol 4), the cells with reduced FSC and SSC represent the apoptotic population, that is, apoptotic cells and apoptotic bodies. On the DNA content histograms, the subdiploid, preG1 nonstained DNA represents the DNA from dead cells. An accumulation of subdiploid DNA content is indicative of cell death. How-

ever, if cell shrinkage is also detected, this indicates that apoptosis is the predominant mode of cell death. The annexin V binding assay (see Basic Protocol 5) is very quick because no fixation or permeabilization of cells is required. Another advantage of this assay is the detection of apoptosis during the early stages of the death process. If a flow cytometer is not available to quantify annexin V–FITC binding during apoptosis, this assay can also be performed by directly visualizing the binding using a fluorescence microscope. Chromatin cleavage assays In both TUNEL protocols (see Basic Protocol 6 and Alternate Protocol 1), very little material is required for quantitative measurement of TUNEL-positive cells. Proportions of apoptotic cells as low as 10% to 15% can be detected by this method. Although the enzymatic labeling methods are time consuming (due to multiple incubation and washing steps), they are very sensitive. The method can also be applied to cryostat sections. In this case, the proteolytic pretreatment (see Alternate Protocol 1, step 4) can be shorter or even be omitted. The whole-cell DNA fragmentation method (see Basic Protocol 7) requires only 4−5 × 105 cells. It can be used for cells growing in suspension as well as attached cells. Usually, the resolution of DNA fragmentation using this protocol is distinct and high. This method does not require DNA purification and is relatively simple and fast. Although Basic Protocol 7 and Alternate Protocol 2 are equally sensitive, the latter is less time consuming. In addition, the dry loading prevents the loss of material. Alternate Protocol 3 is particularly useful for detecting chromatin cleavage in different lymphoid cells. Phenol extraction (see Alternate Protocol 4) is the most commonly used technique for investigating internucleosomal chromatin cleavage. Although it takes much more time than other techniques (almost 21⁄2 working days) and needs many more cells for preparation (2−5 × 106 cells), it yields very clean, protein-free DNA fragments. The quantitative assay of DNA fragmentation described (see Basic Protocol 8) is only one quantitative method for measurement of DNA fragmentation. This method is very simple, and very reproducible results can usually be obtained. Pulsed-field gel electrophoresis can be used to detect the presence of high-molecular-


weight DNA fragments in apoptotic cells (see Basic Protocol 9). The method is very sensitive and requires 5 × 105 cells. Depending on which system is available, one can use either horizontal or vertical gel systems. If necessary, plugs can be kept for several months and run on another gel. Caspase assays Basic Protocol 10 is a highly sensitive, rapid, convenient, and quantitative method for in vitro determination of caspase activity. It is based on the ability of caspases to proteolytically cleave a substrate to form a free fluorescent group that can be measured fluorometrically. A number of caspase substrates conjugated to alternative fluorogenic groups are now commercially available. These include AFC (7-amino-4-trifluoromethyl coumarin), MNA (4-methoxy-2naphthylamine), and pNA (p-nitroaniline). Fluorometric detection of AFC cleaved from its peptide substrate is at excitation 400 nm and emission 505 nm, and that of MNA is at excitation 340 nm and emission 425 nm. The initial rate of increase in the pNA concentration upon release from peptide can be monitored photometrically at 405 nm. Using different substrates, one can discriminate between activities of different caspases. With this method, kinetic study of enzyme activation can also be performed. The immunological method for measuring caspase activation (see Basic Protocol 11) can be used to describe and visualize the process in apoptotic cells. The method is very sensitive, and 1 × 106 cells provide enough sample for several gels. As was mentioned above, procaspases are proteolytically cleaved to become active enzymes. With the antibodies listed in the protocol, large or small subunits (cleavable fragments) of active caspases can be detected. Appearance of these fragments can serve as an early marker of apoptosis. Crude cellular extracts can also be used to detect these fragments. This nonradioactive method does not require prelabeling of proteins.

Time Considerations Morphology assays The time required for preparing solutions for trypan blue exclusion is 20 to 30 min; the solutions can then be kept up to 1 to 2 months at 4°C. Trypan blue staining takes 5 min, and calculation takes 10 min. It takes 30 min to prepare the solutions for differential staining (which can be stored 1 to

2 months at room temperature); this time is not required if the commercial kit is used. 1 hr is required to prepare and analyze samples, and 30 min to evaluate results. It takes about 30 to 40 min to prepare solutions for Hoechst staining and 1 hr to run the test. No more than 30 min are required to evaluate results. It takes 20 to 25 min to prepare solutions for flow cytometry, which can be stored up to 1 month. Sample preparation and analysis take 1 to 1.5 hr, and evaluation of results takes 30 min. The whole annexin V binding test (staining and detection) takes 30 to 40 min, and evaluation of results takes another 30 to 40 min. Chromatin cleavage assays Approximately 3 to 4 hr are required to run TUNEL assays. The evaluation of results requires an additional 1 to 2 hr, depending on the investigator’s experience. About 1 to 2 hr are required to prepare the buffers for assessment of fragmentation in whole cells, which then can be stored for at least 1 month. Usually, gel and sample preparation, which can be done simultaneously, take 2 hr, and gel electrophoresis takes 15 to 16 hr (overnight is better). Incubation with RNase, staining, and destaining take another 6 to 7 hr. Thus, the whole experiment takes almost 1.5 days. Analysis of fragmentation in total genomic DNA takes 1 working day. Preparing solutions for quantitative analysis of DNA fragmentation takes 1.5 to 2 hr; however, the solutions can be stored for up to several weeks. The procedure takes 1.5 working days and 1 hr to evaluate results. It takes about 1 week to perform the whole pulsed-field gel electrophoresis procedure. Vertical gels require a relatively shorter time (1 day less) compared with horizontal gels, with no difference in resolution. Caspase assays Approximately 2 hr are required to perform the whole substrate-cleavage procedure and an additional 2 hr to evaluate results. The immunological analysis is time consuming. Only two tests (on two membranes) may be performed simultaneously. Usually, it takes several hours to prepare solutions, which can, however, be stored for months. Gel electrophoresis, blotting, and staining with antibodies take 1.5 to 2 working days, and evaluation of results takes 1 to 2 hr.



Literature Cited Alnemri, E.S., Livingston, D.J., Nicholson, D.W., Salvesen, G., Thornberry, N.A., Wong, W.W., and Yuan, J. 1996. Human ICE/CED-3 protease nomenclature. Cell 87:171. Anand, R. and Southern, E.M. 1990. Pulsed field gel electrophoresis. In Gel Electrophoresis of Nucleic Acid: A Practical Approach, 2nd ed. (D. Rickwood and B.D. Hames, eds.) pp. 101-123. IRL Press, Oxford. Bowen, I.D. 1980. Techniques for demonstrating cell death. In Cell Death in Biology and Pathology (I.D. Bowen and R.A. Lockshin, eds.) pp. 379-444. Chapman & Hall, London–New York. Burton, K. 1956. A study of the condition and mechanism of the diphenylamine reaction for the colorimetric estimation of deoxyribonucleic acid. Biochem. J. 62:315-323. Cryns, V., and Yuan, J. 1998. Proteases to die for. Genes Devel. 12:1551-1570. Darzynkiewicz, Z., Robinson, J.P., and Crisstman, H.A. (eds.) 1994. Methods Cell Biol., Vols. 41 and 42 (Flow Cytometry: Part A and Part B, 2nd ed.). Academic Press, San Diego. Gavrieli, Y., Sherman, Y., and Ben-Sasson, S.A. 1992. Identification of programmed cell death in situ via specific labelling of nuclear DNA fragmentation. J. Cell Biol. 119:493-501. Kerr, J.F.R., Wylie, A.H., and Currie, A.R. 1972. Apoptosis: A basic biological phenomenon with wide ranging implications in tissue kinetics. Br. J. Cancer 26:239-257. Koopman, G., Reutelingsperger, C.P.M., Kuijten, G.A.M., Keehnen, R.M.J., Pals, S.T., and vanOers, M.H.J. 1994. Annexin V for flow cytometric detection of phosphatidylserine expression on B-cells undergoing apoptosis. Blood 84:14151420. Laemmli, U.K. 1970. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227:680-682 McGahon, A.J., Martin, S.J., Bissonnette, R.P., Mahboubi, A., Shi, Y., Mogil, R.J., Nishioka, W.K., and Green, D.R. 1995. The end of the (cell) line: Methods for the study of apoptosis in vitro. Methods Cell Biol. 46:153-185. Oberhammer, F., Fritsch, G., Scmied, M., Pavelka, M., Printz, D., Purchio, T., Lassman, H., and Schulte-Hermann, R. 1993a. Condensation of the chromatin at the membrane of an apoptotic nucleus is not associated with activation of an endonuclease. J. Cell Sci. 104:317-326. Oberhammer, F., Wilson, J.W., Dive, C., Morris, I.D., Hichman, J.A., Wakeling, A.E., Walker, R.A., and Sikorska, M. 1993b. Apoptotic death in epithelial cells: Cleavage of DNA to 300 and/or 50 kb fragments prior to or in the absence of internucleosomal fragmentation. EMBO J. 12:3679-3684.

Pollak, A. and Ciancio, G. 1990. Cell cycle phase– specific analysis of cell viability using Hoechst 33342 and propidium iodide after ethanol preservation. Methods Cell Biol. 33:19-24. Robinson, J.P., Darzynkiewicz, Z., Dean, P.N., Dressler, L.G., Orfao, A., Rabinovitch, P.S., Stewart, C.S., Tanke, H.J., and Wheeless, L.L. 1999. Current Protocols in Cytometry. John Wiley & Sons, New York. Samali, A., Nordgren, H., Zhivotovsky, B., Peterson, E., and Orrenius, S. 1999. A comparative study of apoptosis and necrosis in HepG2 cells: Oxidant-induced caspase inactivation leads to necrosis. Biochem. Biophys. Res. Commun. 255:611. Skalka, M., Matyasova, J., and Cejkova, M. 1976. DNA in chromatin of irradiated lymphoid tissues degrades in vivo into regular fragments. FEBS Lett. 72:271-275. Sorenson, C.M., Barry, M.A., and Eastman, A. 1990. Analysis of events associated with cell cycle arrest at G2 phase and cell death induced by cisplatin. J. Natl. Cancer Inst. 92:749-755. Thornberry, N.A. 1994. Interleukin-1β-converting enzyme. Methods Enzymol. 244:615-631. Thornberry, N.A., Rano, T., Peterson, E., Rasper, D., Timkey, T., Garcia-Calvo, M., Houtzager, V., Nordstrom, P., Roy, S., Vaillancourt, J., Chapman, K., and Nicholson, D. 1997. A combinatorial approach defines specificities of members of the caspase family and granzyme B: Functional relationships established for key mediators of apoptosis. J. Biol. Chem. 272:17907-17911. Walker, P.R., Smith, C., Youdale, T., Leblamc, J., Whitfield, J.F., and Sikorska, M. 1991. Topoisomerase II–reactive chemotherapeutic drugs induce apoptosis in thymocytes. Cancer Res. 51:1078-1085. Wyllie, A.H. 1980. Glucocorticoid-induced thymocyte apoptosis is associated with endogenous endonuclease activation. Nature 284:555-556. Yamada, T. and Ohyama, H. 1980. Separation of the dead cell fraction from X-irradiated rat thymocyte suspension by density gradient centrifugation. Int. J. Radiat. Biol. 37:695-699. Zhivotovsky, B., Gahm, A., Ankarcrona, M., Nicotera, P., and Orrenius, S. 1995. Multiple proteases are involved in thymocyte apoptosis. Exp. Cell Res. 221:404-412. Zhivotovsky, B., Burgess, D.H., Vanags, D.M., and Orrenius, S. 1997. Involvement of cellular proteolytic machinery in apoptosis. Biochem. Biophys. Res. Commun. 230:481-488.

Contributed by Boris Zhivotovsky, Afshin Samali,and Sten Orrenius Karolinska Institute Stockholm, Sweden



Detection of Covalent Binding

UNIT 2.3

Earlier studies of protein covalent binding were based primarily on detection of covalently bound metabolites from radioactive xenobiotics that were administered in vivo or added to in vitro preparations. Generally, such studies involved repetitive extraction of protein suspensions to remove unbound drug or metabolites. The radioactivity that remained with the protein fraction after the extraction procedure was considered to be covalently bound. Such approaches are useful for quantifying bound metabolites in tissue homogenates or subcellular fractions and for localizing bound metabolites in tissue sections by autoradiography. However, the radiometric approach is generally not practical for studies aimed at the identification of xenobiotics bound to individual protein targets. Mechanistic research in toxicology today is not merely interested in quantification and localization of bound drug. Rather, there is increasing interest in identifying the specifically targeted proteins as a key step to clarifying the mechanistic contributions of the binding to the ensuing toxicity. To this end, immunochemical methods have been developed to detect and identify xenobiotics covalently bound to cellular proteins. Binding to individually targeted proteins as detected immunochemically has been shown to be better associated with toxicity than total covalent binding as assessed radiometrically (Beierschmitt et al., 1989). This unit presents two protocols for the immunochemical detection of xenobiotic covalent binding to proteins. The first (see Basic Protocol 1) outlines the use of specific antibodies for detection of individual xenobiotic-protein adducts on immunoblots, and the second (see Basic Protocol 2) outlines the use of the antibodies in immunohistochemical analysis of fixed tissue sections. Support protocols are included to provide direction for the synthesis of an artificial antigen (see Support Protocol 1) and subsequent immunization (see Support Protocol 2) of a suitable host species (e.g., rabbit or goat), and for affinity purification and characterization of the antibodies in noncompetitive and competitive ELISA (see Support Protocols 3 to 7). The details of antibody development will vary with the nature of the bound drug or metabolite, i.e., the hapten. As an example of this process, a method to elicit production of antibodies against the analgesic/antipyretic acetaminophen (N-acetyl-p-aminophenol, APAP) is described (Support Protocols 1 and 2). The various methods described in this unit emphasize general considerations and salient features. For a more comprehensive review of the theory on individual topics, the reader is referred to additional sources (Taleporos and Ornstein, 1976; Towbin and Gordon, 1984; Tijssen, 1985; Harlow and Lane, 1988; Login and Dvorak, 1988; Ostrove, 1990; Osborn and Isenberg, 1994). DETECTION OF COVALENT BINDING BY IMMUNOBLOTTING The basic principle of immunoblotting is the use of an electrical field to transfer a complex mixture of proteins from a gel matrix to a membrane support (nitrocellulose, nylon, or PVDF) where proteins are thought to be retained by noncovalent hydrophobic interactions. Immunoblotting thus maximizes the resolving power of polyacrylamide gel electrophoresis (SDS, native, or isoelectric focusing; see APPENDIX 3A) to provide a discrete pattern of proteins bound to a membrane for further characterization and identification by immunodetection or protein sequencing. Two types of equipment are most widely used, and there are advantages and disadvantages to each. Wet transfer consumes ∼4 liters of buffer and requires complete immersion of the gel-membrane sandwich. For semidry transfer, the sandwich is placed between absorbent paper that has been soaked in transfer

Contributed by Mary K. Bruno and Steven D. Cohen Current Protocols in Toxicology (1999) 2.3.1-2.3.35 Copyright © 1999 by John Wiley & Sons, Inc.

BASIC PROTOCOL 1


2.3.1

buffer. Wet transfer generally offers greater flexibility with respect to temperature control, voltage settings, and blotting times without buffer depletion; however, reagent consumption, cost, and experimental time are greater. The procedure described in this protocol has been employed for routine immunochemical detection of APAP-protein adducts in tissue homogenates or subcellular fractions (Bartolone et al., 1987, 1988) The following protocol is for wet transfer of proteins from full-size (12 × 14.5–mm) 10% SDS-PAGE gels (0.75-mm thickness)—see APPENDIX 3A for electrophoresis protocols. It incorporates the specific instructions for use of equipment from Bio-Rad, but should be generally applicable. Modifications may be required if different equipment or a semidry blotting method are used. SDS-PAGE and steps 1 to 11 of the immunoblot protocol must be done on the same day. Incubation with primary antibody and subsequent detection of bound antibody may be done at a later time. Inclusion of prestained molecular weight protein standards during electrophoresis is recommended for orientation of the membrane during and after transfer. Gloves should be used throughout these procedures to protect the operator and to prevent contamination of gels with skin proteins. CAUTION: If radioactivity will be used for detection, the user is required to have received institutional training in the handling of radioisotopes, and the experiment must be performed in an area designated for radioactivity use under the supervision of a licensed investigator. All steps must be carried out in accordance with the Nuclear Regulatory Commission (NRC) and institutional guidelines (also see APPENDIX 1A). Materials SDS-PAGE gel with resolved proteins Transfer buffer (see recipe) Methanol Coomassie brilliant blue solution (see recipe) Destaining solution (see recipe) Blocking buffer (see recipe) Primary antibody against protein-xenobiotic adduct (Support Protocol 2), affinity-purified if necessary (see Support Protocol 6) Tris-buffered saline (TBS; see recipe) containing 0.05% (w/v) Tween 20 (store up to 1 week at room temperature) Secondary antibody: 125I-, alkaline phosphatase–, or horseradish peroxidase–conjugated IgG raised against the host animal used for production of the primary antibody 10 mg/ml nitroblue tetrazolium (prepare fresh) 25 mg/ml 5-bromo-4-chloro-3-indolyl-phosphate (BCIP) in N,N-dimethylformamide (prepare fresh) Developing buffer (see recipe) 4-chloro-1-naphthol solution (see recipe) 30% hydrogen peroxide (store up to 1 month at 4°C) Chemiluminescence-based detection system: e.g., ECL Kit (Amersham) or SuperSignal Kit (Pierce Chemical)


Trans-Blot apparatus (Bio-Rad) including: Fiber pads Transfer tank Cassette gel holder Transfer membranes: nitrocellulose (0.2 to 0.45 µm pore size) or polyvinylidene difluoride (PVDF; 0.45 µm pore size) 23 × 33–cm and 21.5 × 21.5–cm glass baking dishes continued


Whatman no. 1 filter paper Platform shaker or rocker Kodak X-Omat AR or equivalent 8 × 10–in. (20.32 × 25.4–cm) X-ray film Lead autoradiography cassette with intensifying screen Additional reagents and equipment for SDS-PAGE (APPENDIX 3A) Transfer proteins 1. Perform SDS-PAGE (APPENDIX 3A), including the appropriate prestained molecular weight standards on the gel. Approximately 30 min prior to the completion of SDS-PAGE, presoak fiber pads (included with Trans-Blot apparatus) in transfer buffer in a 23 × 33–cm glass dish (e.g., a baking dish). Prestained molecular weight standards should always be included during gel electrophoresis to serve as a reference on the membrane. These are readily available from several commercial sources. The pH of the transfer buffer ranges from 8.1 to 8.3, depending on the quality of the reagents used. The inclusion of SDS in the transfer buffer facilitates complete transfer of proteins of 150 kDa) or gels of higher percentage acrylamide and thickness. Decreased blotting times may be necessary for gels of lower protein load, and to prevent the loss of low-molecular-weight proteins by passage completely through the membrane. One can determine if this has occurred by adding a second membrane to the sandwich behind the first. Transfer time should be adjusted accordingly if proteins are detected on the second membrane.

Probe the membrane with antibodies 12. Disassemble the cassette. Stain blotted gel 20 min in Coomassie brilliant blue solution, then destain 1 to 2 hr in destaining solution (until most of the stain has been removed). If transfer was complete, only bands corresponding to higher molecular weight (>150 kDa) proteins should remain.

13. Block the membranes by incubating 1 hr at room temperature in the appropriate blocking buffer (see Reagents and Solutions for alternatives), then pour off the blocking buffer prior to adding the primary antibody. Nitrocellulose membranes may be allowed to dry prior to blocking and probing with antibody. PVDF membranes should be placed in blocking buffer for 15 to 60 min before drying. Alternatively, if PVDF membranes are allowed to dry without soaking in buffer, they must be prewetted again with methanol prior to blocking and probing with antibody. Drying the membranes is preferable to storing them in transfer buffer or blocking buffer overnight. Such storage in aqueous solutions will result in protein desorption, since binding to the membrane is not covalent.


Expose membrane to antibodies 14. Dilute the primary antibody (1:50 to 1:100 for affinity-purified anti-APAP antibody) in 50 ml of blocking buffer (or other volume sufficient to cover the membrane and prevent it from drying out, depending on the size of the membrane and container being used) and apply to the membrane. Incubate 3 hr at room temperature with shaking on a platform shaker.


Antibody dilutions and incubation times will vary depending on the antibody titer and the amount of antigen present. Antibodies of low avidity dissociate more rapidly from the antigen-antibody complex; therefore, lower dilutions and shorter incubation times are recommended. Antibodies of higher avidity can tolerate higher dilutions and longer incubation times (e.g., 18 to 24 hr at 4°C; Towbin and Gordon, 1984).

15. Pour off the antibody solution and wash the membrane three times, each time by immersing for 10 min in 50 to 100 ml TBS containing 0.05% Tween 20, with shaking on a platform shaker. The antibody solution may be reused if stored at 4°C with a preservative (e.g., 0.02% sodium azide or 0.01% merthiolate). Sodium azide inhibits horseradish peroxidase (HRP) and thus cannot be used if the secondary antibody to be used for immunodetection is conjugated with this enzyme.

16. Dilute the secondary antibody in blocking buffer and incubate the membrane for 90 min using the technique described in step 14. Dilution of the secondary antibody will vary depending on the type of conjugate (radiolabel or enzyme for colorimetric or chemiluminescent detection) and the commercial supplier. With anti-APAP, alkaline phosphatase and 125I-conjugated secondary antibodies have been used at 1:2000 and 0.1 ìCi/ml blocking solution, respectively. For chemiluminescent detection systems, e.g., ECL, follow protocols supplied by the manufacturer. If using a chemiluminescence system for detection, the length of the incubation with the secondary antibody may be shortened (e.g., 20 min).

17. Pour off the antibody solution and wash three times as described in step 15. Detect bound antibody 18a. For 125I-labeled secondary antibody: Allow membrane to dry. Enclose membrane in plastic wrap and expose to Kodak X-Omat AR film in a lead autoradiography cassette with an intensifying screen for 24 to 40 hr at −70°C. Bring the cassette to room temperature before removing and developing the film. All manipulations involving film should be done in a darkroom. The detection limit of this method is 10 ng (Promega, 1993). Radioactive and chemiluminescence-based detection systems have the advantage of ease of quantification. They also permit the membrane to be stripped of adsorbed antibodies and then probed again.

18b. For alkaline phosphatase–conjugated secondary antibody: Just before use, combine 0.5 ml of 10 mg/ml nitroblue tetrazolium and 0.5 ml of 25 mg/ml BCIP with 50 ml of developing buffer. Incubate the blotted membrane with this substrate-chromogen solution until color development of specific bands is observed. Stop the reaction by rinsing with water. The 50 ml prepared here is sufficient volume for one full-size gel. The reaction product is an intense purple-black precipitate at the site of enzyme-conjugate binding. The background may be lavender, depending on the development time, but will revert to white upon complete drying overnight. The detection limit of this method is ∼50 to 100 pg (Hoefer, 1994; Promega, 1993).

18c. For horseradish peroxidase–conjugated secondary antibody: Just before use, add 20 µl of 30% hydrogen peroxide to 48 ml 4-chloro-1-naphthol solution. Incubate the blotted membrane with this substrate-chromogen solution until color development of specific bands is observed. Stop the reaction by rinsing with water. The reaction product is a brown-black precipitate at the site of enzyme-conjugate binding. The color intensity tends to fade with time and thus does not provide a good permanent record. The detection limit for this method is 1 ng (according to Promega technical applications guide).



18d. For enzyme-linked secondary antibodies, using chemiluminescence: Follow instructions supplied by the manufacturer of the chemiluminescence detection kit being used. For example, after washing the membrane (step 17), incubate 1 min with the mixed ECL detection reagents when using the Amersham ECL system. Drain off excess solution, then cover the membrane with plastic wrap, remove air bubbles, and place in autoradiography film cassette with the protein side facing toward the film. Expose film for optimal time required for visualization of bands, then develop film. It is best to perform these steps in the darkroom, turning the lights off when exposing and developing film. Work as quickly as possible following the addition of the detection solution. For a given batch of antibody used under standardized conditions, the time of film exposure that gives the greatest signal-to-noise ratio will remain fairly constant. This must be determined empirically when assay conditions and reagents are changed. For anti-APAP, film-exposure times typically range between 2 and 10 min. Because of the stability and sensitivity of the reagents, the same membrane may be exposed sequentially to new film for varied lengths of time. This allows for optimization of signal detection of adducts on the film. The intensity of the signal is maximal between 1 and 5 min, with a half-life of decay of 1 hr. The limit of detection is as low as 1 pg of antigen. Radioactive and chemiluminescence-based detection systems have the advantage of ease of quantification. They also permit the membrane to be stripped of adsorbed antibodies and then probed again. BASIC PROTOCOL 2

DETECTION OF COVALENT BINDING BY IMMUNOHISTOCHEMISTRY Immunohistochemistry has become increasingly important diagnostic and experimental techniques to determine the intracellular localization of a particular antigen. Methods of tissue preparation are versatile and often empirically determined based upon the tissue type and antigen-antibody interaction. Similarly, morphological examination of tissue sections can utilize light, fluorescence, or electron microscopy. The method described below localizes APAP-protein adducts in tissue sections prepared by microwave fixation and processed for immunohistochemistry (Emeigh Hart et al., 1995) Kits other than the AS/AP kit used here, which incorporate enzyme-based detection (alkaline phosphatase or peroxidase) in combination with streptavidin/avidin and biotin interactions offer greater sensitivity and are highly recommended over the traditional enzyme-based detection systems. Such kits are available from BioGenex and Vector Laboratories. Materials Tissue sections (1 mm thickness) containing protein-xenobiotic adducts 0.9% NaCl (APPENDIX 2A) 50%, 95%, and 100% ethanol Xylene Tris-buffered saline (TBS; see recipe) 0.5% pepsin in 0.1 N HCl (prepare fresh), prewarmed to 37°C Nonimmune serum from species in which secondary antibody was raised Primary antibody: anti-APAP (see Support Protocol 2), affinity purified (see Support Protocol 6) TBS (see recipe) containing 0.5% bovine serum albumin (BSA) AS/AP Universal Rabbit Detection System (Bio-Can Scientific; or equivalent detection system employing alkaline phosphatase–conjugated anti-rabbit secondary antibody)


continued


Double-strength (2×) Gill’s hematoxylin (see recipe or purchase from Polyscientific) Ammonium hydroxide Aqueous mounting medium (CrystalMount from BioMedia or equivalent) Coverslips Tissue-processing cassettes Microwave oven with temperature probe Programmable automated tissue processor (e.g., Model LX-120, Innovative Medical Systems) Equipment for paraffin embedding and sectioning (e.g., dedicated histology laboratory) Vacuum desiccator Poly-L-lysine-coated glass microscope slides (see recipe) 60°C and 80°C drying ovens Coplin jars Glass staining pan with metal slide rack Fix tissue 1. Trim tissue samples to 1 mm thickness and place into plastic tissue-processing cassettes. Tissue samples >1 to 2 mm in any dimension may be damaged by conductive heat during microwave fixation. Also, in such sections, chemical fixatives will penetrate too slowly for proper fixation.

2. Place up to five cassettes into a beaker containing 150 ml of 0.9% NaCl and insert into a microwave oven equipped with a temperature probe. Samples must be immersed in the buffer or buffered fixative to prevent dessication and loss of antigenicity during microwave irradiation. If samples are instead contained in individual vials, a beaker containing 300 to 400 ml water must also be placed in the microwave oven. Irradiating a small sample volume without the additional water load will lead to overheating of the samples. The water load will improve the reproducibility of microwave fixation conditions (Login and Dvorak, 1988).

3. Place the temperature probe into the beaker and microwave 5 min at 60°C. The time and temperature for microwave fixation may be varied as appropriate and should be optimized for tissue type, antibody, chamber design, and power output of the microwave oven. Samples immersed in a fixative rather than a saline solution will require much shorter microwave irradiation (e.g., seconds).

4. Transfer cassettes quickly to a beaker containing 10 vol of 50% (v/v) ethanol per vol tissue. Rapid replacement of the hot fixing buffer with the ethanol solution is necessary to prevent overfixation. Samples may be stored in 50% ethanol until further processing is initiated. Duration of storage without loss of antigenicity and architecture may vary and should be determined experimentally.

Process samples for histology 5. Dehydrate tissues using a programmable automated tissue processor, omitting the alcohol-formalin step. Embed tissues in paraffin and store in a vacuum dessicator until ready to section. These operations will normally be performed in a dedicated histology laboratory.



6. Cut 5-µm sections. Mount sections on poly-L-lysine coated glass microscope slides by warming in a drying oven set at 60°C for 30 min. Proper adhesion of sections to slides requires: (1) clean slides, (2) contact of the entire section with the glass surface, and (3) thorough drying. Coating with positively charged poly-L-lysine will prevent detachment of the negatively charged tissue sections. This is necessary if the sections will be subsequently treated with proteases. Tissue sections with folds and wrinkles will not have good contact with the slide surface and will adhere poorly.

7. Rehydrate and deparaffinize tissue sections by immersing sequentially, 5 min each, in Coplin jars containing the following solutions: Xylene Xylene1 100% ethanol 100% ethanol 95% ethanol 95% ethanol 70% ethanol. 8. Transfer the slides to a staining rack and rinse with running tap water in a staining pan for 5 min. The position of the tissue section can be marked with a diamond pen or water-repellent marker. If peroxidase-antiperoxidase will be used for immunodetection, treat slides with 3% hydrogen peroxide for 15 min after the water rinse to quench endogenous peroxidase activity. Slides are then washed with tap water after the peroxide quench. Metal staining racks should not be used to support slides during hydrogen peroxide staining.

9. Immerse slides in TBS for 5 min. Treat slides with pepsin 10. Incubate slides 15 min at 37°C in prewarmed 0.1 N HCl containing 0.5% pepsin. This treatment promotes the accessibility of epitopes for antibody recognition. Avoid metal racks and containers during the digestion step.

11. Rinse slides in running tap water for 5 min, then immerse slides in TBS for 5 min. Blot slides to remove most liquid, leaving the tissue section moist. This will prevent further protease digestion.

Block tissue 12. Place slides in incubation tray with the tissue section facing upward. Block the tissue by covering the section with nonimmune serum (diluted 1:50), collected from the same species used to produce the secondary antibody, then incubating 30 min at room temperature. Use sufficient volume to cover the tissue section. Pretreatment with nonimmune serum will reduce the nonspecific binding of the subsequently applied antibodies.

Incubate in primary antibody 13. Remove blocking serum from the slide by shaking and replace with affinity-purified anti-APAP antibody diluted 1:10 in TBS containing 0.5% BSA. Use sufficient volume to cover the tissue section. Allow antibody to bind for 18 to 21 hr at 4°C. A dilution series is recommended to establish the appropriate concentration of primary antibody. Most polyclonal antibodies can be diluted 1:20 to 1:100; affinity-purified antibodies may be used in the range of 5 to 20 ìg/ml. Incubation of slides in a humidified atmosphere will prevent dehydration. Detection of Covalent Binding

Also include the appropriate controls (see Critical Parameters and Troubleshooting).


14. Wash slides three times with TBS in Coplin jars, each time for 5 min with shaking. Separate jars must be used for slides incubated with different immunoreactants, to prevent cross-contamination. Tween 20 (0.05% w/v) may be included in the wash steps.

Incubate with enzyme-conjugated secondary antibody 15. Blot slides dry and lay flat in a staining tray. Place 2 to 4 drops of alkaline phosphatase–conjugated secondary antibody, provided with the AS/AP Universal Rabbit Detection System from Bio-Can Scientific, on the tissue section and incubate 30 min at room temperature. Wash slides as in step 14 If a kit is not used, the concentration of secondary antibody may be determined experimentally. Typical dilutions range between 1:50 and 1:150. Peroxidase-conjugated secondary antibody may be used instead of alkaline phosphatase.

Visualize staining 16. Blot slides and return to staining tray. Incubate with the chromogenic substrate supplied with the AS/AP kit for 20 min or until color development is visible. 17. Rinse slides thoroughly in running water for 5 min. Counterstain by immersing for 3 min in double-strength (2×) Gill’s hematoxylin, then rinse thoroughly in running water for 5 min and blot dry. It is advisable to purchase prepared staining solutions (e.g., from Polyscientific) as commercial preparations give more uniform results. Conditions for storage (usually room temperature) and expiration dates are provided on the label. Also note that some histology laboratories favor Mayer’s hematoxylin over Gill’s formulation, due to the blue staining of the nucleus and the contrast provided when used with other staining protocols.

18. Dip slides 10 times in dilute ammonium hydroxide (6 to 8 drops ammonium hydroxide in 300 ml water). Rinse thoroughly in running tap water. Slides will turn blue.

19. Place 3 to 4 drops of aqueous mounting medium (e.g., CrystalMount) over each tissue section. Tip slides as required to coat each section uniformly with a thin film. Avoid the use of organic solvents which may solubilize some chromogens.

20. Place slides on a metal tray and incubate in oven 15 min at 80°C. Add coverslips after cooling. Slides may be examined under the microscope at this step. Compare slides from experimental groups to those from positive and negative controls that were run through the process simultaneously (see Commentary). For photography, permanently mount cover slips with Permount (Fisher).

PREPARATION OF APAP ANTIGEN This method involves the use of a linker molecule (p-aminobenzoic acid, PABA) between the hapten (acetaminophen, also called N-acetyl-p-aminophenol or APAP is used as the example here) and the carrier protein, keyhole limpet hemocyanin (KLH). Since the 3 and 5 positions of the aromatic ring of APAP have been identified as sites for protein-adduct formation, the coupling chemistries were selected to retain the natural binding conformation. Construction of the antigen requires a three-step procedure (Fig. 2.3.1) which involves (1) diazotization of the amino group of PABA, (2) conjugation of the diazonium cation of PABA to APAP at the nucleophilic 3 or 5 positions of the APAP ring, and (3) coupling of the hapten-conjugate (APAP-PABA) to the carrier protein KLH by the mixed anhydride method, where an amide bond is introduced between the carboxyl

SUPPORT PROTOCOL 1



group of PABA and the free amino groups on KLH (Bartolone et al., 1988). Radiometric methods are used to monitor formation of the hapten conjugate and to determine epitope density. CAUTION: If radioactivity will be used for detection, the user is required to have received institutional training in the handling of radioisotopes, and the experiment must be performed in an area designated for radioactivity use under the supervision of a licensed investigator. All steps must be carried out in accordance with the Nuclear Regulatory Commission (NRC) and institutional guidelines (also see APPENDIX 1A). NOTE: All aqueous solutions should be made in water of high purity (i.e., double-distilled, or deionized using Milli-Q or equivalent filtration system). Materials p-aminobenzoic acid (PABA) 0.1 N HCl, 4°C 1% (v/v) sodium nitrite

O H N C CH 3 N=N+

NH2

acetaminophen (APAP)

NHC NaNO2

OH O

C C OH O OH O p-aminobenzoic diazonium cation of PABA acid (PABA)

H

N

C CH 3 APAP-PABA

O

CH3 CI-C-O-CH2 -CH CH3 isobutyl chloroformate

OH

N=N

C OH

OH

O H

N

C CH 3 O N=N

mixed anhydride intermediate

O

C-O-C-O-C4H9

OH O keyhole limpet hemocyanin (KLH)

H

N

C CH 3

APAP-PABA-KLH antigen

O N=N OH


C-N-KLH H

Figure 2.3.1 Synthetic scheme for formation of acetaminophen (APAP) antigen.


Source of starch (e.g., 1% starch solution, starch paper, or slice of potato) saturated with freshly prepared 50 mM potassium iodide Acetaminophen (APAP; Sigma; minimum purity, 99%) 0.5 M carbonate/bicarbonate buffer, pH 9.0 (see recipe) 3 [H]acetaminophen ([3H]APAP; 15.5 Ci/mmol, uniformly labeled; must be custom-synthesized) 1.0 N and 0.1 N NaOH N,N-dimethylformamide Keyhole limpet hemocyanin (KLH) Isobutyl chloroformate (Aldrich) Phosphate-buffered saline (PBS), pH 7.0 to 7.2 (see recipe) 6 N perchloric acid 50 mg/ml bovine serum albumin (BSA; store up to 24 hr at 4°C) 80% methanol containing 5 mM unlabeled APAP, ice-cold Nitrogen source Tissue solubilizer (e.g., Solvable from NEN Life Science or Amplify from Amersham; optional) Acetone/dry ice or ethanol/dry ice bath Lyophilizer 6000 to 8000 MWCO dialysis tubing (pretreated; see recipe) and dialysis clamps Diazotize p-aminobenzoic acid (PABA) 1. Dissolve 4 mmol (548 mg) of PABA in 100 ml 0.1 N HCl at 4°C. This is best done with stirring in a large, e.g., 250-ml, beaker.

2. Add 1% (v/v) sodium nitrite solution dropwise with stirring until a yellow-green color appears. Test for completeness of the titration by placing the solution dropwise onto a source of starch, e.g., a slice of potato saturated with freshly made 50 mM KI solution, and observing for the appearance of a blue-black color which should form on the potato in ∼1 min. Allow the solution to stir for an additional 15 min and retest. If starch test remains positive, proceed to step 3. If the starch test is negative, add more sodium nitrite solution, avoiding large excesses, and retest. Repeat as required. PABA is converted to the diazonium ion in the presence of nitrous acid. When diazotization approaches completion, the excess free nitrous acid can be detected by a positive starch test whereupon iodide is oxidized to iodine yielding a blue-black color in the presence of starch. Alternative sources of starch include 1% starch plus 50 mM potassium iodide or starch paper saturated with 50 mM potassium iodide. Free nitrous acid should remain detectable 15 min after the last addition of sodium nitrite, as determined by a positive starch reaction. Since the reaction between PABA and sodium nitrite is stoichiometric, this offers a test for completion of diazotization (Nisonoff, 1967). The reaction should consume ∼27 ml of sodium nitrite solution.

Derivatize APAP to diazotized PABA 3. Dissolve 4 mmol (600 mg) of APAP in 20 ml 0.5 M carbonate/bicarbonate buffer, pH 9.0, with heating at 50° to 55°C. Add 5 µCi 3[H]APAP to determine epitope density (see step 15). Prepare the solution just before use to avoid problems with overheating or crystallization upon cooling. APAP has limited solubility at this concentration (200 mM), and thus it is necessary to heat to 50° to 55°C. At pH 9.0, the solution of APAP will be pink.

4. Using a Pasteur pipet, slowly add the APAP solution to the diazotized PABA from step 2. As the solution approaches neutral pH, the yellow-green solution of PABA will turn red.



5. Quickly adjust the pH to 9.5 with 1.0 N NaOH. The color will deepen to a dark red. It is critical for successful derivatization that the pH not decrease below 9.0.

6. Stir the solution at 4°C, periodically checking the pH. 7. Shell freeze in an acetone/dry ice bath or ethanol/dry ice bath, then lyophilize to dryness. Lyophilization may take 24 to 36 hr.The product should be a fine powder. It is critical that no water be present for the subsequent steps.

Conjugate APAP-PABA to KLH 8. Set up an ice-water bath in a well-ventilated hood. Maintain temperature at 10°C. CAUTION: The following reaction is exothermic. Combine reagents slowly. Take appropriate safety precautions and use chemical-resistant latex gloves.

9. Dissolve the lyophilized APAP-PABA powder in 9.0 ml N,N-dimethylformamide and cool to 10°C. 10. Dissolve 50 mg KLH in 50 ml of 50 mM carbonate/bicarbonate buffer, pH 9.0. KLH dissolves slowly.

11. Add 260 µl isobutyl chloroformate to the solubilized APAP-PABA from step 9 and stir for 30 min at 10°C. IMPORTANT NOTE: It is essential to avoid water. The reaction of isobutyl chloroformate with the carboxyl group of PABA-APAP to form the mixed anhydride is water-sensitive. It is also advisable to conduct the reaction in an environment with low atmospheric humidity. Anhydrides can be rapidly hydrolyzed. If water is present, competition between the acylation and hydrolysis will reduce product yield.

12. Slowly add the KLH solution prepared in step 10 to the acid anhydride of PABAAPAP formed in step 11. Stir for 4 hr while maintaining the temperature at 10°C throughout the reaction. 13. Dialyze the antigen 2 days against PBS, pH 7.0 to 7.2, at 4°C, with at least 5 changes of buffer, using pretreated 6000 to 8000 MWCO dialysis tubing. The dialysate color will decrease in intensity with repeated changes of buffer.

14. Remove two 1.0-ml aliquots for determination of epitope density (see steps 15 to 21) and place in 1.5-ml microcentrifuge tubes. Divide the remaining antigen solution into 1-ml aliquots and store at −70°C. Antigen may be stored for several months at this temperature. Alternatively it may be stored up to 2 weeks at 4°C. The amount of protein adduct used for immunization can be determined by any protein assay presently used in the laboratory—e.g., Lowry or Bradford.

Determine epitope density 15. In each of the two 1.0-ml aliquots taken from the antigen solution (step 14), acid-precipitate the protein with 0.2 ml of 6 N perchloric acid and add 100 µl of 50 mg/ml BSA. Let stand on ice for 10 min, then microcentrifuge 5 min at 14,000 × g, 4°C. The BSA is added as a carrier protein to prevent loss of 3[H]APAP-KLH.

16. Remove supernatants and discard. Resuspend pellets by vortexing in 1.0 ml ice-cold 80% methanol containing 5 mM unlabeled APAP. 17. Add 30 µl of 1.0 N NaOH to the methanol solution/resuspended pellet and confirm that the pH is alkaline with pH paper. Detection of Covalent Binding

This step is necessary since BSA is soluble in acidic methanol. The methanol washes remove residual APAP that is not covalently bound to KLH.


18. Let stand on ice 10 min, and microcentrifuge 5 min at 14,000 × g, 4°C. Repeat methanol wash (removal of supernatants followed by addition of methanol/APAP and microcentrifugation) twice more. 19. Evaporate the residual methanol solution remaining in the pellet under nitrogen. 20. Solubilize the pellet in 0.2 ml of 0.1 N NaOH or a commercially available tissue solubilizer. 21. Determine the amount of 3[H]APAP associated with KLH by liquid scintillation spectrometry. Due to the multisubunit structure and size heterogeneity of KLH, epitope density is best expressed as nmol APAP bound per ìg KLH protein. The following is a sample calculation. Starting material: 4 mmol APAP and 5 ìCi [3H]APAP. Reaction volume: 9.0 ml (step 9) + 50 ml (step 10) containing 50 mg KLH Sample volume: 1.0 ml Specific activity of [3H]APAP: 11.0 × 106 dpm/4 mmol APAP = 2730 dpm/ìmol 1 ml sample volume would contain 847 ìg KLH with a measured amount (x) of [3H]APAP. The number of ìmol APAP bound per ìg KLH is determined as follows: x/2730 (specific activity of [3H]APAP)/847 ìg KLH.

IMMUNIZATION AND SERUM COLLECTION TO PREPARE ANTI-APAP POLYCLONAL ANTIBODIES

SUPPORT PROTOCOL 2

Historically, immunization of female New Zealand rabbits has been done with the synthetic antigen (see, e.g., Support Protocol 1) and Freund’s adjuvant. Freund’s adjuvant results in the formation of granulomas and ulcers at the sites of injection. Alternative adjuvants that are less injurious are now widely available and should be used in accordance with the protocols supplied by the manufacturers (Adjuvax from Alpha-Beta Technology; Hunter’s Titermax from CytRx; and RAS from RIBI Immunochemical Research). Two or more rabbits should be immunized with the hapten conjugate prepared in Support Protocol 1, since not all animals will respond equally to the antigen. New Zealand rabbits that are outbred and commonly used for antibody production exhibit varied response to APAP-PABA-KLH. All personnel should receive prior instruction regarding the care and handling of laboratory animals. NOTE: All protocols using live animals must first be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) or must conform to governmental regulations regarding the care and use of laboratory animals. Materials Female New Zealand white rabbits (3.5 to 4.5 kg) APAP-PABA-KLH conjugate (see Support Protocol 1) Adjuvant (Adjuvax from Alpha-Beta Technology, Hunter’s Titermax from CytRx, or RAS from RIBI Immunochemical Research) 20- and 22-G needles 15- and 30-ml Corex glass centrifuge tubes Refrigerated centrifuge 0.5-ml glass syringes Additional reagents and equipment for detection of anti-APAP antibodies by ELISA (see Support Protocol 4)



Collect preimmune serum 1. One to two weeks prior to immunization, collect 5 to 10 ml of blood from unimmunized rabbit by inserting a 20-G needle into the central vein of the ear and gently holding it in place while allowing blood to drain from the open end of the needle directly into a 15-ml Corex tube. This will serve as the source of preimmune serum, which will be used to determine the specificity of antibody production and to detect any innate immune cross-reactivity with proteins that lack the antigenic adduct. Blood collection is facilitated by wrapping the rabbit in a towel, exposing only the head and ears. Heat generated from a lamp is recommended for keeping the ear warm to ensure adequate blood circulation. Shave a localized area of the ear. Palpation, rather than the use of chemical irritants (i.e., xylene), should be used to dilate the blood vessel. Donovan and Brown (1995) describe the procedure for collection of blood from the auricular vein of the rabbit.

2. Allow blood to clot at room temperature. The clot will retract from the wall of the tube. This can be facilitated by rolling a glass rod or similar device around the inside of the tube.

3. Centrifuge 10 min at 10,000 × g, 4°C. Carefully remove the serum, retain a 1-ml aliquot for initial screening, and store the remainder in 1-ml aliquots at −70°C. Immunize rabbits 4. In a 1.5-ml microcentrifuge tube, mix 1 part APAP-PABA-KLH (containing 400 µg of the conjugate) with 1 part adjuvant. Antigen concentrations can range from 50 to 1000 ìg/ml for rabbits.

5. Vortex 5 to 10 min to emulsify, then inject the immunogen preparation into the rabbit at multiple sites using a 0.5-ml glass syringe and 22-G needle. Plastic syringes are not recommended as they may swell on contact with the adjuvant. Avoid trapping air in the syringe or needle. The volume for injection varies with the site. A typical immunization protocol may include intramuscular injections of 0.3 to 0.5 ml into each hind leg along with two subcutaneous injections (0.1 to 0.25 ml each) at sites along the lower back.

6. Give booster injections 3 to 4 weeks after the initial immunization. Use the same formulation procedure as in step 4, but add only 200 µg antigen. Collect and test antiserum 7. At a point 10 to 14 days after the booster injection, collect 5 ml of blood and process into serum as in steps 1 to 3. Test serum for the presence of anti-APAP antibodies by ELISA (see Support Protocol 4). If ELISA testing of this initial blood sample indicates the presence of specific antibodies, 20 to 25 ml should be drawn during subsequent blood collections. If no antibodies are detected, the rabbit may be given a second booster and the serum retested. If antibodies are not detected at this time, immunization of that particular rabbit should be discontinued. Peak levels of antibody are usually achieved 10 to 14 days following the booster injection and persist for 2 to 4 weeks. Subsequent booster injections may be given as needed, but should not be more frequent than every 4 to 6 weeks. A recommended frequency for blood collection is at 4-month intervals, at which ∼30 to 40 ml of blood is withdrawn. Detection of Covalent Binding


PREPARATION OF NAPQI-PROTEIN ADDUCTS AS COATING ANTIGEN FOR ELISA

SUPPORT PROTOCOL 3

N-acetyl-p-benzoquinoneimine (NAPQI)-protein adducts are used to coat the wells of the plate for the ELISA that will be used to test for antibody specificity (see Support Protocol 4). Antibodies elicited in response to conjugated haptens may be directed against the hapten, the carrier protein, and the linker molecule. To determine whether the antibodies produced following hapten-conjugate immunization can specifically recognize APAP, it is important that ELISA be performed (see Support Protocol 4) using a coating antigen that contains neither the linker, PABA, nor the carrier protein, KLH. Both BSA and aldolase which have been reacted with the APAP electrophile, NAPQI, have been used successfully as coating antigens. The unmodified protein serves as a negative control in the ELISA. This protocol describes the method of Dahlin and Nelson (1982) and Streeter et al. (1984). APAP is converted to NAPQI and allowed to bind to BSA. Materials Acetonitrile Sodium sulfate, anhydrous Acetaminophen (APAP; Sigma; minimum purity 99%) [3H]acetaminophen ([3H]APAP; 15.5 Ci/mmol; must be custom-synthesized) Silver (I) oxide Bovine serum albumin (BSA) or aldolase Phosphate-buffered saline (PBS), pH 7.4 (see recipe) 10 mg/ml BSA in PBS (see recipe for PBS) 1 N perchloric acid 80% methanol, ice-cold 50 mM carbonate/bicarbonate buffer, pH 9.6 (see recipe) 5- to 7-ml glass screw-cap vials 6000 to 8000 MWCO dialysis tubing (pretreated; see recipe) and dialysis clamps Synthesize NAPQI 1. Add the following ingredients sequentially to 625 µl acetonitrile in a 5- to 7-ml glass vial while stirring with a micro stir-bar: 5 mg (35 µmol) anhydrous sodium sulfate 5 mg (33 µmol) APAP 13.2 µCi [3H]APAP 10 mg (43 µmol) silver (I) oxide. Anhydrous sodium sulfate is used to dry organic solvents. Silver oxide is a catalyst in the reaction and is not soluble. Sufficient radioactivity is included to facilitate quantitation of the amount of NAPQI bound to protein using an aliquot of the protein reaction as described below.

2. Stir 15 to 30 min at room temperature. The acetonitrile solution will change from colorless to pale yellow.

3. Transfer the reaction to a 1.5-ml microcentrifuge tube. Microcentrifuge 1 min at 14,000 × g, room temperature, to bring down the silver oxide precipitate. 4. Transfer the supernatant to a clean tube. Assessment of Cell Toxicity


React with BSA or aldolase 5. Solubilize 10 mg BSA or aldolase in 10 ml PBS, pH 7.4 BSA is available as a crystalline powder; aldolase is supplied as a suspension in ammonium sulfate solution and must be dialyzed against PBS to remove the salt.

6. Divide the BSA or aldolase solution into 2-ml aliquots in 7-ml glass screw cap vials. Add the appropriate volume of NAPQI solution (from step 4) to give molar ratios of NAPQI/protein of 25:1, 50:1, and 100:1 in the reaction mixture, assuming 100% conversion of APAP to NAPQI. Prepare additional vials as controls containing the unmodified protein and the unmodified protein plus the maximum volume of acetonitrile used in generating the protein adducts. The reaction of NAPQI with purified proteins in vitro must be carefully monitored to avoid artifactually high binding ratios of NAPQI to protein. Cysteine residues are the primary sites on proteins that react with NAPQI. BSA (66 kDa) has one free -SH residue; therefore the optimal binding of NAPQI to BSA should approximate a 1:1 molar ratio. A ratio in excess may indicate nonspecific binding or polymerization of NAPQI. Aldolase, a tetrameric protein of 160 kDa, may offer greater flexibility as a coating antigen, with 12 and 24 -SH groups available under nondenaturing and denaturing conditions, respectively.

7. React for 30 min at room temperature with shaking. Longer reaction times may result in decomposition of NAPQI.

8. Dialyze overnight against 2.0 liters of PBS to remove unincorporated isotope ([3H]APAP), using pretreated 6000 to 8000 MWCO dialysis tubing. Following dialysis, remove two 100-µl aliquots and proceed to step 9. Store remaining volume of adduct solution at 4°C. It is recommended that the protein adducts, especially aldolase adducts, not be stored for >1 week. Protein denaturation may contribute to decreased sensitivity and greater variability in ELISA.

Determine amount of [3H]APAP bound to the protein 9. Precipitate each of the 100-µl aliquots of [3H]APAP-protein adducts (from step 8) with 1.0 ml of 1 N perchloric acid, then add 50 µl of 50 mg/ml BSA to each aliquot as a cold carrier. The carrier is added to prevent losses of the small amount of radioactive protein.

10. Let stand on ice for 10 min, then centrifuge 5 min at 14,000 × g, 4°C. 11. Remove supernatants and discard. Resuspend pellets by vortexing in 1.0 ml ice-cold 80% methanol. Add 30 µl of 1.0 N NaOH to the resuspended pellet and confirm that the pH is alkaline with pH paper. Repeat the methanol washes twice more using the technique described in Support Protocol 1, step 18. The step for ensuring basic pH is necessary since BSA is soluble in acidic methanol.

12. Determine the amount of bound [3H]APAP (see Support Protocol 1, steps 19 to 21). 13. Calculate the nanomoles of NAPQI bound to the protein by dividing the 3H dpm obtained in step 12 by the specific activity of APAP used in step 1—i.e., 0.4 µCi/µmol. 14. Calculate nanomol of NAPQI bound per nanomol protein in the aliquot sampled. 100 ìl of 1 mg/ml BSA is equivalent to 1.5 nmol BSA.



Dilute NAPQI adducts for ELISA 15. Dilute the protein adducts and corresponding controls in 50 mM carbonate/bicarbonate buffer, pH 9.6, to give 2 µg/ml. Use 100 ng (50 µl) for coating quadruplicate wells of a microtiter plate (see Support Protocol 4). To coat each well of the plate, 100 ng of protein adduct in 50 ìl of 50 mM carbonate/bicarbonate buffer, pH 9.6, is used. From step 5, the starting concentration of protein (BSA or aldolase) is 1 mg/ml. Therefore, to obtain the desired concentration for coating, dilute the 1 mg/ml protein-adduct solution 1:10 by adding 100 ìl of the protein adduct to 900 ìl buffer (the volume of NAPQI solution added to the reaction with protein is 100 kDa), resulting in protein precipitation within the gel. Addition of a low concentration of SDS (0.01% to 0.1%) to the transfer buffer can improve the efficiency of transfer, but this may vary with the properties of the membrane. PVDF will tolerate higher concentrations of SDS, whereas the detergent can reduce protein binding to nitrocellulose. Successful transfer of proteins is also dependent on gel thickness and the percentage acrylamide and cross-linker used. Lower percentages result in more rapid and efficient transfer. For most SDS-PAGE applications, a 10% resolving gel containing 2.7% to 3.4% cross-linker with 0.5 to 0.75 mm thickness will be adequate. Membranes can bind protein noncovalently (nitrocellulose, nylon, PVDF) or covalently (activated papers). Nitrocellulose and PVDF are the most frequently used. PVDF is more hydrophobic than nitrocellulose and offers several advantages including its ease of handling, tolerance to SDS, lower susceptibility to protein loss by desorption during the subsequent incubation steps for immunodetection, and lower background staining with 125I. However, the protein-binding capacity of PVDF is less than that of nitrocellulose (100 µg/cm2) and more stringent blocking conditions are required prior to probing with antibody. The efficiency of transfer can be determined by staining the membrane for total protein prior to immunodetection. Ponceau S is both rapid and reversible but not very sensitive (2 µg detection limit; Harlow and Lane, 1988). India ink staining is sensitive (80 ng), permanent, and generally does not interfere with the antigenantibody reaction (Glenney, 1986). India ink cannot be used with enzyme-based detection methods because it does not provide sufficient background contrast to the colored reaction products. It is important to keep in mind that each antigen-antibody interaction is unique, and that much success associated with immunoassays is derived from the performance of preliminary experiments aimed at optimizing conditions. The problems that may be encountered in immunoblotting can be quite extensive, and a comprehensive discussion of them is not pos-

sible here. Some of these have been addressed as they arise at specific steps in the protocol. The detection sensitivity of immunoblotting is determined by the antibody specificity. Thus, two types of problems generally occur: (1) little or no binding of the antibody to the antigen, and (2) high background (nonspecific) staining. Poor antibody-antigen interactions may be due to low titer of antibody, and this may be evidenced by nonspecific staining of multiple protein bands. Prescreening of immune serum or affinity-purified antibody against an antibody standard by ELISA or immunoblot test strips, respectively, should permit selection of the best available batches of antibody or serum to eliminate this problem. In some cases, binding to specific target antigens will be diminished if the antibody is adsorbed by one or more components of the blocking solution. Comparison of different blocking solutions will identify the problem. Alternatively, the antibody may not recognize the antigen in its denatured form following SDS-PAGE. This can be verified with dot blots using native and denatured cell extracts and can sometimes be overcome by increasing the blocking time to permit renaturation of the proteins. Transfer conditions also may contribute to poor antigen-antibody reactions. Failure to maintain good contact between the membrane and gel during electroblotting, or precipitation of proteins in the gel as a result of high methanol concentrations, result in poor transfer and thus lowers the concentration of antigen present on the membrane. In addition, extended blotting times may result in protein loss as a result of migration completely through the membrane. Staining the gel and/or membrane with a protein stain following transfer will determine if these are contributing factors. Two types of background staining problems are often encountered: (1) a general diffuse staining, or (2) nondiffuse, apparently specific staining of nonantigen proteins. Diffuse staining may result from insufficient blocking or nonspecific interaction of the primary or secondary antibodies with the blocking buffer. This is easily remedied by increasing blocking times or changing the blocking buffer. In some cases greater dilution of the primary or secondary antibodies may also alleviate the problem. Troubleshooting experiments in which the incubation with the primary antibody is omitted or in which different dilutions of the secondary antibody are tested in the presence of a fixed concentration of primary antibody will determine if the secondary antibody alone is contributing to the background-staining problem. Al-



ternatively, the primary antibody may be contributing to the nonspecific background. This is often observed with whole serum of low titer where the relative titers of specific and nonimmune IgG determine the signal-to-noise ratio. In such instances, affinity purification of the antibody may significantly reduce the nonspecific staining. Nondiffuse background appears as distinctly stained protein bands that do not contain the specific antigen of interest. This is most likely attributable to antibody cross-reactivity with the nonantigen. Thus, it is important to include proteins from untreated tissue or cells (negative control) along with tissues or cells known to contain covalently bound xenobiotic adducts (positive control). Any immunostaining of proteins in the negative control samples must be considered as background. Affinity purification of the antibody may also reduce this type of background binding.


Immunohistochemistry A number of factors contribute to the success of immunohistochemical techniques, but common and most critical to all is the preservation of tissue ultrastructure without destruction of antigenic determinants. This, in turn, is dependent on the choice of methods for tissue preparation, fixation, embedding and dehydration. During tissue preparation, tissues undergo rapid decomposition once removed from the animal. To minimize tissue/cell damage and decrease metabolic activity, thereby preserving normal ultrastructure, fixation of samples should occur as rapidly as possible, usually within 15 min following excision. The volume of the tissue sample is also critically important to ensure rapid fixation. Tissue blocks between 1 and 10 mm in at least one dimension are recommended to facilitate penetration of the fixative. Optimal fixation should (1) minimize changes to cellular morphology, (2) result in minimal denaturation of the antigen, (3) retain the antigen in its normal pattern of cellular distribution and localization, and (4) minimize diffusion of endogenous soluble components but allow penetration of the antibody for reaction with antigen. Different tissues may fix at different rates depending on their physical characteristics. Fixation methods that work well for one tissue may not be directly transferable to another. Overfixation may cause the masking or destruction of antigens whereas underfixation may result in incomplete immobilization.

Tissues can be fixed by chemical or physical agents. Organic-solvent or precipitant fixatives, such as 100% methanol or acetone, typically require immersion of the specimen in the solvent for 15 min at −20°C and subsequent washing in buffer to return the pH to neutrality. Such fixatives, although relatively mild, are more susceptible to diffusion artifacts. Crosslinking fixatives (e.g., formaldehyde and glutaraldehyde) are excellent for preserving ultrastructure but frequently destroy antigenicity, although this may be dependent on the concentration and the antigen itself. Microwave fixation has become increasingly popular for its ability to retain both ultrastructure and antigenicity (Moran et al., 1988; Login and Dvorak, 1988, 1994). It has also been used to unmask antigens that previously had no or low immunoreactivity following conventional aldehyde fixation (Login and Dvorak, 1994). Microwave irradiation of tissues at 40° to 55°C for 26 presacral vertebrae No. caudal ossified vertebrae

4.0 ± 2.7 4.3 ± 0.5

1.0 ± 1.0 4.4 ± 0.5

9.1 ± 2.4 3.0 ± 0.4**

74.1 ± 9.3*** 2.0 ± 0.4***

Fused ribs Lumbar ribs

0 12.9 ± 5.6

0 26.7 ± 6.9

0 36.0 ± 6.2*

13.3 ± 7.7 81.3 ± 8.3***

Cervical ribs Wavy/callused ribs

0.7 ± 0.70 1.8 ± 1.2

2.1 ± 1.5 2.6 ± 1.7

0.7 ± 3.6*** 7.0 ± 3.9

6.7 ± 2.6

Fused/duplicated/misarticulated ribs

0.9 ± 0.9

1.3 ± 1.3

2.7 ± 1.6

4.1 ± 2.3

Sternebrae Metacarpals (front limbs)

5.3 ± 0.3 7.5 ± 0.2

5.5 ± 0.4 7.5 ± 0.2

4.7 ± 0.4 6.8 ± 0.3*

3.9 ± 0.5*** 6.4 ± 0.2***

Front proximal phalanges Front distal phalanges

3.0 ± 0.7 9.8 ± 0.1

3.1 ± 0.9 9.8 ± 0.2

1.1 ± 0.5** 9.5 ± 0.2

0.5 ± 0.3*** 8.6 ± 0.8***

Metatarsals (hind limbs) Hind proximal phalanges

8.3 ± 0.2 2.3 ± 0.7

8.6 ± 0.3 3.2 ± 1.0

7.9 ± 0.1 0.5 ± 0.3***

7.5 ± 0.4*** 0.1 ± 0.1***

Hind distal phalanges

8.4 ± 0.9

9.2 ± 0.8

7.3 ± 1.0

7.9 ± 1.0

Percent incidence

Percent incidence

Mean no. ossified

aData from study with valproic acid (Narotsky et al., 1994). Reproduced with permission of the Society of Toxicology. Symbols: one

asterisk (*), significantly different from control (p < 0.05); two asterisks (**), significantly different from control (p < 0.01); three asterisks (***), significantly different from control (p < 0.01). bSUR, small, unossified, or reduced ossification.

Teratology


Supplement 16

Table 13.5.5

Example of Data Summary for Segment II Visceral Findings in Ratsa

Dose (mg/kg/day)

0

100

200

400

No. examined Litters Fetuses

19 124

13 87

19 119

14 87

No. fetuses affected Ventricular septal defect Microphthalmia Micrognathia, cleft palate

1 1 0

0 0 1

0 0 0

0 0 0

Mean score Kidney Ureter Lateral ventricles

2.8 ± 0.1 2.3 ± 0.1 1.0 ± 0.0

2.8 ± 0.1 2.7 ± 0.2 1.1 ± 0.0

3.2 ± 0.1* 2.7 ± 0.2 1.1 ± 0.0

3.6 ± 0.2*** 2.9 ± 0.2 1.1 ± 0.0

Percent incidence Dilated renal pelvis Dilated ureter

5.9 ± 2.9 21.6 ± 4.1

8.1 ± 4.3 31.5 ± 8.0

19.6 ± 4.9* 34.9 ± 5.9

35.9 ± 6.2** 35.4 ± 7.4

aData from study with valproic acid (Narotsky et al, 1995). Reproduced with permission of the Society of

Toxicology. Symbols: one asterisk (*) significantly different from control (p < 0.05); two asterisks (**), significantly different from control (p < 0.01); three asterisks (***), significantly different from control (p < 0.001).

In Vivo Assessment of Prenatal Developmental Toxicity

allowing either microdissection or sectioning of the preserved specimens. Although some regulatory guidelines suggest categorizing fetal examination findings as malformations or variations, or categorizing based on severity (major malformations, minor malformations) it should be noted that there is no generally accepted classification of malformations and variations. Furthermore, there is no scientific justification for using such classification. Instead (or in addition), the authors recommend categorizing findings based on syndrome, organ system, and etiology. When conducting developmental toxicity studies, it is essential to have high pregnancy rates in the test animals. Pregnant animals can be obtained by breeding them in the test facility, or by purchasing timed-pregnant animals from an animal supplier. The latter approach allows some scheduling flexibility in that a larger number of test animals can be bred on the same day. However, disadvantages of buying timed-pregnant animals include the fact that the animals are subject to the stress of shipping during the preimplantation period of pregnancy (possibly contributing to low pregnancy rates) and that a quarantine period may be impossible, in addition to the increased cost.

Anticipated Results Uterine findings Table 13.5.3 presents an example of uterinefindings data from a Segment II study (Narotsky et al., 1994). In this example, no significant differences between groups were noted for the numbers of corpora lutea, implantation sites, or live fetuses. More importantly, the values for pre- and post-implantation loss were also comparable between groups. The values for fetal weight, however, indicated significant dose-related reductions at dosages of 200 and 400 mg/kg. Therefore, based on the fetal weight data, the test agent in this example was developmentally toxic at 200 and 400 mg/kg. On the basis of this study alone, it is unclear whether the isolated case of full-litter resorption at 400 mg/kg was treatment-related. Skeletal findings An example of findings from fetal skeletal examinations in a Segment II study (Narotsky et al., 1994) is presented in Table 13.5.4. Findings consistent with delayed development at dosages of 200 and 400 mg/kg include reductions in ossification (e.g., of the calvaria, hyoid, supraoccipital, sternebrae, metacarpals, phalanges, and caudal vertebrae) and an increased fontanel score. Findings indicating a toxic, and




9 12 10 10 7c

0 5.1 6.8 9.0 12.0

GD 6-20 (adjusted)b 16.5 ± 1.2 16.4 ± 1.7 15.6 ± 1.8 7.3 ± 2.2* –2.3± 6.5*

GD 6-8 0.3 ± 0.8 0.8 ± 0.7 0.2 ± 0.3 –1.4± 0.5 –2.3±0.9* 11 0 0 0 0

Dams with full-litter resorption (%) 14.7 ± 10.8 4.1 ± 1.8 10.5 ± 3.7 5.0 ± 2.9 3.7 ± 2.3

Prenatal 1.4 ± 1.4 8.3 ± 8.3 23.8 ± 10.0 63.3 ± 13.8*** 59.7 ± 18.6***

Postnatal

Percent loss

5.4 ± 0.1 5.2 ± 0.1 5.1 ± 0.1* 5.4 ± 0.1 5.1 ± 0.3*

10.0 ± 0.3 9.5 ± 0.1 8.3 ±0.4*** 9.0 ± 0.3 8.7 ±0.6***

PD 6

Pup weight (g) PD 1

Mean ± SE per litter

0 0 0.9 ± 0.9 0 2.1 ± 2.1

Pups with eye defects (%)

cIncludes one dam that died at term and two that died during lactation.

bAdjusted for weight of live litter on PD 1.

aData from study with heptachlor (Narotsky et al., 1995). Symbols: one asterisk (*), significantly different from control (p < 0.05); three asterisks (***), significantly different from control (p < 0.001).

No. of dams

Mean ± SE weight gain (g)

Example of Data Summary for a Chernoff/Kavlock Assaya

Daily dose (mg/kg)

Table 13.5.6

Teratology

13.5.21

Supplement 16

dose-related, effect on the development of the axial skeleton include lumbar ribs at dosages of 200 and 400 mg/kg, extra cervical ribs, extra (>26) presacral vertebrae, and rib and vertebral malformations at dosages of 400 mg/kg. Although the incidences of rib and vertebral malformations (fusion, duplication, misarticulation) did not reach statistical significance, they were attributed to treatment. Visceral findings Table 13.5.5 gives an example of fetal visceral examination findings in a Segment II study (Narotsky et al., 1994). Significant, doserelated increased incidences of dilated renal pelvis (also reflected by kidney scores) indicated a toxic effect at 200 and 400 mg/kg. Sporadic, non-dose-related malformations (ventricular septal defect, microphthalmia, micrognathia, cleft palate) were not attributed to treatment. Chernoff/Kavlock assay An example of a data summary from a Chernoff/Kavlock assay (Narotsky et al., 1995) is presented in Table 13.5.6. In this example, dose-related maternal toxicity at 9 and 12 mg/kg was evidenced by adjusted gestational weight gains (weight gain from GD 6 to 20, minus the live PD 1 litter weight). Maternal toxicity is also evident at 12 mg/kg, as indicated by significant weight losses early in the treatment period (GD 6 to 8) and by maternal deaths at term and during lactation (see footnote to Table 13.5.6). Developmental toxicity was evidenced by significantly increased postnatal loss at both 9 and 12 mg/kg dosages. Although pup weights were unaffected at dosages of 9 mg/kg, reduced pup weights at 6.8 and 12 mg/kg were attributed to treatment. Therefore, in this example, maternal toxicity was evident at 9 and 12 mg/kg; whereas developmental toxicity was evident at 6.8, 9, and 12 mg/kg. Pups with eye defects (anophthalmia or microphthalmia) occurred in one litter in each of the 6.8- and 12-mg/kg groups; based on this study alone, it was unclear whether this finding was treatment related.

Time Consideration

In Vivo Assessment of Prenatal Developmental Toxicity

Typically, the in-life phase of a Segment II stu dy (see Basic Protocol 1) or a Chernoff/Kavlock assay (see Basic Protocol 2) is ∼3 to 5 weeks for mice and ∼4 to 6 weeks for rats, depending on the time required to obtain sufficient mated animals for the study. The number of mated animals obtained on a given

day should not exceed the number of animals that the available workforce can accommodate in one day, particularly for labor-intensive tasks such as cesarean sections. The time required to complete the cesarean sections (see Support Protocol 1) in a Segment II study may range from 1 day to 2 weeks, depending on the time required to obtain sufficient mated animals for the study. Examinations of apparently nongravid uteri (see Support Protocol 2) are done in conjunction with cesarean sections as well as Chernoff/Kavlock assays and generally require 3 weeks, and spermatid development also requiring ∼3 weeks. The arrangement of the different

Male Reproductive Toxicology


populations of germ cells within the seminiferous tubule is also similar between species, with Sertoli cells providing a supporting structural framework for discrete layers of the different germ cell types. Spermatogonia are always at the base of the tubule and progressively more mature germ cells are found in layers moving toward the lumen. In all species, 3 or 4 generations of germ cells are developing within a tubule at any given time. The development of each of these generations occurs in synchrony with each other, giving rise to specific and predictable cellular associations (Fig. 16.4.1). The complete sequence of cellular associations is referred to as the cycle of spermatogenesis while the individual cell associations form the stages of the cycle. Each stage is therefore defined by its germ cell complement and consequently, identifying the stage defines what cells should be in that tubule (and what cells are missing). In order for a pathologist to detect subtle changes in germ cell loss or, in the case of spermatid retention, the inappropriate presence of a population of germ cells, a thorough understanding of the cellular makeup of the spermatogenic cycle is essential.

Species-Specific Variations in the Organization of Spermatogenesis Although the fundamentals of the spermatogenic cycle are similar between species, there are certain details that vary. These can have a significant impact on histopathologic evaluation. Number of stages The number of stages and their cellular makeup varies between species and depends on the morphological criteria used by the classification system. It is important that the pathologist is familiar with the germ cell development and the stage map of the species under investigation. A highly recommended text that explains in detail how to stage tubules in a number of common species and provides stage maps for each species is provided by Russell et al. (1990).

Histopathology of the Male Reproductive System II: Interpretation

Duration The duration of spermatogenesis (the time taken for a spermatogonium to develop into sperm) and the duration of the spermatogenic cycle (the time taken to complete a cycle of cell associations) varies between species. This information is important so that the pathologist can predict how long it should take for any particular cell to reach a later cell type (e.g., if

a toxicant affects leptotene spermatocytes, how long will it take before the animal becomes infertile?). Alternatively, if a particular cell type is missing at some defined period after dosing, knowing the dynamics of the spermatogenic cycle will allow extrapolation as to what stage of development that cell was in when dosing began. Software programs have also been developed to calculate this type of information (Hess and Chen, 1992). Cell associations The organization of cell associations along the length of the tubule is linear for most mammalian species, including most species of monkey used in toxicological studies. This means that a tubular cross section normally contains only one cell association and that the adjoining length of tubule (which is often the adjacent tubule in a cross section of testis) will generally contain the consecutive stage. This is not the case in humans where cell associations are arranged in a helical pattern resulting in a mosaic of cell associations in a single cross section. In dogs, although only one stage is present in a cross section, adjoining lengths of tubule do not necessarily contain consecutive stages.

Disturbances in Spermatogenesis Almost regardless of the cellular target of toxicity within the reproductive system, the most common morphological consequence of injury is a disturbance in spermatogenesis. This is because spermatogenesis is dependent on, or sensitive to, functional perturbations in most other parts of the reproductive tract. Spermatogenic disruption may reflect a direct effect on the seminiferous epithelium, affecting either the Sertoli cell or any one of the germ cell populations, or it may occur as a secondary response to altered hormone levels, altered vascular supply or altered fluid balance, either within the testis or within the epididymis. It is therefore extremely important that disturbances in spermatogenesis are detected. The pattern of disturbance can be very specific and diagnostic of the mechanism of toxicity, but generally, this is only seen during the early development of the lesion. With longer periods of dosing, the development of maturation depletion (whereby death of a specific precursor germ cell causes the progressive loss of its descendant generations), reduces the specificity of the pattern of spermatogenic disturbance as the tubules become depleted of more and more germ cells.



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XIV

I

Figure 16.4.1 Stages of the spermatogenic cycle of the rat. Cell associations for 4 of the 14 stages of the spermatogenic cycle of the rat (stages I, VIII, XI, XIV). Spermatogonia: A, type A; In, intermediate; Spermatocytes: PL, preleptotene; L, leptotene; Z, zygotene; EP, early pachytene; MP, mid pachytene; LP, late pachytene; D, dividing. Spermatids: 1, 8, 11, 14, 15, and 19 indicates steps 1 to 19 of spermatid development. The tubular cross sections (stages I, VIII, XI, and XIV) show the arrangement of cells within the seminiferous epithelium. The columns of cells at the base of the figure show the maturation of the cells during one spermatogenic cycle. Each generation of cells develops sequentially. During stage VIII, the mature step 19 spermatids are shed into the lumen (arrows) while a new generation develops from stem cell spermatogonia. As spermatocytes undergo meiotic division (D) in stage XIV, they produce step 1 spermatids and the cell association returns to stage I to begin another cycle. (Reproduced from Creasy, 1997, with permission.)



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Table 16.4.1 Specific EPA and OECD Recommendations for Histopathological Examination of the Testes and Epididymides in Studies to Detect Effects on Reproduction and Fertility

Testis

Epididymis

Detailed histopathological examination of testes should be conducted in order to identify treatment-related effects such as: Retained spermatids Missing germ cell layers or types Multinucleate giant cells Sloughing of spermatogenic cells into the lumen

Examination of the intact epididymis should include the caput, corpus, and cauda. The epididymis should be evaluated for: Leukocyte infiltration Sperm granulomas Change in prevalence of cell types Absence of clear cells in the cauda epithelium Aberrent cell types in the lumen Phagocytosis of sperm


Regulatory Guidelines and the Role of “Staging” in Histopathologic Examination of the Testis

COMMON TOXICOLOGICALLY INDUCED FINDINGS AND THEIR POSSIBLE SIGNIFICANCE

Recently revised regulatory guidelines have placed increased emphasis on the importance of histopathology for detecting toxicological effects in the male reproductive system. Recommendations have been made, not only for fixation and staining procedures but also for the microscopic examination of tissues, providing examples of findings that should be recorded (Table 16.4.1). During the drafting of these guidelines, there was much discussion relating to the subject of “staging” of testes, and although there is no mention of staging in the final issued guidelines, the issue has become surrounded by confusion. The ability to recognize stages of the spermatogenic cycle is important in order for the pathologist to recognize when cells are missing or are inappropriately present. Due to the lack of understanding of this concept, there has been a move to expect the pathologist to produce a quantitative assessment of stages—e.g., a frequency distribution of tubules for individual stages of the spermatogenic cycle. While this may be useful information in an investigative study to determine whether the dynamics of the spermatogenic cycle have been disturbed (Hess, 1990), it is inappropriate to carry out in a regulatory study, which is designed as a screening study to detect effects on spermatogenesis. Knowledge of staging should be used in a qualitative way to evaluate the normality of the cellular makeup of the seminiferous tubules. In other words, the testis should be examined with an understanding of the normal progression of the stages of the spermatogenic cycle. This approach is explained below. For a more detailed discussion of this issue see Creasy (1997) and Chapin and Conner (1999).

As with any tissue, the cellular response to injury is limited and at times, nonspecific. However, certain aspects of the early pathogenesis of toxicologically induced lesions in the testis and accessory tissues can provide important information on the mechanism of injury. Additional information can be found in Nolte et al. (1995), Creasy (2001), and Creasy and Foster (2001).

Testes Germ cell degeneration/multinucleate aggregates Whether spontaneous or induced, death of germ cells appears to occur predominantly through apoptosis, a process that is closely regulated by the Sertoli cell (Lee et al., 1997, 1999). This is particularly true for spermatogonia, which may be seen apoptosing in occasional stage XII tubules. However, many of the dying cells do not have the classic morphological appearance of apoptotic cells. Dying spermatocytes generally develop cytoplasmic eosinophilia and nuclear pyknosis while round spermatids show chromatin margination. If cell death progresses rapidly, then the apoptotic cell is rapidly phagocytized by the surrounding Sertoli cell cytoplasm and all evidence of cell death is rapidly removed. Cell death and phagocytosis of the remains can be complete within 24 hr, so if the process is not examined during this time span, the only evidence of cell death will be an absence of the cell (cell depletion). If the degenerative process is slow, then adjacent germ cells belonging to the same cohort, may form a multinucleate syncitium (symplast, multinucleate giant cells) probably due to the



ES ES

RS PS

RS

Sg

Sg

day of dosing

1 week

ES Sg

Sg 2 weeks

4 weeks

Figure 16.4.2 Development of maturation depletion following daily dosing with a spermatocyte toxicant. Time-dependent progression of maturation depletion following cell-specific damage to pachytene spermatocytes (PS). If the tubule is examined on the day of dosing, spermatocyte degeneration and necrosis will be seen (top left). Phagocytosis of the necrotic cells by Sertoli cells results in their rapid disappearance and because dosing continues, newly formed pachytene spermatocytes will also be killed. Examination of the same stage tubule after 1 week of dosing (top right) will reveal an absence of pachytene spermatocytes. After 2 weeks of dosing (one spermatogenic cycle duration) pachytene spermatocytes will still be missing but round spermatids will also be absent because their precursor cells were destroyed in the previous cycle (bottom left). Similarly, after 4 weeks of dosing (bottom right), pachytene spermatocytes, round spermatids, and elongated spermatids will be absent, leaving only spermatogonia. This progressive loss of subsequent germ cells following injury to a previous cell type is termed maturation depletion. ES, elongated spermatids; RS, round spermatids; PS, pachytene spermatocytes; Sg, spermatogonia. (Reproduced from Creasy, 1997, with permission.)

breakdown of the cytoskeletal fibers supporting the interconnecting cytoplasmic bridges. Multinucleate aggregates are less readily phagocytized by Sertoli cells and are present for longer periods and therefore more frequently seen. They are most often composed of round spermatids, but can also be formed by fusion of neighboring spermatocytes or elongating spermatids. Germ cell depletion This is the most common sequel to spermatogenic disturbance and is generally a consequence of germ cell death rather than exfoliation. It may be seen as a generalized or partial depletion of the germ cells or it may only affect a specific cell type (e.g., spermatogonia). Sometimes a specific cell type within specific stages may be affected (e.g., pachytene spermatocytes in stages XII and XIII). Once the cell has been phagocytized, the only way of recog-

nizing the lesion is by the abnormal cellular association of individual stages of the spermatogenic cycle and the progressive development of maturation depletion with time (Fig. 16.4.2). The appearance of the testis, in terms of what cells are missing, will depend largely on how severe the initial effect was and how long after dosing the testis is examined. Instead of a specific cell type being killed, a focal cohort of cells within a tubule may be affected and result in a focal “blow out” of the epithelium. This may be due to an effect on a few adjacent spermatogonia, which then fail to produce their cohort of spermatocytes and spermatids, or on one or two Sertoli cells, which are then unable to support spermatogenesis. Partial or generalized germ cell depletion may affect only a small number of tubular profiles or a large proportion of the tubules. When only a few scattered tubules are affected, it is not possible to determine whether they



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represent multiple convolutions of the same tubule or focal segments of multiple affected tubules. Prolonged dosing with a number of testicular toxicants may cause generalized germ cell depletion, affecting a large proportion of the tubules. It generally represents an advanced or end-stage lesion, and in order to elucidate the primary target cell, a time course study needs to be carried out. It is not possible to say whether spermatogenic depletion is or is not reversible without carrying out an appropriate study. If spermatogonia are still present, then the lesion is potentially reversible but if the Sertoli cells are functionally compromised, spermatogenesis may not be supportable. The chronic effects of 2,5hexanedione on the rat testis exemplify this. Although spermatogonia remain and are seen to divide, spermatogenesis does not recover. This is thought to be due to the inhibition of a critical Sertoli cell factor (Blanchard et al., 1998). Conversely, spermatogonia may be significantly depleted, but if the Sertoli cells are functionally intact and sufficient time is allowed for stem cell renewal and repopulation (and this may require several spermatogenic periods), substantial recovery may be seen (Meistrich, 1986). Germ cell exfoliation Loss of adhesion between Sertoli cell and germ cell, or shearing of Sertoli cell cytoplasm (as seen with cytoskeletal disrupting agents) will result in exfoliation of germ cells into the lumen of the seminiferous tubule and subsequent transport of the cells to the rete testis and the epididymis. The exfoliated cells may appear morphologically normal but are rapidly removed from the testis. Once the cells have been removed, cell depletion is the only recognizable finding. Lumenal germ cells may also be present as a result of handling trauma at necropsy. Care must be taken to distinguish between real and artifactual exfoliation (Foley, 2001). Abnormal residual bodies shed into the lumen can sometimes be mistaken for exfoliated germ cells. These generally occur as a result of disturbances in spermiation (see below).


Tubular vacuolation Vacuolation within or between Sertoli cells is a common early sign of Sertoli cell damage. The vacuoles may be solitary and situated amongst the germ cells at varying depths throughout the epithelium. It is generally not possible by light microscopy to determine

whether they are intra- or extra-cellular. In other cases, intracellular microvacuolation or swelling may be seen affecting the basal area of the Sertoli cell cytoplasm and causing germ cell displacement and disorganization. Such findings are suggestive of disturbances within the Sertoli cell and may represent alterations in the smooth endoplasmic reticulum or in fluid homeostasis. Vacuolation may also be seen in end-stage lesions, associated with extensive germ cell loss. In this situation, it should not be regarded as a primary effect on the Sertoli cell. Occasional solitary vacuoles are sometimes seen in tubules from normal testes but these are generally few in number. Vacuoles in the basal compartment of the tubule, surrounding spermatogonia are generally fixation-induced artifacts due to osmotic shrinkage. Tubular contraction Reduction in the overall diameter of the seminiferous tubule will occur as a result of germ cell depletion and/or as a result of reduced secretion of seminiferous tubule fluid. Seminiferous tubule fluid is secreted by the Sertoli cell and maintains the lumenal size, which varies with the stage of spermatogenesis. This is an androgen-dependent function of the Sertoli cell and will be affected by altered testosterone secretion. Another major regulatory factor for fluid secretion is the presence of elongating and elongated spermatids. Therefore, if these cells are depleted, fluid production and consequently lumenal size are decreased. Germ cell loss and decreased fluid will have a significant effect on testis weight. Tubular dilatation Dilatation of the tubular lumen will occur as a result of increased lumenal fluid volume. This can occur through increased secretion by the Sertoli cell or decreased expulsion of fluid from the tubule, which is thought to be a function of the contractile peritubular cells. Also decreased resorption of fluid by the epithelial cells of the rete and efferent ducts or obstruction of the outflow (e.g., a sperm granuloma) can cause increased tubular fluid. The increased fluid volume will generally be reflected by an increased weight of the testis unless there is an accompanying significant cell loss. The pathological consequences of the finding depend on the severity and duration of the effect. Prolonged increased pressure on the seminiferous epithelium will generally result in pressure atrophy of varying degrees and may also lead to inspissated sperm and granulomatous inflammation.



Spermatid retention This is a subtle but relatively common finding that may be caused by a number of chemicals as well as by hormonal disturbance. It is characterized by the retention of step 19 spermatids (which should be released during stage VIII) through stages VIII to XII. The position of the retained spermatids varies with different chemicals. In some cases, e.g., boric acid (Chapin and Ku, 1994), the retained spermatids remain in a predominantly lumenal position through stages VIII to XI and are then pulled down into the basal cytoplasm of stage XII tubules where they are phagocytized. With other chemicals the step 19 spermatids are rapidly pulled down into the basal cytoplasm and phagocytized during stages VIII to XI, leaving very few in a lumenal position. The formation and behavior of the residual bodies is often also disturbed with residual bodies of abnormal shape and size being seen in the tubular or epididymal lumen. Descent and phagocytosis of residual bodies normally occurs during stages IX to XI but in cases of spermatid retention this may be delayed into stage XII. The pathological significance of spermatid retention can be varied. It is often associated with abnormal sperm parameters (number, motility, or morphology) and it may be associated with alterations in fertility parameters. If homogenization resistant spermatids are measured, the retained spermatids should be reflected by an increase in this parameter. However, identification by histopathology is a much more sensitive endpoint since it can detect very small numbers of phagocytized spermatids. Tubular necrosis While germ-cell necrosis proceeds by apoptosis, tubular necrosis is characterized by coagulative (oncotic) necrosis of Sertoli and germ cells. Sertoli cells are normally highly resistant to cell death even though they may be very sensitive to functional perturbations. Consequently, they are often the only cell left lining severely damaged tubules (Sertoli cell–only tubules). Ischemia is one of the few situations where they are killed. The effects of this can be seen with cadmium toxicity, which damages the testicular capillary endothelium. It can also be seen following administration of vasoactive compounds such as serotonin. Necrosis and loss of the Sertoli cells from tubules is the major characteristic of the lesion, and this is associated with gross disorganization and necrosis of the germ cells as well as stasis of sperm in the tubular lumen. Due to the loss of the Sertoli cell

blood-tubule barrier, the changes are also accompanied by an inflammatory infiltrate, which is an otherwise rare accompaniment to toxic injury. Tubular necrosis is a serious irreversible lesion because Sertoli cells are unable to proliferate and the affected tubules are likely to involute and be replaced by scar tissue. Dilated rete Both ends of the seminiferous tubules empty into the rete. Most of the tubule fluid is reabsorbed in the epithelium of the rete and efferent ducts. If there is an obstruction in the efferent ducts or in the epididymis, the fluid back-pressure will cause the rete to dilate and if the obstruction is severe, the back pressure will progressively dilate the seminiferous tubules. The tubules in the area of the rete also appear to be a preferential location for some testicular toxicants, but this should not be confused with the transitional tubuli rectii that join the seminiferous tubules to the rete and can be mistaken for depleted tubules. Leydig cell atrophy/hypertrophy/ hyperplasia/adenoma Testosterone secretion is the major function of the Leydig cell and the abundance of smooth endoplasmic reticulum in the cell reflects this activity. Increased stimulation by luteinizing hormone results in functional hypertrophy and hyperplasia. With prolonged gonadotropin stimulation in the rat, Leydig cell hyperplasia will usually progress to adenoma formation. Many classes of compounds with diverse chemical structures have been shown to produce this effect in the rat but the significance to man is considered limited (Clegg et al., 1997). Decreased secretion of testosterone, whether through inhibition of biosynthesis or decreased gonadotropin stimulation, will lead to atrophic changes in the Leydig cell. Recognition of atrophy, hypertrophy, and hyperplasia on a qualitative basis is not easy unless the changes are marked. Contraction of tubules due to cell loss will result in an apparent increase in the volume of the interstitial space. This may or may not be contributed to by a real increase in size and number of Leydig cells, but quantitative analysis may be necessary to separate real from apparent effects.

Epididymis Lumenal germ cells/debris Cells and residual bodies exfoliated from the testis will be transported into the epididymis.



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This can serve as useful confirmatory evidence for changes seen in the testis. It can also alert the pathologist to changes that may have been overlooked in the testis. Occasional exfoliated germ cells are sometimes seen in normal animals, and in immature and peripubertal animals this number is significantly increased. Abnormal residual bodies may also be detected in the epididymis as a consequence of disturbed spermiation in the testis. The presence or absence of germ cells in the luminal contents can also aid the pathologist in evaluating whether apparently exfoliated germ cells in the lumina of seminiferous tubules are real effects or artifacts of trimming; such artifacts will not be present in epididymal lumena. Epithelial vacuolation Microvacuolation of the epididymal epithelium can be seen as a specific chemically induced finding. Macrovacuolation and cribriform change (infolding of the epithelium within itself) is often seen accompanying contraction of the atrophic aspermic epididymis. This may represent a normal mechanism of surface area reduction but has also been reported as a toxicologic change (Foley, 2001). Epithelial vacuoles are also sometimes seen as a normal finding in some species at the junction of the corpus and cauda. Since fluid absorption and secretion are both major functions of the epididymal epithelium, vacuolation is a likely sequel to disturbance of either function.


Epithelial inflammation and sperm granuloma The antigenically foreign sperm in the epididymal lumen and in the seminiferous tubule are in an immunologically protected environment. The protection is afforded by the lumenal tight junctions between epithelial cells in the excurrent ducts and by the basal occlusive junctions between Sertoli cells in the testis. If these barriers are damaged, then an inflammatory response against the sperm develops and generally progresses to form a sperm granuloma. This is a chronic, progressive lesion and in the coiled epididymal duct has the added complication of causing obstruction to the passage of sperm. Furthermore, the oxidative free radicals produced by inflammatory cells in contact with sperm can lead to genotoxic damage, which may have implications for male-mediated congenital defects and post-implantation losses (Chellman et al., 1986). The efferent ducts, which join the caput epididymis with the rete testis, are a particular site for damage.

Certain chemicals, e.g., carbamates, cause sperm stasis and inflammation of these ducts resulting in partial or complete obstruction to sperm transit and secondary dilatation of seminiferous tubules. The mechanism may be through increased fluid absorption resulting in sperm stasis and inflammation (Hess, 1998). The efferent ducts are also a frequent site for the occurrence of spontaneous sperm granulomas. In species such as the dog, they often form blind ending tubules that contain inspissated sperm, which can develop inflammation and progress to sperm granulomas. Ductular dilatation/interstitial edema This can occur as a result of fluid imbalance mediated through the vasculature or inhibited fluid reabsorption by the epithelial cells. Inflammatory infiltrate and sperm granulomas are a frequent consequence.

Prostate/Seminal Vesicles Acinar atrophy Secretory activity by the prostate and seminal vesicles is a sensitive, androgen-dependent function. Decreased circulating testosterone levels, or interference with androgen receptors in these two tissues will result in reduced secretion leading to atrophic changes. These may be detected by organ weight changes as well as by microscopic changes

PRACTICAL APPROACH FOR EXAMINATION OF THE TESTIS AND EPIDIDYMIS FOR TOXICOLOGICAL EFFECTS The approach used is influenced by the duration of the study. Cell- and stage-specific disturbances in spermatogenesis are usually only seen in short duration studies of 1 ml blood at each time point. Remove the minimal volume required to complete the work, and reinfuse resuspended RBCs whenever possible. By reinfusing the RBCs resuspended in saline, the animal’s blood volume and hematocrit remain somewhat constant even though large amounts of blood and plasma have been removed over the total sampling period. RBCs can be damaged, however, as a result of all the processing. Thus, it is advisable to check hematocrit levels of the animals periodically. Hematocrit levels will not change immediately following blood transfusions or blood loss. Thus checking the hematocrit at the beginning and end of a sampling period will yield little difference in levels. Check the hematocrit on the first blood sample collected and again 24 hr later. Rats rarely show adverse clinical signs until their hematocrits are quite low (40% to 50% of normal). It would, however, be advis-



able to make sure that the hematocrits are within 10% to 20% of the original values before initiating a second sampling period (e.g., 1 or 2 days later). Additional do’s and don’ts during blood sampling include (1) making sure there is no air in the cannula and/or syringe when reinfusing RBCs or flushing through heparinized saline, (2) flushing saline or reinfusing RBCs relatively slowly to avoid a vascular system overload with a large bolus of solution, (3) avoiding leaving blood in the cannula for more than a few seconds (clotting occurs rather rapidly), and (4) avoiding using infusion solutions colder than room temperature. Whether the permanent tether system or the vascular access port is used, if the cannula is patent for infusion but blood cannot be withdrawn, then there is the possibility that a small clot or flap may have formed at the tip of the cannula. Filling the cannula with a fibrinolytic agent, such as streptokinase or trypsin, for 24 hr may dissolve the clot. Often a fibrous sheath will form around the cannula. This has been determined to be composed of vascularized fibrous connective tissue and not fibrin (O’Farrell et al., 1996). Thus, fibrinolytic agents have little to no effect on it. It is best to flush the cannula with heparinized saline (in the case of the tether system) as often as possible, or to use a continuous infusion pump, to keep clots from forming. With the vascular access port, flushing the port every other day for the first week postsurgery is recommended. Generally, it is not necessary to check patency and flush the port and cannula more than once or twice a week after that. Depending upon study design, the blood sampling procedure can be very labor intensive. If blood samples (with RBC reinfusion) are to be collected every 10 min, one person should be able to sample up to three animals (if all goes well). If 30 animals are being sampled at one time, then blood sampling personnel will number at least 10. At least two additional persons will be needed to continually process the blood samples during the sampling period. One or two people may be used to transport blood samples and reinfusion samples back and forth between the animal room(s) and the processing room. Another individual can provide technical and troubleshooting support to all other staff (backup). The use of cannulated animals is particularly important when (1) stress hormones are being studied, (2) the effects of anesthesia are unde-

sirable, (3) only one person is available for sampling, or (4) infusions are administered via computerized or mechanical pumps. With either approach (permanent tether or vascular port), blood samples can be withdrawn and/or treatments administered without handling the animal. This minimizes the stress response of hormones such as prolactin, corticosterone, oxytocin, and/or epinephrine, which are released when animals are restrained, anesthetized, frightened, or in pain (Suzuki et al., 1997). The collection of samples from cannulated animals minimizes the release of these stress hormones and their impact on the hormones being studied. For example, norepinephrine and epinephrine concentrations were 2- to 8fold lower in blood samples collected from jugular cannulas than in those collected from decapitated animals (Paulose and Dakshinamurti, 1987). Corticosterone response due to handling or stress was minimized (MacLeod and Shapiro, 1988). Cardiac response to the cannulation is resolved within 24 hr (Paulose and Dakshinamurti, 1987), and LH response to GnRH is normal within 2 days (Fail et al., 1992, 1998). FSH response may not be normal until 5 to 7 days after cannulation (P. Fail, unpub. observ.). Replacement of RBCs is used to maintain hematocrit in the normal range (MacLeod and Shapiro, 1988) for single samples taken over several days or during intense sampling windows such as for the ECT or LEBA (Fail et al., 1992).

Anticipated Results The use of intra-atrial cannulation for serial sampling of unrestrained, untreated animals to illustrate endocrine secretion patterns is well established in the reproductive endocrinology literature and has significantly contributed to our understanding of endocrine signaling. These LEBAs are designed based on knowledge of the endocrine system being evaluated. For example, samples collected every 10 to 15 min during a 2- to 4-hr period can be used to determine profiles of a single hormone such as LH or thyroid stimulating hormone (TSH) and/or the relationship of two or more hormones (such as LH and testosterone, or TSH and thyroxine). The authors (Fail et al., 1992, 1995, 1998, 1999) have collected endocrine data using both of these designs. The relationship of secretory patterns of LH to those of testosterone for untreated male rats in a LEBA is shown (Fig.



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A 5.50

2.00 LH testosterone 1.50 LH (ng/ml)

3.50 1.00

2.50

1.50

0.50

Testosterone (ng/ml)

4.50

0.50 0.00

10:00

11:00

12:00

13:00

14:00

B 4.50

20.00

3.50

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LH (ng/ml)

3.00 2.50 10.00 2.00 1.50

Testosterone (ng/ml)

LH testosterone

4.00

5.00

1.00 0.50 0.00

0.00 10:00

11:00

12:00

13:00

14:00

Sampling time (time of day)

Figure 16.5.3 Longitudinal endocrine baseline assessment (LEBA) in two animals. Plasma luteinizing hormone (LH) and testosterone concentrations in a male Sprague Dawley rat were collected every 10 min during a 4-hr period. (A) Data are typical of a control (vehicle-treated) animal used for repeated sampling without endocrine challenge. The relationship of the two hormones within each time point and the prolonged response of testosterone to the LH spike is evident. In control animals, the LH spike lasted 2 hr. (B) In animals treated with flutamide, LH concentrations were increased in amplitude and the spikes lasted longer than in controls. Testosterone response to LH, although it occurred, was of a shorter duration than for the control rat. Ten cannulated animals were prepared for each treatment group in this experiment, but the data were examined for individual animals, and typical curves are shown here. Data are from the authors’ laboratory and are used with client’s permission.

Monitoring Endocrine Function in Males



SD

0 TRH (n = 4) 10 TRH (n = 6-7) 25 TRH (n = 7-8) 100 TRH (n = 7) 250 TRH (n = 8) 1000 TRH (n = 4-5)

F344

0 TRH (n = 15-16) 10 TRH (n = 3-4) 25 TRH (n = 2-3) 100 TRH (n = 14-15) 250 TRH (n = 13-14) 1000 TRH (n = 15-17) 2500 TRH (n = 8-9) 5000 TRH (n = 6-7)

20

TSH (ng/ml)

15

10

5

0 20

TSH (ng/ml)

15

10

5

0 0

15

30

45

60

Time (min)

Figure 16.5.4 An endocrine challenge test (ECT) summarized for groups of animals. The response of thyroid stimulating hormone (TSH) to different concentrations of thyrotropin releasing hormone (TRH; dosage given in nanograms per 100 g) in male Sprague Dawley (SD) and Fisher 344 (F344) rats. In this ECT, an effective dose and duration of response was established. The difference in effective dose 50 (ED50) for the two strains was established. Occasionally, samples could not be obtained from all animals for a specific time point. This is indicated by a range of n values. The lower number is the minimal number of animals represented in all points for that treatment. Asterisks indicate a statistically significant difference from controls within treatments (p ≤ 0.05). Reprinted from Fail et al. (1999) with permission from Oxford University Press.

16.5.3A), and the data agree with those reported elsewhere (Ellis and Desjardins, 1982). The secretory patterns were changed for LH, FSH, and testosterone in males given flutamide (Fig. 16.5.3B). The authors also have ED50 data for LH, FSH, and testosterone after GnRH and hCG challenge in Sprague Dawley males, and for TSH, triodothyronine (T3), and thyroxine (T4) after thyrotropin releasing hormone (TRH) challenge (Fig. 16.5.4) in male Sprague Dawley and F344 rats.

Time Considerations The total time for the cannulation protocol is short compared with that for the entire experiment. Planning should include the overall experimental plan. The total time will equal the sum of the following parts. 1. The development and design of the experiment requires 1 to 4 weeks depending on the available preliminary data and the knowledge of the endocrine system being evaluated. (An endocrine consultant should be used as needed.)



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2. Animal Use Committee approval requires 1 to 4 weeks based on the frequency with which the committee meets. Also plan for Quality Assurance Unit approval of the experimental protocol if relevant. 3. Allow time for ordering supplies and materials; some materials are difficult to obtain or require assembly. The design will dictate whether vascular access ports are needed or if noncoring needles should be used for repeated access. 4. The surgery time per animal is based on the experience of the surgeon. Some animals will be more difficult to complete than others. If 30 animals are being used, allow 2 or 3 days. Sterilization of materials requires 1 to 7 days. 5. A recovery time of 7 days is recommended for indwelling (vascular access) ports. For an acute or short-term protocol, 2 days may be sufficient. The cannula should be flushed every other day during the recovery period. 6. The time required for sample collection depends on the experimental design. Allow 1 day for ECT and 1 day for LEBA studies. These times may be stretched out if all animals cannot be sampled during the same day (and same time regimen). Each technician can handle two to four animals simultaneously depending on the sampling regimen. The authors routinely handle 30 animals in a protocol using 10 to 14 technical staff. Allow 6 hr for a 4-hr LEBA. 7. Processing of the blood samples occurs simultaneously with sample collection. This includes the time needed to label and store the samples. Two technical staff are required. Subsequent time considerations beyond the scope of these protocols include the sample radioimmunoassays (which require 2 days to 1 week per hormone and are hormone and sample number dependent), data management and statistical analysis (which require 2 to 4 weeks), and the generation of a study report (which requires 1 to 4 weeks, depending on the extent of the project and the use of good laboratory practices regulatory oversight).

Literature Cited Coquelin, A. and Desjardins, C. 1982. Luteinizing hormone and testosterone secretion in young and old male mice. Am. J. Physiol. 243:E257-E263. Dons, R.F. and Havlik, R. 1986. A multilayered cannula for long-term blood sampling of unrestrained rats. Lab. Anim. Sci. 36:544-547. Ellis, G.B. and Desjardins, C. 1982. Male rats secrete luteinizing hormone and testosterone episodically. Endocrinology 110:1618-1627. Monitoring Endocrine Function in Males

Fail, P.A., Sauls, H.R., Pearce, S.W., Anderson, S.A., and Izard, M.K. 1992. Measures of pituitary and testicular function evaluated with an endocrine

challenge test (ECT) in cannulated male rats. Toxicologist 12:436 (abstr. 1725). Fail, P.A., Pearce, S.W., Anderson, S.A., Tyl, R.W., and Gray, L.E. Jr. 1995. Endocrine and reproductive toxicity of vinclozolin (VIN) in male LongEvans hooded rats. Toxicologist 15:293 (abstr. 1570). Fail, P.A., Chapin, R.E., Price, C.J., and Heindel, J.J. 1998. General, reproductive, developmental, and endocrine toxicity of boric acid and inorganic boron-containing compounds: A review. Reprod. Toxicol. 12:1-18. Fail, P.A., Anderson, S.A., and Friedman, M.A. 1999. Response of the pituitary and thyroid to tropic hormones in Sprague-Dawley versus Fischer 344 male rats. Toxicol. Sci. 52:107-121. Harms, P.G. and Ojeda, S.R. 1974. A rapid and simple procedure for chronic cannulation of the rat jugular vein. J. Appl. Physiol. 36:391-392. MacLeod, J.N. and Shapiro, B.H. 1988. Repetitive blood sampling in unrestrained and unstressed mice using a chronic indwelling right atrial catheterization apparatus. Lab. Anim. Sci. 38:603-608. McKenna, M.L. and Bier, J.G. 1984. Multilayer cannula for long-term infusion of unrestrained rats. Lab. Anim. Sci. 34:308-310. O’Farrell, L., Griffith, J.W., and Lang, C.M. 1996. Histological development of the sheath that forms around long-term implanted central venous catheters. J. Parenter. Enteral Nutr. 20:156158. Paulose, C.S. and Dakshinamurti, K. 1987. Chronic catheterization using vascular-access-port in rats: Blood sampling with minimal stress for plasma catecholamine determination. J. Neurosci. Methods 22:141-146. Simard, J., Luthy, L., Guay, J., Belanger, A., and Labrie, F. 1986. Characteristics of interaction of the antiandrogen flutamide with the androgen receptor in various target tissues. Mol. Cell. Endocrinol. 44:261-270. Suzuki, K., Koizumi, N., Hirose, H., Hokao, R., Takemura, N., and Motoyoshi, S. 1997. Blood sampling technique for measurement of plasma arginine vasopressin concentration in conscious and unrestrained rats. Lab. Anim. Sci. 47:190193. Tyl, R.W. 2002. In vivo models for male reproductive toxicity. In Current Protocols in Toxicology (M. Maines, L.G. Costa, E. Hodgson, and D.J. Reed, eds.) pp. 16.1.1-16.1.15. John Wiley & Sons, New York. Viguier-Martinez, M.C., Hochereau de Reviers, M.T., Barenton, B., and Perreau, C. 1983a. Effect of a nonsteroidal antiandrogen, flutamide, on the hypothalamo-pituitary axis, genital tract and testis in immature rats: Endocrinological and histological data. Acta Endocrinol. 102:299-306. Viguier-Martinez, M.C., Hochereau de Reviers, M.T., Barenton, B., and Perreau, C. 1983b. Endocrinological and histological changes induced by flutamide treatment on the hypothalamo-hy-



pophyseal testicular axis of the adult male rat and their incidences on fertility. Acta Endocrinol. 104:246-252. Weeks, J.R. and Davis, J.D. 1964. Chronic intravenous cannulas for rats. J. Appl. Physiol. 19:540541.

Key Reference Anonymous, 1993. Preclinical Mini-Port Implantable Vascular Access Device, pamphlet 40-2931-

01A. Pharmacia-Deltec (now SIMS Deltec), St. Paul, Minnesota. This pamphlet is the basis for the surgical implantation and the use of the vascular access port.

Contributed by Patricia A. Fail and Stephanie A. Anderson Research Triangle Institute Research Triangle Park, North Carolina



Supplement 13

Epididymal Sperm Count

UNIT 16.6

Epididymal sperm count is a widely used, simple, and sensitive method for assessing the effects of male reproductive toxicants on the epididymis and/or testis. Usually the epididymis is divided into three parts: the caput, corpus, and cauda (Fig. 16.6.1). It is the epididymal cauda that is extensively used to perform epididymal sperm count. The method described in this unit is principally used for fresh tissue from rodents; however, the method can also be used for frozen epididymides with modifications.

BASIC PROTOCOL

Materials PBS (APPENDIX 2A) Necropsied male rats or mice, treated and control, euthanized 15 × 60–mm (rats) or 15 × 30–mm (mice) and 1.5 × 2.0–cm petri dishes Slide warmer 13 × 100–mm test tubes and appropriate rack Microscope: 10× and 40× objectives and 10× or 15× eyepieces Necropsy board 0.1-mm-deep improved Neubauer hemacytometer and coverslips No. 11 scalpel blades 60°C water bath Gauze Hand tally counter Data collection system (i.e., hand-generated data form or computer-generated data collection system) Additional reagents and equipment for using a hemacytometer (APPENDIX 3B) NOTE: All protocols using live animals must first be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) and must follow officially approved procedures for the care and use of laboratory animals.

caput

vas deferens corpus

cauda

Figure 16.6.1 A rat epididymis and its three anatomic parts: caput, corpus, and cauda. Male Reproductive Toxicology Contributed by Yefan Wang Current Protocols in Toxicology (2002) 16.6.1-16.6.5 Copyright © 2002 by John Wiley & Sons, Inc.


Prepare equipment 1. Label 15 × 60–mm (rats) or 15 × 30–mm (mice) petri dishes one for each animal, with the corresponding animal numbers. Fill with PBS (2 or 10 ml for mice or rats, respectively), and place on a slide warmer set at 35° to 37°C. 2. Equilibrate a water bath to ∼60°C. If using a hot plate, use a glass beaker (100 ml) containing ∼50 ml water. The purpose of the water bath is to kill motile sperm and allow an accurate count.

3. Label 13 × 100–mm test tubes, one for each animal, with the corresponding animal numbers. Add 2 ml PBS to each tube. 4. Turn the microscope on. 5. Clean a 0.1-mm-deep improved Neubauer hemocytometer and coverslips (both need to be cleaned between animals). Prepare samples 6. Acquire the first necropsied male rats or mice, treated or control, euthanized by an approved method. 7. Place the animal on a necropsy board with the ventral side up. Open the abdominal cavity by making an ∼5 cm incision in the lower right abdominal area. Pull out the right epididymis and testis by grasping the epididymal fat with a pair of forceps, leaving the vas deferens attached as a reference point (Fig. 16.6.1). 8. Isolate the epididymis from the testis and trim away fat. Place the entire epididymis on a flat surface. Procure the epididymal cauda by cutting ∼1 mm down from the junction of the vas deferens and the epididymis cauda (see Fig. 16.6.1). 9. Measure and record the weight of the cauda in milligrams. 10. Place the cauda in a petri dish containing a known volume of PBS (2 or 10 ml for mice and rats, respectively) prewarmed to 35° to 37°C, and mince using two no. 11 scalpel blades to open the epididymal duct and release its contents. Swirl the petri dish several times to achieve a uniform sperm suspension. 11. After incubating at least 15 min at 35° to 37°C, mix the sperm suspension several times using a Pasteur pipet. 12. Using a 1-ml pipet, transfer 0.5 ml sperm suspension from the petri dish to a corresponding test tube containing 2 ml PBS, room temperature, avoiding transferring the epididymal tissue. Mix the tube by lightly tapping the bottom. 13. Place the test tube in a ∼60°C water bath for ∼1 min. Return the tube to the test tube rack and cool to room temperature. Count sperm 14. Mix the diluted suspension, measure 10 µl with a micropipet, and load one side of the hemacytometer. Next, mix the suspension again by vortexing and load the other chamber. Place the loaded hemacytometer onto a clean 1.5 × 20–cm petri dish, bedded with a portion of moist gauze, for 2 min to allow the sperm to settle. 15. Place the hemacytometer on the microscope stage and focus on the top left secondary square (square no. 1; see Fig. A.3B.1 in APPENDIX 3B) of the tertiary square. Epididymal Sperm Count

16. Count the number of sperm (entire structure) lying in the five secondary squares (nos. 1 to 5) and lying on or touching the top and left lines of the secondary squares (Fig.



Figure 16.6.2 Actual microscopic photograph of the secondary square (area within the triple line border) of a hemacytometer (see Fig. A.3B.1 in APPENDIX 3B). The black arrows indicate the nine sperm that will be counted; five white arrows identify the sperm that will not be counted.

16.6.2). If there are less than 25 sperm in those five secondary squares, count the sperm in the entire tertiary square (all 25 secondary squares). If any part of the sperm lies within or over the right and/or bottom lines, crossing right and bottom lines, it must be excluded from the count. If any part of the sperm lies within or over the left and/or top line, crossing either top or left line, that sperm must be counted. There is one tertiary square on each side of the hemacytometer.

17. If clumped sperm are noted, reload and recount. 18. Record two counts for each animal on a hand tally counter and data collection system. 19. Calculate the total number of sperm per cauda epididymis based on the secondary count: Total sperm = mean count × dilution factor where: Mean count = (count 1 + count 2)/2 and dilution factor =

total vol. PBS in dish total vol. test tube × transferred vol. vol. secondary square × no. squares

For the protocol listed above, the transferred volume is 0.5 ml and the volume of a secondary square is 4 × 10−6 ml.

20. Calculate the epididymal sperm count per milligram cauda (sperm density): Sperm density = (mean count × dilution factor)/cauda weight (mg) or Sperm density = total number sperm per cauda/cauda weight (mg).



Supplement 14

COMMENTARY Background Information The epididymis is a single, highly-coiled duct with functions that include reabsorption of the rete testis fluid, metabolism, epithelial cell secretions, sperm maturation, and sperm storage. The caput and the corpus play important roles in sperm maturation, with the cauda regarded as the site of sperm storage. The sperm require 1.8 to 4.9 days to move through the caput to the corpus epididymis and the travel time of sperm through the cauda ranges from 3.7 to 9.7 days among different species (Klassen, 1996). The first appearance of spermatozoa is detected in the caput at 45 days of age and in the cauda at 52 days of age, while the sperm population reaches the maximum in the caput at 72 days of age and in the cauda at 100 days of age. The sperm population is maintained beyond 450 days of age (Saksena et al., 1979). Table 16.6.1

The number of sperm in the corpus and the caput epididymis is similar in sexually rested males and in males ejaculating daily. The number of sperm in the cauda epididymis may vary depending on the male’s sexual activity (see Table 16.6.1). Epididymal tissues are often obtained after a terminal sacrifice. This method can be modified and used on other species by adjusting the dilution. W.G. Kempinas (1988) recommends treating sperm suspensions with 0.05% collagenase for 20 to 60 min or 0.025% trypsin for 1 to 2 min at 34° to 37°C, which was found to result in a consistently homogeneous sperm sample. The Hamilton Thorne Biosciences Integrated Visual Optical System provides a fluorescent method of determining sperm concentration without the interference of detritus by

Sperm Count in the Reproductive Track of Albino Ratsa

Sexually rested rats Mated rats

Number of animals

Number of Sperm in vas deferensb

Number of sperm in caudab

Number of sperm in caput and corpusb

13 12

28.2 ± 3.4 3.4 ± 1.6

299.3 ± 19.7 110 ± 5.8

153.9 ± 8.2 129.8 ± 11.1

aRats are the Wistar strain of Rattus norwegicus. See Ratnasooiya and Wadsworth (1987) for details. bSperm count for the paired tract, given as mean ± s.e.m. All values are given in millions.

Table 16.6.2

Troubleshooting Guide for Sperm Count Errors

Problem

Possible cause

Solution

Count too low

Animals are too young (125% of the lower count), the count should be repeated (steps 18 to 23). As an example, if the first count was 67 and the second count was 77, the second count is 15% higher than the first count: (77/67) × 100 = 1.149 or 15% higher. The count should also be repeated if any of the counts are 110. In the case of a repeated count only the two most recent counts are used to generate an average. Sperm samples with very high count (>150) can be diluted with DMSO/saline before recounting. Counts will generally range from 60 to 100 spermatids for a typical rat or mouse testis. Various factors such as low testis weight or incorrect homogenization volume can change the counts.

CALCULATING TESTICULAR SPERMATID HEAD CONCENTRATION Each chamber of the hemacytometer is designed to hold 0.1 µl of solution in its tertiary square. The tertiary square is divided into 25 secondary squares, which are in turn divided into 16 primary squares. Thus each secondary square contains 0.004 µl. Because ten secondary squares are used for counting, 0.04 µl of suspension is used per chamber. The total volume of rat testis suspension is 100.5 ml, and mouse testis is 10.05 ml. Thus, the total number of spermatid heads in a rat testis is (mean count/0.00004 ml) × 100.5 ml. The total number of spermatid heads in a mouse testis is (mean count/0.00004 ml) × 10.05 ml.

SUPPORT PROTOCOL

Example 1: If counts for a rat were 75 and 79, or a mean count of (75 + 79)/2 = 77, then the total number of spermatid heads in the testis is (77/0.00004) × 100.5. This is equal to ∼193 million spermatid heads. If the testis weighed 1.529 g, then there are 127 million spermatid heads per gram testis. The daily sperm production is then 20.7 million spermatids per gram testis per day or 31.7 million spermatids per testis per day, using 6.10 days as the time divisor (see Background Information; Oakberg, 1956). Example 2: If the counts for a mouse testis were 69 and 76, or a mean count of (69 + 76)/2 = 72.5, then the total number of spermatid heads in the testes is (72.5/0.00004) × 10.05. This is equal to ∼18.2 million spermatid heads. If the testis weighed 0.1016 g, then there are 179 million spermatid heads per gram testis. The daily sperm production is then 37.0 million spermatids per gram testis per day or 3.8 million spermatids per testis per day, using 4.84 days as the time divisor (see Background Information; Robb et al., 1978). Male Reproductive Toxicology


Supplement 16

REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.

Dimethyl sulfoxide (DMSO)/saline solution Place 18.00 g NaCl (0.9% final) in a 2-liter container. Add 1.60 liter water. Mix vigorously to dissolve NaCl. Add 200 ml DMSO (10% final). Bring to 2 liters with water. Store up to 30 days at room temperature. This volume should be enough to suspend ∼20 samples of rat testes or 200 samples of mice testes. CAUTION: DMSO can be very toxic because of its permeability.

Trypan blue stain, 0.1% (w/v) Mix together 10 mg trypan blue and 10 ml water. Stir until all crystals are dissolved. Store up to 1 year at room temperature. The solution is a dark navy blue color. It may take a few minutes of stirring for all crystals to dissolve completely. This volume should be enough for ∼20 samples of rat testes or 200 samples of mice testes.


Performing a Testicular Spermatid Head Count

The testis is a multicompartmental organ packed with highly convoluted tubules called seminiferous tubules. These tubules meet to form the rete testis. From the rete testis, ductuli efferentes connect to the epididymis where sperm mature and are stored. These tubules contain Leydig cells, Sertoli cells, spermatogonia, and other cells. Leydig cells secrete hormones such as testosterone. Sertoli cells secrete fluid into the lumen of the tubules. The testis is wrapped in a tough connective tissue called the tunica, which holds the seminiferous tubules together. The seminiferous tubules are lined with spermatogonial stem cells known as A0. A mitotic division of these cells gives rise to A1 spermatogonial cells. These cells gradually displace toward the lumen by subsequent mitotic divisions. After six mitotic divisions, spermatogonial cells divide to produce primary spermatocytes. Further meiotic division produces secondary spermatocytes, and after another meiotic division, they form spermatids. At the beginning, the heads of spermatids are round without remarkable morphology. However, they undergo a dramatic metamorphosis and become spermatozoa, which look like hooks. Toward the end of this metamorphosis, the nucleoprotein condenses and the cells become resistant to homogenization and detergent. The period during which cells are resistant to homogenization and detergent is referred to as the time divisor and is used to calculate the

daily production of spermatids in a testis. The time divisor for rats is 6.10 and that of mice is 4.84 according to Oakberg (1956) and Robb et al. (1978), respectively. The overall duration of spermatogenesis within a testis is 52 days in rats and 36 days in mice. An alternative method of estimating daily sperm production is by counting A1 cells. A1 cells are the spermatogonial cells derived from spermatogonial stem cells. By counting the A1 cells, one can estimate the theoretical concentration of spermatids in a testis. Not all cells arising from A1 cells, however, will become mature sperm. Also, this requires histological sampling. This method requires more time and materials and is more prone to error during preparation without better accuracy, as compared with the method of counting spermatid heads in testicular homogenates described here. The epididymal sperm count (UNIT 16.6) provides quantitation of sperm that is stored and thus is available for ejaculation and fertility. Sperm is stored over a period of time, however, masking acute changes in the sperm production rate. Unlike epididymal sperm count, the testicular spermatid head count provides better quantitation of acute changes in sperm production, thus providing a better snapshot of changes in spermatogenesis within testes. This parameter can provide crucial information regarding reproductive toxicity of a test compound. Either an increase or decrease in the concentration of spermatid heads can indicate pathology. An increase in the spermatid head



concentration suggests that there is sperm retention (or inhibited sperm release; Ku et al., 1993; Linder et al., 1997). A decrease in the spermatid head concentration may be due to atrophy of the testis, changes in circulating hormones, loss of germ cell line, or physiological changes to Sertoli cells or Leydig cells. Histological examination may help clarify the underlying pathologic process (UNITS 16.3 & 16.4). Spermatid head count will not, however, provide any information about fertility without other parameters such as motility, morphology, and epididymal sperm counts. There are two mixtures commonly used for homogenization. The authors prefer using DMSO/saline because of its shorter preparation time and longer expiration date. Another solution to suspend the spermatid heads is 0.9% (w/v) saline/0.1% (v/v) Triton X-100. This solution requires several hours to prepare and expires in weeks. Unlike saline/Triton X-100, DMSO/saline can be prepared within minutes, so as not to lose the capability of counting spermatid heads, and expires in 1 month. The Basic Protocol describes a method that uses a hemacytometer to count testicular spermatid heads. An alternative method is to use a computer-assisted counting machine, such as the Hamilton Thorne 2030 (Hamilton Thorne Biosciences) or IVOS (Hamilton Thorne Biosciences). According to Centola (1996), the variability between samples is less when counts by the computer system are compared with manual ones, and counting error within a sample is comparable between the two methods.

Critical Parameters and Troubleshooting Storage conditions It is critical that the testes not be stored at −70°C for >6 months; for long-term storage (up to 12 months), they should be stored at −80°C or below. If the testes are not stored at −80°C or lower, they will be under risk of desiccation (freezer burn), a drying of the sample during storage in a freezer. Desiccation changes testicular weight, thus changing data for spermatids per gram testis. Thawing The purpose of thawing is to remove the parenchyma surrounding the testis. If the testis is not thawed sufficiently, it will make removal of the parenchyma difficult. If the testis is thawed too much, then there is a risk of remov-

ing a portion of the testis, decreasing the total number of spermatids per testis. Homogenization All the materials used during homogenization must be clean. Otherwise there will be cross-contamination between samples. Rinsing with deionized water three to four times will prevent cross-contamination between samples. Consistency of counts It is imperative for the person who counts the spermatids to be consistent. All the counts must be done in exactly the same manner. For example, if right and lower borders of secondary squares are included and not upper and left borders, then all the counts must include any spermatid heads touching lower and right borders, and they must exclude any spermatid heads touching upper and left borders of secondary squares. Because only a small amount of the solution is used to count, the stained spermatids must be thoroughly suspended before loading the hemacytometer. Small deviations in the suspension will result in a great variability in the concentration of spermatids. Also, it is important to focus in and out during counting. Because of the high magnification power of the microscope, the focal plane is rather shallow. Most of the spermatid heads will settle down on the hemacytometer, but some may stay suspended between the grids and the coverslip. Exclusion of these spermatid heads will underestimate the true concentration and renders the data inaccurate. Data analysis Repeating a spermatid head count if the higher count is >125% of the lower count is to prevent random abnormal sampling. Repeated counts ensure that an abnormally higher or lower count does not affect the mean of the group being analyzed. Counts of 110 spermatids are repeated to ensure accuracy of a dense field. It is harder to count when there are many overlapping spermatids in the field.

Anticipated Results A typical count using the hemacytometer method described in this unit will yield between 60 to 100 testicular spermatid heads for rat or



Supplement 16

mouse testis. According to a study conducted by Blazak et al. (1985) on the daily sperm production rate of Fischer 344 rats, a typical 20-week-old male rat has a testicular weight of 1.2 to 1.6 g/testis and produces 16 to 30 million sperm a day. With a time divisor of 6.10, a typical total spermatid head count for a rat testis will contain 185 million sperm. In contrast, according to the same study, the daily sperm production rate of a typical 13week-old male B6C3F1 mouse is 3.4 to 6.8 million per day, and its testis weighs 0.08 to 0.11 g. With a time divisor of 4.84 days, a typical total spermatid head count from a mouse testis is 24.1 million sperm.

Time Considerations The whole procedure, including making solutions, will require 10 hr to perform spermatid head counts on 20 rat testes. The approximate time required for each step is as follows: solution preparation, 0.5 hr; setup and labeling flasks or tubes, 0.5 hr; sample preparation, 4 hr; spermatid head counts, 5 hr.

Literature Cited Blazak, W.F., Ernst, T.L., and Stewart, B.E. 1985. Potential indicators of reproductive toxicity: Testicular sperm production and epididymal sperm number. Transit time, and motility in fischer 344 rats. Fundam. Appl. Toxicol. 5:1097-1103. Centola, G.M. 1996. Comparison of manual microscopic and computer-assisted methods for analysis of sperm count and motility. Arch. Androl. 36:1-7.

Feldman, D.B. and Seeley, J.C. 1988. Necropsy Guide: Rodents and the Rabbit. CRC Press, Boca Raton, Fla. Ku, W.W., Chapin, R.E., Wine, R.N., and Gladen, B.C. 1993. Testicular toxicity of boric acid (BA): Relationship of dose to lesion development and recovery in the F344 rat. Reprod. Toxicol. 7:305319. Linder, R.E., Klinefelter, G.R., Strader, L.F., Suarez, J.D., and Roberts, N.L. 1997. Spermatotoxicity of dichloroacetic acid. Reprod. Toxicol. 11:681688. Oakberg, E.F. 1956. Duration of spermatogenesis in the mouse and timing of stages of the cycle of the seminiferous epithelium. Am. J. Anat. 99:507-516. Robb, G.W., Amann, R.P., and Killian, G.J. 1978. Daily sperm production and epididymal sperm reserves of pubertal and adult rats. J. Reprod. Fertil. 54:103-107.

Key Reference Zenick, H., Clegg, E., Perreault, D., Klinefelter, G., and Gray, L. 1994. Assessment of male reproductive toxicity: A risk assessment approach. In Principles and Methods of Toxicology, 3rd ed. (A.W. Hayes, ed.) pp. 937-988. Raven Press, New York. Useful and detailed background information.

Contributed by Han Seung, Gary Wolfe, and Meredith Rocca TherImmune Research Corporation Gaithersburg, Maryland

Performing a Testicular Spermatid Head Count



Transgenerational (In Utero/Lactational) Exposure to Investigate the Effects of Endocrine Disrupting Compounds (EDCs) in Rats

UNIT 16.8

In this unit, timed-pregnant Sprague-Dawley rats are dosed daily during pregnancy and in some instances through lactation. At birth the number of pups/litter are counted and at 2 days of age, anogenital distance and body weight are recorded for all pups. When the pups are 13 days old, nipple and areolar retention are evaluated in females exposed to potential androgens or males exposed to potential antiandrogens. Pups are weaned at 21 to 28 days and housed in unisexual littermate groups of two to three pups/cage; the dams are sacrificed, and body weight and the number of implantation scars are recorded at this time. The onset of puberty is monitored by observing the age of vaginal opening in the females and the age of preputial separation in the males. Other endpoints that may be collected after puberty, if indicated, include estrous cyclicity and fertility in females after puberty, and ejaculated sperm counts, mating behavior, and fertility in males after sexual maturity (after 3 months). As soon as all necessary data have been collected, sexually mature rat offspring are necropsied and an extensive evaluation of the reproductive system is conducted. The basic protocols recommended for transgenerational studies investigating the effects of EDCs on the reproductive system are: study design and exposure (see Basic Protocol 1); anogenital distance measurement (see Basic Protocol 2); assessment of areolae/nipple development (see Basic Protocol 3); detection of puberty (vaginal opening and preputial separation; see Basic Protocol 4); and necropsy, histology, and radioimmunoassays (see Basic Protocol 5). NOTE: All protocols using live animals must first be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) and must conform to governmental regulations regarding the care and use of laboratory animals. STUDY DESIGN This protocol describes basic techniques for assigning timed-pregnant rats to treatment groups and administrating test compounds.

BASIC PROTOCOL 1

Materials Individually housed timed-pregnant rats Test compounds Laboratory-grade corn oil (Sigma) Balance, accurate to 0.1 g, that integrates the weight over several measurements (e.g., Sartorius IP65) 1-ml glass tuberculin syringe (BD) 1.5-in. × 18- or 20-G curved gavage needle (Popper and Sons) 25-G × 5/8-in. or 1/2-in. sterile needle (BD) Saturated picric acid solution and envelope moistener or cotton swabs to apply picric acid to dams for treatment identification Software such as Microsoft Excel or SAS (PC SAS or IBM Host on Demand) for calculating means and standard errors by treatment FTP95 for transferring SAS output data sets to WordPerfect or Microsoft Word Contributed by Joseph S. Ostby and L. Earl Gray Jr. Current Protocols in Toxicology (2004) 16.8.1-16.8.16 Copyright © 2004 by John Wiley & Sons, Inc.



Assign rats to treatment groups 1. On the morning when dosing is initiated, give a temporary cage number (cage cards numbered consecutively in pencil) to individually housed timed-pregnant rats and record their body weights. 2. After weighing, sort the rats by body weight from the lightest to the heaviest rat and assign treatment groups in the following manner: Randomly assign lightest rat to one treatment group Randomly assign next lightest rat to one of the remaining treatment groups Continue assigning rats in this manner until each treatment group contains one rat to form a block (block = a group of rats with similar body weights consisting of 1 rat/treatment group), which contains the lightest rats in the experiment. 3. Randomly assign a permanent cage number ranging from 1 through the total number of treatments to each rat in the first block. Assign the second block from the next lightest group of rats to treatment groups in a similar manner and give a permanent cage number ranging from the number equal to 1 plus the total number of treatment groups through the number equal to two times the total number of treatment groups in a random fashion. Thus, if the study has five treatment groups, the first block consists of the five lightest rats housed in cages 1 through 5 and the next block consists of the next five lightest rats housed in cages 6 through 10. Continue this procedure until all rats have been assigned to treatment groups and the number of blocks equals the number of rats/treatment group. This procedure minimizes potentially confounding experimental variables across treatment groups. The first block of rats (one rat/treatment group) representing the lightest rats in the experiment are placed on the first row(s) of the rack so that each rat is exposed to similar environmental conditions such as temperature, humidity, air flow, and light intensity. Likewise, the second block of rats will be placed on the next row(s) of the rack ensuring all rats in this block will also be exposed to similar environmental conditions.

4. After treatment assignment, calculate the mean body weights and standard errors for each treatment group to ensure that they are as similar as possible and not significantly different. 5. Once the rats have been assigned to treatment groups, give them an i.d. to identify their treatment group. Use picric acid to identify the treatment group. For example, rats could be marked as follows: left rear leg (LR), right rear leg (RR), left front leg (LF), right front leg (RF), back (B), both hind limbs (LRRR), both forelegs (LFRF), both left legs (LRLF), and both right legs (RRRF). Other means of marking the animals include ear tags, tail tattoos, or implantable transponders.

Prepare and administer dosing solutions 6. Prepare dosing formulations on the basis of microgram or milligram test compound/kilogram body weight/milliliter vehicle/day. Tare the dosing solution vial on a balance, add the calculated quantity of the test compound and record its weight, then add the necessary volume of the vehicle (laboratory-grade corn oil). After thorough mixing, further dilutions can be made from the original stock solution. Effects of Endocrine Disrupting Compounds (EDCs) in Rats

7. Weigh timed-pregnant rats and prepare the dose for gavage at 2.5 µl of the vehicle per gram body weight (2.5 ml corn oil/kg body weight; therefore, the volume administered to a 300- to 400-g pregnant rat would be 0.75 to 1 ml). Accurately measure this volume with a 1-ml glass tuberculin syringe.



A 1-ml dose is sufficient to allow most compounds to go into solution but does not exceed the stomach capacity of the rat. Dose the timed-pregnant rats orally or inject subcutaneously (s.c.) with test compounds in a vehicle (e.g., laboratory-grade corn oil) depending on the chemical and its concentration (whether a solution or suspension). Some compounds readily go into solution while others may require mixing for several days or even 1 week to form uniform suspensions. Suspensions must be stirred constantly while dosing and can be difficult to administer s.c. through small gauge needles. To avoid degradation of test compounds, avoid heating (to facilitate mixing) of dosing solutions unless chemical stability is assured. Depending upon solubility, a wide variety of other vehicles, other than corn oil, may be used.

8a. For oral administration: Dose orally by gently grasping the loose skin on the back and neck of the rat in such a manner that the head, neck, and back form a straight line. Gently insert a 1.5-in. × 18- or 20-G curved gavage needle into the left side of the oral cavity and down into the esophagus past the epiglottis (if any resistance is encountered, withdraw the needle and re-insert). To avoid discomfort and startling the rat, administer the dosing solution at a moderate rate and withdraw the needle along a path similar to the path used to insert the needle. Prior to oral dosing, the rats should be gently held in one hand and then held as described above to acclimate them to this type of restraint (since they are normally only picked up by their tails during cage changing). Administration of compounds typically requires a health and safety research protocol. Prior to administration of these compounds the rats should be dosed for several days with water to acclimate them to this procedure and to reduce the possibility of the rat jumping while the compound is being administered. Rats undergoing rapid weight changes such as timed-pregnant rats should be weighed daily and the dose adjusted to body weight.

8b. For subcutaneous administration: Administer subcutaneous injections as microgram or milligram of the test compound/rat in a constant vehicle volume/day or microgram or milligram of the test compound/kilogram body weight/milliliter vehicle/day. Administer injections by placing the rat on a flat surface and gently grasping the loose skin on the back of the rat between the last three fingers and the heel of the hand and raise the skin on the neck with the thumb and index finger. Insert a 25-G × 5/8-in. needle into the raised skin on the neck below the thumb and index finger. Once this is accomplished, the tip of the needle can be felt between the fingers, and injection can be confirmed by feeling the fluid flow into the injection site (the pressure produced even by microliter quantities can easily be detected in this manner). No more than ten rats should be injected with a single needle. Needles that are 25 or 27 G permit a moderate flow rate with corn oil (be aware that excessive plunger pressure may forcefully eject the needle and syringe contents) but are small enough to prevent excessive leakage at the injection site. For example, potent steroidal androgens are administered at 0.1 to 10 mg per rat per day. The optimum injection volume (∼0.1 ml of the vehicle) can be accurately administered with a gas-tight microliter or a 1-ml tuberculin syringe, does not readily leak from the injection site due to excessive pressure, and is readily absorbed within 24 hours.

9. For the dosing described in steps 8a and 8b, use different dosing syringes and needles for each dose group to avoid contamination from residual material. Discard syringes on a daily basis, clean the gavage needles daily during the study and reuse but only for the same dosing solution. Male Reproductive Toxicology


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10. Select exposure regime. Exposure may be initiated on gestational day 8 (GD 8) prior to onset of fetal gonadal differentiation or on GD 14 near the onset of fetal testis steroid hormone synthesis (GD 1 = day of sperm positive smear). Exposure should continue through at least GD 18 to cover the primary period of reproductive tract development, but pups may also be exposed via the mother’s milk through postnatal day (PND) 3 to encompass the period of sexual differentiation of the brain and central nervous system, or exposure may continue through the period of lactation. One final option would be to continue exposures to the pups after weaning using the same route and dosage levels given the dams. BASIC PROTOCOL 2

MEASURING ANOGENITAL DISTANCE The sexually dimorphic secondary sex characteristic in many mammalian species, anogenital distance (AGD; see Fig. 16.8.1), may be used to measure the degree of demasculinization of males as a consequence of developmental exposure to androgen receptor (AR) antagonists (e.g., vinclozolin, procymidone, flutamide, linuron, prochloraz, etc.), 5-alpha reductase inhibitors (finasteride), or compounds that inhibit steroidogenesis (some phthalates). Likewise, AGD is useful in measuring the degree of masculinization of females exposed during sexual differentiation to androgenic compounds such as testosterone or the anabolic growth stimulant trenbolone used in cattle. AGD and body weight are normally measured in rats at birth or 2 days of age. As the animal grows, increased activity and variability in growth rates increase the variability of this measurement. However, decreased male AGD and increased female AGD observed in adult rats at necropsy demonstrates that endocrine disrupting compounds do permanently alter AGD.

Effects of Endocrine Disrupting Compounds (EDCs) in Rats

Figure 16.8.1 Anogenital distance in neonatal male and female SD rats. Photographs taken at the same magnification of perineal area of male and female siblings demonstrating the normal developmental effects of testicular androgens on AGD (the sexually dimorphic distance between the sex papilla and the anus). This black and white facsimile is intended only as a placeholder; for full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm



Studies conducted in the authors’ laboratory that followed individual pups from birth to necropsy as adults have demonstrated there is a statistically significant correlation between AGD observed at birth and the presence of nipples in infant males, AGD at necropsy, as well as malformations observed when the rats were necropsied as adults. These results demonstrate that AGD is a reliable predictor of permanent alterations of the reproductive system that are not often apparent until the animal reaches sexual maturity. Similarly, increased AGD in female pups is associated with decreased numbers of nipples (as infants and adults) and malformations of the reproductive tract. Materials Sprague-Dawley or Long Evans pups at birth or on PND 2 and as adults Dissecting microscope, 0.63× to 4× and 10-mm (0.1-mm divisions) ocular reticle (e.g., Leica MZ6) 1-mm stage micrometer (0.01-mm divisions) Vernier caliper (0 to 150 mm with 0.1-mm divisions) Two-place balance (e.g., Mettler PM2000) 1. Set a dissecting microscope fitted with a 10-mm ocular reticle (with graduated 0.1-mm divisions) at a magnification of ∼1.5× and calibrate with a 1-mm stage micrometer (with graduated 0.01-mm divisions). 2. After the microscope has been set at ∼1.5×, carefully adjust the magnification so that the 1-mm stage micrometer is exactly equal to 1.5 mm on the ocular reticle. A magnification of 1.5× has been determined to be the optimum magnification for obtaining the most accurate AGD measurement without being distracted by subtle pup movements.

3. After calibrating, raise the microscope ∼4 to 6 in. using the coarse focus knob to allow the hands of the observer to rest comfortably on the stage while holding the pup. Position the perineal area of the pup in the focal plane and measure the distance between the posterior base of the sex papilla and the anterior anus using the ocular reticle. Record AGD along with the sex and body weight of the pup. Record the body weight to the nearest 0.01 g and record AGD to the nearest 0.1 scale on the micrometer (divide this measurement by 1.5, magnification = 1.5×, to obtain the actual AGD). At birth, AGD in male Sprague-Dawley or Long Evans rats is ∼3 mm, while female AGD is ∼50% smaller than the males or 1.5 mm.

4. At necropsy, measure AGD by placing the animal with the base of the tail on the edge of a table. Use the thumb to secure the tail to the side of the table and place the index finger above the phallus to maximally stretch the skin in the perineal area. Then, use a vernier caliper to measure the distance between the posterior base of the phallus and the anterior rim of the anus. 5. If pup body weight is significantly reduced by treatment, then adjust the AGD for body weight using body weight as a covariate in the statistical analysis. Since litter effects can occur particularly at this age, the data analyzed typically are litter means or n = number of litters/treatment unless it can be demonstrated that litter effects are not present in the data.



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BASIC PROTOCOL 3

ASSESSING AREOLAE/NIPPLE DEVELOPMENT IN INFANT MALE AND FEMALE RATS In the absence of testicular androgens, female rats develop areolae/nipples, while dihydrotestosterone induces regression or apoptosis of the nipple anlage. Thus, nipple development in male rats is a sensitive endpoint for evaluating the degree of demasculinization produced by in utero exposure to antiandrogenic compounds. Conversely, the partial or complete attenuation of areolae/nipples in female rats is useful in evaluating the masculinizing effects of androgenic compounds (see Fig. 16.8.2). Although this endpoint is a sensitive indicator of endocrine disruptions, in 13-day-olds a small percent of the control males may have faint areolae. Therefore, it is recommended that prior to experimental data collection for this technique, and AGD as well, in experimental situations, researchers should practice collecting these data in pilot studies with both control and treated (testosterone propionate, TP, for females and vinclozolin for males) animals from several litters to establish a baseline for recording these data and to assure acceptable levels of reproducibility. If more than one observer is used in a study, then one must assure that there is a high degree of inter-observer reliability on these measures, and the study should be designed in a manner such that the treatment groups are split among different observers. Due to the subjective nature of these endpoints, it is critical that the data are collected in a “blinded” manner, ensuring that the observer is not aware of the treatment group of each litter. Because of the rapid development at this age, AGD and nipple data should be collected on all offspring at the same age in days over one 8-hr interval, by a single observer, if possible.

Control Male


Control Female

DEHP Male

Figure 16.8.2 Areolas and nipples in infant male and female rats. Photographs taken when the pups were 14 days old showing a normal male without nipples, a normal female with twelve evenly spaced nipples (no. 9 not visible in photograph) and a male exposed to 750 mg of diethylhexyl phthalate from gestational 14 through postnatal 3. Some phthalate diesters such as diethylhexyl, benzylbutyl, and dibutyl phthalate inhibit fetal testosterone synthesis and thus prevent dihydrotestosterone from inhibiting development of the nipple anlage. This black and white facsimile is intended only as a placeholder; for full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm



Materials Infant male and female rats (e.g., Sprague-Dawley) Bouins’s fixative (see recipe) Balance (e.g., Mettler PM2000) Small animal clippers with size no. 40 clipper blade (Oster) 1. Collect areolae/nipple data shortly after control females develop prominent areolae/nipple buds but before the nipples are partially or completely hidden by rapid ventral hair growth (in Sprague-Dawley rats, this normally occurs when they are ∼13 days old). 2. On PND 13, have an observer who is blind to treatment separate each litter by sex, determine the number of areolae/nipples, their position, and the body weight of the pup for each male or female in the litter, depending on the type of compound administered. If antiandrogens are administered, males exposed to antiandrogens are checked for nipple retention and females exposed to androgens are checked for nipple regression. The sex not checked for nipple development at 13 days can be counted and weighed together to determine average pup weight for that sex. Nipple data collection normally requires three technicians: an observer, a data recorder, and a technician responsible for removing and returning litters to their home cage.

3. Number the nipples to identify their location (female rats normally have twelve nipples, six symmetrical pairs, with the pairs distributed identically on all rats). For example, give nipples on the right side odd numbers (1, 3, 5, 7, 9, and 11) with no. 1 caudal and no. 11 cranial, and assign nipples on the left side even numbers (2, 4, 6, 8, 10, and 12), with no. 2 caudal and no. 12 cranial. Rats with nipple retention or regression can be given a unique picric acid i.d. when 13 days old and shaved and checked at necropsy to determine if abnormal nipple development observed persists into adulthood. Also, retained areolae/nipples in males may be scored as absent, faint, moderately developed, or prominent (female-like) and regressed areolae/nipples in females can be scored as normal, moderately faint, very faint, or absent.

4. At necropsy, shave the rats with small animal clippers and record the number of nipples/areolae. Fix retained or regressed nipples in Bouin’s fixative and save for histological evaluation. DETECTING PUBERTY—VAGINAL OPENING (VO) AND PREPUTIAL SEPARATION (PPS)

BASIC PROTOCOL 4

The onset of puberty in the rat can be evaluated by recording the body weight and age of the rat at vaginal opening (VO) or preputial separation (PPS). Although direct exposure from weaning through puberty may be the optimum period of time to alter this endpoint, in utero/lactational exposure can also impact puberty and consequently should be checked in transgenerational studies. VO occurs when the rise in circulating ovarian hormones induces apoptosis of the vaginal membrane cells and development of the vaginal canal. The day of vaginal opening normally coincides with the day of first estrus and thus marks the initiation of the onset of estrous cyclicity in rats. The first few estrous cycles are typically longer and irregular after VO. VO is useful to identify/evaluate EDCs that accelerate (estrogen) or delay (antiestrogens, inhibitors of aromatase or inhibitors of hypothalamic-pituitary maturation) puberty in female rats. On occasion, females display a transient thread of tissue along the midline of the vaginal opening, which persists for several days or occasionally is



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permanent. For this reason, the age and body weight at the onset of VO should be noted as well as the completion of VO, with dissolution of the thread, if present. The authors have found that female rats exposed in utero to Ah receptor agonists often display a permanent thread across the vaginal opening. PPS in rats (complete manual retraction of the prepuce) occurs when increasing testicular androgen production produces cornification of the balano-preputial epithelial cells permitting the prepuce to retract along the length of the glans penis. The age at PPS is useful to identify antiandrogens which can delay puberty through altered development of the balano-preputial membrane (permanent incomplete PPS) or interfere with the normal rise in circulating androgens. Malformations such as incomplete PPS or hypospadias, which prevent accurate PPS data collection, are reported as a malformation when the adults are necropsied. Occasionally, males display a transient thread of tissue along the frenulum of the glans, which persists for several days or, on rare occasions, is permanent. For this reason, the age and weight at the onset of PPS should be noted as well as the completion of PPS, with the dissolution of the thread, if present. Also male rats with hypospadias may display a permanent thread on the glans or they are so malformed that PPS does not occur at all. If time does not permit examining a large number of rats daily for VO or PPS, then the rats can be examined two or three times per week and the age of VO or PPS and body weight at puberty recorded. These data can then be analyzed and reported as the percentage of females or males which opened or separated on a given day for each treatment group. Materials Rats from litters of dams in treatment groups, approaching puberty Balance for animal weight recorded to the nearest 0.01 g (e.g., Sartorius IP65) with integration capability to accurately weigh moving animals Detect vaginal opening (VO) 1a. Check female rats daily for VO beginning at about PND 24, ∼5 days prior to the onset of this landmark to ensure the day of VO is detected in all rats. Monitor each rat daily from then until the process is complete. VO normally occurs in Sprague-Dawley and Long Evans rats between 29 and 35 days old (DOB = PND 1). The index finger may be gently rubbed across the vaginal area to determine if the vaginal membrane is present or if the layer of dying cells are simply obscuring the vaginal opening.

2a. When VO is initiated and when complete, record the ages and body weights of each rat and calculate the mean ages and body weights for each treatment group, then check for statistical significance. Detect preputial separation (PPS) 1b. Check male rats daily beginning at 35 to 37 days of age to ensure no rats have preputial separation. Apply gentle pressure to the prepuce to retract the prepuce and expose the glans penis. PPS normally occurs in Sprague-Dawley and Long Evans rats between 39 and 45 days old (DOB = PND 1). PPS is complete when the entire perimeter of the prepuce can be retracted evenly around the base of the glans penis. Effects of Endocrine Disrupting Compounds (EDCs) in Rats

2b. When PPS begins and when it is complete, record the ages and body weights of each rat and calculate the mean age, body weight, and standard errors by treatment group, and check for statistical significance.



NECROPSY At 90 to 120 days of age, male and female Sprague-Dawley rats have reached sexual and physical maturity, therefore, this time frame represents the ideal time to evaluate their reproductive systems. Although the reproductive system of both males and females should be thoroughly evaluated when administering test compounds for the first time, the primary emphasis of each necropsy can be designed to focus on endpoints known or expected to be affected based upon previous in vitro/in vivo studies or on the results of data collected earlier in this study.

BASIC PROTOCOL 5

Materials Male and female rats, 90 to 120 days old CO2 Bouin’s fixative (see recipe) Balance accurate to 0.1 g with integration capability for weighing rats (e.g., Sartorius IP65) Rodent restraint cones (Harvard Apparatus) Rodent guillotine Vernier caliper (0 to 150 mm with 0.1-mm divisions) Small animal clippers with size # 40 clipper blade (Oster) for shaving ventral surface to check areolae/nipples Surgical instruments (Roboz): micro dissecting forceps (RS-5230 for grasping delicate tissues such as ventral prostate and RS-5236 for holding tougher tissue such as skin), micro dissecting scissors (RS-5852 for delicate dissections and RS-6814 for cutting tougher tissue such as skin) and hemostats (RS-7172 for clamping seminal vesicles) Balance accurate to 0.0001 g for weighing tissues (e.g., Sartorius BP 121 S) Perform male necropsy The order of tissue dissection listed below is designed to minimize tissue dehydration and should facilitate accurate dissection in a timely manner. 1a. Weigh male rats, place in a restraint cone, and decapitate within 60 sec of removal from their cage. Collect the trunk blood for serum hormone analysis by radioimmunoassay. 2a. For external examination, shave the rat and examine for the presence of areolae/nipples and malformations of the external genitalia such as cleft prepuce, incomplete preputial separation, hypospadias (urethra opens along the underside of phallus rather than at the tip), or vaginal pouch. Measure anogenital distance at this time with a Vernier caliper, especially if it was affected at birth. Weigh the glans penis. Note if there are undescended, ectopic testes. 3a. Begin internal examination. Check for bladder stones (antiandrogens such as vinclozolin or procymidone increase the incidence of bladder stones), hydroureter, and obvious hydronephrosis resulting from back pressure on the kidneys. Check for undescended ectopic testes, which could be attached just below the kidneys to abdominal muscle wall by a cranial gonadal suspensory ligament or testes embedded in the abdominal muscle wall (ectopic). In the process of dissecting and weighing the tissues listed below, note any additional malformations. 4a. Dissect and weigh the ventral prostate; dissect (weighing is optional) the dorsolateral prostate from around the base of the seminal vesicles; and remove and weigh the seminal vesicles and coagulating glands after clamping the base of the seminal vesicles with hemostats to avoid leakage of seminal fluid. Note abnormalities such as infection, unusual coloration/texture, tissue adhesions, or missing or poorly



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developed lobes of the seminal vesicles, coagulating glands, ventral or dorsolateral prostate and photograph if time permits. The seminal vesicles can also be weighed dry after the seminal fluid is expressed and the tissue blotted dry.

5a. Withdraw the testes and epididymides from the scrotum and examine the gubernacular cords to ensure they are present and not elongated or filamentous (typically 4 to 8 mm in length), and dissect and weigh the testes and epididymides (see Fig. 16.8.3 and Fig. 16.8.4). Note any testicular/epididymal abnormalities such as undescended (also check for the presence of a cranial gonadal suspensory ligament) or ectopic testes (testis located in abdominal muscle wall), fluid-filled or hypoplastic or atrophic testes and epididymal hypoplasia or agenesis. The testes can be saved for sonication resistant testicular sperm head counts or histology or both and the epididymides can be saved for whole, caudal, and caput plus corpus epididymal sperm counts or fixed for histology.

6a. Remove and weigh the Cowper’s glands, and dissect and weigh the levator ani/bulbocavernosus (LABC), which loops around the colon and is attached at each end to the penile bulbs. 7a. Dissect and weigh nonreproductive tissues, which may be affected by EDCs. Include the pituitary, kidneys, adrenals, and liver. Weigh the brain and other tissues at this time. It may also be helpful to slice the kidneys longitudinally to check for hydronephrosis and kidney stones.


Figure 16.8.3 Testes and epididymides from control and in utero linuron-treated male rat offspring, necropsied as adults. Photographs in panels on the left show a control testis, epididymis and epididymal fat pad and the epididymis dissected from the testis with the fat removed. Panels on the right show a testis, epididymis and epididymal fat pad and the epididymis dissected from the testis with the fat removed, from a rat exposed in utero from gestational day 14 through 18 to 100 mg linuron/kg. Agenesis of the caput produces fluid pressure atrophy of the testis. This black and white facsimile is intended only as a placeholder; for full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm



8a. Save all tissues weighed, especially tissues with gross malformations, and fix in Bouin’s fixative for histological evaluation or process for other endpoints such as immunohistochemical staining. Perform female necropsy 1b. Weigh females, anesthetize in CO2, decapitate, and save the trunk blood for serum hormone analysis by radioimmunoassay. 2b. For external examination, shave the female rats and check for the absence/prominence of areolae/nipples (DHT inhibits development of nipples) and malformations of the external genitalia such as cleft phallus, vaginal threads, and absence of external vaginal opening (DHT inhibits development of lower vagina). Measure anogenital (AGD) and anovaginal distance (AVD) to calculate vaginal-genital distance (AGDAVD), which decreases with exposure to androgens. 3b. Begin internal examination. Since androgens inhibit the development of the ovarian ligament, measure the kidney to ovary distance when females are exposed perinatally to suspected androgens. Androgens may also promote the development of male reproductive tissues such as prostatic tissue, seminal vesicles, penile bulbs with attached levator ani muscle, and Cowper’s glands; note the presence of these tissues and photograph, if possible, and save for histological confirmation. 4b. Dissect and weigh the ovaries, and weigh the uterus with and without fluid (remove fluid by cutting each uterine horn open and blotting all fluid). 5b. Dissect and weigh the nonreproductive tissues—the pituitary, adrenals, kidneys (section each kidney to check for hydronephrosis), and liver. 6b. Save all tissues weighed, especially tissues with gross malformations, and fix in Bouin’s fixative for histological evaluation or process for other endpoints such as immunohistochemical staining.

Figure 16.8.4 Gubernacular ligaments in control and phthalate ester–treated male rat offspring, necropsied as adults. Photographs showing a control rat with normal gubernacular cords (left panel) and a rat whose mother received received 750 mg diethylhexyl phthalate/kg from gestational day 14 through postnatal day 3 with a poorly developed gubernacular cord (right panel). Diethylhexyl, dibutyl, and benzylbutyl phthalate produced partial or complete agenesis of the gubernacular cord resulting in undescended testes in some rats, while other rats were observed with scrotal testes but absent or poorly developed, thin and elongated gubernacular cords. This black and white facsimile is intended only as a placeholder; for full-color version of figure go to http://www.interscience.wiley.com/c_p/colorfigures.htm



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SUPPORT PROTOCOL

DATA INTERPRETATION AND SUMMARY Analyze AGD, areolar, nipple, pubertal landmark (VO or PPS), and organ weight data using litter means or a nested analysis that accounts for litter effects. If an overall ANOVA model analysis is significant, then use posthoc t-tests to compare dose groups to one another and the control group. Alternatively, use a one-tailed Dunnett’s test to compare each treated group to the control group. As these developmental reproductive toxicants (EDCs) produce characteristic syndromes, each containing a variety of effects, the data should be compiled in a manner that enables the investigator to determine what percentage of the offspring are normal and what percentage display any abnormality associated with the particular syndrome. For example, phthalate ester exposure in utero can induce any number of epididymal, testicular, gubernacular, internal sex organ, or external affects. Any animal displaying a morphological or histological alteration in any one of these tissues should be considered affected. If one of ten animals displayed epididymal agenesis, a second had a histological lesion of the epididymis, a third had testis agenesis, a fourth had seminiferous tubular agenesis, a fifth had agenesis of the gubernacular ligament, a sixth had agenesis of the ventral prostate, a seventh displayed hypoplasia of the right horn of the seminal vesicle, and an eighth had six permanent nipples, then it should be reported that 80% of the animals are adversely affected in this group. See Gray et al. 1994, 1997a or b, 2000; Lambright et al., 2000, and McIntyre et al. 2002 for examples of data presentation. REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.

Bouin’s fixative 3 liters saturated picric acid solution (Sigma) 1 liter 37% formaldehyde solution (Sigma) 200 ml glacial acetic acid (Sigma) Store at room temperature in a fume hood; replace if a precipitate forms or at 4 months CAUTION: Upon drying, the picric acid solution will form a crystal residue that will ignite if sufficient heat or friction is applied. Any residue that forms on storage containers may be safely removed with a damp sponge.



Identification and comparison of the potency of endocrine disrupting compounds (EDCs) to natural ligands may be best accomplished using rapid in vitro screening assays such as receptor binding and transcriptional activation assays or short-term in vivo assays such as the Hershberger and uterotropic assay (Gray et al., 2002). In the authors’ laboratory, the transgenerational protocol described has been used primarily to quantify and characterize the demasculinizing effects of antiandrogens in males or the masculinizing effects of androgens in females for compounds identified in these screening assays. Additionally, transgenerational exposure to EDCs such as androgen receptor (AR) antagonists or fetal testos-

terone synthesis inhibitors display unique profiles of malformations characteristic of their specific mechanism(s) of action (Gray et al., 1994, 1997a,b, 2000; Lambright et al. 2000; McIntyre et al., 2002). Accurate oral or subcutaneous exposure is accomplished by administering the test compound on a milligram or microgram/kilogram body weight/milliliter vehicle/day basis and adjusting the dose daily according to body weight, which increases dramatically during the latter stages of gestation. Daily weighing also serves as a sensitive indicator of toxicity and consequently is monitored daily both on an individual basis and analyzed on a treatment group basis.



The sexually dimorphic androgen-sensitive perineal area between the anus and sex papilla (anogenital distance—AGD) provides a useful biomarker for quantitative evaluation of the fetal androgen status of a rat. Immunohistochemical staining demonstrates that androgen receptor expression in mesenchymal cells in the perineal and prepubic area first occurs on GD 19 (copulatory vaginal plug = GD 1) in male rats (Bentvelsen, 1995). AGD can be thought of as a phenotypic continuum between females (∼1.5 mm) and males (∼3 mm) in which developmental exposure of females to androgens masculinizes or increases female AGD, while developmental exposure of males to antiandrogens, feminizes or decreases male AGD. AGD at birth or PND 2 has been shown to correlate significantly with adult male urogenital malformations when the data analyzed are litter means. With the aid of a dissecting microscope fitted with an ocular reticle, neonatal AGD can be accurately measured to 0.1 mm and, if effected, this measurement should be repeated in the adult necropsy with the aid of vernier calipers. Androgen receptor expression in mesenchymal cells of the nipple anlage is first demonstrated by immunohistochemical staining on GD 15 (vaginal plug = GD 1) in male rats and th e presence of androgens (dihydrotestosterone) at this time promotes regression of the nipple anlage in both sexes (Bentvelsen, 1995). Consequently, recording the number of areolae/nipples at 13 days of age provides another sensitive endpoint for evaluating endocrine status of males potentially exposed to antiandrogens or females exposed to androgens during development. If the normal areolae/nipple phenotype is altered at this age, then at necropsy, the adults should be shaved, examined for the presence of areolae/nipples, and the tissues saved for histological evaluation. Vaginal opening (VO) and preputial separation (PPS), which coincide with the rise in circulating estrogens or androgens, serve as indices of puberty in female and male rats, respectively. VO and PPS can be accelerated or delayed by hormonal manipulation and consequently provide useful endpoints for evaluating the hormonal status of animals exposed to potential EDCs (Korenbrot et al., 1977: Ojeda et al., 1976). For example, perinatal oral exposure to 200 mg of the AR antagonist, vinclozolin, from GD 14 through PND 3 (sperm + = GD 1) delayed preputial separation by 3 days (Gray et al., 1994), a single oral dose of 0.2 and 0.8 µg o f 2 ,3 ,7 ,8 -tetrachlorodibenzo-p-dioxin

(TCDD) administered on GD 16 (sperm + = GD 1) delayed preputial separation by 1.5 and 3 days, respectively (Gray et al., 1997a), while 0.8 µg of 2,3,7,8 TCDD delayed vaginal opening by 1 day (Gray et al., 1997b). At necropsy, sexually mature adult rats exposed perinatally to EDCs are examined externally to evaluate development of androgen-dependent tissues such as the perineal area and genital tubercle and tissues whose development are inhibited by androgens such as areolae/nipples. Permanent malformations of external tissues stimulated or inhibited developmentally by androgens in males include cleft prepuce/phallus, hypospadias, incomplete preputial separation, vaginal pouch, reduced glans penis weights, decreased anogenital distance, and nipples (Clark et al., 1990, 1993; ImperatoMcGinley et al., 1992). Externally in females, androgens increase AGD, decrease the distance between the vagina and phallus, inhibit normal areolae/nipple development and prevent development of the lower vagina (Wolf et al., 2002). EDCs such as 2,3,7,8 TCDD decrease the distance between the urethral and vaginal openings, produce permanent vaginal threads (Gray et al., 1997b) and reduce glans penis weights (Gray et al., 1997a). Internally, androgen-dependent tissues are examined for abnormal development, their weights are recorded and the tissues may be saved for histological evaluation. In male rats, these tissues should include the ventral prostate, seminal vesicles plus coagulating glands, testes, epididymides, and the levator ani/bulbocavernosus muscle complex. Other tissues that have been affected by EDCs in previous experiments and may be weighed, measured, or examined and saved for histology include the bulbourethral glands, gubernacular ligament, liver, kidneys, adrenals, dorsolateral prostate, bladder, and ureters (Gray et al., 1999; 2000). Additionally, malformations such as undescended or ectopic testis or the presence of cranial gonadal suspensory ligaments should be noted. EDCs operating through different mechanisms of action produce different malformation profiles. AR antagonists such as vinclozolin target DHT-dependent tissue development including the ventral prostate and external genitalia and produce malformations such as permanent nipples, urinary bladder stones, ectopic testes, hypospadias, and vaginal pouches (Gray et al., 1994). In contrast, some EDCs, which inhibit fetal Leydig cell steroid and peptide hormone synthesis such as diethylhexyl phtha-



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late (Parks et al., 2000), also induce malformations such as epididymal hypoplasia/agenesis and subsequent fluid pressure atrophy of the testes and undescended testes as a consequence of partial or complete agenesis of the gubernacular ligament (Gray et al., 2000; Kubota et al., 2001). Other EDCs such as linuron (Lambright et al., 2000) and prochloraz may alter reproductive development through several mechanisms of action (AR antagonism and steroid inhibition) and produce a profile of malformations intermediate between an AR antagonist and steroidogenesis inhibitor.



Ideally, test compound dosages should be selected to cover the full range of the dose-response curve with the lowest dose producing no or only minimal effects and the highest dose eliciting slightly less than maximal response without producing nonspecific, systemic maternal or neonatal toxicity. Generally, the highest dosage level should not induce a reduction in maternal weight >10%. Generally, litters are not culled and data collected on all offspring for each specific endpoint are used to provide the most accurate representation of the dose-response curve. If one is interested in subtle treatment-related changes in organ weights and somatic growth, the litters could be standardized to a litter size of 8 to 10 pups per litter. Because most data are analyzed as litter means (n = number of litters/treatment group), the number of litters/treatment group must be of sufficient size to ensure adequate statistical power. A typical study might consist of five treatment groups (a control and four different treatment groups) with eight dams/treatment group; this would potentially generate 500 offspring (100/treatment group). Data for protocols in which data can be collected in 1 day, such as anogenital distance or nipple development at 13 days of age, should be collected one block at a time to minimize the effect of time on treatment and the data should be collected by only one observer who is blind to treatment. Likewise, data collection such as necropsy, which requires >1 day to complete, can be collected on one animal/litter for each treatment group in one block, again to minimize the effect of time/age on treatment and each technician should conduct the same tasks throughout the entire necropsy in order to reduce technician-to-technician variability for each specific endpoint.

Considerable attention should be given to assuring that the individuals involved are all adequately trained and follow similar procedures. If data on an endpoint like AGD or areolas must be collected by more than one technician, then the study director must ensure that the inter-observer reliability is high from training exercises; he also must avoid having one technician collect the data from the control group, while another collects the data from a treated group. All treatment/dose groups in a study should be run concurrently. If the whole experiment cannot be run at once, then it should be split into balanced complete block designs. While these comments might seem obvious, it has been known that investigators have had one technician collect the data on all the controls 1 month, while others collected the data on the treated groups at other times.

Anticipated Results Perinatal exposure to EDCs such as antiandrogens or steroidogenic inhibitors may demasculinize males and reduce anogenital distance, induce female-like nipple development, delay preputial separation and inhibit normal development of the testes, epididymis, and sex accessory tissues such as the glans penis, ventral prostate, seminal vesicles/coagulating glands, bulbourethral glands, and levator ani/bulbocavernosus muscles in a dose-related fashion (Table 16.8.1). In contrast, perinatal exposure to androgens can masculinize and defeminize the female offpsring resulting in increased anogenital distance, agenesis of areolae and nipples, agenesis of the lower vagina, absence of vaginal opening, hydrometrocolpus, and retained male tissues including the ventral prostate, seminal vesicles, and levator ani.

Time Considerations Data collection for some endpoints such as anogenital distance at 2 days of age and areolae/nipple retention at 13 days of age, ideally, should be collected in 1 day by only one observer, while other protocols such as necropsy may be conducted over a considerably longer period such as 1 week or several weeks. Consequently the sample size (n = number of litters) should permit sufficient statistical power without jeopardizing data collection on all offspring at one time point. A study with five treatment groups and 8 to 10 dams/treatment or 100 to 120 offspring/treatment group should satisfy these criteria.



Table 16.8.1

Effects of Perinatal Vinclozolin on Male Offspringa

Observation

Percent affectedb

Dose (mg/kg/day) AGD reductionc Permanent nipples Areolae/nipples in infantsd Ventral protstate gland size reductione Hypospadiasise Vaginal pouche Ectopic testese

0 (mean) 0 (3.43 mm) 0 5 0 (564 mg)

3 7 1 17 11

6.25 4.2 2.6 33 26

12.5 9.1 3.6 55 22

25 14.7 5.4 49 32

50 24.9 50 100 46

100 35 100 100 88

0 0 0

0 0 0

0 0 0

0 0 0

0 0 0

45 15 0

100 100 100 100 20 50

200 50 100 100 95

aCombined data from Gray et al., 1994 and 1997. bPerinatal oral maternal treatment with vinclozolin. cAnogenital distance (AGD) was measured at PND 2. dAreolae were observed at PND 13. eMales were necropsied at full maturity as adults and organs were weighed and examined for structural malformations.

Disclaimer The research described in this unit has been reviewed by the National Health and Ecological Effects Research Laboratory, U.S. Environmental Protection Agency, and approved for publication. Approval does not signify that the contents necessarily reflect the views and policies of the Agency nor does mention of trade names or commercial products constitute endorsement or recommendation for use.

Literature Cited Bentvelsen, F.M., Brinkmann, A.O., van der Schoot, P., van der Linden, J.E., van der Kwast, T.H., Boersma, W.J., Schroder, F.H., and Nijman, J.M. 1995. Developmental pattern and regulation by androgens of androgen receptor expression in the urogential tract of the rat. Mol. Cell Endocrinol. 113:245-253 Clark, R.L., Antonello, J.M., Grossman, S.J., Wise, L.D., Anderson, C., Bagdon, W.J., Prahalada, S., MacDonald, J.S., and Robertson, R.T. 1990. External genitalia abnormalities in male rats exposed in utero to finasteride, a 5 alpha-reductase inhibitor. Teratology 42:91-100. Clark, R.L., Anderson, C.A., Prahalada, S., Robertson, R.T., Lochry, E.A., Leonard, Y.M., Stevens, J.L., and Hoberman, A.M. 1993. Critical developmental periods for effects on male rat genitalia induced by finasteride, a 5 alpha–reductase inhibitor. Toxicol. Appl. Pharmacol. 119:34-40. Gray, L.E. Jr., Ostby, J.S., and Kelce, W.R. 1994. Developmental effects of an environmental antiandrogen: The fungicide vinclozolin alters sex differentiation of the male rat. Toxicol. Appl. Pharmacol. 129:46-52.

Gray, L.E. Jr., Ostby, J.S., and Kelce, W.R. 1997a. A dose-response analysis of the reproductive effects of a single gestational dose of 2,3,7,8tetreachlorodibenzo-p-dioxin in male Long Evans hooded rat offspring. Toxicol. Appl. Pharmacol. 146:11-20. Gray, L.E. Jr., Ostby, J.S., and Kelce, W.R. 1997b. In utero exposure to low doses of 2,3,7,8tetreachlorodibenzo-p-dioxin alters reproductive development of female Long Evans hooded rat offsp ring. Toxicol. Appl. Pharmacol. 146:237-244. Gray, L.E. Jr., Wolf, C., Lambright, C., Mann, P., Price, M., Cooper, R.L., and Ostby, J. 1999. Administration of potentially antiandrogenic pesticides (procymidone, linuron, iprodione, chlozolinate, p,p′-DDE, and ketoconazole) and toxic substances (dibutyl- and diethylhexyl phthalate, PCB 169, and ethane dimethane sulphonate) during sexual differentiation produces diverse profiles of reproductive malformations in the male rat. Toxicol. Ind. Health 15:94-118. Gray, L.E. Jr., Ostby, J., Furr, J., Price, M., Veeramachaneni, D.N., and Parks, L. 2000. Perinatal exposure to the phthalates DEHP, BBP, and DINP, but not DEP, DMP, or DOTP, alters sexual differentiation of the male rat. Toxicol. Sci. 58:350-365. Gray, L.E. Jr., Ostby, J., Wilson, V., Lambright, C., Bobseine, K., Hartig, P., Hotchkiss, A., Wolf, C., Furr, J., Price, M., Parks, L., Cooper, R.L., Stoker, T.E., Laws, S.C., Degitz, S.J., Jensen, K.M., Kahl, M.D., Korte, J.J., Makynen, E.A., Tietge, J.E., and Ankley, G.T. 2002. Xenoendocrine disrupters_tiered screening and testing: Filling key data gaps. Toxicology 182:371-382. Imperato-McGinley, J., Sanchez, R.S., Spencer, J.R., Yee, B., and Vaughan, E.D. 1992. Comparison of the effects of the 5 alpha-reductase inhibitor finasteride and the antiandrogen flutamide on prostate and genital differentiation: Dose-response studies. Endocrinology 131:1149-1156.



Supplement 19

Korenbrot, C.C., Huhtaniemi, I.T., and Weiner, R.I. 1977. Preputial separation as an external sign of pubertal development in the male rat. Biol. Reprod. 17:298-303. Kubota, Y., Nef, S., Farmer, P.J., Temelcos, C., Parada, L.F., and Hutson, J.M. 2001. Leydig insulin-like hormone, gubernacular development and testicular descent. J. Urol. 165:16731675. Lambright, C., Ostby, J., Bobseine, K., Wilson, V., Hotchkiss, A.K., Mann, P.C., and Gray, L.E. Jr. 2000. Cellular and molecular mechanisms of action of linuron: An antiandrogenic herbicide that produces reproductive malformations in male rats. Toxicol. Sci. 56:389-399. McIntyre, B.S., Barlow, N.J., and Foster, P.M. 2002. Male rats exposed to linuron in utero exhibit permanent changes in anogenital distance, nipple retention, and epididymal malformations that result in subsequent testicular atrophy. Toxicol. Sci. 65:62-70.

Ojeda, S.R., Wheaton, J.E., Jameson, H.E., and McCann, S.M. 1976. The onset of puberty in the female rat: Changes in plasma prolactin, gonadotropins, luteinizing hormone-releasing hormone (LHRH), and hypothalamic LHRH content. Endocrinology 98:630-638. Parks, L.G., Ostby, J.S., Lambright, C.R., Abbott, B.D., Klinefelter, G.R., Barlow, N.J., and Gray, L.E. Jr. 2000. The plasticizer diethylhexyl phthalate induces malformations by decreasing fetal testosterone synthesis during sexual differentiation in the male rat. Toxicol. Sci. 58:339-349. Wolf, C.J., Hotchkiss, A., Ostby, J.S., LeBlanc, G.A., and Gray, L.E. Jr. 2002. Effects of prenatal testosterone propionate on the sexual development of male and female rats: A dose-response study. Toxicol. Sci. 65:71-86.

Contributed by Joseph S. Ostby and L. Earl Gray Jr. United States Environmental Protection Agency Research Triangle Park, North Carolina




CHAPTER 17 Oxidative Stress INTRODUCTION any chemical-induced toxic insults involve oxidative events as major components. Yet, we know that certain oxidative events are essential components of complex signaling processes in cell growth and regulation. Although natural antioxidants have essential roles in optima cellular functions and health, proper balance in both oxidative events and antioxidart functions remains to be fully elucidated. In this chapter, efforts are directed towards understanding and analyzing the oxidative events that appear crucial to toxicology. The formation and functions of protein sulfenic acids that are formed by oxidative processes are covered in an overview (UNIT 17.1). Sulfenic acids in proteins are formed upon reaction of cysteinyl thiols with oxidizing agents including mild oxidizing agents such as molecular oxygen and hydrogen peroxide. Topics covered in this overview include peroxide sensitivity, detection of nitrosative stress, participation of protein sulfenic acids in redox catalysis by antioxidant enzymes, in other catalytic functions and in transcriptional regulators. Examples of the role of protein sulfenic acids in catalysis and regulation are given including redox sensing mechanisms.

M

The measurement of protein sulfenic acid content is presented in UNIT 17.2. Notoriously difficult to identify due to their high reactivity, particularly outside the native protein environment, protein sulfenic acids are measured with techniques that are based upon this high degree of reactivity. Chemical modification methods are useful for sulfenic acid identification. Sulfenic acid can be trapped and detected using 7-chloro-4-nitrobenzo-2oxa-1,3-diazole (NBD chloride), which reacts with both thiol groups and sulfenic acids in proteins but gives unique products that can be detected by their UV-visible spectra and mass spectrometry. An alternative utilizes a more reactive and less stable reagent 4-fluoro7-nitrobenz-2-oxa-1,3-diazole (NBD fluoride). Another reagent, 2-nitro-5-thiobenzoic acid (TNB), reacts stoichiometrically with sulfenic acids for quantification. A fourth reagent, 5,5-dimethyl-1,3-cyclohexanedione (dimedone) reacts specifically with sulfenic acids but not thiol groups on proteins and relies on ESI-MS analysis for identification. Oxidative stress and lipid peroxidation are implicated in chemical-induced cell injury/ death. UNIT 17.3 describes the adaptation of a fluorescence microplate reader for measuring tissue susceptibility to lipid peroxidation. Lipid peroxidation is initiated by Fe3+ -ADP in the presence ascorbate yielding oxidative products, including 4-hydroxy-2-nonenal, which form fluorescent products that are measured with a fluorescence microplate reader. Real-time measurement of a large number of samples for their susceptibility to lipid peroxidation is possible with this procedure. Oxidative stress is a part of many disease states and may result from the accumulation of metals by tissues. Transition metals, including iron and copper, are widely studied for the many types of associated oxidative reactions. UNIT 17.4 describes methods for in situ localization of nonenzymatic peroxidase-like activity of tissue-bound transition metals. A significant improvement over existing methods is an enhanced sensitivity based on the application of hydrogen peroxide and diaminobenzidine (DAB) for the detection of iron. An alternate protocol for the detection of redox-active transition metals is given

Oxidative Stress Contributed by Donald J. Reed Current Protocols in Toxicology (2005) 17.0.1-17.0.2 C 2005 by John Wiley & Sons, Inc. Copyright


as is a support procedure that utilizes selective chelating agents to evaluate the relative contributions of iron and copper to the redox activity. An important measure of oxidant stress is the quantification of lipid peroxidation products. UNIT 17.5 is an overview of the formation of lipid peroxidation products with emphasis on those products formed from the oxidation of ubiquitous arachidonic acid. This overview includes a discussion of the biochemistry of F2-isoprostanes as well as their utility as markers of oxidant stress. Mechanisms of formation of isoprostanes, their quantification in vitro, in vivo, and in animal models of oxidant stress are described. A major topic is the formation of F2-Isoprostanes in human diseases including atherosclerotic cardiovascular disease and Alzheimer’s disease. The quantification of F2-isoprostanes in tissue lipids by gas chromatography/mass spectrometry as a measure of oxidant stress is the main protocol in UNIT 17.6. An alternative protocol is the extraction of lipids from phospholipid-containing biological fluids. The protocol for quantification F2-isoprostanes by GC/negative ionization MS is extremely sensitive with a lower limit of detection in the range of 1 to 5 picograms. An example is given for the analysis of F2-isoprostanes in plasma obtained from a rat 4 hour after treatment with carbon tetrachloride. A discussion of critical parameters and troubleshooting along with anticipated results are included in this protocol. Oxidant stress leading to biological oxidative damage includes the formation of proteincentered radicals. UNIT 17.7 describes a new technology that utilizes several protocols of immuno-spin trapping for the detection of protein-centered radicals. The spin trap being utilized is 5,5-dimethyl-1-pyrroline N-oxide (DMPO which, under proper conditions, can form stable DMPO-protein radical-derived nitrone adducts that are detected and identified by immuno-spin trapping with an antiserum against DMPO. A basic protocol describes the procedure for the production and detection of hemoprotein-centered radicals by immunospin trapping ELISA. A second basic describes the preparation and immunoblot analysis of DMPO-protein radical-derived nitrone adducts. Because UNIT 17.7 focuses on a new technology, a detailed discussion of critical parameters and troubleshooting along with anticipated results are included in this protocol. Donald J. Reed

Introduction



Formation and Functions of Protein Sulfenic Acids Sulfenic acids (R-SOH) are the simplest oxy-acids of organic sulfur and, unlike with higher sulfur oxidation states, sulfinic (RSO2H) and sulfonic (R-SO3H) acids, they are inherently unstable and highly reactive (Hogg, 1990). Considerable insight has been gained through studies of small molecules wherein the sulfenic acid moieties have been stabilized by steric, hydrogen-bonding, and/or electronic factors. Recent progress in the identification and analysis of cysteine-sulfenic acids formed within proteins has also added to the understanding of the forces that contribute to their stabilization (Claiborne et al., 2001). Cysteine sulfenic acids, as reversibly oxidized products of peroxide, peroxynitrite, or NO-mediated oxidation of cysteinyl thiols, are outstanding candidates as players in redox regulation, an area of intense investigation in recent years (Poole et al., 2004). Indeed, protein sulfenic acids are involved in multiple ways in oxidative and nitrosative stress as (1) redox centers or catalytic intermediates in enzymes, (2) reversibly oxidized sensors of peroxide or NO levels in regulatory proteins, and (3) intermediates in the irreversible damage to cysteinyl residues caused by overoxidation of these amino acids. Sulfenic acids in proteins have been notoriously difficult to identify due to their reactivity, particularly outside of their native protein environment. Early evidence supporting the formation and stabilization of sulfenic acids in two proteins, glyceraldehyde-3-phosphate dehydrogenase and papain, relied only on indirect methods for their identification (Allison, 1976). Over the last decade, the growing number of proteins known to form sulfenic acids of functional significance and the improvement in definitive identification via structural and chemical techniques have greatly enhanced recognition of their importance in biological systems. Hopefully, with the tools now available, the widespread biological significance of these cysteine modifications will be further clarified.

FORMATION, REACTIVITY, AND DETECTION OF CYSTEINE SULFENIC ACIDS IN PROTEINS Sulfenic acids in proteins are formed upon reaction of cysteinyl thiols with mild oxidizing agents. Even molecular oxygen can act as an Contributed by Leslie B. Poole Current Protocols in Toxicology (2003) 17.1.1-17.1.15 Copyright © 2003 by John Wiley & Sons, Inc.

UNIT 17.1

oxidant of protein thiols (termed autooxidation), although the rate of this reaction is insignificant in the absence of catalytic quantities of metals (particularly iron and copper; Torchinsky, 1981). Oxidizing agents of greatest biological significance are peroxides, including hydrogen peroxide, organic hydroperoxides, and peroxynitrite, and nitric oxide and/or its derivatives. All of these agents participate in cell signaling and have been shown to convert protein thiols to sulfenic acids. The reactivity of particular cysteinyl thiols toward these molecules is controlled by their respective protein environments. Important factors include accessibility of the thiol toward these agents and the proximity of other important amino acid side chains, which may interact with the thiol group or with the oxidant or its product. Most reactive are cysteine residues with relatively low pKa values (below the typical value of ~8.5 for an unperturbed cysteine thiol), which are stabilized as the nucleophilic thiolate anion at neutral pH. Proximity to basic amino acids that stabilize the negative charge on the thiolate therefore promotes the oxidation of given cysteine residues.

Peroxide Sensitivity Other factors are clearly involved, as well. Documented reactivities of thiolate anions with hydrogen peroxide, for example, range from ∼20 M−1 sec−1 for protein tyrosine phosphatases to 105 to 106 M−1 sec−1 for cysteinedependent peroxidases, such as peroxiredoxins and enterococcal NADH peroxidase (Poole et al., 2004). OxyR, a bacterial transcriptional regulator with exquisite sensitivity toward hydrogen peroxide, also exhibits peroxide reactivity within this range, at ∼2 × 105 M−1 sec−1 (Åslund et al., 1999). A likely important active site feature in these peroxide-sensitive sites is the proximity of a general acid catalyst to protonate the poor RO− leaving group of the hydroperoxides, although no such catalyst has as yet been identified in these proteins. Although some of these features in proteins are coming to light, understanding the molecular basis for redox sensitivity toward oxidants of given cysteinyl residues is still at an early stage. Another mechanism for enhancement of cysteine reactivity toward peroxides, coordination of the cysteine sulfur to a metal, has re-

Oxidative Stress


(1)

R-SOH + HNO

R-S-N=O + H2O

H O R S

(2)

2 R-SOH

(3)

R-SH

(4)

R-SOH + R′-SH

[O]

R-SOH

[O]

H O S R

R-SO2H

O R S S R + H2O [O]

R-SO3H

R-S-S-R′ +

H2O

O

t - Bu-S

(5)

t - Bu-SOH + HC

C

CO2CH3

H C C

H O

O

CH3

(6)

CH3

R-SOH +

+ H2O

R-S

CH3

CH3

O

O COO−

(7)

R-SOH + −S

CO2CH3

NO2 + H+

COO− R-S-S

NO2 + H2O

Figure 17.1.1 (continues on right) Chemical reactions relevant to the redox chemistry of sulfenic acids.

Formation and Functions of Protein Sulfenic Acids

cently been characterized as well. In one heat shock protein from Escherichia coli, Hsp33 (Graumann et al., 2001), cysteinyl residues coordinated to zinc were shown to react with hydrogen peroxide, resulting in disulfide-bond formation. Furthermore, release of the bound metal concomitant with disulfide bond formation is an essential part of the activation of this protein. This mechanism likely also applies to the iron-coordinated cysteinyl residues within the peroxide-sensitive transcriptional repressor PerR from Bacillus subtilis (Mongkolsuk and Helmann, 2002). Although cysteine sulfenic acid formation was not demonstrated in these cases, its formation as an intermediate leading to disulfide bond formation is a plausible hypothesis. Such metal-enhanced reactivity of cysteine thiols may also be relevant to the sensitivity of Fe-S clusters toward oxidative damage (e.g., in aconitase and fumarase; Costa Seaver and Imlay, 2001). The release of the Fe(II) under conditions where H2O2 is already present undoubtedly exacerbates the deleterious effects of Fe-S center breakdown. Such cysteine-coordinated metal centers can thus

serve as peroxide sensors, as well (Mongkolsuk and Helmann, 2002).

Nitrosative Stress and Sulfenic Acids Several important players in nitrosative stress, including nitric oxide, S-nitrosoglutathione, and the product of nitric oxide and superoxide, peroxynitrite (OONO−), deserve special attention as modifiers of cysteine thiols. Although the pathways through which nitric oxide modifies cysteinyl residues are complex, NO and its derivatives have been shown to modify free cysteine residues to give the S-nitrosothiol (R-SNO) and/or sulfenic acid (RSOH) products or, in the presence of additional thiols, mixed disulfide bonds (Stamler and Hausladen, 1998; Marnett et al., 2003). Interestingly, when human glutathione reductase was inactivated with S-nitrosoglutathione (GSNO), the active-site cysteine residue proximal to the flavin (Cys63) was oxidized to a stable sulfenic acid and the interchange cysteine (Cys58) was linked through a mixed disulfide bond to glutathione (Becker et al., 1998). Oxidation of protein thiol groups to sulfenic,



R

Cl

S N

N

(8)

R-SH +

O + H+ + Cl−

O N

N NO2

NO2

R

Cl

S

O N

N

(9)

R-SOH +

O

O

+ H+ + Cl−

N

N NO2

NO2

+ H+

(10)

H2O2 + NADH

(11)

O2 + 2 NADH + 2 H +

(12)

ROOH + XH 2

(13)

Met S

2 H2O + NAD+

2 H2O + 2 NAD+

ROH + H2O + X

O + Trx (SH2)

Met S

CH3

+

H2O

+ Trx (S-S)

CH3 O

(14)

R

C

N + H2O

R

C NH2

Figure 17.1.1 (continued)

sulfinic, and sulfonic acids has also been observed to occur subsequent to S-nitrosothiol generation, suggesting that these oxyacids are produced by initial hydrolysis of the nitrosothiol groups to generate the protein sulfenic acid and HNO, the one electron reduced form of NO (Fig. 17.1.1, reaction 1; Poole et al., 2004). The sulfenic acid can then be further oxidized or condensed with reduced glutathione (GSH) or other thiol groups to form a mixed disulfide. Here again, factors stabilizing S-nitrosothiols, particularly those which preferentially yield either Cys–SNO or Cys–SOH products, are relatively poorly understood.

Chemistry of Sulfenic Acids Once formed, sulfenic acids are intrinsically unstable due to their chemical reactivity but can

be stabilized in the protein environment. The chemistry of sulfenic acids in small molecules and proteins, which has been reviewed elsewhere (Allison, 1976; Hogg, 1990; Claiborne et al., 2001), is summarized below in terms of the relevant reactions in which they participate and the forces which stabilize cysteine sulfenic acids in proteins. Self condensation Based on small molecules as model compounds, the greatest propensity of sulfenic acids is to undergo self-condensation via a hydrogen-bonded dimeric intermediate to yield a thiosulfinate linkage (Fig. 17.1.1, reaction 2). This irreversible condensation reaction highlights a remarkable feature of sulfenic acids: the displacement of −OH from one sulfur by the direct attack of the other demonstrates

Oxidative Stress


Supplement 18

their participation as both nucleophiles and electrophiles in chemical reactions. Given the intermediate which must be formed to promote this reaction, steric hindrance (preventing the close approach of two sulfenic acid moieties) and sulfenate anion stabilization (disfavoring generation of the hydrogen bonding network) are thus important factors in sulfenic acid stability in these molecules. Oxidation, reduction, and trapping Sulfenic acids have generally been considered transient intermediates in the oxidation of thiols to sulfinic (R-SO2H) or sulfonic (RSO3H) acids (Fig. 17.1.1; reaction 3) and in the formation of disulfide bonds from free thiols (Fig. 17.1.1, reaction 4). While sulfenic acids are readily reduced back to the thiol state with the chemical reductants dithiothreitol (DTT) or arsenite (which is reportedly unable to reduce disulfide bonds; Torchinsky, 1981), sulfinic acids are only reducible by such agents at very low pH (i.e., 0.1.

3. Read the A412 of a blank consisting of neutral pH buffer containing the same concentration of TNB as the sample. 4. Calculate the sulfenic-acid content of the protein as follows. a. Subtract the final A412 value of the modified sample (step 2) from the A412 of the blank (step 3). b. Using the extinction coefficient 14,150 M−1 cm−1 (Riddles et al., 1979), convert this value to concentration of sulfenic acid–containing residues. c. Divide by the protein concentration to obtain the sulfenic acid content of the protein. Excess peroxide and at least some other oxidants also react with TNB over time; if present, these oxidants should be added to control reactions without protein to assess the contribution of this reaction.

Isolate TNB-labeled protein 5. Transfer the reaction mixture to an ultrafiltration unit of appropriate molecular weight cutoff (MWCO). Remove unreacted TNB by washing with neutral pH buffer through repeated cycles of concentration and dilution (e.g., concentrate to 50 µl and redilute with 5 ml buffer two to three times) according to manufacturer’s instructions. Attention must be made to the stated MWCO of the device so that the protein of interest will be retained. With each concentration cycle, the flow-through solution (filtrate) is removed and fresh neutral pH buffer is added to dilute the protein again, for a total of two to three concentration/redilution cycles. Either Centricon ultrafiltration units, which allow 2 ml to be concentrated to 40 to 50 ìl, or larger units offered by several other suppliers (e.g., Orbital Biosciences or Vivascience) which allow 6 to 7 ml to be concentrated to ~10 to 50 ìl, can be used. In the case of the latter, the ability to concentrate more solution at once and to use swinging bucket rotors rather than fixed-angle rotors can speed up the washing process.

6. Check for the presence of free TNB in the filtrate by determining the absorbance at 412 nm. Repeat steps 5 and 6 until no free reagent is detected (i.e., A412 is 300

13.4 ± 1.4 — 12.7 ± 0.3 —

Mitochondria Vehicle α-Tocopherol α-Tocopheryl acetate α-Tocopheryl succinate

0.07 ± 0.01 0.08 ± 0.01 0.08 ± 0.01 0.20 ± 0.01

— — ND 1.81 ± 0.23

41 ± 8 31 ± 7 49 ± 6 >300

15.0 ± 0.3 15.1 ± 0.6 13.3 ± 0.1 —

Inner mitochondrial membrane Vehicle α-Tocopherol α-Tocopheryl acetate α-Tocopheryl succinate

0.06 ± 0.01 0.06 ± 0.01 0.06 ± 0.01 0.22 ± 0.05

— — ND 3.86 ± 0.79

12 ± 2 22 ± 3 18 ± 1 106 ± 49

23.3 ± 2.0 20.5 ± 0.6 21.7 ± 1.2 15.3 ± 0.2

Tissue/treatment

Lag time (min) Rate (AFU/min)

aAbbreviation: ND, not detectable. bRats were administered a single dose (i.p.) of vehicle (olive oil) or 0.19 mmol/kg of α-tocopherol, α-tocopheryl acetate, or α-tocopheryl

succinate. After 18 hr, liver homogenates and subcellular fractions were isolated as described by Hovius et al. (1990) with modifications. Tocopherol and tocopheryl ester levels were then determined by HPLC assay (Fariss et al., 1989). Data are the mean ± std. error; the sample size is 3 to 5 rats.

authors conclude that the enrichment of mitochondrial and inner mitochondrial membrane with functional tochopherol is a critical event in tochopheryl succinate cytoprotection against toxic oxidative stress. Antioxidant capacity can be estimated by direct measurement of the ability of biological samples to scavenge free radicals or by their susceptibility to lipid peroxidation. Several methods have been developed to assess the total antioxidant capacities of biological samples such as blood plasma, other body fluids, and plant extracts; the methods are mainly based on the ability of these biological samples to scavenge free radicals, or to chelate the transition metals (Ghiselli et al., 2000). These methods include the ferric reducing ability of plasma (FRAP; Benzie and Strain, 1996), total radical trapping parameter (TRAP; Wayner et al., 1985), trolox equivalent antioxidant capacity (TEAC; Re et al., 1999), oxygen radical absorbance capacity (ORAC), total oxyradical scavenging capacity (TOSC), and cycle voltammetry (CV) assays (Chevion et al., 2000). Although these methods can provide useful information, especially with the assessment of low-molecular-weight water-soluble antioxidants in particular, they may have some diffi-

culties in assessing certain antioxidants. For example, protein antioxidants like superoxide dismutase were not suitable for the assessment with such assays (Yu and Ong, 1999). In addition, such assays may underestimate the antioxidant capability for antioxidants acting as iron chelators, such as albumin (Yu and Ong, 1999). Furthermore, most of these assays are limited to determining the total antioxidant capacity in plasma or serum, but not in tissue. In contrast, the fluorescence microplate reader assay reported here has the distinct advantage of assessing the antioxidant capacity of tissues, cells, and subcellular fractions. However, this fluorescence assay is not useful for serum or plasma, as the peroxidation of this tissue does not produce a strong fluorescent signal (apparently insufficient polyunsaturated fatty acids are present to produce reactive aldehydes and fluorescence). Another important advantage of this fluorescence microplate reader assay is in determining the functional antioxidant contributions of lipophilic antioxidants such as tocopherol. Unfortunately, this fluorescence assay’s ability to assess the antioxidant contributions of water-soluble, non-enzymatic and enzymatic antioxidants has not been determined.

Oxidative Stress


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Measurement of Tissue Susceptibility to Lipid Peroxidation

Other methods for evaluation of antioxidant capacity include the assessment of the susceptibility to lipid peroxidation (Tirmenstein et al., 1998; Cervato et al., 1999), which is the strength of the fluorescent assay reported here. Lipid peroxidation is probably the most extensively investigated process induced by free radicals. A number of assays exist for determining lipid peroxidation based on several different biochemical endpoints. One of the common methods is the thiobarbituric acid reactive substances (TBARS) assay, which is known to detect malondialdehyde (MDA) and other monofunctional aldehydes. However, most of these assays require considerable sample preparation and analysis time. Samples either need to be extracted with organic solvents or dissolved in SDS; furthermore, the assay is not in real-time. The fluorescence microplate reader assay reported here has numerous advantages. First, assay procedures and sample preparations are rapid, inexpensive, and simple. Second, it allows the rapid monitoring of lipid peroxidation in real time using a large number of biological samples, including tissue, cells, and subcellular fractions. To clearly illustrate the advantages of this fluorescence microplate reader assay over the traditional TBARS assay, consider the time and expense required to analyze the samples in Table 17.3.2 for antioxidant capacity or susceptibility to lipid peroxidation. Using the microplate reader assay, these samples can be analyzed in a single 8-hr day by placing each sample (∼5 samples per treatment and subcellular fraction for a total of 60 samples to be analyzed) in individual wells of the 96-well microplate, adding the appropriate buffer and initiator of lipid peroxidation, and reading each well for fluorescence in the plate reader for 300 min. (The microplate reader automatically reads the fluorescence of each well every 5 min.) In contrast, these same samples would take ∼6 days to analyze for susceptibility to lipid peroxidation by the TBARS assay. Each sample (only 20 samples/day can be analyzed) is incubated in a glass scintillation vial with the appropriate buffer and initiators of lipid peroxidation, and an aliquot of this solution is collected every 15 min for 3 hr for TBARS analysis (12 aliquots for TBARS analysis per tissue sample). Accordingly, using the TBARS assay, it would take 3 full days to analyze all 60 tissue samples in this experiment (Table 17.3.2) and at least another 3 days to analyze the 720 aliquots for TBARS levels. Thus the advantages of this fluorescence microplate reader assay for

determining a tissues’ susceptibility to lipid peroxidation are substantial. For the most part, the authors have found good agreement between the TBARS assay and the fluorescence microplate reader assay in assessing the susceptibility of biological samples to lipid peroxidation (Tirmenstein et al., 1998). However, the assay has disadvantages as well. For example, the microplate reader assay cannot be used for quantification of amounts of lipid peroxidation produced, since the identity of the fluorophore is not known. Regardless of the methods used for measuring lipid peroxidation, the susceptibility of biological samples to lipid peroxidation can be estimated by adding lipid peroxidation initiators in the assay system and measuring the commencement time of lipid peroxidation or maximal rate of second propagation phase. To initiate the production of pro-oxidants in antioxidant capacity assays, numerous initiator systems have been used, such as hydrogen peroxide alone or coupled with horseradish peroxidase, 2,2′-azobis(2-amidinopropane)dihydrochloride (AAPH), 2,2′-azinobis(3-ethylbenzothiazoline 6-sulfonate) (ABTS), 2′-azobis(2,4-dimethylvaleronitrile) (AMVN), and inorganic iron or copper, given alone or in a chelated form. After examining several of these initiators, Fe3+/ADP was selected for the susceptibility to lipid peroxidation assay. It is known that inorganic iron in certain chelated forms is capable of promoting free-radical-mediated peroxidation of membrane lipids. In addition, Fe2+/ADP or Fe3+/ADP-ascorbate has been extensively studied and used as an in vitro model. The mechanism of iron-induced lipid peroxidation is hydroxylradical–production related via the Fenton reaction. The transition metal iron reacts with oxygen to produce superoxides, which can be dismutased by superoxide dismutase to form hydrogen peroxides. Hydrogen peroxides undergo Fenton reaction in the presence of ferrous iron (Fe2+) to form hydroxyl radicals (OH⋅; Fig. 17.3.2). Lipid peroxidation is initiated by this free radical and the chain reaction is promoted to the generation of lipid alkyl radicals (L⋅), followed by the formation of a lipid peroxyl radical (LOO⋅) due to the interaction with oxygen. LOO⋅ eventually converts to LOOH via hydrogen abstraction from neighboring allylic bonds (Fig. 17.3.2). Lipid peroxides in the presence of ferrous iron are converted to lipid alkoxyl radicals (again by the Fenton reaction) which degrade and fragment (again with the help of iron) to form a wide variety of reactive aldehyde products, including



Fe3+-ADP + ascorbate

−⋅⋅

Fe2+ + O 2 −⋅⋅

O2−⋅⋅ + O 2 + 2H+ Fe2+ + H2O2 LH + OH

H2O2 + O2

Fe 3+ + OH− + OH

⋅

⋅

⋅

⋅

LOO (O2 addition)

⋅

⋅

LOO + LH

LOOH + L (propagation)

LOOH + Fe 2+ ⋅ Fragmentation

SOD

L + H2 O (initiation)

L + O2

LO

Fe3++ O2

Fenton

⋅

Fe 2+ -ADP

⋅

LO + Fe 3+ + OH −

Lipid Aldehydes (4-HNE, MDA Acrolein)

Figure 17.3.2 Fe2+/Fe3+ initiation of generation of hydroxyl radicals and lipid peroxidation products.

4-hydroxy-2-nonenal, MDA, and acrolein. Amino acids, especially basic amino acids such as glycine, histidine, lysine, and arginine, can strongly react with the α, β-unsaturated carbonyl groups of these by-products to form fluorescence adducts. Such a process is believed to cause or at least be associated with numerous clinical conditions, including the formation of lipofuscin and ceroid pigments during aging, amyloid β-peptide plaques during Alzheimer’s disease, Lewy bodies in Parkinson’s disease, and the formation of advanced glycation end products (AGE)–associated protein adducts in diabetic complications such as nephropathy. The authors have experimented briefly with the use of other initiators in the assay, such as AAPH and Fe/H2O2, but found that the initiator Fe/ADP results in the greatest fluorescence signal and reproducibility in a wide variety of biological specimens (Tirmenstein, pers. comm.).

Critical Parameters and Troubleshooting Both the lag time and maximal rate of propagation phase can be used for estimation of susceptibility to lipid peroxidation. Extended lag time or lowered maximal rate suggests increased tissue antioxidant capacity or decreased susceptibility to lipid peroxidation. Lag time is the most valuable parameter used in this assay. The area under the curve (AUC)

may be an alternative way to describe the antioxidant capacity since both changes in lag time and maximal rate will significantly affect this parameter. All these values can be readily obtained from the array (Fig. 17.3.3). This method was originally developed and used for the determination of the susceptibility of rat liver and its subcellular organelles to lipid peroxidation. In terms of use for other biological samples, a few factors should be considered. First, the quantity of antioxidants in plasma, brain, kidney, heart, lung, and liver, as well as their peroxidizable lipids are different. For example, the formation of MDA in rat liver mitochondria was 12-fold higher than those of cardiac mitochondria with the same iron/ascorbate system (Wiswedel et al., 1989). Therefore, selecting the correct amount of tissue and the concentrations of lipid peroxidation initiator are critical to ensure that the sample is peroxidized in the time frame of the assay. For example, with heart tissue, a greater amount of tissue and higher concentrations of lipid peroxidation initiator are required (see Table 17.3.1). The use of plasma or serum in this assay has not been successful, presumably due to the low level of unsaturated lipid in this biological specimen, thus limiting the ability to generate a detectable fluorescence signal. Background fluorescence is increased slightly after 5 hr of incubation with the lipid peroxidation initiator as described (see Basic Oxidative Stress


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Figure 17.3.3 Kinetic traces of fluorescence for lipid peroxidation in isolated hepatic mitochondrial membrane induced by Fe3+-ADP/ascorbate. The mitochondrial samples were isolated from rats administered (i.p.) either vehicle (traces A1-A3, A10-A12, B7-B9, C4-C6), tocopheryl acetate (A4-A6, B1-B3, B10-B12, C7-C12), or the Tris salt of tocopheryl succinate (A7-A9, B4-B6, C1-C3), 18 hr prior to isolation.


Protocol) but lacking biological samples. The results also indicate that the increase in fluorescence with incubation time only occurs in incubations containing ascorbate, ADP, ferric chloride, and glycine. If any one of these compounds is omitted from the incubation solution, no increase in fluorescence is detected. In general, the concentration of ascorbate is also important in determining whether it is pro-oxidant (reducing iron) or antioxidant. Ascorbate at higher concentrations acts as an antioxidant, but lipid peroxidation (MDA) production by Fe3+/ADP was largely potentiated at lower concentrations of ascorbate. In the present assay, a final concentration of 450 mM ascorbate was used. The lag time can be shortened by increasing the concentration of ferric chloride/ADP, and thus varying the concentration of ferric chloride is an effective way of controlling lag time. Metal chelators such as EDTA or EGTA are commonly used during sample preparation, such as fractionation of homogenates, mitochondria, and microsomes from tissue. These

chelators can form complexes with iron and inhibit lipid peroxidation. Therefore, isolated fractions should be completely washed with non-EDTA or -EGTA buffer. Other considerations should also be taken into account during the studies of the antioxidant capacity of vitamin E esters, such as tocopheryl succinate. It has been shown that tocopheryl succinate can be taken up rapidly by the cells and incorporated into the membrane where it is hydrolyzed into tocopherol by membrane-bound esterase (Fariss et al., 2001). Thus, during the microplate reader assay, esterase inhibitors should be added in order to avoid any hydrolysis of tocopheryl succinate in the samples. A recent study by the authors showed that thenoyltrifluoroacetone is a potent inhibitor of esterase activity and it effectively inhibits the hydrolysis of tocopheryl succinate without affecting the measurement of susceptibility to lipid peroxidation (Zhang and Fariss, 2002). The addition of 25 µM thenoyltrifluoroacetone (final concentration) is recommended in these studies.



Anticipated Results Figure 17.3.1 shows the kinetic profile of lipid peroxidation with a 5-hr incubation time. Typically, three phases are observed: (I) the initial latency phase, followed by (II) a second propagation phase, and finally (III) a termination phase. Lag time can vary depending on the source of tissues and concentration of lipid peroxidation initiator used. With the current assay method (see Basic Protocol), the lag time for different tissues ranged from 10 to 40 min and maximal rates from 10 to 25 AFU/min (Table 17.3.1). As mentioned previously, the authors’ laboratory has a long-term interest in the protective role of vitamin E derivatives, including tocopherol, tocopheryl acetate, and tocopheryl succinate. Tocopherol is a chain-breaking antioxidant and is known to react with a variety of free radicals, in particular peroxyl radicals Due to its lipophilic property, tocopherol does not rapidly accumulate in hepatic mitochondria. The authors and others have found that tocopheryl succinate is far more effective than tocopherol in protecting isolated hepatocytes against many different toxic oxidative changes, presumably via enriching mitochondria with tocopherol (Fariss et al., 2001). Recent in vivo studies by the authors showed that a single dose of 0.19 mmol/kg (~100 mg/kg) tocopheryl succinate i.p. can rapidly enrich not only rat hepatocytes, but most importantly, mitochondria and inner mitochondrial membranes, which consequently results in a decrease in their susceptibility to lipid peroxidation as compared with samples from vehicle control animals. The decrease in susceptibility to lipid peroxidation with this treatment provides important evidence that the measured enrichment of tocopherol in these mitochondrial fractions is functioning as an effective antioxidant. For example, as illustrated in Table 17.3.2 for mitochondria, the lag time (elapsed time between addition of initiator to the sample and the onset of lipid peroxidation, as measured by an increase in fluorescence in this assay) for samples isolated from vehicle control rats is very short, ∼41 min. with a rate of 15.0 AFU/min. This rapid onset of lipid peroxidation (fluorescence) indicates that this sample is highly susceptible to lipid peroxidation. By contrast, hepatic mitochondria isolated from rats pretreated with tocopheryl succinate were completely resistant to the induction of lipid peroxidation, as indicated by a lag time of >300 min (maximum incubation time) and no measurable rate (an increase in fluorescence was not observed in

this sample). Interestingly, pretreatment of rats with tocopherol or tocopheryl acetate did not significantly alter the susceptibility of their hepatic mitochondria to lipid peroxidation (with lag times and rates similar to those observed for vehicle control, Table 17.3.2). Decreased tissue susceptibility to lipid peroxidation in these studies correlated well with enhanced tocopherol levels in these samples. These results confirm that the assay is a reliable way to assess the total antioxidant status in tissue. It is interesting to note however, that unlike mitochondria, the inner mitochondrial membrane was not completely protected from the susceptibility to lipid peroxidation following tocopheryl succinate administration in vivo (Table 17.3.2), even though these two mitochondrial fractions were enriched with identical levels of tocopherol. These results can be explained by the greater cardiolipin content of inner mitochondrial membrane (as compared with mitochondria) and the well-known enhanced susceptibility of this inner mitochondrial membrane–specific phospholipid to lipid peroxidation (Lesnefsky et al., 2001).

Time Consideration

The total time required for this assay is ∼6 hr. Sonication of samples and preparation of lipid peroxidation initiator can be finished within 1 hr.

Literature Cited Benzie, I.F.F. and Strain, J.J. 1996. The ferric reducing ability of plasma (FRAP) as a measure of “antioxidant power”: The FRAP assay. Anal. Biochem. 239:70-76. Cervato, G., Viani, P., Cazzola, R., and Cestaro, B. 1999. A fluorescence method for the determination of plasma susceptibility to lipid peroxidation. Clin. Biochem. 32:171-177. Chevion, S., Roberts, M.A., and Chevion, M. 2000. The use of cyclic voltammetry for the evaluation of antioxidant capacity. Free Radic. Biol. Med. 28:860-870. Fariss, M.W., Merson, M.H., and O’Hara, T.M. 1989. Alpha tocopheryl succinate protects hepatocytes from chemical-induced toxicity under physiological calcium conditions. Toxicol. Lett. 47: 61-75. Fariss, M.W., Nicholls-Grzemski, F.A., Tirmenstein, M.A., and Zhang, J-G. 2001. Enhanced antioxidant and cytoprotective abilities of vitamin E succinate is associated with a rapid uptake advantage in rat hepatocytes and mitochondria. Free Rad. Biol. Med. 31:530-541. Ghiselli, A., Serafini, M., Natella, F., and Scaccini, C. 2000. Total antioxidant capacity as a tool to assess redox status: Critical view and experimental data. Free Rad. Biol. Med. 29:1106-1114.

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Hovius, R., Lambrechts, H., Nicolay, K., and De Kruijff, B. 1990. Improved methods to isolate and subfractionate rat liver mitochondria. Lipid composition of the inner and outer membrane. Biochim. Biophys. Acta. 1021:217-226. Lesnefsky, E.J., Slabe, T.J., Stoll, M.S.K., Minkler, P.E., and Hoppel, C.L., 2001. Myocardial ischemia selectively depletes cardiolipin in rabbit heart subsarcolemmal mitochondria. Am. J. Physiol. Heart Circ. Physiol. 280:H2770-H2778. Niki, E. and Noguchi, N., 2000. Evaluation of antioxidant capacity. What capacity is being measured by which method? IUBMB Life 50:323-329. Re, R., Pellegrini, N., Proteggente, A, Pannala, A., Yang, M., and Rice-Evans, C. 1999. Antioxidant activity applying an improved ABTS radical cation decolorization assay. Free Rad. Biol. Med. 26:1231-1237. Tirmenstein, M.A., Pierce, C.A., Leraas, T.L., and Fariss, M.W. 1998. A fluorescence plate reader assay for monitoring the susceptibility of biological samples to lipid peroxidation. Anal. Biochem. 265:246-252. Wayner, D.D.M., Burton, G.W., Ingold, K.U., and Locke, S. 1985. Quantitative measurement of the total, peroxyl radical-trapping antioxidant capacity of human blood plasma by controlled peroxidation. FEBS Lett. 187:33-37.

Wiswedel, I., Ulbricht, O., and Augustin, W. 1989. Studies of lipid peroxidation in isolated rat heart mitochondria. Biomed. Biochim. Acta 48:S73S76. Woodford, F.P. and Whitehead, T.P. 1998. Is measuring serum antioxidant capacity clinically useful? Ann. Clin. Biochem. 35:48-56. Yu, T-W. and Ong, C.N. 1999. Lag-time measurement of antioxidant capacity using myoglobin and 2,2′-azino-bis(3-ethylbenzthiazoline-6-sulfonic acid): Rationale, application, and limitation. Anal. Biochem. 275:217-223. Zhang, J.G. and Fariss, M.W. 2002. Thenoyltrifluoroacetone, a potent inhibitor of carboxylesterase activity. Biochem. Pharmacol. 63:753-757.

Contributed by Jin-Gang Zhang BD Biosciences Woburn, Massachusetts Marc W. Fariss University of Colorado Health Sciences Center Denver, Colorado




In Situ Localization of Nonenzymatic Peroxidase-Like Activity of Tissue-Bound Transition Metals

UNIT 17.4

The protocols described below pertain to detection of tissue iron using the DAB-enhanced Perls method (see Basic Protocol) as well as the general detection of redox-active transition metal ions without the introduction of any metal-containing reagents (see Alternate Protocol). In the latter case, use of the MBTH/o-phenylenediamine method in the absence of H2 O2 pretreatment permits the selective visualization of endogenous enzymatic peroxidase-like activity, whereas use of DAB following the H2 O2 pretreatment permits the visualization of endogenous nonenzymatic redox-active metals. Whether the staining reflects mainly iron or mainly copper can be ascertained by noting whether it is attenuated more by preincubation with increasing concentrations of the iron-selective chelator deferoxamine or by increasing concentrations of the copper-selective chelator diethylenetriaminepentaacetic acid (DTPA), as described in the Support Protocol. These methods do not permit one to distinguish other redox-active metals such as chromium, and possibly manganese, whose contributions cannot be excluded. Noting an inhibitory effect of prior incubation with chelating agents is further evidence that the DAB staining reflects adventitiously bound redox-active metals rather than enzymatic activity, because the latter is refractory to chelators. A key advance represented by the protocols provided below is an enhanced sensitivity arising from the recognition that standard formalin fixation detracts from detection of metal-based redox activity, which is not the case with the fixative used in the protocols below (methacarn; 60:30:10 methanol/chloroform/acetic acid; see Reagents and Solutions), as verified by limited studies using nonfixed tissue.

DETECTION OF IRON Iron can be detected in freshly isolated, paraffin-embedded tissue sections fixed in the absence of formalin, using potassium ferrocyanide and 3,3 -diaminobenzidine tetrahydrochloride (DAB).

BASIC PROTOCOL

Materials Fresh tissue of interest Fixative solution (see recipe) 30%, 50%, 70%, 80%, 95%, and 100% (v/v) ethanol Xylene Paraffin 50 mM Tris·Cl, pH 7.6 (APPENDIX 2A) 7% (w/v) potassium ferrocyanide solution (see recipe), freshly prepared DAB/H2 O2 solution (for Basic Protocol; see recipe), freshly prepared Permount mounting medium (e.g., Fisher) Tissue-embedding cassettes (Omnisette; Fisher) Paraffin-embedding system or machine (e.g., TBS ATPI Automated Tissue Processor; Fisher) Microtome Superfrost Plus microscope slides (Fisher) Coplin jars Oxidative Stress Contributed by Lawrence M. Sayre, Peggy L.R. Harris, George Perry, and Mark A. Smith Current Protocols in Toxicology (2004) 17.4.1-17.4.8 C 2004 by John Wiley & Sons, Inc. Copyright


Humified chamber (storage container with sealable lid, lined with moistened paper towels and containing elevated surface on which to place slides) or container designed for immunohistochemical incubations 3-cc BD disposable syringes (Fisher) 0.22-µm Millex-GS syringe filters (Millipore) Coverslips Prepare tissue 1. Dissect the tissue sample. 2. Place an ∼0.5-cm3 sample of tissue in a tissue-embedding cassette and immerse in 50 ml of fixative solution. Incubate overnight at 4◦ C 3. Remove sample from fixative and place in 50% ethanol. 4. Using a histology paraffin-embedding system/machine, dehydrate sample through an ascending ethanol series and carry out paraffin embedding as follows:

70% ethanol for 40 min at 40◦ C 80% ethanol for 40 min at 40◦ C 95% ethanol for 40 min at 40◦ C 100% ethanol for 40 min at 40◦ C 100% ethanol for 40 min at 40◦ C 100% ethanol for 1 hr 10 min at 40◦ C xylene for 40 min at 40◦ C xylene for 40 min at 40◦ C paraffin for 40 min at 60◦ C paraffin for 1 hr at 60◦ C paraffin for 40 min at 60◦ C. Follow the manufacturer’s suggested operating guidelines, but run the machine without formalin, thus avoiding the standard formalin fixative step at the beginning. 5. Once a paraffin block is obtained, cut 6-µm sections using a microtome and place the sections on Superfrost Plus microscope slides. Dry sections on the slide overnight at 37◦ C 6. Rehydrate sections by immersing in the following solutions in Coplin jars at room temperature for the indicated lengths of time.

xylene for 10 min xylene (fresh) for 10 min 100% ethanol for 10 min 95% ethanol for 10 min 70% ethanol for 10 min 50% ethanol for 10 min 30% ethanol for 10 min 50 mM Tris·Cl, pH 7.6, for 10 min 50 mM Tris·Cl, pH 7.6 (fresh) for 10 min.

Nonenzymatic Peroxidase-Like Activity of Tissue-Bound Transition Metals

Stain for iron 7. Apply enough freshly made 7% potassium ferrocyanide solution to cover the section. Place slide in humidified chamber and incubate 1.5 to 2 hr at 37◦ C or overnight at 4◦ C. 8. Rinse slide in 50 mM Tris·Cl, pH 7.6, for 10 min.



Figure 17.4.1 Histochemical detection of iron in Alzheimer’s disease (A) compared with control case (B) shows striking association of iron with neurofibrillary tangles (arrowheads) and senile plaques (arrows) characteristic of the Alzheimer’s disease brain. Scale bar = 200 µm.

9. Develop signal by covering the section with freshly prepared DAB/H2 O2 solution (using a 3-cc syringe and 0.22-µm syringe filter to filter the DAB/H2 O2 ) and leaving the solution on the section for 5 min. 10. Dehydrate sections by immersing in the following solutions (ascending ethanol series) in Coplin jars at room temperature for the indicated lengths of time:

30% ethanol for 10 min 50% ethanol for 10 min 70% ethanol for 10 min 95% ethanol for 10 min 100% ethanol for 10 min xylene for 10 min. 11. Apply coverslip to slide using Permount. A dark brown precipitate, easily visualized by light microscopy, will form at sites containing iron (Fig. 17.4.1). Time consideration—approximately 4.5 to 18 hr.

DETECTION OF REDOX-ACTIVE TRANSITION METALS Redox activity, a key feature of non-sequestered transition metals (e.g., iron, copper) provides a key enhancement to metal localization studies.

ALTERNATE PROTOCOL

Additional Materials (also see Basic Protocol) Peroxide/methanol: add 4 ml of 30% hydrogen peroxide to 36 ml of 100% methanol; prepare fresh daily DAB/H2 O2 solution (for Alternate Protocol; see recipe), freshly prepared MBTH/o-phenylenediamine/H2 O2 solution (see recipe), freshly prepared 1. Dissect, fix, dehydrate, embed, and section sample; place sections on slides and dry overnight (see Basic Protocol, step 1 to 5). 2. Rehydrate sections (and optionally inactivate endogenous peroxidases) by immersing in the following solutions in Coplin jars at room temperature for the indicated lengths of time:

xylene for 10 min xylene (fresh) for 10 min 100% ethanol for 10 min

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95% ethanol for 10 min peroxide/methanol for 30 min (optional; to inactivate endogenous peroxidase activity) 70% ethanol for 10 min 50% ethanol for 10 min 30% ethanol for 10 min 50 mM Tris·Cl, pH 7.6, for 10 min 50 mM Tris·Cl, pH 7.6, for 10 min. 3. Optional: Perform chelation to titrate away endogenous metals to determine the source of activity (i.e., metal) and to verify the specificity of the finding (see Support Protocol). 4. Using a 3-cc syringe and 0.22-µm syringe, apply enough freshly made DAB/H2 O2 solution to cover the section. 5. To an adjacent section on the same slide or a section on a second slide, using a 3-cc syringe and 0.22-µm syringe filter to filter the MBTH/o-phenylenediamine solution, apply MBTH/o-phenylenediamine solution for comparison purposes. 6. Place slides in a humidified chamber and incubate 1.5 to 2 hr at room temperature. 7. Rinse slides in 50 mM Tris·Cl, pH 7.6, for 10 min. 8. Dehydrate sections by immersing in the following solutions (ascending ethanol series) in Coplin jars at room temperature for the indicated lengths of time:

30% ethanol for 10 min 50% ethanol for 10 min 70% ethanol for 10 min 95% ethanol for 10 min 100% ethanol for 10 min xylene for 10 min. 9. Apply coverslip to slide using Permount. A dark brown precipitate will form at sites containing redox activity. SUPPORT PROTOCOL

CHELATION OF METALS FOR CONTROLS After the tissue sections have been hydrated, a chelation step may be performed to selectively remove individual endogenous metals. A serial dilution of DEF (which selectively removes iron) and DTPA (which selectively removes copper) can be used, starting with a 0.1 M concentration and doing successive 1:10 dilutions with water (0.01 M and 0.001 M). Although higher concentrations of either chelator will remove all metals, a determination of the relative effectiveness of the chelators at lower concentration may permit an assessment of which metal ions are most responsible for the redox activity.

Additional Materials (also see Basic Protocol and Alternate Protocol)


0.1 M DEF stock: Weigh 65.68 mg deferoxamine (DEF; Sigma) and add 1 ml H2 O 0.1 M DTPA stock: Weigh 39.33 mg diethylenetriaminepentaacetic acid (DTPA; Sigma) and add 1 ml H2 O 1. After performing the rehydration series in step 2 of the Alternate Protocol, remove the slides from the Tris·Cl buffer, and apply either DEF or DTPA to appropriate sections at the desired concentration (e.g., serial dilutions of 0.1, 0.01, and 0.001 M).



Figure 17.4.2 Lesion-associated iron (see Fig. 17.4.1) could be completely stripped with deferoxamine (A) but readily rebound to the same sites following incubation with iron(III) citrate and iron(II) chloride (B). Asterisks indicate landmark blood vessels in adjacent sections. Scale bar = 100 µm.

2. Incubate overnight at room temperature in humidified chamber. 3. Rinse extensively with 50 mM Tris·Cl, pH 7.6, prior to continuing the detection procedure (see Alternate Protocol, step 4). Typical results are shown in Figure 17.4.2.

REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for the preparation of all solutions and in all protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.

DAB/H2 O2 solution (for Alternate Protocol) Weigh 5 mg of 3,3 -diaminobenzidine tetrahydrochloride (DAB). Add 6 ml 50 mM Tris·Cl, pH 7.6 (APPENDIX 2A), and 672 µl of 30% hydrogen peroxide, then mix. Prepare immediately before use. Final concentrations: 0.75 mg/ml DAB/3% H2 O2 .

DAB/H2 O2 solution (for Basic Protocol) Weigh 5 mg of 3,3 -diaminobenzidine tetrahydrochloride (DAB). Add 6.67 ml 50 mM Tris·Cl, pH 7.6 (APPENDIX 2A), and 3.36 µl of 30% hydrogen peroxide, then mix. Prepare immediately before use. Final concentrations: 0.75 mg/ml DAB/0.015% H2 O2 .

Fixative solution Mix 300 ml methanol, 150 ml chloroform, and 50 ml glacial acetic acid to give a final volume of 500 ml. Store in a glass bottle at 4◦ C until needed. Final concentrations: 60:30:10 (v/v/v) methanol/chloroform/acetic acid.

MBTH/o-phenylenediamine/H2 O2 solution Stock solutions: 3 mg/ml MBTH: Weigh 3 mg of 3-methyl-2-benzothiazolinone hydrazone (MBTH; Sigma) and add 1 ml of 50 mM Tris·Cl, pH 7.6 (APPENDIX 2A). 2 mg/ml o-phenylenediamine: Weigh 2 mg of o-phenylenediamine and add 1 ml of 50 mM Tris·Cl, pH 7.6 (APPENDIX 2A). continued

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Working solution: Mix the following immediately before use: 125 µl of 3 mg/ml MBTH 125 µl of 2 mg/ml o-phenylenediamine 50 µl of 30% hydrogen peroxide 200 µl of 50 mM Tris·Cl, pH 7.6 (APPENDIX 2A)

Potassium ferrocyanide solution, 7% Weigh 0.7 g K4 Fe(CN)6 ·3H2 O into a graduated cylinder. Add 5 ml water and 300 µl concentrated hydrochloric acid. Mix, then bring to a final volume of 10 ml. Prepare immediately before use. COMMENTARY Background Information


Many disease states are associated with tissue accumulation of metals. This accumulation can reflect unusually high tissue levels of metal-storage proteins such as ferritin (iron) and ceruloplasmin (copper), or metal transporters such as transferrin (iron) and the copper-transporting ATPase implicated in Wilson’s disease. When such proteins become saturated, free metal ions are released into plasma and, at least in the case of iron, may result in deposition of simple inorganic forms of the metal, such as Fe2 O3 . A number of techniques have been used for quantitation of tissue metals, including atomic absorption spectroscopy, magnetic resonance imaging, Xray microanalysis, and laser microprobe mass analysis (LAMMA). For semiquantitative determination of tissue metals, however, various histochemical methods have always seen greater use on account of their convenience (Bancroft and Stevens, 1982; Pearse, 1985). Specificity of the latter methods for individual metals relies on the use of metal ion-specific chelating agents and often requires prior release of metals from their protein-binding sites by treatment with trichloroacetic acid or HCl. The most popular method for histochemical detection of iron has been Perls’ method (Perls, 1867; Thompson, 1966), which relies on the formation of the deep blue mixed-valent (Fe2+ Fe3+ ) cyano complex known as Prussian blue. Usually, this technique localizes ferric iron that can be mobilized by treatment with ferrocyanide, though reaction of ferricyanide with Fe2+ gives the same complex (Parmley et al., 1978). A later development of this method was to take advantage of the fact that Prussian blue is especially active in catalyzing the H2 O2 -dependent oxidation of 3,3 -diaminobenzidine (DAB), the latter serving as the most commonly used chromogenic

substrate for peroxidase. This so-called enhanced Perls’ method (Nyguyen-Legros et al., 1980) has been widely applied to the histochemical localization of total iron (ferritin, transferrin, and free iron) in brain and other tissues as a function of aging and disease (Conner et al., 1990). The advantage of using DAB is that its oxidation yields an insoluble brown polymer deposited at the sites of enzyme localization and readily detected by microscopy. Nonetheless, with application of H2 O2 and DAB becoming the most generally used chromogenic method for immunohistochemical studies that involve the (horseradish) peroxidase-antiperoxidase (PAP) method or direct use of horseradish peroxidase–linked secondary antibodies, it has been important to eliminate any endogenous peroxidase activity in the tissue by prior treatment with H2 O2 at a concentration (usually one part 3% H2 O2 diluted with four parts methanol) that inactivates such enzymatic activity (Sternberger, 1986). Without the H2 O2 pretreatment, any endogenous peroxidase-like activity would interfere. Indeed, the H2 O2 -dependent oxidation of peroxidase substrates such as DAB or 3,3 ,5,5 -tetramethylbenzidine (TMB) has been directly used to localize endogenous peroxidase activity in neural tissue (Svensson et al., 1984). The key point here is that such activity can represent either authentic enzymatic (peroxidase or catalase) activity or nonenzymatic “pseudo”-peroxidase activity of adventitiously bound redox-active transition metal ions. A distinction between the two can be ascertained only by studies that probe the conditions under which the peroxidase-like activity persists. Thus, when the activity shows a pH dependence reminiscent of a particular heme enzyme, can be blocked by prior treatment with sodium azide (Schaermeyer, 1992) or 3-amino-1,2,4-triazole, or can be eliminated



by heating, such as occurs in the case of staining of peroxisomes (Schipper et al., 1990a), the interpretation has been in terms of endogenous enzyme activity. On the other hand, when staining is independent of pH, heat, or the presence of inhibitors (Schipper et al., 1990b), as has been found for astrocyte mitochondria in the aging brain (Schipper et al., 1998), the interpretation has been in terms of nonenzymatic redox activity. In the latter regard, concomitant use of X-ray microanalysis has provided evidence for localization of both iron (presumed to be nonenzymatic heme iron on the basis of lack of reaction with Prussian blue; Schipper et al., 1990a) and copper (Brawer et al., 1994). The authors’ studies revealed the presence of H2 O2 -dependent DAB oxidizing activity in the neurofibrillary pathology of Alzheimer disease (Sayre et al., 2000). Since this activity was not attenuated by prior exposure of the tissue sections to peroxidase-inactivating levels of H2 O2 , the interpretation was in terms of nonenzymatic redox-active transition metals. Nonetheless, to demonstrate that the level of H2 O2 being used was sufficient to inactivate endogenous enzymatic peroxidase activity, a method was needed for detecting the latter which was independent of the use of DAB. A range of studies on the effect of added iron- and copper-selective chelating agents on the intrinsic ability of these metals to mediate H2 O2 -dependent DAB oxidation suggested, at least in the case of copper, that a key facet of this reaction was the ability of DAB to enter into bidendate coordination with Cu2+ . Thus, the authors found that simple transition metal complexes were incapable of mediating the H2 O2 -dependent oxidation of noncoordinating peroxidase substrates such as TMB. This would suggest that TMB is an ideal reagent for selective detection of enzymatic peroxidase activity in the face of redox-active free metal ions that are detected by DAB (Trojanowski et al., 1983). Indeed, the discrepancy between DAB staining and TMB staining observed by others (Svensson et al., 1984) most likely reflects the redox activity of nonenzymatic transition metals. The authors, however, were unable to observe a reproducible tissue staining using TMB. On the other hand, they found that a 1:1 mixture of 3-methyl-2-benzothiazolinone hydrazone (MBTH) and o-phenylenediamine, which, like TMB, gave a negative result with H2 O2 in the presence of simple transition metal complexes, permitted the observation of endogenous enzymatic peroxidase

activity. The reaction in this case involved an oxidative coupling (Ngo and Lenhoff, 1980) to generate an insoluble material similar to that seen using DAB. Staining using the MBTH/ophenylenediamine reagent, albeit weak, was eliminated by the H2 O2 pretreatment, unlike the DAB staining, confirming that the latter reflected nonenzymatic redox activity (Sayre et al., 2000).

Critical Parameters and Troubleshooting The most important factor to ensure success is the fixative. The tissue should not be exposed to any aldehyde-containing fixative, e.g., formalin or formaldehyde. If this does occur, the methods will only show a limited stain or not work at all. Also, the potassium ferrocyanide, MBTH, and DTPA solutions should be made just prior to use so that they do not become oxidized and lose any of their strength. In any event, if the staining of the sample is weak, a longer incubation with the solutions may be needed. It is possible to incubate the samples overnight in the working solutions at 4◦ C to increase the stain intensity, or even at room temperature overnight, although this may increase the background of the stain. Occasionally, a precipitate forms when the DAB is applied to the samples. Once on the sections, it is difficult to remove; however, a more thorough filtering of freshly made DAB solution should correct this for future stains.

Anticipated Results A brown precipitate, easily visualized by light microscopic techniques, will form at the sites containing iron (see Basic Protocol) or redox activity (see Alternate Protocol). Representative results are shown in Figures 17.4.1 and 17.4.2.

Time Considerations After the tissue samples have been embedded in paraffin and sections cut (which will take 1 day), the procedure should take ∼5 hr (or 48 hr if an overnight incubation is required).

Literature Cited Bancroft, J.D. and Stevens, A. (eds.) 1982. Theory and Practice of Histological Techniques, 2nd ed. Churchill Livingstone, New York. Brawer, J.R., Stein, R., Small, L., Cisse, S., and Schipper, H.M. 1994. Composition of Gomoripositive inclusions in astrocytes of the hypothalamic arcuate nucleus. Anat. Record 240:407415.

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Connor, J.R., Menzies, S.M., St. Martin, S.M., and Mufson, E.J. 1990. Cellular distribution of transferrin, ferritin, and iron in normal and aged human brains. J. Neurosci. Res. 27:595-611. Ngo, T.T. and Lenhoff, H.M. 1980. A sensitive and versatile chromogenic assay for peroxidase and peroxidase-coupled reactions. Anal. Biochem. 105:389-397. Nguyen-Legros, J., Bizot, J., Bolesse, M., and Pulicani, J.-P. 1980. Noir de diaminobenzidine: Une nouvelle methode histochimique de revelation du fer exogene. Histochem. 66:239-244. Parmley, R.T., Spicer, S.S., and Alvarez, C.J. 1978. Ultrastructural localization of nonheme cellular iron with ferrocyanide. J. Histochem. Cytochem. 26:729-741. Pearse, A.G.E. 1985. Histochemistry. Theoretical and Applied. Vol. 2. Analytical Technology, 4th ed., Churchill Livingstone, New York. Perls, M. 1867. Nachweis von Eisenoxyd in gewissen Pigmenten. Virchows Arch. Pathol. Anat. 39:42-48. Sayre, L.M., Perry, G., Harris, P.L.R., Liu, Y., Schubert, K., and Smith, M.A. 2000. In situ oxidative catalysis by neurofibrillary tangles and plaques in Alzheimer’s disease: A central role for bound transition metals. J. Neurochem. 74:270-279. Schaermeyer, U. 1992. Localization of peroxidase activity in the retina and the retinal pigment epithelium of the Syrian golden hamster (Mesocricetus auratus). Comp. Biochem. Physiol. 103:139-145.

Schipper, H.M., Lechan, R.M., and Reichlin, S. 1990b. Glial peroxidase activity in the hypothalamic arcuate nucleus: Effects of estradiol valerate-induced persistent estrus. Brain Res. 507:200-207. Schipper, H.M., Vininsky, R., Brull, R., Small, L., and Brawer, J.R. 1998. Astrocyte mitochondria: A substrate for iron deposition in the aging rat substantia nigra. Exp. Neurol. 152:188-196. Sternberger, L.A. 1986. Immunocytochemistry, 3rd ed. John Wiley & Sons, New York. Svensson, B.A., Rastad, J., and Westman, J. 1984. Endogenous peroxidase-like activity in the feline dorsal column nuclei and spinal cord. Exp. Brain Res. 55:325-332 Thompson, S.W. 1966. Histochemical and Histopathological Methods, pp. 592-594. Charles C. Thomas, Springfield, Ill. Trojanowski, J.Q., Obrocka, M.A., and Lee, V.M.-Y. 1983. A comparison of eight different chromogen protocols for the demonstration of immunoreactive neurofilaments or glial filaments in rat cerebellum using the peroxidaseantiperoxidase method and monoclonal antibodies. J. Histochem. Cytochem. 31:1217-1223.

Contributed by Lawrence M. Sayre, Peggy L.R. Harris, George Perry, and Mark A. Smith Case Western Reserve University Cleveland, Ohio

Schipper, H.M., Scarborough, D.E., Lechan, R.M., and Reichlin, S., 1990a. Gomori-positive astrocytes in primary culture: Effects of in vitro age and cysteamine exposure. Dev. Brain Res. 54:7179.




F2-Isoprostanes as Markers of Oxidant Stress: An Overview Lipid peroxidation, the oxidation of cellular lipids, is a central feature of oxidant stress, a phenomenon that has been increasingly implicated as causative in numerous pathological conditions. Lipid peroxidation products are frequently used to quantify oxidative injury, and can be assessed by a number of methods that include the measurement of either primary or secondary peroxidation end products. The development of specific, reliable, and non-invasive methods for measuring oxidative stress in humans is of fundamental importance for establishing the role of free radicals in human diseases. Primary end products of lipid peroxidation include conjugated dienes and lipid hydroperoxides, while secondary end products include thiobarbituric reactive substances (TBARS), gaseous alkanes, and a group of prostaglandin (PG) F2 -like products termed F2 -isoprostanes (F2 -IsoPs; Halliwell and Grootveld, 1987; Morrow et al., 1990a). Quantification of these various compounds has proven highly useful for the study of free radical–mediated lipid peroxidation in in vitro model systems. However, the F2 -IsoPs appear to be a significantly more accurate marker of oxidative stress in vivo in humans and animals than other compounds (Liu et al., 1999; Morrow and Roberts, 1999; Fam and Morrow, 2003). The biochemistry of F2 -IsoPs, as well as considerations regarding their utility as markers of oxidative stress will be discussed herein.

MECHANISM OF FORMATION OF ISOPROSTANES IsoPs are PG-like compounds formed from the peroxidation of arachidonic acid, a ubiquitous polyunsaturated fatty acid (Morrow et al., 1990a; Morrow and Roberts, 1999). Unlike PGs, which are formed via the action of the cyclooxygenase enzymes, F2 -IsoPs are formed non-enzymatically as a result of the free radical–mediated peroxidation of arachidonic acid. Figure 17.5.1 outlines the mechanism by which IsoPs are generated. Following abstraction of a bis-allylic hydrogen atom and the addition of a molecule of oxygen to arachidonic acid to form a peroxyl radical, endocyclization occurs and an additional molecule of oxygen is added to form PGG2 -like compounds. These unstable bicycloendoperoxide intermediates

UNIT 17.5

are then reduced to the F2 -IsoPs. Based on this mechanism of formation, four F2 IsoP regioisomers are generated (see Fig. 17.5.1; Morrow et al., 1990a,b; Morrow and Roberts, 1999). Compounds are denoted as 5-, 12-, 8-, or 15-series regioisomers depending on the carbon atom to which the side chain hydroxyl is attached; the authors have proposed a nomenclature system based on this feature (Taber et al., 1997). An alternative system for the IsoPs has been proposed by FitzGerald and colleagues in which the abbreviation iP is used for isoprostane, and regioisomers are denoted as III to VI based upon their structure (Rokach et al., 1997). IsoPs that contain F-type prostane rings are isomeric to PGF2α and are thus referred to as F2 -IsoPs. It should be noted that IsoPs containing alternative ring structures (such as those resembling PGD2 /E2 and PGA2 /J2 ) can also be formed by this mechanism (Chen et al., 1999; Reich et al., 2001a). F2 -IsoPs, however, have been the most studied class of IsoPs and, because of their stability, afford the most accurate measure of oxidative stress (Liu et al., 1999). An important structural distinction between IsoPs and cyclooxygenase-derived PGs is that the former contain side chains that are predominantly oriented cis to the prostane ring while the latter possess exclusively trans side chains (Morrow et al., 1990a,b). A second important difference between IsoPs and PGs is that IsoPs are formed in situ esterified to phospholipids and are subsequently released by a phospholipase(s) (Liu et al., 1999; Morrow and Roberts, 1999), while PGs are generated only from free arachidonic acid.

QUANTIFICATION OF F2 -ISOPROSTANES Over the past 12 years, several methods have been developed to quantify the F2 -IsoPs. The authors’ laboratory uses a gas chromatographic/negative ion chemical ionization mass spectrometric (GC/NICI MS) approach employing stable isotope dilution (Morrow and Roberts, 1999). For quantification purposes, the F2 -IsoP, 15-F2t -IsoP, and other F2 -IsoPs that co-elute with this compound are measured. Other investigators quantify different F2 -IsoP isomers, as discussed subsequently Oxidative Stress

Contributed by Erik S. Musiek and Jason D. Morrow Current Protocols in Toxicology (2005) 17.5.1-17.5.10 C 2005 by John Wiley & Sons, Inc. Copyright


Figure 17.5.1 Mechanism of formation of the F2 -isoprostanes. Four regioisomers are formed, each consisting of eight racemic diastereomers. Stereochemistry is not indicated.

F2 -Isoprostanes as Markers of Oxidative Stress

(Rokach et al., 1997). Several internal standards are available from commercial sources to quantify the IsoPs. These include [2 H4 ]15-F2t IsoP ([2 H4 ] 8-iso-PGF2∝ ) and [2 H4 ]-PGF2∝ . The advantages of mass spectrometry over other approaches include its high sensitivity and specificity, which yield quantitative results in the low picogram range. Its drawbacks are that it is labor intensive and requires considerable expenditures on equipment. Several alternative mass spectrometric assays have been developed by different investigators including FitzGerald and colleagues (Rokach et al., 1997; Pratico et al. 1998a). As with the authors’ assay, these methods require solid-phase extraction with a C18 column, TLC purification, and chemical derivatization. Furthermore, IsoPs are quantified using isotope dilution NICI GC/MS but the assay measures F2 -IsoP isomers other than 15-F2 -IsoP. These methods appear to be comparable to the authors’ in terms of utility. In addition, a number of liquid chromatographic MS methods for F2 -IsoPs have been recently developed; these require less sample preparation (Liang et al., 2000;

Bohnstedt et al., 2003), but the sensitivity and reliability of these for the analysis of IsoPs in complex biological samples is unknown. Alternative methods have been developed to quantify IsoPs with immunological approaches (Morrow et al., 1999a). Antibodies have been generated against 15-F2t -IsoP and at least three immunoassay kits are commercially available. A potential drawback of these methods is that limited information is currently available regarding their precision and accuracy. In addition, little data exist comparing IsoP levels determined by immunoassay to those determined by mass spectrometry. Analogous to immunological methods to quantify cyclooxygenase-derived PGs, it might be predicted that immunoassays for IsoPs will suffer from a lack of specificity (Roberts and Morrow, 2000). Furthermore, the sensitivity and/or specificity of these kits may vary substantially between manufacturers. However, while mass spectrometric methods of IsoP quantification are considered the “gold standard,” immunoassays have expanded research in this area because of their low cost and relative ease of use.



F2 -ISOPROSTANES AS AN INDEX OF OXIDANT STRESS Measurement of F2 -IsoPs In Vitro To demonstrate the utility of quantifying F2 -IsoPs as an index of oxidant stress, it is necessary to compare the formation of these compounds with other known indices of oxidant stress using established in vitro models of oxidant stress. The formation of F2 -IsoPs has been compared to malondialdehyde (MDA), one of the most commonly used measures of lipid peroxidation that utilizes Fe/ADP/ascorbate-induced peroxidation of rat liver microsomes (Longmire et al., 1994). Both F2 -IsoP and MDA (measured as thiobarbituric acid reacting substances) formation increased in parallel in a time-dependent manner and correlated with the loss of arachidonic acid and with increasing oxygen concentrations up to 21%. Although the formation of F2 -IsoPs correlated with other measures of lipid peroxidation in this in vitro model, the authors have reported that quantification of F2 -IsoPs is far superior to measurements of MDA as an index of lipid peroxidation in vivo (Longmire et al., 1994). It is hypothesized that the oxidation of low density lipoprotein (LDL) in vivo converts it to an atherogenic form, which is taken up by macrophages in the vessel wall. Subsequent activation of these cells likely plays an important role in the development and progression of atherosclerotic lesions in humans (Steinberg et al., 1989). Thus, the authors have performed studies examining the formation of F2 IsoPs in LDL that is oxidized to determine whether measurement of F2 -IsoPs esterified to lipoproteins may provide an approach to assess lipoprotein oxidation in vivo (Lynch et al., 1994). In these studies, either plasma lipids or purified LDL from humans was peroxidized with Cu2+ or the water-soluble oxidizing agent 2,2-azo-bis(2-amidinopropane) (AAPH; Lynch et al., 1994), and the formation of F2 -IsoPs was compared to other markers of lipid peroxidation including formation of cholesterol ester hydroperoxides, phospholipid hydroperoxides, loss of antioxidants, and changes in the electrophoretic mobility of LDL. In plasma oxidized with AAPH, increases in the formation of F2 -IsoP paralleled increases in lipid hydroperoxide formation and occurred only after depletion of the antioxidants ascorbate and ubiquinol-10. In purified LDL that was oxidized, formation of F2 -IsoPs again correlated with increases in lipid hydroperoxides and increases in the

electrophoretic mobility of LDL and occurred only after depletion of the antioxidants αtocopherol and ubiquinol-10. Similar findings have been reported by others (Gopaul et al., 1994; Pratico and FitzGerald, 1996). Taken together, these in vitro studies suggest that quantification of F2 -IsoP correlates with other established indices of oxidative stress, and serves as a useful marker of lipid peroxidation.

Measurement of F2 -IsoPs In Vivo It has been previously recognized that one of the greatest needs in the field of free radical research is a reliable non-invasive method to assess lipid peroxidation in vivo in humans (Roberts and Morrow, 2000). In this respect, most methods available to assess oxidant stress, which are adequate for in vitro purposes, have suffered from a lack of sensitivity and/or specificity or are unreliable when applied to complex biological fluids and tissues. However, a substantial body of evidence indicates that measurement of F2 -IsoPs in body fluids such as plasma provides a reliable approach to assess lipid peroxidation in vivo and represents a major advance in the ability to assess oxidative stress status in animals and humans (Morrow and Roberts, 1997; Roberts and Morrow, 2000). Normal levels of F2 -IsoPs have been defined in human biological fluids such as plasma and urine. (Liu et al., 1999; Morrow et al., 1999a; Morrow and Roberts, 1999). It is important to note that quantities of these compounds exceed those of cyclooxygenase-derived PGs by at least an order of magnitude, suggesting that IsoPs are a major pathway of arachidonic acid disposition. Further, levels of F2 -IsoPs are sufficient to be detected in every normal biological fluid that has been assayed including plasma, urine, bronchoalveolar lavage fluid, cerebrospinal fluid, and bile (Morrow et al., 1999a). The finding of significant levels of F2 -IsoPs in normal animal and human biological fluids and tissues indicates there is ongoing lipid peroxidation that is incompletely suppressed by antioxidant defenses, even in normal humans and animals, lending support to the hypothesis that the normal aging process is due to enhanced oxidant damage of relevant biological molecules over time. In this regard, previous studies have suggested that IsoP levels in normal mice and humans increase with age (Rokach et al. 1997; Roberts and Reckelhoff, 2001), although another report refutes this (Feillet-Coudray et al., 1999).

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An attractive possibility suggested by these findings is the measurement of F2 -IsoPs in urine as an index of systemic or whole body oxidant stress integrated over time. However, the measurement of free F2 -IsoPs in urine can be confounded by the potential contribution of local IsoP production in the kidney, although the extent to which this occurs is unclear (Morrow and Roberts, 1997; Morrow et al., 1999a; Roberts and Morrow, 2000). In light of this issue, the primary urinary metabolite of 15-F2t -IsoP has previously been identified to be 2,3-dinor-5,6-dihydro-15-F2t -IsoP, and a highly sensitive and accurate mass spectrometric assay to quantify this molecule has been developed (Roberts et al. 1996; Morrow et al., 1999b; Morales et al., 2001). Thus, the quantification of 2,3-dinor-5,6-dihydro-15F2t -IsoP may represent a truly non-invasive, time-integrated measurement of systemic oxidation status that can be applied to living subjects.

Formation of Isoprostanes in Animal Models of Oxidant Stress


F2 -IsoPs have also proven highly valuable in studying oxidative injury in vivo in many animal models of disease. Administration of carbon tetrachloride (CC14 ) intragastrically to rats is a well-established model of oxidative injury, causing severe free radical–induced damage to the liver and other organs. Esterified levels of F2 -IsoPs in liver tissue increase by 200fold within 1 hr of CCl4 treatment and subsequently decline over 24 hr (Morrow et al., 1992), while plasma free and lipid-esterified IsoP concentrations increased after liver levels up to 50-fold in a dose-dependent manner. Administration of the antioxidant lazaroid U78517 to CCl4 -treated animals significantly blunted the enhanced formation of F2 -IsoPs in this model (Morrow and Roberts, 1997). As a second example, F2 -IsoP formation has been employed to study the toxicity of diquat, a dipyridyl herbicide. Diaquat undergoes redox cycling in vivo, generating large amounts of the superoxide anion and causing hepatic and renal injury in rats. This effect is markedly augmented in animals deficient in selenium (Se), a trace element that is required for the enzymatic activities of glutathione peroxidase and other antioxidant proteins. To study whether lipid peroxidation occurs in this model, levels of F2 -IsoPs were quantified in plasma and tissues from Se-deficient rats following diquat administration. Se-deficient rats administered diquat showed 10- to 200-fold increases in plasma F2 -IsoPs, with the primary

sites of IsoP generation being the kidney and liver (Awad et al., 1994). Further studies disclosed that the extent of tissue injury and IsoP formation directly correlated with the degree of Se depletion (Burk et al., 1995). The measurement of F2 -IsoPs can also be employed to examine the oxidation status of transgenic animals. For instance, Pratico et al. (1998b) demonstrated that apolipoprotein E–deficient mice, which developed severe atherosclerotic disease, also showed marked increases in plasma F2 -IsoP levels. Dietary supplementation with the antioxidant αtocopherol prevented the increase in plasma F2 -IsoP levels, and reduced atherosclerosis in these animals. Increases in F2 -IsoPs have been observed in animal models of disease in nearly every organ. Using the brain as an example, increased F2 -IsoP levels have been described in mice subjected to a vast array of neurological insults, including amyloid precursor protein overexpression (Pratico et al. 2001), intracerebroventricular lipopolysacharide injection (Montine et al., 2002a), kainate-induce seizures (Patel et al., 2001; Montine et al., 2002a), and cerebral ischemic injury (Marin et al., 2000). Taken together, these studies suggest that quantification of F2 -IsoPs in animal models of oxidant injury represents an accurate method to assess lipid peroxidation in vivo.

F2 -ISOPROSTANE FORMATION IN HUMAN DISEASES From the above examples, measurement of IsoPs appears to be a reliable index of lipid peroxidation in vivo and thus potentially provides a tool to assess the role of oxidative stress in the pathophysiology of human disease. Elevations of IsoPs in human body fluids and tissues have been found in a diverse array of human disorders (Table 17.5.1). For purposes of this brief review, the authors have chosen to discuss IsoP formation in several human diseases in which their generation has been examined in some detail, namely atherosclerosis and Alzheimer’s disease. For a more detailed discussion of these and other disorders, the reader is referred to the cited references in Table 17.5.1.

Atherosclerotic Cardiovascular Disease The association between various risk factors for atherosclerosis and enhanced IsoP generation has been extensively explored, and it was found that IsoP formation is increased in humans with these risk factors. These data suggest that enhanced oxidant stress may play a role in the development of atherosclerosis, although the mechanisms involved have not been



Table 17.5.1 Disorders in Which F2 -IsoPs Have Implicated a Role for Free Radicals in Human Diseases

Disease

Reference

Acute chest syndrome of sickle cell disease

Klings et al., 2001

Acute cholestasis

Leo et al., 1997

Adult respiratory distress syndrome

Carpenter et al., 1998

Alcohol-induced liver injury

Aleynik et al., 1999

Allergic asthma

Dworski et al., 1999

Alzheimer’s disease

Montine et al., 1999a

Chronic obstructive lung disease

Pratico et al., 1999

Crohn’s disease

Wendland et al., 2001

Diabetes

Gopaul et al., 1995

Hemodialysis

Spittle et al., 2001

Hepatorenal syndrome

Morrow et al., 1993

Huntington’s disease

Montine et al., 1999c

Hypercholesterolemia

Davi et al., 1997

Hyperhomocysteinemia

Voutilainen et al., 1999

Interstitial lung disease

Montuschi et al., 1998

Ischemia/reperfusion injury

Reilly et al., 1997

Muscular side effects of statins

Sinzinger et al., 2001

Pulmonary hypertension

Cracowski et al., 2001

Rhabdomyolysis renal injury

Holt et al., 1999

Rheumatic inflammatory response

Basu et al., 2001

Scleroderma

Stein et al., 1996

Sepsis

Ekmekcioglu et al., 2002

Smoking

Morrow et al., 1995

elucidated. Although trials with antioxidants have generally failed to show benefit in the prevention of heart disease in humans, there is still considerable evidence to support the hypothesis that oxidative stress is intimately involved in atherosclerosis. A link between cigarette smoking and the risk of cardiovascular disease is well established (Kannel, 1981). However, the underlying mechanism(s) for this effect is not fully understood. The gaseous phase of cigarette smoke contains a number of oxidants and exposure of LDL to the gaseous phase of cigarette smoke in vitro induces oxidation of the LDL lipids (Frei et al., 1991). Thus, the hypothesis that smoking induces an oxidative stress by examining F2 -IsoP levels in plasma from smokers was explored. Ten individuals who smoked heavily (>30 cigarettes/day) and ten age and sex matched non-smoking normal volunteers were studied (Morrow et al., 1995).

Plasma concentrations of free and esterified F2 -IsoPs were significantly elevated in the smokers compared to the non-smokers (p = 0.02 and p = 0.03, respectively). In all subjects, levels of F2 -IsoPs both free in the circulation and esterified to plasma lipoproteins were significantly reduced following 2 weeks of abstinence from smoking (p = 0.03 and p = 0.02, respectively). The occurrence of enhanced formation of IsoPs in smokers has also subsequently been confirmed in studies by other groups (Reilly et al., 1996). Collectively, these findings suggest strongly that smoking causes an oxidative stress and the observation that smokers have elevated levels of F2 -IsoPs esterified in plasma lipids also supports the hypothesis that the link between smoking and risk of cardiovascular disease may be attributed to enhanced oxidation of lipoproteins. It has been well established that patients with hypercholesterolemia have an increased

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risk for the development of atherosclerosis. Thus, it was of interest to determine whether levels of F2 -IsoPs are increased in patients with this condition. Levels of F2 -IsoPs esterified in plasma lipids were determined in patients with polygenic hypercholesterolemia (Roberts and Morrow, 1999). Levels in patients with hypercholesterolemia were found to be significantly increased by a mean of 3.4-fold (range 1.7to 7.5-fold) above levels measured in normal controls (p < 0.001). Interestingly, in these patients, there was no correlation between levels of F2 -IsoPs and serum cholesterol, triglycerides or LDL-cholesterol. In addition, plasma arachidonic acid content was measured in these patients and normal controls. Again, no correlation between IsoP and arachidonate levels was found. Thus, these data suggest that the finding of high levels of F2 -IsoPs in patients with hypercholesterolemia is not due simply to the presence of more lipid, i.e., arachidonic acid substrate. Rather, it is suggested that hypercholesterolemia is associated with enhanced oxidative stress. The underlying basis for this observation, however, remains unclear. Interestingly, a report also found that the urinary excretion of F2 -IsoPs was also increased in patients with Type II hypercholesterolemia by a mean of 2.5-fold, which was suppressed by ∼60% with vitamin E treatment (600 mg/day; Davi et al., 1997). Patients with diabetes are known to have an increased incidence of atherosclerotic vascular disease. Furthermore, the formation of F2 -IsoPs has been found to be induced in vascular smooth muscle cells by hyperglycemia (Natarajan et al., 1996). Thus, a link between enhanced oxidative stress in vivo and diabetes has been explored. Gopaul et al. (1995) reported a mean 3.3-fold increase in free F2 -IsoPs in plasma from diabetic patients compared to non-diabetic healthy control subjects. A later report demonstrated that plasma F2 -IsoP levels increased significantly within 90 min during acute hyperglycemia in type 2 diabetic patients (Sampson et al., 2002). In addition, Davi et al. (1999) reported that elevated urinary F2 -IsoP levels in diabetic patients could be suppressed by vitamin E supplementation or glycemic control. High plasma levels of homocysteine are an independent risk factor for cardiovascular disease (Boushey et al., 1995). The mechanism by which hyperhomocysteinemia induces atherosclerosis is not fully understood but promotion of LDL oxidation has been sug-

gested. The relationship between total plasma concentrations of homocysteine and F2 -IsoPs in 100 Finnish male participants in the Antioxidant Supplementation in Atherosclerosis Prevention study has been explored (Voutilainen et al., 1999). The mean plasma total homocysteine and F2 -IsoP concentrations were 11.1 µmol/liter and 29.6 ng/liter, respectively. The simple correlation coefficient for the association between plasma concentrations of homocysteine and F2 -IsoPs was 0.40 (p < 0.0001). Plasma concentrations of F2 -IsoPs increased linearly across quintiles of homocysteine levels. The finding of a positive correlation between plasma concentrations of F2 -IsoPs and homocysteine supports the suggestion that the mechanism underlying the link between high homocysteine levels and risk for cardiovascular disease may be attributed to enhanced lipid peroxidation. In accordance with the LDL oxidation hypothesis of atherosclerosis, levels of F2 -IsoPs should be higher in atherosclerotic plaques than in normal vascular tissue. To address this issue, levels of F2 -IsoPs were measured in fresh advanced atherosclerotic plaque tissue removed during arterial thrombarterectomy (n = 10) and compared with levels measured in normal human umbilical veins removed from the placenta immediately after delivery (n = 10) (Gniwotta et al., 1997). Levels of F2 -IsoPs esterified in vascular tissue normalized to both wet weight and dry weight were significantly higher in atherosclerotic plaques compared to normal vascular tissue. When the data was normalized to arachidonic acid content, the F2 -IsoP/arachidonic acid ratio was about four-fold higher than the ratio in normal vascular tissue (p = 0.009). This finding indicates that unsaturated fatty acids in atherosclerotic plaques are more extensively oxidized than lipids in normal vascular tissue. These observations are also in accord with data from FitzGerald and colleagues who have shown increased amounts of F2 -IsoPs in human atherosclerotic lesions including the localization of F2 -IsoPs in atherosclerotic plaque tissue to foam cells and vascular smooth muscle cells (Pratico et al., 1997).

Alzheimer’s Disease Oxidative stress has been implicated in the pathogenesis of numerous neurodegenerative conditions, including Alzheimer’s Disease (AD). Regional increases in oxidative damage and lipid peroxidation have been described in brain tissue obtained post mortem from patients with AD (Markesbery, 1997). Similarly,



F2 -IsoP levels are significantly elevated in affected regions of post-mortem brain samples from AD patients as compared to controls (Reich et al., 2001a). However, an objective index of oxidative damage associated with AD that can be assessed in living patients is lacking. Such a biomarker could be vital for understanding the role of oxidative damage in AD patients by permitting repeated evaluation of disease progression or responses to therapeutic interventions. Toward such a goal, the authors obtained post-mortem ventricular fluid from eleven patients with a pathological diagnosis of AD and eleven control patients, in order to evaluate F2 -IsoP levels in cerebrospinal fluid (CSF; Montine et al., 1998). All subjects participated in a rapid autopsy protocol such that fluid was collected within 3 hr of death. F2 IsoP levels were significantly increased in ventricular fluid from AD patients (72 ± 7 pg /ml, mean ± S.E.M.) compared to CSF from control individuals (46 ± 4 pg/ml, p < 0.01), and correlations were identified between increases in IsoP levels and higher Braak stage and decreased brain weight, two indices of AD severity. In a larger study, the authors have shown that CSF F2 -IsoP level correlates with the extent of pathological neurodegeneration but not with density of neuritic plaques or neurofibrillary tangles (Montine et al., 1999a; Reich et al., 2001b). Subsequently, the authors undertook a study to examine CSF F2 -IsoP levels in living patients with probable AD (Montine et al., 1999b). CSF was obtained from the lumbar cistern in 27 patients with AD and 25 controls without neurodegenerative disorders matched for age and gender. In keeping with postmortem studies, lumbar CSF levels of F2 IsoPs were significantly increased (31.0 ± 2.6 pg/ml) in AD patients compared to control subjects (22.9 ± 1.0 pg/ml, p < 0.05). Pratico et al. (2000) have also observed increased F2 IsoPs in CSF of patients with probable AD, as well as in patients with mild cognitive impairment (MCI), a condition which precedes symptomatic dementia in AD (Pratico et al., 2002). However, this group also reports increased F2 -IsoPs in plasma and urine of both MCI and AD patients, though the authors’ laboratory and others have not been able to detect these changes in peripheral F2 -IsoPs (Montine et al., 2002b; Bohnstedt et al., 2003). Taken together, these studies suggest that quantification of IsoPs in cerebrospinal fluid of patients with Alzheimer’s disease may be of use as an intra vitum index of disease progression or as a tool to monitor response to therapy.

CONCLUSIONS The discovery of IsoPs as products of nonenzymatic lipid peroxidation has been a major breakthrough regarding the quantification of oxidant stress in vivo. The quantification of these molecules has opened up new areas of investigation regarding the role of free radicals in human physiology and pathophysiology, and appears to be the most useful tool currently available to explore the role of free radicals in the pathogenesis of human disease. Although considerable information has been obtained since the initial discovery of IsoPs, much remains to be understood about the role of these molecules as markers of oxidant stress. It is anticipated that additional research in this area will continue to provide important insights into the role of oxidative stress in human disease.

LITERATURE CITED Aleynik, S.I., Leo, M.A., Aleynik, M.K., and Lieber, C.S. 1999. Increased circulating products of lipid peroxidation in patients with alcoholic liver disease. Alcohol Clin. Exp. Res. 22:192-196. Awad, J.A., Morrow, J.D., Hill, K.E., Roberts, L.J., and Burk, R.F. 1994. Detection and localization of lipid peroxidation in selenium- and vitaminE deficient rats using F2 -isoprostanes, J. Nutr. 124:810-816. Basu, S., Whiteman, M., Mattey, D.L., and Halliwell, B. 2001. Raised levels of F(2)isoprostanes and prostaglandin F(2alpha) in different rheumatic diseases. Ann. Rheum. Dis. 60:627-631. Bohnstedt, K.C., Karlberg, B., Wahlund, L.O., Jonhagen, M.E., Basun, H., and Schmidt, S. 2003. Determination of isoprostanes in urine samples from Alzheimer patients using porous graphitic carbon liquid chromatography-tandem mass spectrometry. J. Chromatogr. B. 796:1119. Boushey, C.J., Beresford, S.A., Omenn, G.S., and Motulsky, A.G. 1995. A quantitative assessment of plasma homocysteine as a risk factor for vascular disease. Probable benefits of increasing folic acid intakes. J. Am. Med. Assoc. 274:10491057. Burk, R.F., Hill, K.E., Awad, J.A., Morrow, J.D., Kato, T., Cockell, K.A., and Lyons, P.R. 1995. Pathogenesis of diquat-induced liver necrosis in selenium-deficient rats: Assessment of the roles of lipid peroxidation and selenoprotein P. Hepatology 21:561-569. Carpenter, C.T., Price, P.V., and Christman, B.W. 1998. Exhaled breath condensate isoprostanes are elevated in patients with acute lung injury or ARDS. Chest 114:1653-1659. Chen, Y., Morrow, J.D., and Roberts, L.J. 2nd 1999. Formation of reactive cyclopentenone compounds in vivo as products of the isoprostane pathway. J. Biol. Chem. 274:10863-10868.

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Cracowski, J.L., Cracowski, C., Bessard, G., Pepin, J.L., Bessard, J., Schwebel, C., StankeLabesque, F., and Pison, C. 2001. Increased lipid peroxidation in patients with pulmonary hypertension. Am. J. Resp. Crit. Care Med. 164:10381042. Davi, G., Alessandrini, P, Mezzetti, A., Minotti, G., Bucciarelli, T., Costantini, F., Cipollone, F., Bon, G.B., Ciabattoni, G., and Patrono, C. 1997. In vivo formation of 8-Epi-prostaglandin F2 alpha is increased in hypercholesterolemia. Atheroscler. Thromb. Vasc. Biol. 117:32303235. Davi, G., Ciabattoni, G., Consoli, A., Mezzetti, A., Falco, A., Santarone, S., Pennese, E., Vitacolonna, E., Bucciarelli, T., Costantini, F., Capani, F., and Patrono, C. 1999. In vivo formation of 8-iso-prostaglandin F2alpha and platelet activation in diabetes mellitus: Effects of improved metabolic control and vitamin E supplementation. Circulation 99:224-229. Dworski, R., Murray, J.J., Roberts, L.J., Oates, J.A., Morrow, J.D., Fisher, L., and Sheller, J.R. 1999. Allergen-induced synthesis of F(2)-isoprostanes in atopic asthmatics. Evidence for oxidant stress. Am. J. Respir. Crit. Care Med. 160:1947-1951. Ekmekcioglu, C., Schweiger, B., Strauss-Blasche, G., Mundigler, G., Siostrzonek, P., and Marktl, W. 2002. Urinary excretion of 8-iso-PGF(2 alpha) in three patients during sepsis, recovery and state of health. Prostaglandins, Leukot. Essent. Fatty Acids 66:441-442. Fam, S.S. and Morrow, J.D. 2003. The isoprostanes: Unique products of arachidonic acid oxidation—A review. Curr. Med. Chem. 10:1723-1740. Feillet-Coudray, C., Tourtauchaux, R., Niculescu, N., Rock, E., Tauveron, I., Alexandre-Gouabau, M., Rayssiguier, R., Jalenques, Y.I., and Mazur, A. 1999. Plasma levels of 8-epi-PGF2α, an in vivo marker of oxidative stress, are not affected by aging or Alzheimer’s disease. Free Rad. Biol. Med. 27:463-469. Frei, B., Forte, T.M., Ames, B.N., and Cross, C.E. 1991. Gas phase oxidants of cigarette smoke induce lipid peroxidation and changes in lipoprotein properties in human blood plasma. Protective effects of ascorbic acid. Biochem. J. 277:133-138. Gniwotta, C., Morrow, J.D., Roberts, L.J., and Kuhn, H. 1997. Prostaglandin F2-like compounds, F2-isoprostanes, are present in increased amounts in human atherosclerotic lesions. Arterioscler. Thromb. Vasc. Biol. 17:32363241. Gopaul, N.K., Nourooz-Zadeh, J., Malle, A.I., and Anggard, E.E. 1994. Formation of F2 isoprostanes during aortic endothelial cellmediated oxidation of low density lipoprotein. FEBS Lett. 348:297-300.


Gopaul, N.K., Anggard, E.E., Mallet, A.I., Beteridge, D.J., Wolff, S.P., and Nourooz-Zadey, J. 1995. Plasma 8-epi-PGF2 alpha levels are elevated in individuals with non-insulin dependent diabetes mellitus. FEBS Lett. 368:225-229.

Halliwell, B. and Grootveld, M. 1987. The measurement of free radical reactions in humans. FEBS Lett. 213:9-14. Holt, S., Reeder, B., Wilson, M., Harvey, S., Morrow, J.D., Roberts, L.J. II, and Moore, K. 1999. Increased lipid peroxidation in patients with rhabdomyolysis. Lancet 353:1241. Kannel, W.B. 1981. Update on the role of cigarette smoking in coronary artery disease. Am. Heart J. 101:319-328. Klings, E.S., Christman, B.W., McClung, J., Stucchi, A.F., McMahon, L., Brauer, M., and Farber, H.W. 2001. Increased F2 isoprostanes in the acute chest syndrome of sickle cell disease as a marker of oxidative stress. Am. J. Resp. Crit. Care Med. 164:1248-1252. Leo, M.A., Aleynik, S.I., Siegel, J.H., Kasmin, F.E., Aleynik, M.K., and Lieber, C.S. 1997. F2isoprostane and 4-hydroxynonenal excretion in human bile of patients with biliary tract and pancreatic disorders. Am. J. Gastroenterol. 92:20692072. Liang, Y., Wei, P., Duke, R.W., Reaven, P.D., Harman, S.M., Cutler, R.G., and Heward, C.B. 2000. Quantification of 8-iso-prostaglandinF(2alpha) and 2,3-dinor-8-iso-prostaglandinF(2alpha) in human urine using liquid chromatography-tandem mass spectrometry. Free Radic. Biol. Med. 34:409-418. Liu, T., Stern, A., Roberts, L.J., and Morrow, J.D. 1999. The isoprostanes: Novel prostaglandinlike products of the free radical-catalyzed peroxidation of arachidonic acid. J. Biomed. Sci. 6:226-235. Longmire, A.W., Swift, L.L., Roberts, L.J., Awad, J.A., Burk, R.F., and Morrow, J.D. 1994. Effect of oxygen tension on the generation of F2 -isoprostanes and malondialdehyde in peroxidizing rat liver microsomes. Biochem. Pharm. 47:1173-1177. Lynch, S.M., Morrow, J.D., Roberts, L.J., and Frei, B. 1994. Formation of non-cyclooxygenasederived prostanoids (F2 -isoprostanes) in plasma and low density lipoprotein exposed to oxidative stress in vitro. J. Clin. Invest. 93:998-1004. Marin, J.G., Cornet, S., Spinnewyn, B., DemerlePallardy, C., Auguet, M., and Chabrier, P.E. 2000. BN 80933 inhibits F2-isoprostane elevation in focal cerebral ischaemia and hypoxic neuronal cultures. Neuroreport. 11:1357-1360. Markesbery, W.R. 1997. Oxidative stress hypothesis in Alzheimer’s disease. Free Rad. Biol. Med. 23:137-147. Montine, T.J., Markesbery, W.R., Morrow, J.D., and Roberts, L.J. 1998. Cerebrospinal fluid F2isoprostane levels are increased in Alzheimer’s disease. Ann. Neurol. 44:410-413. Montine, T.J., Markesbery, W.R., Zackert, W., Sanchez, S.C., Roberts, L.J. 2nd, and Morrow, J.D. 1999a. The magnitude of brain lipid peroxidation correlates with the extent of degeneration but not with density of neuritic plaques or neurofibrillary tangles or with APOE genotype



in Alzheimer’s disease patients. Am. J. Pathol. 155:863-868. Montine, T.J., Beal, M.F., Cudkowicz, M.E., O’Donnell, H., Margolin, R.A., McFarland, L., Bachrach, A.F., Zackert, W.E., Roberts, L.J., and Morrow, J.D. 1999b. Increased CSF F2isoprostane concentration in probable AD. Neurology. 52:562. Montine, T.J., Beal, M.F., Robertson, D., Cudkowicz, M.E., Biaggioni, I., Brown, R.H., O’Donnel, H., Zackert, W.E., Roberts, L.J., and Morrow, J.D. 1999c. Cerebrospinal fluid F2-isoprostanes are elevated in Huntington’s disease. Neurology 52:1104-1105. Montine, T.J., Milatovic, D., Gupta, R.C., ValyiNagy, T., Morrow, J.D., and Breyer, R.M. 2002a. Neuronal oxidative damage from activated innate immunity is EP2 receptor-dependent. J. Neurochem. 83:463-470. Montine, T.J., Quinn, J.F., Milatovic, D., Silbert, L.C., Dang, T., Sanchez, S., Terry, E., Roberts, L.J. 2nd, Kaye, J.A., and Morrow, J.D. 2002b. Peripheral F2-isoprostanes and F4neuroprostanes are not increased in Alzheimer’s disease. Ann. Neurol. 52:175-179. Montuschi, P., Ciabattoni, G., Paredi, P., Pantelidis, P., DuBois, R.M., Kharitoniv, S.A., and Barnes, P.J. 1998. 8-Isoprostane as a biomarker of oxidative stress in interstitial lung diseases. Am. J. Resp. Crit. Care Med. 158:1524-1527. Morales, C.R., Terry, E.S., Zackert, W.E., Montine, T.J., and Morrow, J.D. 2001. Improved assay for the quantification of the isoprostane 15-F2t isoprostane (8-iso-PGF2α ) by a stable isotope dilution mass spectrometric assay. Clin. Chim. Acta 314:93-99. Morrow, J.D. and Roberts, L.J. 1997. The isoprostanes: Unique bioactive products of lipid peroxidation. Prog. Lipid Res. 36:1-21. Morrow, J.D. and Roberts, L.J. 1999. Mass spectrometric quantification of F2 -isoprostanes in biological fluids and tissues as measure of oxidant stress. Meth. Enzymol. 300:3-12. Morrow, J.D., Hill, K.E., Burk, R.F., Nammour, T.M., Badr, K.F., and Roberts, L.J. 1990a. A series of prostaglandin F2 -like compounds are produced in vivo in humans by a noncyclooxygenase, free radical-catalyzed mechanism. Proc. Natl. Acad. Sci. U.S.A. 87:93839397. Morrow, J.D., Harris, T.M., and Roberts, L.J. 1990b. Noncyclooxygenase oxidative formation of a series of novel prostaglandins: Analytical ramifications for measurement of eicosanoids. Analyt. Biochem. 184:1-10. Morrow, J.D., Awad, J.A., Kato, T., Takahashi, K., Badr, K.F., Roberts, L.J., and Burk, R.F. 1992. Formation of novel non-cyclooxygenasederived prostanoids (F2 -isoprostanes) in carbon tetrachloride hepatotoxicity. J. Clin. Invest. 90:2502-2507. Morrow, J.D., Moore, K.P., Awad, J.A., Ravenscraft, M.D., Marini, G., Badr, K.F., Williams R., and Roberts, L.J. 1993. Marked overproduc-

tion of non-cyclooxygenase derived prostanoids (F2-isoprostanes) in the hepatorenal syndrome. J. Lipid Mediators 6:417-420. Morrow, J.D, Frei, B., Longmire, A.W., Gaziano, M., Lynch, S.M., Shyr, Y., Strauss, W.E., Oates, J.A., and Roberts, L.J. 1995. Increase in circulating products of lipid peroxidation (F2isoprostanes) in smokers. Smoking as a cause of oxidative damage. N. Engl. J. Med. 332:11981203. Morrow, J.D., Chen, Y., Brame, C.J., Yang, J., Sanchez, S.C., Xu, J., Zackert, W.E., Awad, J.A., and Roberts, L.J. 1999a. The isoprostanes: Unique prostaglandin-like products of free radical-initiated lipid peroxidation. Drug Metab. Rev. 31:117-139. Morrow, J.D., Zackert, W.E., Yang, J.P., Kurhts, E.H., Callawaert, D., Dworski, R., Kanai, K., Taber, D., Moore, K., Oates, J.A., and Roberts, L.J. 1999b. Quantification of the major urinary metabolite of 15-F2t -isoprostane (8-iso-PGF2α ) by a stable isotope dilution mass spectrometric assay. Analyt. Biochem. 269:326-331. Natarajan, R., Lanting, L., Gonzales, N., and Nadler, J. 1996. Formation of an F2-isoprostane in vascular smooth muscle cells by elevated glucose and growth factors. Am. J. Physiol. 271:H159-165. Patel, M., Liang, L.P., and Roberts, L.J. 2nd. 2001. Enhanced hippocampal F2-isoprostane formation following kainate-induced seizures. J. Neurochem. 79:1065-1069. Pratico, D. and FitzGerald, G.A. 1996. Generation of 8-epi-prostaglandin F2α by human monocytes. Discriminate production by reactive oxygen species and prostaglandin endoperoxide synthase-2. J. Biol. Chem. 271:89198924. Pratico, D., Iuliano, L., Mauriello, A., Spagnoli, L., Lawson, J.A., Rokach, J., Maclouf, J., Violi, F., and FitzGerald, G.A. 1997. Localization of distinct F2-isoprostanes in human atherosclerotic lesions. J. Clin. Invest. 100:2028-2034. Pratico, D., Barry, O.P., Lawson, J.A., Adiyaman, M., Hwang, S.W., Khanapure, S.P., Iuliano, L., Rokach, J., and FitzGerald, G.A. 1998a. IPF2alpha-I: An index of lipid peroxidation in humans. Proc. Natl. Acad. Sci U.S.A. 95:34493454. Pratico, D., Tangirala, R.K., Rader, D.J., Rokach, J., and FitzGerald, G.A. 1998b. Vitamin E suppresses isoprostane generation in vivo and reduces atherosclerosis in ApoE-deficient mice. Nat. Med. 4:1189-1192. Pratico, D., Basili, S., Vieri, M., Cordova, C., Violi, V., and FitzGerald, G.A. 1999. Chronic obstructive pulmonary disease is associated with an increase in urinary levels of isoprostane F2alphaIII, an index of oxidant stress. Am. J. Resp. Crit. Care Med. 158:1709-1714. Pratico, D., Clark, C.M., Lee, V.M., Trojanowski, J.Q., Rokach, J., and FitzGerald, G.A. 2000. Increased 8,12-iso-iPF2alpha-VI in Alzheimer’s disease: Correlation of a noninvasive index of

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lipid peroxidation with disease severity. Ann. Neurol. 48:809-812.

8-iso-prostaglandin F2α in humans. J. Biol. Chem. 271:20617-20620.

Pratico, D., Uryu, K., Leight, S., Trojanoswki, J.Q., and Lee, V.M. 2001. Increased lipid peroxidation precedes amyloid plaque formation in an animal model of Alzheimer amyloidosis. J. Neurosci. 21:4183-4187.

Rokach, J., Khanapure, S.P., Hwang, S.-W., Adiyaman, M., Lawson, J.A., and FitzGerald, G.A. 1997. The isoprostanes: A perspective. Prostaglandins 54:823-851.

Pratico, D., Clark, C.M., Liun, F., Rokach, J., Lee, V.Y., and Trojanowski, J.Q. 2002. Increase of brain oxidative stress in mild cognitive impairment: A possible predictor of Alzheimer disease. Arch. Neurol. 59:972-976. Reich, E.E., Markesbery, W.R., Roberts, L.J. 2nd, Swift, L.L., Morrow, J.D., and Montine, T.J. 2001a. Brain regional quantification of F-ring and D-/E-ring isoprostanes and neuroprostanes in Alzheimer’s disease. Am. J. Pathol. 158:293297. Reich, E.E., Markesbery, W.R., Roberts, L.J., Zackert, W.E., Swift, L.L., Morrow, J.D, and Montine, T.J. 2001b. Quantification of F-ring and D-/E-ring isoprostanes and neuroprostanes in Alzheimer’s disease. Adv. Exp. Med. Biol. 500:253-256. Reilly, M., Delanty, N., Lawson, J.A., and FitzGerald, G.A. 1996. Modulation of oxidant stress in vivo in chronic cigarette smokers. Circulation 94:19-25. Reilly, M.P., Delanty, M., Roy, L., Rokach, J., Callaghan, P.O., Crean, P., Lawson, J.A., and FitzGerald, G.A. 1997. Increased formation of the isoprostanes IPF2alpha-I and 8-epiprostaglandin F2alpha in acute coronary angioplasty: Evidence for oxidant stress during coronary reperfusion in humans. Circulation 96:3314-3320. Roberts, L.J. and Morrow, J.D. 1999. Isoprostanes as markers of lipid peroxidation in atherosclerosis. In Molecular and Cellular Basis of Inflammation. (C.N. Serhan and P.A. Ward, eds.) pp. 141. Humana Press, Totowa, N.J. Roberts, L.J. II and Morrow, J.D. 2000. Measurement of F2-Isoprostanes as an index of oxidative stress in vivo. Free Radic. Biol. Med. 28:505513. Roberts, L.J. and Reckelhoff, J.F. 2001. Measurement of F(2)-isoprostanes unveils profound oxidative stress in aged rats. Biochem. Biophys. Res. Commun. 287:254-256. Roberts, L.J., Moore, K.P., Zackert, W.E., Oates, J.A., and Morrow, J.D. 1996. Identification of the major urinary metabolite of the F2 -isoprostane

Sampson, M.J., Gopaul, N., Davies, I.R., Hughes, D.A., and Carrier, M.J. 2002. Plasma F2 isoprostanes: Direct evidence of increased free radical damage during acute hyperglycemia in type 2 diabetes. Diabetes Care 25:537-541. Sinzinger, H., Lupatelli, G., Chehne, F., Oguogho, A., and Furberg, C.D. 2001. Isoprostane 8-epiPGF2alpha is frequently increased in patients with muscle pain and/or CK-elevation after HMG-Co-enzyme-A-reductase inhibitor therapy. J. Clin. Pharm. Ther. 26:303-310. Spittle, M.A., Hoenich, N.A., Handelman, G.J., Adhikarla, R., Homel, P., and Levin, N.W. 2001. Oxidative stress and inflammation in hemodialysis patients. Am. J. Kidney Dis. 38:14081413. Stein, C.M., Tanner, S.B., Awad, J.A., Roberts, L.J. II, and Morrow, J.D. 1996. Evidence of free radical-mediated injury (isoprostane overproduction) in scleroderma. Arth. Rheum. 39:11461150. Steinberg, D., Parthasarathy, S., Carew, T.E., Khoo, J.C., and Witztum, J.L. 1989. Beyond cholesterol: Modifications of low density lipoprotein that increase its atherogenicity. N. Engl. J. Med. 86:915-924. Taber, D.F., Morrow, J.D., and Roberts, L.J. 1997. A nomenclature system for the isoprostanes. Prostaglandins 53:63-67. Voutilainen, S., Morrow, J.D., Roberts, L.J., Alfthan, G., Alho, H., Nyyssonen, K., and Salonen, J.T. 1999. Enhanced in vivo lipid peroxidation at elevated plasma total homocysteine levels. Arterioscler. Thromb. Vasc. Biol. 19:12631266. Wendland, B.E., Aghdassi, E., Tam, C., Carrier, J., Steinhart, A.H., Wolman, S.L., Baron, D., and Allard, J.P. 2001. Lipid peroxidation and plasma antioxidant micronutrients in Crohn disease. Am. J. Clin. Nutr. 74:259-264.

Contributed by Erik S. Musiek and Jason D. Morrow Vanderbilt University School of Medicine Nashville, Tennessee




Quantification of F2-Isoprostanes by Gas Chromatography/Mass Spectrometry as a Measure of Oxidant Stress

UNIT 17.6

This unit outlines methods to assess lipid peroxidation associated with oxidant injury in vivo by quantifying concentrations of either esterified or free F2 -isoprostanes (F2 IsoPs). Quantification of these compounds overcomes many of the shortcomings associated with other assays of oxidant stress, especially in humans in vivo. Thus, measurement of F2 -IsoPs likely represents an important advancement in the ability to assess the role of oxidative stress in human disease (Fam and Morrow, 2003). The techniques employed for the analysis of F2 -IsoPs from biological sources that are detailed herein utilize mass spectrometric approaches. Procedures are outlined for the analysis of both F2 -IsoPs esterified to tissue phospholipids (see Basic Protocol 1) and free F2 -IsoPs (see Basic Protocol 2). In addition, quantification of F2 -IsoPs esterified in plasma lipids is also detailed (see Alternate Protocol). NOTE: All inorganic solvents should be high purity HPLC or GC grade. All reagents/ chemicals should be ACS certified or equivalent grade.

QUANTIFICATION OF F2 -ISOPROSTANES IN TISSUE LIPIDS F2 -IsoPs from biological sources can only be quantified as free compounds using gas chromatography (GC)/mass spectrometry (MS) (Morrow and Roberts, 1999) Thus, to measure levels of these compounds esterified to phospholipids, the phospholipids are first extracted from the tissue sample and subjected to alkaline hydrolysis to release free F2 -IsoPs. Free F2 -IsoPs are then quantified using the same procedure for the measurement of free compounds in biological fluids (see Alternate Protocol). This protocol outlines the extraction and hydrolysis of F2 -IsoPs from tissue lipids.

BASIC PROTOCOL 1

The assay described herein is applicable to all animal and human tissues and cell samples. Preparation of phospholipid-containing biological fluids, such as plasma, differs slightly (see Alternate Protocol). To measure free F2 -IsoPs from low-protein fluids (such as urine or low-serum cell medium) that do not require lipid extraction, see Basic Protocol 2.

Materials Fresh or frozen tissue Folch solution (see recipe) Butylated hydroxytoluene (BHT; Aldrich) Nitrogen or argon source 0.9% (w/v) aqueous sodium chloride (NaCl; high purity; EMD Biosciences) prepared in ultrapure water Methanol (high-quality; Burdick and Jackson or VWR Scientific) containing 0.005% (v/v) BHT 15% (w/v) aqueous potassium hydroxide (KOH pellets; high purity; EMD Biosciences) prepared in ultrapure water 1 N HCl (American Chemical Society certified or equivalent grade) 50-ml conical, polypropylene centrifuge tubes Blade homogenizer (PTA 10S generator, Brinkman Instruments) Table-top centrifuge Analytical evaporation unit (Organomation Associates) 37◦ C water bath Oxidative Stress Contributed by Erik S. Musiek and Jason D. Morrow Current Protocols in Toxicology (2005) 17.6.1-17.6.9 C 2005 by John Wiley & Sons, Inc. Copyright


1. Weigh out 0.05 to 1 g of either fresh tissue or tissue frozen at −70◦ C. If the sample weighs 1 ng/ml), 0.2 ml of urine was found to be more than adequate to quantify urinary F2 -IsoPs (Morrow et al., 2001). The following assay procedure that is described and summarized in Figure 17.6.1 is the method used for analysis of free F2 -IsoPs in plasma but is equally adaptable to other biological fluids and hydrolyzed lipid extracts of tissues (see Basic Protocol 1).

BASIC PROTOCOL 2

Materials Fluid or tissue extract, hydrolyzed 1 N HCl [2 H4 ] 15-F2 -IsoP (8-iso-PGF2α ) internal standard (Cayman Chemical) C18 Sep-Pak column (Waters) Methanol (high purity; Burdick and Jackson, VWR Scientific) pH 3 water (ultrapure filtered, adjusted to pH 3 with ACS-grade HCl) Heptane (high purity; Burdick and Jackson, VWR Scientific) Ethyl acetate (high purity; Burdick and Jackson, VWR Scientific)

Figure 17.6.1 Outline of the procedures used for the extraction, purification, derivatization, and mass spectrometric analysis of F2 -IsoPs from biological sources.

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Na2 SO4 (anhydrous) Silica Sep-Pak column (Waters) 10% (v/v) pentafluorobenzyl bromide (PFBB; Sigma-Aldrich) in acetonitrile 10% (v/v) N,N -Diisopropylethylamine (DIPE; Sigma-Aldrich) in acetonitrile Chloroform with ethanol (high purity; Burdick and Jackson, VWR Scientific) Ethanol (high purity; Burdick and Jackson, VWR Scientific) PGF2α methyl ester (Cayman Chemical) 10% phosphomolybdic acid (Sigma) solution in ethanol N,O-bis(trimethylsilyl)trifluoroacetamide (BSTFA; Supelco) Dimethylformamide (DMF; Aldrich) Undecane (Aldrich) 10-ml disposable plastic syringes 20-ml glass scintillation vials 5-ml Reacti-Vials (Pierce Scientific) Analytical evaporation unit (Organomation Associates) 37◦ C water bath Glass TLC tank and TLC paper TLC plates (LK6B silica; Whatman) 90◦ C oven Dessicator 1.5-ml microcentrifuge tubes Gas chromatograph/mass spectrometer with capabilities for negative ion chemical ionization (NICI) mass spectrometry (model 6890N, Agilent Technologies or equivalent) Capillary gas chromatography column (DB-1701, Fisons) Prepare sample and apply on column 1. Acidify hydrolyzed extract to pH 3 with 1 N HCl. 2. Add 1 to 5 nmol of deuterated [2 H4 ] 15-F2 -IsoP (8-iso-PGF2α ) internal standard and vortex. 3. Precondition one C18 Sep-Pak column connected to a 10-ml disposable plastic syringe per sample with 5 ml methanol and then 7 ml pH 3 water. 4. Apply acidified sample mixture to the preconditioned Sep-Pak column. Care must be taken when applying the sample to the column to avoid loss of IsoPs. The sample should be pushed through the column at ∼1 to 2 ml/min, such that individual drops emerge from the Sep-Pak. Slightly more vigorous fluid flow is acceptable during the subsequent wash and elution steps, though a steady stream is not recommended.

Wash and elute column 5. Wash column first with 10 ml pH 3 water and then with 10 ml heptane. 6. Elute F2 -IsoPs from column with 10 ml of 50:50 (v/v) ethyl acetate/heptane into a 20-ml glass scintillation vial. 7. Add 5 g anhydrous Na2 SO4 to the vial and swirl gently. This step removes residual water from the eluate.

8. Precondition silica Sep-Pak column with 5 ml ethyl acetate. 9. Apply eluate to silica Sep-Pak in the same manner as in step 3.

Quantification of F2 -Isoprostanes

10. Wash cartridge with 5 ml ethyl acetate, then elute F2 -IsoPs from silica Sep-Pak with 5 ml of 1:1 (v/v) ethyl acetate/methanol into a 5-ml Reacti-Vial.



11. Evaporate eluate under nitrogen in an analytical evaporation unit. Samples can be safely stored for up to 1 week at −20◦ C.

Prepare sample for TLC 12. Convert F2 -IsoPs to pentafluorobenzyl esters (which facilitates GC/MS analysis) by adding 40 µl of 10% PFBB in acetonitrile and 20 µl 10% DIPE in acetonitrile to the Reacti-Vial, vortex briefly, and incubate for 20 min in a 37◦ C water bath. 13. Prepare TLC tank by adding 97 ml chloroform, 3 ml ethanol, and TLC paper to saturate the tank, and allow to equilibrate for 30 min. 14. Dry sample thoroughly under nitrogen in an analytical evaporation unit in a fume hood and resuspend material in 50 µl methanol. Vortex briefly.

Perform TLC 15. Prewash a silica TLC plate with 90:10 (v/v) ethyl acetate/ethanol. Dry the plate 10 min in a 90◦ C oven then cool in a dessicator prior to sample application. Apply sample mixture to a lane on the prewashed silica TLC plate. For a TLC standard, apply ∼2 to 5 µg of the methyl ester of PGF2α to another TLC lane. Avoid applying sample to the first 1 cm of the plate (preloading area).

16. Chromatograph to 13 cm. Visualize the TLC standard by spraying the lane with a 10% phosphomolybdic acid solution in ethanol and heating on a hot plate. Only spray and heat the plate containing the PGF2α standard, not the sample-containing plates. A single band will appear, which can be measured to determine the area to be scraped on the sample-containing plates.

17. Scrape silica from the TLC plate in the region of the methyl ester of PGF2α (Rf = 0.18) and adjacent areas 1 cm above and below.

Extract isoprostanes 18. Place silica in a 1.5-ml microcentrifuge tube and add 1 ml ethyl acetate to extract isoprostanes. Vortex vigorously for 30 sec, then microcentrifuge 3 min at 13,000 rpm at room temperature. 19. Remove the ethyl acetate and place it in a new 1.5-ml microcentrifuge tube, taking care not to disrupt the silica pellet. Samples can be safely stored for up to 1 week at −20◦ C.

20. Dry under nitrogen, then add 20 µl BSTFA and 7 µl DMF to residue to covert to trimethylsilyl ether derivatives for GC/MS analysis. 21. Vortex well and incubate sample 20 min at 37◦ C, then dry reagents under nitrogen. 22. Resuspend in 20 µl undecane and vortex briefly. Undecane is stored over calcium hydride to prevent water accumulation. Sample is now ready for mass spectrometric analysis and should be analyzed within 24 hr.

Perform mass spectrometry 23. For quantification of F2 -lsoPs by GC/NICI MS analysis, use an Agilent 6890N mass spectrometer with a computer interface. Chromatograph F2 -IsoPs on a 15-meter DB1701 fused silica capillary column. Program the column temperature from 190◦ C to 300◦ C at 20◦ C/mm. Use methane as the carrier gas for NICI at a flow rate of

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1 µl/min. Set the ion source temperature to 250◦ C, the electron energy to 70 eV, and the filament current to 0.25 mA. Monitor for endogenous F2 -IsoPs using the carboxylate anion m/z 569 (M-181, loss of CH2 C6 F5 ). The corresponding carboxylate anion for the deuterated intemal standard is m/z 573. In the authors’ experience, the indicated column produces excellent separation of individual regioisomers compared to other columns.

REAGENTS AND SOLUTIONS Use ultrapure (triply distilled) water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.

Folch solution Combine 2 volumes of high-purity chloroform (containing ethanol as a preservative) with 1 volume of high-purity methanol, both available from Burdick and Jackson or VWR Scientific. Cool to 4◦ C. Store in dark in a brown bottle to prevent light degradation. Store up to 1 month at −20◦ C. COMMENTARY Background Information


Free radicals derived primarily from oxygen have been implicated in the pathophysiology of a wide variety of human diseases including atherosclerosis, cancer, neurodegenerative disorders, and even the normal aging process (Halliwell and Gutteridge, 1990). Much of the evidence for this, however, is indirect or circumstantial, largely because of limitations of methods that are available to quantify free radicals or the products they produce in biological systems. This is a particular problem when non-invasive approaches are used to assess oxidant injury in animals or humans (Halliwell and Grootveld, 1987; DeZwart et al., 1999). Measures of lipid peroxidation are frequently utilized to implicate free radicals in pathophysiological processess. These measurements include quantification of shortchain alkanes, malondialdehyde, or conjugated dienes. Each of these, however, suffers from problems related to specificity and sensitivity, especially when utilized to quantify oxidant stress in vivo. Further, artifactual generation of these lipid peroxidation products can occur ex vivo, and factors such as endogenous metabolism can affect levels of compounds measured (DeZwart et al., 1999). In 1990, the authors reported that a series of prostaglandin (PG)F2 -like compounds, termed F2 -isoprostanes (F2 -IsoPs), are produced in vivo in humans by a non-cyclooxygenase free radical–catalyzed mechanism involving the peroxidation of arachidonic acid (Morrow et al., 1990). Formation of these compounds initially involves the generation of four positional peroxyl radical isomers of arachi-

donic acid; these undergo endocyclization to PGG2 -like compounds that are subsequently reduced to PGF2 -like compounds. Four F2 -IsoP regioisomers are formed, each of which can theoretically be comprised of eight racemic diastereomers. The authors have accumulated a large body of evidence that suggests that quantification of F2 -IsoPs represents a reliable and useful approach to assess lipid peroxidation and oxidant stress in vivo (see also UNIT 17.5). They readily increase in animal models of oxidant stress. Further, concentrations of F2 -IsoPs are present that are easily detected in normal human and animal biological fluids such as plasma and urine (Roberts and Morrow, 2000; Fam and Morrow, 2003). This allows the definition of a normal range, which permits an assessment of small increases in the formation of F2 -IsoPs in settings of mild oxidant stress. In addition, F2 -IsoPs can be detected in virtually every type of biological fluid analyzed thus far including plasma, urine, cerebrospinal fluid, bile, lymph, bronchoalveolar lavage fluid, and synovial fluid (Morrow et al., 2001). Levels of these compounds are also detectable in all types of tissues examined to date. These include liver, kidney, stomach, brain, lung, vascular tissue, muscle, and heart. Thus, the fact that F2 -IsoPs are detectable in various tissues and fluids provides the opportunity to assess the formation of these compounds at local sites of oxidant injury. Finally, F2 -IsoPs are increased in a number of human disorders associated with enhanced oxidant stress (Roberts and Morrow, 2000; Fam and Morrow, 2003).



The precursor of the F2 -IsoPs is arachidonic acid. The vast majority of arachidonic acid present in vivo exists esterified to phospholipids. Previously, the authors reported the novel finding that F2 -IsoPs are initially formed in situ from arachidonic acid esterified to phospholipids and then subsequently released in the free form by phospholipases (Morrow et al., 1992). This observation provides the basis for an important concept regarding the assessment of isoprostane formation in vivo in that when isoprostane generation is quantified, total production of F2 -IsoPs may be more accurately assessed by measuring levels of both free and esterified compounds. Furthermore, the fact that F2 -IsoPs are formed in situ in phospholipids can be utilized in an advantageous way to assess oxidant injury in specific animal or human organs by analyzing levels of these compounds esterified in phospholipids from tissue biopsy specimens obtained for diagnostic purposes.

Critical Parameters and Troubleshooting Considerable care must be taken when harvesting and handling samples to prevent the artifactual ex vivo generation of F2 -IsoPs (Morrow and Roberts, 1997). F2 -IsoPs can be generated artifactually in arachidonatecontaining biological fluids that remain >2 hr at room temperature, or stored at −20◦ C. However, storage of samples at −70◦ C prevents ex vivo oxidation and maintains sample integrity for at least 5 years. Ex vivo formation of F2 -IsoPs does not occur if samples are processed immediately after procurement or if BHT (and TPP in the case of arachidonate-rich fluids such as plasma) is added to the organic solvent during lipid extraction (Morrow and Roberts, 1997). Hydrolysis of lipids in tissues or biological fluids should be performed immediately after lipid extraction to avoid potential autoxidation of arachidonate contained in the phospholipids. In addition, artifactual generation of F2 -IsoP can also occur during sample preparation if reagents contain significant trace metal contamination. Thus, reagents should be of very high purity. The quality of water used during lipid extraction is extremely important. The authors routinely use triply distilled water or its equivalent. Finally, all glassware and plasticware should be rinsed with ultrapure (triply distilled) water prior to use (Morrow and Roberts, 2002). During sample workup, there are several points in the assay during which tissue or fluid extracts can be safely stored overnight at −70◦ C. These include (1) following elution

from the silica Sep-Pak in Basic Protocol 2, step 10, and (2) following extraction from silica after TLC purification in Basic Protocol 2, step 19. A common problem encountered when performing this assay occurs after derivatization of F2 -IsoPs to trimethylsilyl ethers. These derivatives are readily hydrolyzed upon exposure to very small amounts of water. Loss of the trimethylsilyl ethers will result in a lack of detectable signal at the correct molecular weight when analyzed by MS. Should this occur, the sample should be dried under nitrogen and rederivatized and analyzed immediately. Quantification by GC/negative ion chemical ionization (NICI) MS is an extremely sensitive method to quantify the F2 -IsoPs (Morrow and Roberts, 1999). The lower limits of detection of F2 -IsoPs is in the range of 1 to 5 pg using a deuterated standard with a blank of 1 to 3 ml of a fluid such as plasma, 0.2 ml urine, or 50 to 100 mg tissue. It should be noted that the method outlined herein is one of several GC/MS techniques that have been developed by investigators to quantify F2 -IsoPs. The other assays utilize very similar purification and derivatization approaches although they quantify different F2 -IsoP regioisomers including iPF2α -I and iPF2α -VI (Lawson et al., 1998; Pratico et al., 1998). In addition, IsoPs can be quantified by liquid chromatography/MS and immunoassay although neither of these methods may be as sensitive or accurate as GC/MS (Fam and Morrow, 2003; Liang et al., 2003).

Anticipated Results Figure 17.6.2 shows the selected ion current chromatogram obtained from the analysis of F2 -IsoPs in plasma from a rat after treatment with CCl4 to induce an oxidant stress. The series of peaks in the upper m/z 569 selected ion current chromatogram represents different endogenous F2 -IsoPs. This pattern of peaks is virtually identical to that obtained from all other biological fluids and tissues that the authors have examined to date. In the lower m/z 573 chromatogram, the single peak represents the [2 H4 ] 15-F2t -IsoP internal standard that was added to the plasma sample. For quantification purposes, the peak denoted by an asterisk (∗ ), which co-elutes with the 15-F2t -IsoP internal standard, is routinely measured. Using the ratio of the intensity of this peak to that of the internal standard, the concentration of F2 -IsoPs was calculated to 742 pg/ml, ∼36-fold above normal. The normal plasma

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Figure 17.6.2 Analysis of F2 -IsoPs in plasma obtained from a rat 4 hr after treatment with CCl4 (2 ml/kg orogastrically) to induce endogenous lipid peroxidation. The m/z 569 ion current chromatogram represents endogenous F2 -IsoPs. The m/z 573 ion current chromatogram represents the [2 H4 ] 15-F2t -IsoP internal standard. The peak in the upper chromatogram represented by the asterisk (∗ ) is the one routinely used for quantification of the F2 -IsoPs. The concentration of F2 IsoPs in the plasma using the asterisk (∗ ) peak for quantification was calculated to be 742 pg/ml.

concentration of F2 -IsoPs in rats is 22 ± 4 pg/ml (mean ± 1 SD) and in normal human plasma is 35 ± 6 pg/ml (mean ± 1 SD) The reader is referred to previously published data for normal levels of F2 -IsoPs in other biological fluids and tissues (Morrow et al., 2001). Quantification of the F2 -IsoPs based on the intensity of the asterisk (∗ ) peak shown in Figure 17.6.2 is highly precise and accurate. The precision is ± 6% and the accuracy is 96%.

Time Considerations In general, 15 to 20 samples per day can be processed by an experienced investigator. Lipid extraction and hydrolysis of this number of samples requires ∼3 hr, Sep-Pak purification requires ∼1.5 hr, drying, derivatization, and TLC purification require ∼2 hr, and extraction, drying, and silylation require 1 hr. Mass spectrometric analysis is automated and each sample requires ∼15 min of machine time.

Literature Cited


DeZwart, L.L., Meerman, J.H.N., Commandeur, J.N.M., and Vermeulen, P.E. 1999. Biomarkers of free radical damage applications in experimental animals and in humans. Free Rad. Biol. Med. 26:202-226.

Fam, S.S. and Morrow, J.D. 2003. The isoprostanes: Unique products of arachidonic acid oxidation. Curr. Med. Chem. 10:1723-1740. Halliwell, B. and Grootveld, M. 1987. The measurement of free radical reactions in humans. Some thoughts for future experimentation. FEBS Lett. 213:9-14. Halliwell, B. and Gutteridge, J.M.C. 1990. Role of free radicals and catalytic metal ions in human disease: An overview. Methods Enzymol. 186: 1-85. Lawson, J.A., Li, H., Rokach, J., Adiyaman, M., Hwang, S.W., Khanapure, S.P., and FitzGerald, G.A. 1998. Identification of two major F2 isoprostanes, 8,12-iso- and 5-epi-8,12-isoisoprostane F2α VI, in human urine. J. Biol. Chem. 293:29295-29301. Liang, Y., Wei, P., Duke, R.W., Reaven, P.D., Harman, S.M., Cutler, R.G., and Heward, C.B. 2003. Quantification of 8-iso-prostaglandin F2α and 2,3-dinor-8-iso-prostaglandin F2α in human urine using liquid chromatography-tandem mass spectrometry. Free Radic. Biol. Med. 34:409418. Morrow, J.D. and Roberts, L.J. 1997. The isoprostanes: Unique bioactive products of lipid peroxidation. Prog. Lipid Res. 36:1-21. Morrow, J.D. and Roberts, L.J. 1999. Mass spectrometric quantification of F2 -isoprostanes in biological fluids and tissues. Meth. Enzymol. 300:312.



Morrow, J.D. and Roberts, L.J. 2002. Mass spectrometric quantification of F2 -isoprostanes as indicators of oxidant stress. Meth. Molec. Biol. 186:57-66. Morrow, J.D., Hill, K.E., Burk, R.F, Nammour, T.M., Badr, K.F, and Roberts, L.J. 1990. A series of prostaglandin F2 -like compounds are produced in vivo in humans by a noncyclooxygenase, free radical catalyzed mechanism. Proc. Natl. Acad. Sci. U.S.A. 87:93839387. Morrow, J.D., Awad, J.A., Boss, H.J., Blair, I.A., and Roberts, L.J. 1992. Non-cyclooxygenasederived prostanoids (F2 -isoprostanes) are formed in situ on phospholipids. Proc. Natl. Acad. Sci. U.S.A. 89:10721-10725. Morrow, J.D., Zackert, W.E., Van der Ende, D.S., Reich, E.E., Terry, E.S., Cox, B., Sanchez, S.C., Montine, T.J., and Roberts, L.J. 2001. Quantification of isoprostanes as indicators of oxidant stress in vivo. In Handbook of Antioxidants 2nd ed. (E. Cadenas and L. Packer, eds.) pp. 57-74. Marcel Dekker, New York.

Pratico, D., Barry, O.P., Lawson, J.A., Adiyaman, M., Hwang, S.W., Khanapure, S.P., Iuliano, L., Rokach, J., and FitzGerald, G.A. 1998. IPF2α -1: An index of lipid peroxidation in humans. Proc. Natl. Acad. Sci. U.S.A. 95:3449-3454. Roberts, L.J. and Morrow, J.D. 2000. Measurement of F2 -isoprostanes as an index of oxidative stress in vivo. Free Radic. Biol. Med. 28:505-513.

Key References Fam and Morrow, 2003. See above. An up-to-date review of the isoprostane field. Morrow et al., 1990. See above. The initial report of isoprostane formation in vivo and the potential use of these compounds as an index of oxidant stress in vivo.

Contributed by Erik S. Musiek and Jason D. Morrow Vanderbilt University School of Medicine Nashville, Tennessee

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Immuno-Spin Trapping: Detection of Protein-Centered Radicals

UNIT 17.7

Protein-centered radicals are involved in biological oxidative damage induced by drugs, environmental hazards, and cellular reactive oxygen species. They can be generated by, among others, hydrogen peroxide (H2 O2 )-induced hemoprotein- or metal-catalyzed protein oxidations (Fig. 17.7.1). Because of their paramagnetic properties, these radicals can be detected and analyzed by direct electron spin resonance (ESR; also known as electron paramagnetic resonance, EPR). After a few microseconds or seconds, via mechanisms that are not completely understood, they decay to diamagnetic species (ESR-silent species). In the presence of spin traps, such as 5,5-dimethyl-1-pyrroline N-oxide (DMPO), proteincentered radicals can be trapped to form DMPO-protein radical adducts in which the formation of a new covalent linkage between the spin trap and the protein-centered radical preserves their free radical character. These radical adducts have a longer halflife than their parent radicals, thus DMPO-protein nitroxide radical adducts can also be characterized and studied by ESR. However, after a few seconds or minutes these species also decay to ESR-silent species by reduction or oxidation reactions to hydroxylamine or nitrone adducts, respectively, with preservation of the covalent bridge but not their paramagnetic properties. In an oxidizing environment, DMPO-protein nitroxide radical adducts decay to stable DMPO-protein radical–derived nitrone adducts, which can be detected and identified by immuno-spin trapping. This new technology combines the specificity and sensitivity of both spin trapping and antigen-antibody interaction and is based on the detection of the nitrone moiety in DMPO-protein radical–derived nitrone adducts by using an antiserum against DMPO (Fig. 17.7.1). Some of the advantages of using immuno-spin trapping rather than ESR and ESR-spin trapping to detect protein-centered radicals are: (1) the feasibility of applying this technology in any research, clinic, or academic laboratory without needing complicated equipment or specialists in physical chemistry and quantum mechanics, (2) the small quantity of sample required (microgram), and (3) the possibility of detecting more than one protein-centered radical in the same system and determining their approximate molecular weights. Moreover, day-to-day immuno-spin trapping is proving to be a

Figure 17.7.1

Detection of hydrogen peroxide (H2 O2 )-induced protein-centered radicals. Oxidative Stress

Contributed by Dario C. Ramirez and Ronald P. Mason Current Protocols in Toxicology (2005) 17.7.1-17.7.18 C 2005 by John Wiley & Sons, Inc. Copyright


simple, reliable, affordable, sensitive, and specific approach to detecting protein-centered radicals in situ and in real time in vitro, ex vivo, and in vivo, and it could be a powerful tool in toxicological research. In this unit, protocols for the production and immuno-spin trapping detection of H2 O2 induced horse heart metmyoglobin (metMb)- and human oxyhemoglobin (oxyHb)DMPO radical–derived nitrone adducts are described (see Basic Protocols 1 and 2). Immuno-spin trapping assays include enzyme-linked immunosorbent assay (ELISA; UNIT 18.7) and immunoblotting to detect and characterize protein-centered radicals as their DMPO-protein radical–derived nitrone adducts. BASIC PROTOCOL 1

PRODUCTION AND DETECTION OF HEMOPROTEIN-CENTERED RADICALS BY IMMUNO-SPIN TRAPPING ELISA Enzyme-linked immunosorbent assay (ELISA; UNIT 18.7) is a heterogeneous—i.e., requires separation between bound and unbound material in some step(s)—immunochemical technique that allows simultaneous screening of many samples (Tijssen, 1985). In this technique, antigens (nitrone adducts) are bound under alkaline conditions at the bottom of a 96-well microtiter plate through hydrophobic interactions. After coating, unbound material is removed by a wash step with a buffer containing detergent and an inert protein to remove any nonspecific interactions. This step allows removal of any non-protein nitrone adducts that can interfere with the detection of protein nitrone adducts. This represents one of the most important advantages of heterogeneous assays like ELISA compared to homogeneous assays such as ESR-spin trapping. After the wash step, the first antibody is added. After incubation, nonspecific bound and unbound antiserum is removed by two or three wash steps followed by addition of the secondary antibody labeled with an enzyme. Then, after further incubation, unbound and weak nonspecific interactions are removed by two or three wash steps. Finally, immuno-complexes are detected by adding enzyme substrates that produce luminescence or soluble-colored products that can be detected and measured.

Materials

Immuno-Spin Trapping

Metmyoglobin (metMb; USB) Oxyhemoglobin (oxyHb; Apex Biochemicals) H2 O2 (Alfa Aesar) Chelex-treated sodium phosphate buffer, pH 7.7 (see recipe) 5,5-Dimethyl-1-pyrroline N-oxide (DMPO; Alexis Biochemicals) Chelex-treated sodium phosphate buffer, pH 7.4, containing 1 mM diethylenetriaminepentaacetic acid (DTPA) PD-10 desalting columns (Amersham Biosciences) Catalase solution (100 or 500 IU, Roche Applied Bioscience) Coating buffer (see recipe) Wash buffer (see recipe) Blocking buffer (see recipe) Primary antibody solution (see recipe) Secondary antibody solution (see recipe) Chemiluminescence substrate for ELISA (see recipe) 37◦ C incubator with agitator 96-well white microtiter plates (e.g., Greiner Bio-One, PGC) Automated microtiter plate washer (optional) Multichannel pipettor Luminescence reader for microtiter plates or strips (optional)



Table 17.7.1 Extinction Coefficients of Reagents

Reagent (abbreviation)

Supplier

Extinction coefficient

Alexis Biochemicals

ε228 = 7800 M−1 cm−1

H2 O2

Alfa Aesar

ε240 = 43.6 M−1 cm−1

Metmyoglobin (metMb)

USB

ε406 = 154 mM−1 cm−1

Oxyhemoglobin (oxyHb)b

Apex Biochemicals

ε541 = 13.8 mM−1 cm−1

DMPO

a

a Purified by vacuum sublimation at room temperature and stored under an argon atmosphere at −70◦ C. b A gift from Apex Biochemicals.

Prepare DMPO-protein radical-derived nitrone adducts (i.e., sample to analyze) 1. Prepare hemoprotein [metmyoglobin (metMb) and oxyhemoglobin (oxyHb)] solutions by completely dissolving metMb or diluting oxyHb in 500 µl of Chelex-treated sodium phosphate buffer, pH 7.7, at room temperature. Pour solutions of metMb and oxyHb through PD-10 desalting columns to eliminate any preservatives and collect the colored eluate. Immediately before use, prepare 10× stock solutions of the reagents necessary to produce DMPO-protein radical-derived nitrone adducts (e.g., hemoprotein, H2 O2 , and DMPO) by using known extinction coefficients (see Table 17.7.1). Prepare all stock solutions in Chelex-treated sodium phosphate buffer and keep on ice until use. MetMb and oxyHb solutions are brown and red solutions, respectively. Hemoprotein stock solutions can be stored for 1 day at 4◦ C before use without significant variation in results. Typical reagent concentrations used in immuno-spin trapping experiments can be found in Figures 17.7.2 to 17.7.6 (also see Anticipated Results).

2. Prepare the reaction mixture by adding 30 µl of a 10× stock solution of hemoprotein (metMb or oxyHb) to 210 µl Chelex-treated sodium phosphate buffer containing 1 mM DTPA, at room temperature. Then, add 30 µl of 10× DMPO stock solution and start reaction by adding 30 µl of freshly prepared 10× H2 O2 stock solution. 3. Incubate reaction mixture 1 hr at 37◦ C with agitation. Stop reaction by removing excess H2 O2 with 10 µl of 100 IU/ml catalase solution. Under these conditions DMPO-protein radical-derived nitrone adducts are formed and are ready to be measured by ELISA. After the chemical reaction and before analysis, it is prudent, but not necessary, to eliminate excess reagent (e.g., DMPO and H2 O2 ) by dialysis using a membrane with a cut-off appropriate for the molecular weight of the protein under analysis. Dialysis of the reaction mixture (300 µl) may be performed using a 0.1- to 0.5-ml Slide-A-Lyzer dialysis cassette (Pierce) against 2 liters of Chelex-treated sodium phosphate buffer, pH 7.4, containing 1 mM DTPA at 4◦ C with three changes of buffer. After stopping the reaction with catalase or elimination of reagent excess by dialysis, DMPO-protein radical-derived nitrone adducts can be stored up to 3 weeks at 4◦ C or up to 6 months at –20◦ C. Protein-centered radicals have been detected in many systems containing protein-redox active metals and peroxides. Consequently, careful selection of the conditions for preparing the nitrone adducts is a key factor for success.

Bind DMPO-protein radical-derived nitrone adducts to ELISA plates 4. Add 190 µl coating buffer and 10 µl (∼0.5 to 3 µg proteins) experimental and control samples to each well of a 96-well white microtiter plate. Incubate plate 60 min at room temperature to allow samples to bind to the plate. Oxidative Stress


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High protein binding microtiter plates are recommended. The authors use 96-well white microtiter plates. Typically, 0.5 to 3 µg of protein are used for ELISA analysis of DMPO-protein radical– derived nitrone adducts.

5. Wash plate one time with 300 µl wash buffer in each well and incubate plate for 10 min at room temperature. Remove wash buffer. For more reproducible results, the authors recommend using multichannel pipettors and an automated microtiter plate washer. However, if an automated plate washer is not available, washing steps can be performed as follows without significant variations in the results: add 300 µl of wash buffer to each well and incubate the plate for 5 min in an orbital shaker. Remove the wash buffer by aspirating with a multichannel pipettor, being careful not to touch the bottom of the wells, or by inverting and shaking the plate. Repeat wash buffer addition, incubation, and evacuation two or three additional times. Remove the excess wash buffer from the last wash step by inverting the plate on paper towels. Do not let the plate dry.

6. Add 200 µl blocking buffer to each well and incubate the microtiter plate 60 min at room temperature. 7. Repeat step 5.

Detect antigen 8. Add 200 µl primary antibody solution to each well and incubate 60 min at 37◦ C to allow the antibodies to recognize and bind to the DMPO-protein radical–derived nitrone adducts. 9. Repeat step 5 three times. This step removes the unbound primary antibody.

10. Add 200 µl of the secondary antibody solution and incubate microtiter plate 60 min at 37◦ C to allow the secondary antibody to recognize and bind to the primary antibody. 11. Repeat step 5 three times.

Detect antigen-antibody complexes 12. Add 200 µl chemiluminescence substrate for ELISA and read immediately in a microtiter plate reader equipped with a luminometer. Alternatively, other chromogenic substrates for alkaline phosphatase can be used and the absorbance detected by studying the absorbance of its soluble product. BASIC PROTOCOL 2


PREPARATION AND IMMUNOBLOT ANALYSIS OF DMPO-PROTEIN RADICAL–DERIVED NITRONE ADDUCTS The authors use ELISA to quantify protein radical–derived nitrone adduct generation, but to study and identify any structural effect on the protein (e.g., aggregation or fragmentation), immunoblot analysis, which also provides an approximate molecular weight for the DMPO-protein radical–derived nitrone adduct(s), is used. Although less sensitive than ELISA, immunoblotting is the only immunochemical analysis that allows for the investigation of more than one protein-centered radical adduct in the same system (see Anticipated Results) or in complex mixtures such as cellular environments (Ramirez et al., 2003) or tissue homogenates (Detweiler et al., 2002). Once protein-centered radicals are detected, the corresponding band in gels stained by Coomassie blue can be cut, digested, and analyzed by mass spectrometry to identify the protein and even the residue where DMPO is bound, i.e., where the radical was produced (Deterding et al., 2004).



Materials Metmyoglobin (metMb; USB) H2 O2 (Alfa Aesar) Oxyhemoglobin (oxyHb; Apex Biochemicals) 5,5-Dimethyl-1-pyrroline N-oxide (DMPO; Alexis Biochemicals) Chelex-treated sodium phosphate buffer, pH 7.4, containing 1 mM diethylenetriaminepentaacetic acid (DTPA) PD-10 desalting columns (Amersham Biosciences) Catalase solution (100 or 500 IU, Roche Applied Bioscience) 4× NuPage LDS sample buffer (Invitrogen) 10× NuPAGE sample reducing agent (Invitrogen) 4% to 12% NuPAGE Novex Bis-Tris gel (1.0-mm, 10-well; Invitrogen) Running buffer (see recipe) Nitrocellulose membranes (0.45-µm pore size, Invitrogen) Transfer buffer (see recipe), cold 0.1% (w/v) Ponceau S red in 4% (v/v) acetic acid Blocking buffer (see recipe) Wash buffer (see recipe) Primary antibody solution (see recipe) Secondary antibody solution (see recipe) TBS, pH 9.6 (see recipe) Chemiluminescence substrate for westernblots (see recipe) 1-Step NBT/BCIP reagent (Pierce) 37◦ C incubator with agitator 70◦ to 80◦ C water bath Mini electrophoresis system (e.g., XCell SureLock Mini-Cell system, Invitrogen) Mini-gel blot module (e.g., XCell II Blot Module, Invitrogen) X-ray film (e.g., CL-XPosure film, Pierce) Prepare DMPO-protein radical-derived nitrone adducts (i.e., the sample to analyze) 1. Prepare 10× stock solutions of reagents [i.e., metmyoglobin (metMb), oxyhemoglobin (oxyHb), DMPO, and H2 O2 ]. Completely dissolve metMb or dilute oxyHb solution in 500 µl of Chelex-treated sodium phosphate buffer, pH 7.4, at room temperature. Pour solutions of metMb and oxyHb through PD-10 desalting columns (equilibrated with distilled water) to eliminate any preservatives and collect the colored eluate. Immediately before use, prepare ten times concentrated stock reagent (hemoproteins, H2 O2 , and DMPO) solutions than the final concentrations required in the reaction mixture (see Fig. 17.7.4) in Chelex-treated sodium phosphate buffer by using known extinction coefficients (see Table 17.7.1) and keep on ice until used. MetMb and oxyHb solutions are brown and red, respectively. These hemoprotein solutions can be stored for 1 day at 4◦ C before use without significant variation in results.

2. Prepare the reaction mixture in disposable 1.5-ml microcentrifuge tubes. Add 30 µl of 10× stock solution of hemoprotein (metMb or oxyHb) to 210 µl Chelex-treated phosphate buffer containing 1 mM DTPA at room temperature. Then add 30 µl of 10× stock DMPO solution and start reaction by adding 30 µl of freshly prepared 10× H2 O2 stock solution. 3. Incubate reaction mixture 1 hr at 37◦ C with agitation. Stop reaction by removing excess H2 O2 with 10 µl of 500 IU/ml catalase solution. Under these conditions, DMPO-protein radical-derived nitrone adducts are formed and are ready to be resolved by SDS-PAGE.

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It is recommended to eliminate excess reagent (i.e., DMPO and H2 O2 ) by dialysis. Dialysis of the reaction mixture (300 µl) is performed using a 0.1- to 0.5-ml Slide-A-Lyzer dialysis cassette (Pierce) against 2 liters of Chelex-treated sodium phosphate buffer, pH 7.4, containing 1 mM DTPA at 4◦ C with three changes of buffer. After stopping the reaction with catalase or eliminating reagent excess by dialysis, DMPOprotein radical-derived nitrone adducts can be stored for 3 weeks at 4◦ C or up to 6 months at –20◦ C. Protein-centered radicals have been detected in many systems containing protein-redox active metals and peroxides. Consequently, careful selection of the conditions for preparing the nitrone adducts is a key factor for success.

Prepare sample to resolve by SDS-PAGE 4. Mix 30 µl of reaction mixture, 10 µl of 4× NuPage LDS sample buffer, and 4 µl of 10× NuPAGE sample reducing agent. In place of the NuPAGE sample reducing agent, 4 µl of fresh 0.5 M dithiotreitol (DTT) solution can be used with similar results.

5. Heat the sample 7 to 10 min in a 70◦ to 80◦ C water bath, and then let it cool to room temperature. 6. Load 10 µl sample per lane of a 4% to 12% NuPAGE Novex Bis-Tris gel. The ten-well, 1-mm, 4% to 12% NuPAGE Novex Bis-Tris gels are appropriate for separation of proteins between 3 and 100 kDa and resolve well for proteins used in this protocol. Careful selection of gels is a key factor in successful immuno-spin trapping. For better standard-molecular-weight band visualization, the authors add 1 to 3 µl of 1× SeeBlue Plus2 pre-stained standard (Invitrogen) to the first and middle lanes of the gel.

7. Perform protein separation using an electrophoresis system under continuous voltage conditions for 40 min at 200 V with running buffer. Remove the running buffer and add distilled water to cool the gel. This step in the procedure permits easier handling of the gel during the following steps.

8. Remove glass or plastic from the gel, cut a piece of nitrocellulose membrane for orientation, and place on top of the gel. Remove any bubbles between membrane and gel by submerging under cool (0◦ to 4◦ C) transfer buffer. CAUTION: Acrylamide compounds are neurotoxic and carcinogenic for laboratory animals and humans; consequently, gloves and adequate safety practices must be followed to avoid exposure.

9. Blot proteins onto nitrocellulose membrane by using a nitrocellulose membrane filter paper sandwich. Perform the transfer in a mini-gel blot module for 20 to 30 min at 40 V. Under the buffer conditions used in this process, proteins will migrate from the negative electrode to the positive electrode. Those who are unfamiliar with blotting may refer to the NuPAGE Bis-Tris gel instruction manual, where clear instructions on how to assemble the blotting sandwich are provided. This booklet is included in every box of NuPAGE gels and can also be found on the manufacturer’s Web site (see Internet Resources).

Stain protein on membrane 10. Transfer the membrane to a clean tray or dish (e.g., weigh boats, pipet tip boxes, or other) and rinse with distilled water. Immuno-Spin Trapping

To minimize the volume of stains, buffers, and antibody solutions, it is convenient that the tray or dish is only slightly larger than the membrane.



11. Incubate the membrane with 0.1% Ponceau S red in 4% acetic acid for 10 to 30 sec. Wash the membrane with distilled water until red bands (i.e., indication of proteins) appear on a white background, which can then be scanned or photographed. The red bands on the membrane will disappear during the blocking process. This staining allows for observation of any defects in the blotting process such as poor blotting or diffusion of proteins on the membrane. Usually, the latter is due to the poor quality of sponges used in the transfer sandwich. If this occurs, the authors recommend using new sponges. Sponges should not be used for >20 blotting processes. Before immunoblotting, membranes can be stored dry for weeks between pages of a book, or in water if the immunostaining is started the next morning. If stored dry, it is important to rehydrate the membrane for at least 30 min in distilled water at room temperature.

Detect DMPO-protein radical-derived nitrone adducts on membrane by immunoblotting 12. Incubate the membrane with enough volume of blocking buffer to cover the membrane 60 min at room temperature. Remove blocking buffer. Typically, the authors use 25 ml of reagents. In the following steps, use enough volume of buffers and solutions to cover the membrane.

13. Cover the membrane with wash buffer and incubate for 10 min at room temperature. Remove wash buffer. 14. Incubate the membrane with primary antibody solution 60 min at room temperature. 15. Repeat step 13 three times. 16. Incubate the membrane with secondary antibody solution 60 min at room temperature. 17. Repeat step 13 three times. After removal of the last wash buffer, cover the membrane with TBS, pH 9.6. Incubation of the membrane after the last wash with TBS, pH 9.6, will increase the activity of alkaline phosphatase with a consequent increase in detection sensitivity.

Develop antigen-antibody complexes by using ECL 18. Remove the membrane, place it face up onto a piece of plastic, add 5 ml chemiluminescence substrate for western blots, and incubate for 2 min in the dark. 19. Remove excess reagent by wicking with paper towels without touching the membrane and place the membrane between two pieces of plastic. The authors use office plastic folders.

20. Remove the bubbles remaining between the plastic and membrane by rolling with a pipet or pressing them onto a paper towel. This procedure also ensures the complete removal of excess liquid.

21. In a dark room, expose membrane to X-ray film. Exposure time may vary from a few seconds to a few minutes (or longer), depending upon the amount of antigen to be detected.

22. Develop the X-ray film. If desired, the same membrane can be used for colorimetric development of the antigenantibody complexes (optional).

23. Scan or photograph the membrane to make a permanent record of the results. Oxidative Stress


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Develop antigen-antibody complex by using one-step NBT/BCIP reagent (optional) Sometimes unexpected problems can occur due to failure of the X-ray film, equipment, or developing reagents. NBT/BCIP development is also a way to keep a good record, but is not necessary. 24. (Optional) For membrane used for luminescent development, wash three times, 5 min each, in distilled water with agitation. Finally, wash with 25 ml TBS, pH 9.6, for 5 min. 25. (Optional) Add 10 ml 1-Step NBT/BCIP reagent, incubate 15 min or until purple bands are clearly visible, then stop the reaction by washing three times, 5 min each, with distilled water. 26. Scan or photograph the membrane to make a permanent record of the results. CAUTION: Use gloves and laboratory clothes when developing antigen-antibody complexes by either method as the reagents can be toxic and carcinogenic.

REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.

Blocking buffer 100 ml coating buffer (see recipe) 2 ml gelatin from cold water fish skin (45% v/v, Sigma) Mix until total dissolution. Prepare 1 hr before use to ensure total solubilization Alternatively, 2.5% (w/v) bovine serum albumin and 2.5% (w/v) casein in coating buffer or SeaBlock (Pierce) can be used without significant variation in results.

Chelex-treated sodium phosphate buffer, 0.1 M (pH 7.4) Weigh 5 g of Chelex 100 resin (Bio-Rad) in a 1-liter beaker and add 1 liter of water with a magnetic stirrer, stir the resin at low speed (to avoid damaging the resin) for 2 hr at 4◦ C. Allow the resin to sediment and remove the water. Resuspend the resin in 1 liter of distilled water and allow the resin to wash overnight at 4◦ C. Repeat this procedure one more time for 2 hr. Then filter the suspension using a 0.45-µm filter and wash the resin with 1 liter of 0.1 M sodium phosphate buffer, pH 7.4, to adjust pH. Filter the excess buffer and add the washed resin to 1 liter phosphate buffer and stir with a magnetic stirrer at slow speed overnight at 4◦ C. On the next day, filter the treated phosphate buffer to eliminate the resin and add 1 ml of 100 mM DTPA solution (stock) to the filtrate to make 1 liter of Chelex-treated sodium phosphate buffer, pH 7.4, containing 1 mM DTPA. Chemiluminescence substrate For ELISA: Add 50 µl CDP-star (Roche Applied Science) to 50 ml TBS, pH 9.6 (see recipe). Prepare immediately before use in buffer at room temperature and mix well. For western blot: Add 50 µl CDP-star to 50 ml TBS, pH 9.6 (see recipe) and 100 µl Nitro-Block II (Tropix). Prepare immediately before use in buffer at room temperature and mix well.


Coating buffer 1.59 g sodium carbonate (Na2 CO3 ) 2.93 g sodium bicarbonate (NaHCO3 ) continued



0.20 g sodium azide (NaN3 ) Dissolve in 900 ml distilled water, adjust to pH 9.6 with NaOH or HCl and make up to 1 liter Store up to 1 month at room temperature In some cases, antigens bound under neutral conditions, such as distilled water or phosphate buffer, give lower backgrounds than bicarbonate buffer. The authors recommend that the coating buffer be optimized to produce the highest signal/background ratio.

Primary antibody solution Dilute primary anti-DMPO (Alexis Biochemicals, Cayman Chemicals, and Oxford Biomedical Research) antibody 1:5000 in wash buffer (see recipe). Dilute the antiserum immediately before use. Running buffer 50 ml 20× NuPAGE MES SDS running buffer (Invitrogen) 850 ml distilled water Prepare immediately before use Secondary antibody solution Dilute secondary ImmunoPure goat anti-rabbit IgG (specific to Fc) antibody, alkaline phosphatase–conjugated (Pierce) 1:5000 in wash buffer. Dilute the antiserum immediately before use. IMPORTANT NOTE: Hemoproteins and metal-protein complexes can catalyze the reduction of H2 O2 and oxidation of peroxidase substrates with the production of nonspecific signals in ELISA and western blot analysis. Consequently, secondary antiserum conjugated to peroxidases will produce high unspecific signals that are not associated with antigenantibody interaction, but related to the peroxidase activity of the hemoprotein or metalprotein under analysis.

Tris-buffered saline, 10× (pH 7.4) 30 g TRIZMA base 80 g NaCl 2 g KCl 800 ml distilled water Adjust to pH 7.4 with concentrated HCl. Bring to 1 liter, and filter sterilize into 100-ml flasks Store up to 1 month at room temperature Tris-buffered saline, pH 9.6 (TBS, pH 9.6) In 800 ml distilled H2 O, add the following: 32 g NaCl 0.8 g KCl 12 g Tris Adjust to pH 9.6 with 1 M HCl Bring up to 1 liter with H2 O Sterilize by filtering through a 0.45-µm filter Store up to 1 month at room temperature Transfer buffer 800 ml distilled water 50 ml 20× NuPAGE transfer buffer (Invitrogen) 100 ml methanol 1 ml antioxidant Add distilled water to make 1 liter Prepare immediately before use

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Wash buffer 100 ml 10× TBS (see recipe) 900 ml distilled water 2 ml cold-water fish gelatin 0.5 ml Tween 20 (USB) Mix until total dissolution Store at 4◦ C and let it reach room temperature before use Alternatively, 0.25% (w/v) BSA plus 0.25% (w/v) casein acid hydrolyzate in 1× TBS and 0.05% (v/v) Tween 20 can be used in place of fish gelatin.

COMMENTARY Background Information A growing field of evidence suggests that free radicals and particularly protein-centered radicals are involved in aging, pathophysiology, and toxicological processes (Berlett and Stadtman, 1997; Dean et al., 1997; Mason, 2000; Hawkins and Davies, 2001). A free radical is an atom or group of atoms possessing one unpaired electron. The unpaired electron gives these species paramagnetic properties that make them suitable for detection by electron spin resonance (ESR). However, because of the high reactivity of protein-centered radicals, they are generally stable for only microseconds to seconds before they decay to ESR-silent species. In the spin-trapping technique, a reactive radical (R• ) adds across the double bond of a diamagnetic spin trap (S = T) to form a much more stable free radical, a radical adduct (R-S-T• ), which can then be examined by electron spin resonance (ESR; Mason, 2000). R• + S = T → R-S-T•


The most popular of these spin traps is 5,5dimethyl-1-pyrroline N-oxide (DMPO). When DMPO traps a protein-centered radical, a nitroxide radical adduct is formed with a covalent bond between DMPO and the protein (Davies and Hawkins, 2004). This species conserves the unpaired electron of the parental protein-centered radical, resulting in another paramagnetic species that can be studied by ESR (Davies and Hawkins, 2004). The specificity of the reaction between a nitrone spin trap and protein-centered radicals has already made spin trapping with ESR detection the most universal, specific tool for the detection of free radicals (Mason, 2000) and protein-centered radicals (Clement et al., 2001; Davies and Hawkins, 2004) in biological systems. However, ESR and ESR-spin trapping require extensive knowledge of mathematics and physical chemistry and also require complicated

equipment, which has limited their application in biomedical laboratories. The radical adduct is a paramagnetic species that decays (seconds to minutes) by oxidation or reduction to ESR-silent species by mechanisms still not well understood. Although the oxidation of nitroxide radical adducts to the corresponding ESR-silent nitrone adduct has not been well studied (Mason, 2004), once the ESR signal has decayed, nitrone adducts are evidence of earlier radical reactions (Deterding et al., 2004). Clearly, it would be advantageous to detect markers of protein radical formation without using ESR. A potential alternative to ESR was suggested by flow injection electrospray ionization mass spectrometry (ESI/MS) of the reaction products of myoglobin with H2 O2 and DMPO. This analysis, performed after the ESR signal disappeared, demonstrated the presence of a myoglobin-derived product with a mass increase of 111 Da. This species corresponded to the addition of DMPO minus two hydrogen ions as expected for the formation of a covalent bond between Mb and DMPO (Detweiler et al., 2002). The persistence of the covalent bond after the radical character of the adduct was lost suggests that the oxidized form, the nitrone moiety, could form the basis of an antigen (Mason, 2004). The nitrone group does not exist in nature, thus eliminating the possibility of a false-positive from endogenous cellular components. It should be highly antigenic, as is the related nitro group (Mason, 2004). Moreover, the site of DMPO attachment to the protein is a specific marker for where the radical resided. Consequently, an antiserum that recognizes DMPO was developed in the authors’ laboratory (Detweiler et al., 2002). To facilitate antiserum production, a DMPO derivative containing an octanoic acid (OA) tail was synthesized, and this DMPO-OA was conjugated to a carrier protein chicken egg albumin for the immunization of rabbits. After the immunization



Table 17.7.2 Troubleshooting Guide to Immuno-Spin Trapping

Problem

Possible cause

Solution

No nitrone adducts are detected by No amino acid-centered radical was ELISA in a solution of a hemoprotein formed as is the case with horseradish with peroxidase activity, DMPO, and peroxidase H2 O2

Check if the protein radical has previously been detected by ESR and/or ESR-spin trapping with DMPO. Study a known system to obtain training.

High background/signal ratio in ELISA

Inappropriate blocking buffer

Probe with another blocking protein such as dry milk, casein/BSA, or SeaBlock solution (Pierce).

DMPO was not purified

Use purified DMPO.

High amount of protein added to well, Add a lower amount of protein to each producing high nonspecific antiserum well (typically 0.5 to 3 µg protein/well are enough). binding. High nonspecific binding

Increase the number of washes or increase Tween-20 concentration in the washing buffer to 0.1%.

H2 O2 -concentration dependence in the nitrone adducts was observed by ELISA but not by immunoblot analysis

It is possible that DMPO binds nonspecifically to protein radicals formed at the high temperatures used to prepare samples for reducing blots

Dialyze samples before mixing with the sample buffer.

No protein-centered radicals are observed in immunoblot analysis but they were detected by ELISA

Nitrone epitope was modified by the reducing agent

Run the SDS-PAGE under non-reducing conditions or change the reducing agent. The use of 2-ME as a reducing agent (see Anticipated Results; Fig. 17.7.2B) is not recommended.

Diffuse band pattern was observed

If SDS-PAGE was well done, as Change sponges, check blotting confirmed by protein staining of the apparatus in special electrodes. gel, it could be due to a low blot quality; this must be checked immediately after the transfer by using the temporary stain procedure described in the immunoblot protocol

regime, the rabbits were bled and the serum containing anti-DMPO polyclonal antibodies was collected (Detweiler et al., 2002). This antiserum has been used to develop immuno-spin trapping (Ramirez et al., 2003) and it is commercially available from Alexis Biochemicals, Cayman Chemicals, and Oxford Biomedical Research. Initially, this antiserum was successfully used to detect myoglobin-centered radicals in supernatant obtained from rat heart homogenate (Detweiler et al., 2002). Later immuno-spin trapping was applied in the study of protein-centered radicals generated in hemoglobin treated with peroxynitrite (Romero et al., 2003), ketoprofen/UV-A irradiation (He et al., 2003), and H2 O2 (Ramirez et al., 2003; Deterding et al., 2004). The H2 O2 induced DMPO-hemoglobin adducts were de-

tected by immuno-spin trapping (Ramirez et al., 2003), and then the exact localization of the radical sites where DMPO was bound to Hb was determined using LC/MS and MS/MS analysis of proteolytic fragments (Deterding et al., 2004). Recently, a lactoperoxidasecentered, thiol-dependent radical was also detected by immuno-spin trapping in incubations containing lactoperoxidase, thiols, and DMPO (Guo et al., 2004). Hypochloriteinduced DMPO-cytochrome c adducts have also been reported (Chen et al., 2004).

Critical Parameters and Troubleshooting Mixing unpurified DMPO with oxyHb or metHb, in the absence of H2 O2 produced protein aggregation detectable as dimers (∼32 kDa) in reducing SDS-PAGE and high

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background in ELISA and immunoblot experiments. These background and aggregation patterns were abolished when catalase was added before the addition of DMPO, suggesting that H2 O2 is a contaminant present in commercial DMPO preparations (Ramirez and Mason, unpub. observ.). Before DMPO is used as a spin trap, it is appropriate to purify it by vacuum sublimation and store it in 50-µl aliquots under an argon atmosphere at −70◦ C. It is important that no protein radical–derived nitrone adducts or aggregation be observed when purified DMPO is incubated for 1 to 2 hr with hemoproteins.


On the other hand, when compounds such as reducing thiols are added to the analysis mixture, they have effects not associated with their metal-chelating abilities, and in these systems, they can modify nitrone adducts to immuno-spin trapping silent species (see the effect of 2-ME on DMPO-Hb radical–derived nitrone adducts) or even produce DMPOtrappable protein-centered radicals (Guo et al., 2004). Because of the heating procedure required before SDS-PAGE separation, the nitrone adducts or the spin trap may react with the protein and thiols, i.e., under reducing

Figure 17.7.2 Detection of myoglobin (Mb) and hemoglobin (Hb) radical–derived nitrone adducts by immuno-spin trapping. (A) Reaction mixture contained 50 mM DMPO, 10 µM H2 O2 , and varying concentrations of metMb or oxyHb (as heme); (B) Reaction mixture contained 5 µM metMb (as heme), 5 µM H2 O2 , and varying concentrations of DMPO. Reactions were performed in 100 mM Chelex-treated sodium phosphate buffer, pH 7.4, containing 1 mM DTPA. After a 1-hr incubation at 37◦ C the excess H2 O2 was removed by adding 50 IU catalase. Protein-radical–derived nitrone adducts were determined by ELISA (see Basic Protocol 1). Data show mean values ± S.E. from three experiments performed in triplicate.



conditions. In this case, sample dialysis prior to electrophoresis is sufficient to prevent false-positive results. Detailed analysis of parameters that can affect immunoassays can be found in immunoassay manuals (Tijssen, 1985). See Table 17.7.2 for a guide on troubleshooting immuno-spin trapping.

Anticipated Results The incubation of heme proteins with weak peroxidase activity such as hemoglobin (Hb),

myoglobin (Mb), and cytochrome c with H2 O2 produces protein-centered radicals that have been detected and characterized by direct ESR and ESR-spin trapping. With a very low concentration of H2 O2 , i.e., 10 µM, Hb- or Mb-centered radicals were produced and detected as DMPO-protein radical–derived nitrone adducts by ELISA (Fig. 17.7.2A). Even a low concentration of metMb (0.5 µM) produced significant amounts of DMPO-protein radical–derived nitrone adduct as detected by ELISA. As hemoprotein concentration

Figure 17.7.3 Detection of Hb- and Mb-centered radicals by immuno-spin trapping assays. (A) 5 µM hemoproteins (HP, as heme) were incubated with 50 mM DMPO and varying concentrations of H2 O2 in 100 mM Chelex-treated sodium phosphate buffer, pH 7.4, containing 1 mM DTPA. (B) 10 µM oxyHb was reacted with 10 µM H2 O2 and 50 mM DMPO in 100 mM Chelex-treated phosphate buffer containing 100 µM DTPA. Reaction was performed 1 hr at 37◦ C and stopped by adding 50 IU catalase. Then the reaction mixture was incubated 10 min at 37◦ C with different percentages of 2-mercaptoethanol (2-ME, stock 14 M), its chemical structural analog ethanol, DTT (1 M), or phosphate buffer to correct for changes in volume. Nitrone adducts were determined in 10 µl of the reaction mixture by ELISA (see Basic Protocol 1). Data show mean values ± S.E. from three experiments carried out in triplicate and in three separate experiments.

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increases, generation of nitrone adducts rises steadily (Fig. 17.7.2A). Mb always produces more nitrone adducts than Hb or cytochrome c (data not shown), and metHb produces more adducts than oxyHb (Fig. 17.72A). The amount of nitrone adducts detected by ELISA depends on the concentration of DMPO available to trap radical sites in the protein (Fig. 17.7.2B). The authors do not recommend the use of higher concentrations of DMPO because it may suppress hydrophobic binding of the protein to the ELISA microtiter plate or bind non-covalently to the protein during sample preparation for SDS-PAGE. If higher concentrations of DMPO must be used, try dialyzing the sample before processing by ELISA or immunoblotting. If purified DMPO is used, no hemoprotein radical–derived nitrone adducts should be detected when H2 O2 is omitted (Fig. 17.7.2B, closed bars). It is also important to conduct control experiments in which one of the components is omitted. Previously, hemoglobin-centered radicals were detected by direct ESR and ESR-


spin trapping using millimolar concentrations of H2 O2 , which do not represent physiological conditions. With the immunospin trapping assay and ELISA, a significant amount of Mb radical–derived nitrone adduct can be detected with H2 O2 concentrations as low as 1 µM. No DMPO-protein radical– derived nitrone adducts were detected when hemoprotein, DMPO, or H2 O2 was omitted in the reaction mixture. In general, the production rate of DMPO-protein radical–derived nitrone adduct increases with hemoprotein (Fig. 17.7.2A), DMPO (Fig. 17.7.2B) and H2 O2 (Fig. 17.7.3A) concentration. Preliminary observations showed that immunoblot analysis of Hb radical–derived nitrone adducts obtained after protein separation under non-reducing conditions (i.e., without thiols) gave a higher signal/background ratio than those obtained after protein separation under reducing conditions (i.e., adding 10% v/v 2-mercaptoethanol during sample preparation). The authors hypothesized that this excessive background was due to the reaction

Figure 17.7.4 Immuno-spin trapping detection of hemoprotein radical–derived nitrone adducts by immunoblot analysis. The reaction contained 10 µM hemoprotein (metMb or oxyHb), with 0, 0.5, 1, 5, 10, and 50 µM H2 O2 (lanes a through f, respectively) in the presence of 100 mM DMPO in 100 mM Chelex-treated sodium phosphate buffer, pH 7.4. Reactions were performed 1 hr at 37◦ C and stopped by adding 50 IU catalase. The reaction mixture was separated by SDSPAGE and nitrone adducts were detected by immunoblotting and enhanced chemiluminescence (ECL) development (left membranes). The membranes were then washed with distilled water and incubated with 5 ml BCIP/NBT One-Step reagent until purple bands appeared on a white background (right membranes). The membranes were washed with distilled water and placed in a plastic office folder. X-ray films and membranes were scanned and acquired using an office scanner. M indicates molecular-weight marker.



of the nitrone adducts with the added thiols during the preparation for reducing gels. To investigate this, ELISA experiments, which do not require heating steps (Fig. 17.7.3B), were performed. Apparently, 2-ME changes the DMPO-Hb radical–derived nitrone adducts to an immuno-spin trapping silent species, and in this process, the thiol group of 2-ME affects the nitrone moiety of the adduct, the epitope recognized by the anti-DMPO antiserum. 2ME did not affect the yield of DMPO-Mb or

SOD1-radical derived nitrone adducts. Based on these results, the authors do not recommend the use of 2-ME as a reducing agent for reducing gels, especially when the system under study is unknown. The authors have also found that DTT can replace 2-ME without causing this effect (Fig. 17.7.3B). Figure 17.7.4 shows typical immunoblot results obtained when metHb or oxyHb is incubated with H2 O2 and DMPO; the blot is less sensitive than ELISA. The choice of enhanced

Figure 17.7.5 Simultaneous detection of two protein-centered radicals in the same system. Reaction mixtures contained 15 µM SOD1, 7.5 µM human serum albumin (HSA, ε280 nm = 35,700 M−1 cm−1 ), 100 mM DMPO and/or 1 mM H2 O2 (as indicated in the figure) in 100 mM Chelex-treated (bi)carbonate buffer, pH 7.4, containing 1 mM DTPA. Reactions were incubated for 2 hr at 37◦ C and stopped by adding 10 IU catalase. Then 30 µl of the reaction mixtures were mixed with 10 µl sample buffer and 4 µl of 10× reducing agent. After heating 5 min at 70◦ C, the mixture was run on two 4% to 12% SDS-PAGE gels; one gel was stained with Coomassie blue (upper gel) and the other was blotted to a nitrocellulose membrane. Nitrone adducts were detected by immunoblotting (lower gel) as described in Basic Protocol 2. M indicates molecular-weight marker.

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chemiluminescence (ECL) or colorimetric development (NBT/BCIP reagent, Pierce) depends on the sensitivity and amount of adducts to be detected and on the availability of the reagents and equipment. These results show that immuno-spin trapping is sufficiently flexible to fit academic, biomedical, or research laboratory needs in detecting protein-centered radicals. Previously, the authors reported that DMPO-bovine Cu, Zn superoxide dismutase (SOD1) radical-derived nitrone adducts are induced by H2 O2 only when (bi)carbonate, i.e., CO2 , HCO3 – , or CO3 –2 , is present in the reaction medium (Ramirez et al., 2005). Under conditions described in that report, the generation of DMPO trappable radical sites in SOD1 was most probably due to oxidation promoted by the carbonate radical anion. When human serum albumin (HSA) is included in the SOD1/H2 O2 /DTPA/ (bi)carbonate system, both SOD1- and HSA-centered radicals are produced and they can be simultaneously detected as DMPOprotein radical derived nitrone adducts by immunoblot analysis (Fig. 17.7.5). By comparison of the Coomassie blue stain (Fig.


17.7.5, upper gel) with the corresponding immunoblot (Fig. 17.7.5, lower gel), it is evident that only SOD1 monomer and dimer are detected as DMPO-protein radical derived nitrone adducts, even when fragments of SOD1 were also observed in the gel (compare lanes 2 and 6 in the upper and lower panels, Fig. 17.7.5). The origin of each one of these radicals is difficult, and sometimes impossible, to determine by direct ESR or ESR-spin trapping, due to the overlapping of ESR spectra of protein. The application of ESR to detect protein radicals in vivo has been pursued for many years. The stability of the DMPO-protein radical–derived nitrone adducts made it possible to analyze them in red blood cells (RBC) of rats exposed to tert-buthyl hydroperoxide (Fig. 17.7.6). As little as 15 min after the administration of tert-butyl hydroperoxide and DMPO, the authors detected protein-centered radicals in RBC whose signal increased with time, demonstrating that the production of proteincentered radicals is related to the toxicokinetics of tert-butyl hydroperoxide. In toxicological studies, the amount and accessibility of the sample is very important. The authors used

Figure 17.7.6 Detection of protein radical–derived nitrone adducts in blood of rats exposed to tert-buthyl hydroperoxide (tert-BOOH). Male Sprague Dawley rats (150 to 200 g) received 50 mg Nembutal/kg body weight (i.p.) and 50 µl blood was obtained by orbital bleeding. Then rats were injected with 1 g/kg DMPO (i.p.) and 0.5 ml/kg tert-BOOH or saline (8.5 g/liter NaCl) by intragastric intubation. Blood (10 to 50 µl) was obtained 15 min after the administration of tert-BOOH, from the orbital plexus, and red blood cells (RBC) were washed three times by centrifugation (5 min at 400 × g, each wash) and diluted to 108 cells/ml with 100 mM Chelex-treated sodium phosphate buffer, pH 7.4. Washed RBCs were broken by three freeze-thaw cycles and 15 µl (∼1 µg proteins) was used to detect protein-centered radicals by ELISA as described in Basic Protocol 1. Animal studies mentioned in this protocol were performed according to an approved Animal Study Proposal, National Institute of Environmental Health Sciences.



10 to 50 µl of blood in these experiments, and this was adequate to run the sample in triplicate. In theory, any compound that can generate oxygen-active species in vivo, such as H2 O2 or lipid peroxides, can induce protein-centered radicals that can be detected by immuno-spin trapping using very small amounts of sample.

Time Considerations Depending on the equipment available (e.g., multichannel pipets, automated microtiter plate washer and reader), an ELISA experiment can take ∼4 hr. Under the conditions described in this protocol, the authors did not find any difference between blocking overnight at 4◦ C and incubating for 40 to 90 min at 37◦ C with agitation. When an overnight experiment is required, it is important to let the plate and wash buffer stand at room temperature for at least 30 min to avoid gel formation, especially when gelatin is used as the blocking agent. Immunoblot analysis of DMPO-protein radical–derived nitrone adducts usually takes 5 to 7 hr including sample preparation if precast gels are available.

Literature Cited Berlett, B.S. and Stadtman, E.R. 1997. Protein oxidation in aging, disease, and oxidative stress. J. Biol. Chem. 272:20313-20316. Chen, Y.-R., Chen, C.-L., Liu, X., Li, H., Zweier, J.L., and Mason, R.P. 2004. Involvement of protein radical, protein aggregation, and effects on NO metabolism in the hypochlorite-mediated oxidation of mitochondrial cytochrome c. Free Radic. Biol. Med. 37:1591-1603. Clement, J.-L., Gilbert, B.C., Rockenbauer, A., and Tordo, P. 2001. Radical damage to proteins studied by EPR spin trapping techniques. J. Chem. Soc. Perkin Trans. 9:1463-1470. Davies, M.J. and Hawkins, C.L. 2004. EPR spin trapping of protein radicals. Free Radic. Biol. Med. 36:1072-1086. Dean, R.T., Fu, S., Stocker, R., and Davies, M.J. 1997. Biochemistry and pathology of radicalmediated protein oxidation. Biochem. J. 324:118. Deterding, L.J., Ramirez, D.C., Dubin, J.R., Mason, R.P., and Tomer, K.B. 2004. Identification of free radicals on hemoglobin from its self-peroxidation using mass spectrometry and immuno-spin trapping. J. Biol. Chem. 279:11600-11607. Detweiler, C.D., Deterding, L.J., Tomer, K.B., Chignell, C.F., Germolec, D., and Mason, R.P. 2002. Immunological identification of the heart myoglobin radical formed by hydrogen peroxide. Free Radic. Biol. Med. 33:364-369.

Guo, Q., Detweiler, C.D., and Mason, R.P. 2004. Protein radical formation during lactoperoxidase-mediated oxidation of the suicide substrate glutathione. Immunochemical detection of a lactoperoxidase radical-derived 5,5-dimethyl-1-pyrroline N-oxide. J. Biol. Chem. 279:13272-13283. Hawkins, C.L. and Davies, M.J. 2001. Generation and propagation of radical reactions on proteins. Biochim. Biophys. Acta 1504:196-219. He, Y.Y., Ramirez, D.C., Detweiler, C.D., Mason, R.P., and Chignell, C.F. 2003. UVAketoprofen-induced hemoglobin radicals detected by immuno-spin trapping. Photochem. Photobiol. 77:585-591. Mason, R.P. 2000. In vivo spin trapping-from chemistry to toxicology. In Toxicology of the Human Environment. The Critical Role of Free Radicals (C.J. Rhodes, ed.) pp. 49-70. Taylor and Francis, London. Mason, R.P. 2004. Using anti-5,5-dimethyl-1pyrroline N-oxide (anti-DMPO) to detect trapped protein radicals in time and space with immuno-spin trapping. Free Radic. Biol. Med. 36:1214-1223. Ramirez, D.C., Chen, Y.R., and Mason, R.P. 2003. Immunochemical detection of hemoglobinderived radicals formed by reaction with hydrogen peroxide: Involvement of a protein-tyrosyl radical. Free Radic. Biol. Med. 34:830-839. Ramirez, D.C., Gomez-Mejiba, S.E., and Mason, R.P. 2005. Mechanism of hydrogen peroxide– induced Cu,Zn-superoxide dismutase-centered radical formation as explored by immuno-spin trapping. The role of copper- and carbonate radical anion-mediated oxidations. Free Radic. Biol. Med. 38:201-214. Romero, N., Radi, R., Linares, E., Augusto, O., Detweiler, C.D., Mason, R.P., and Denicola, A. 2003. Reaction of human hemoglobin with peroxinitrite: Isomerization to nitrate and secondary formation of protein radicals. J. Biol. Chem. 278:44049-44057. Tijssen, P. 1985. Practice and theory of enzyme immunoassays. In Laboratory Techniques in Biochemistry and Molecular Biology, Vol. 15 (R.H. Burdon and P.H. van Knippenberg, eds.). Elsevier, New York.

Key References Detweiler et al., 2002. See above. Describes the production and validation of the antiDMPO antiserum and its application to the detection of DMPO-myoglobin radical-derived nitrone adducts. Mason, 2004. See above. Reviews all published literature relevant to immunospin trapping. Ramirez et al., 2003. See above. This report describes immuno-spin trapping and its validation for the detection of hemoglobin radical adducts in red blood cells. Oxidative Stress


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Internet Resources

http://www.alexis-corp.com

http://epr.niehs.nih.gov

http://www.oxfordbiomed.com

Laboratory where the anti-DMPO antiserum was developed

http://www.caymanchem.com

https://catalog.invitrogen.com/ Commercial supplier of gels and electrophoresis equipment http://www.piercenet.com Commercial supplier for protein staining, dialysis cassettes, and anti-rabbit (IgG)–alkaline phosphatase conjugate

Commercial suppliers of the rabbit anti-DMPO antiserum

Contributed by Dario C. Ramirez and Ronald P. Mason National Institute of Environmental Health Science, National Institutes of Health Research Triangle Park, North Carolina




CHAPTER 18 Immunotoxicology INTRODUCTION s evolutionary principles would predict, the development of multicellular organisms with specialized cells has lead to highly complex, but efficient, compartmentalization (separate organ systems) for select processes. This expanded developmental organization has allowed specialized activities to increase, as is most evident with the nervous system. However, along with increased structural adaptation of an organ for a particular function, which improves upon an organism’s adjustment to its environment, comes inter-organ dependency. The immune system also has been expanded in its specialization to assist with this interdependency. Immune cells and their factors (e.g., antibodies and cytokines) are critical for the homeostasis of each organ and maintenance of health.

A

The immune system has evolved to protect each organ against attack by pathogens and to accelerate the clearance of toxins and aberrant self cells, including those cells and cellular products altered by toxicants, drugs, carcinogenic transformations, and senescence. In performing the clearance of altered-self or non-self constituents, the immune system is unique, because somatic changes are constantly and consistently evolving to expand cellular clones to better recognize antigens. Furthermore, the antigen-specific cells (lymphocytes: B and T cells) and the innate, non-antigen-specific leukocytes (neutrophils, monocytes/macrophages, and NK cells) are continuously trafficking in search of altered-self and non-self antigens (immunosurveillance). Some immune cells also are relatively sessile cells in different organs: for example, the monocytic lineage includes the Langerhans cells of the skin, the Kupffer cells of the liver, the microglial cells of the brain, the mesangial cells of the kidney, and the osteoclasts of bones. Each of these cell types assist with normal organ functions as well as local defensive actions when necessary. The importance of the immune system is obvious when an individual is born without a particular immune component. Lack of T cells because of a deficit in thymus development (the primary immune organ of T cells), as with DiGeorge Syndrome, causes loss of cell-mediated immunity, which manifests as the inability to eliminate numerous opportunistic infections, such as Candida, Listeria, and Mycobacteria infections. Lack of B cells because of a deficit in bone marrow development (the primary immune organ of B cells), as with Bruton’s X-linked hypogammaglobulinemia, causes loss of humoral or antibody-mediated immunity, which manifests as the inability to clear exogenous pathogens, such as staphylococcal and streptococcal infections. Loss of both B and T cells (severe combined immunodeficiency disease, SCID) is a rapidly fatal disease unless major interventions are employed. Since the 1980’s, the HIV epidemic has created public awareness of the consequences of HIV-mediated depletion of CD4+ helper T cells (cells needed to promote both cell-mediated and humoral immunity), which can lead to acquired immunodeficiency syndrome (AIDS). It is now apparent that environmental agents depleting or altering the function of just one type of immune cell can induce major illness. Although usually more subtle than HIV in their mechanisms of immunomodulation, many environmental agents can modify immune functions. Since the immune system is composed of multiple cell types, which localize Immunotoxicology Contributed by David A. Lawrence Current Protocols in Toxicology (2005) 18.0.1-18.0.3 C 2005 by John Wiley & Sons, Inc. Copyright


in multiple organs and traffic throughout the organism, it is often arduous to delineate the cell type or cellular function being modulated, especially for clinical investigations when the only specimen usually available is blood. A further complication is age, in that early in life the immune system has not fully developed and late in life the immune system is intrinsically declining in function. Thus, the two most vulnerable times for the immunotoxic effects of environmental agents are during the neonatal and elderly stages. Additionally, immunomodulators are not restricted to pathogens and chemicals; physical and psychological stressors alone or combined with each other and/or chemical stressors can elicit changes of immune functions. Unfortunately, immune responses, which develop to foreign antigens, can subsequently lead to pathologies, such as allergies/asthma. Additionally, immune responses can develop to self-antigens, giving rise to autoimmune responses with major immunopathologies and possibly fatal outcomes. Besides chemical, physical, and/or psychological stressors causing immunosuppression, they also may heighten immune reactivities, which could generate an overly zealous immune response that causes pathology, or induce a loss of tolerance to self. Cell-mediated and antibody-mediated immunities to self constituents are differentially involved in autoimmune diseases. Thus, both immunosuppression and immunoenhancement can lead to various diseases. This emphasizes the need to assess potential immunotoxicological outcomes for an imbalance in either direction. The units in this chapter describe in vitro and in vivo immunological assays to qualitatively and quantitatively determine various immune system changes that could accompany exposure to environmental agents or drugs. provides a commentary on the various types of immunotoxicological analyses necessary to fully understand the importance of immune components and immunoregulation in maintenance of a healthy state. As alluded to earlier, the immune system is a defense network sending its cellular components to multiple sites including the skin. UNIT 18.2 describes the local lymph node assay. This is an assay to assess allergic contact dermatitis, which is a delayed-type hypersensitivity involving antigen-specific activation of T lymphocytes. Whereas the local lymph node assay evaluates allergic reactions originating in the skin, allergic hyperimmune reactions in the lungs can lead to airway hyperresponsiveness, which is referred to as asthma. UNIT 18.3 provides an overview of the initiators of asthma, the resultant pathology, and the methods to evaluate sentization and elicitation of the asthmatic response. Respiratory distress occurs with asthma as well as with respiratory infections or septicemia. In experimental models, immune infiltrates into lung tissue can be evaluated to determine the extent of the pathology and the involved mechanisms. However, good estimates of ongoing reactions also can be achieved with investigation of the immune cells and their products in bronchoalveolar lavages. Although these lavages are invasive and can be dangerous under some circumstances, they are an efficient means to assess immune reactivities in animals as well as humans. UNIT 18.4 describes how these lavages can be obtained and the methodologies utilized to assist in the determination of the associated lung injury. UNIT 18.1

The preceding units described more systemic approaches to evaluation of the immune system. UNIT 18.5 delves into the molecular mechanisms by which immune cells are regulated for the proliferative and maturational changes signaled through their surface antigenspecific receptors and accessory molecules. The means to quantify the activators of transcription by enzyme-linked immunoadsorbent assays (ELISAs) are described. Although gel-shift assays have long been the standard for analysis of transcription factors, the ELISA method described outlines substantially more efficient process for screening of multiple specimens. Introduction

provides the methodology to measure the cytolytic activity of natural killer (NK) cells and CD8+ T cells. Although these cell types kill by similar mechanisms, they

UNIT 18.6



represent the innate and adaptive arms of the immune system, respectively. The NK cells, often referred to as large granular lymphocytes, are not antigen-specific; whereas, CD8+ T cells are activated through their antigen-specific receptor (TCR) by specific peptide presented by major histocompatability complex (MHC) class I molecules. Toxicant interference with either of these cytolytic lymphoid subsets can substantially lessen host defenses. provide protocols that are integral to most immunotoxicological investigations as well as evaluation of any antigenic or structural changes of non-immune cells and factors. UNIT 18.7 deals with protocols that rely on antibody antigen interactions for quantification of either. The enzyme-linked immunoabsorbant assay (ELISA) has become a very standard method for multiple disciplines. For toxicological studies, however, it must be remembered that the assay relies on maintenance of the specificity of the antibody and conservation of the epitope of the antigen recognized by this specificity. A toxicant could potentially interfere with either compromising the quantitative nature of the assay. This possible complication exists for all assays that rely on complementary binding of proteins. The ELISPOT assay takes the ELISA to the level of the cells producing the antibody or antigen allowing quantification of the number of cells producing such factors. UNIT 18.7 additionally presents a new methodology with substantial promise for future endeavors; grating-coupled surface plasmon resonance imaging (GCSPRI) takes the ELISA and transforms it into a rapid screening method. With GCSPRI, hundreds of antibodies or antigens can be simultaneously quantified with a minimum of specimen. UNIT 18.8 describes the basics of flow cytometry procedures. Flow cytometry has become an important tool for biomedical and clinical studies. This unit provides critical methodologies that must be considered when using this method. Inaccurate results can be obtained if inappropriate gating or fluorescence compensation are not properly adjusted. Although the method can be useful for both quantification of the number of cells expressing a particular combination of markers and the quantification of these markers per cell, it must be realized that the instrument must be quality controlled for linearity and adjusted for optimal CV assessment of cell populations. This unit summarizes the important aspects that need to be considered when using flow cytometry for toxicological parameters. UNITS 18.7 & 18.8

embarks on methodologies capable of assessing whether a toxicant can alter the development of an antigen-specific naive T cell and/or modify the activity of a select subclass of T cell. The approach to assay antigen-specific naive T cell utilizes transgenic T cells genetically engineered to express the α and β chains of an antigen-specific T cell receptor (TCR). Thus, most of the T cells in these transgenic mice will have many CD4+ T cells or CD8+ T cells dependent on whether the TCR is specific for antigenic peptide in association with major histocompatibility complex (MHC) class II or class I molecules, respectively. However, to ensure that the T cells are naive, the researcher must isolate resting, non-memory T cells (CD62Lhi /CD44lo /CD45RBhi T cells). Alternatively, one can be assured of having naive T cells if the TCR transgenes are bred to Ragdeficient mice (mice with no antigen-specific lymphocytes except the TCR-transgene expressing T cells). In either case the mice need to be kept in specified pathogen free (SPF) conditions to maintain their health and lessen odds of activation by cross-reactive antigens. For analysis of differentiated T cells with specific effector functions, cloned T cells are utilized. Numerous Th1, Th2, Tc1 and Tc2 clones now exist, and can be obtained from various researchers. UNIT 18.9 describes some of these clones and their use for toxicological assessments. UNIT 18.9

David A. Lawrence Immunotoxicology


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Associating Changes in the Immune System with Clinical Diseases for Interpretation in Risk Assessment While it is well established that immunosuppression (see Terminology) can lead to an increased incidence and/or severity of infectious and neoplastic diseases, interpreting immunosuppression data from experimental immunotoxicology studies, or even epidemiological studies, for use in quantitative risk assessment is problematic. This is mostly due to a paucity of information on the health consequence of minimal-to-moderate immunosuppression, as might be expected to occur from inadvertent exposure to immunotoxic agents in humans. It is important that a scientifically sound framework be established that allows for the accurate quantitative interpretation of such data for use in the risk assessment process. In immunotoxicology, this may require, for example, development of a model to equate moderate changes in leukocyte counts, CD4 cell numbers, and/or immunoglobulin levels, tests which can be readily performed in human populations, to potential changes in the incidence or severity of infectious diseases, as well as establishing the social and economic impact of the incidence change. Although experimental animal models usually provide an opportunity to establish more reliable exposure estimates and conduct more informative immune tests than those that can be conducted in humans, extrapolating these findings across species is always of concern. The development of a framework to perform such extrapolations is currently being addressed by the authors’ workgroup. As an integral process to this undertaking, a review of studies that address the qualitative and quantitative relationships between immune parameters and disease is contained herein. Initially, the most likely clinical consequences that may occur as a result of chronic mild-to-moderate immunosuppression are described, as well as nonimmune factors that may modify these disease outcomes. Clinical and experimental animal studies that have examined immunosuppression disease relationships are reviewed and quantitative relationships, when available, are delineated to help address gaps in human health risk assessment. To address the potential social and economic consequences

that could result from immunotoxicity, a brief description of the general impact of infectious disease is provided on these parameters. The most comprehensive data bases that address immunodeficiency disease relationships, specifically primary (genetic) immunodeficiency and AIDS, are not discussed, as these represent extreme examples of immunosuppression and neither the specific clinical diseases that result nor the eventual outcomes have much in common to that which occurs in individuals with chronic mild-to-moderate immunosuppression.

TERMINOLOGY Before continuing, it is useful to provide clarification of certain terminology. Immunosuppression, immunodeficiency, and immunocompromised are nonquantitative terms that reflect a reduced capacity of the immune system to respond to antigens, and the terms are often used interchangeably in immunotoxicology. For the purpose of risk assessment, immunosuppression can be defined as a loss in the ability of the immune system to respond to a challenge from a level considered normal, regardless of whether clinical disease ensues. Immunodeficiency often represents an alteration in the immune system that can potentially lead to clinical disease, whether primary (i.e., genetic etiology) or secondary (epigenetic) in nature. The term, immunocompromised, like immunosuppression, indicates a deficient immune response, independent of whether or not it is maladaptive. Immunotoxicity encompasses each of these terms, but it specifies that the effect on the immune system originates from xenobiotic exposure.

DISEASES ASSOCIATED WITH IMMUNOSUPPRESSION As immunotoxicology testing is increasingly included in toxicological evaluations (House, 2003), there is added impetus to better associate detected changes with these tests to potential clinical outcomes. Although infectious disease is the most obvious, the consequences of maladaptive immunity can

Contributed by Michael I. Luster, Dori R. Germolec, Christine G. Parks, Laura Blanciforti, Michael Kashon, and Robert Luebke Current Protocols in Toxicology (2004) 18.1.1-18.1.20 C 2004 by John Wiley & Sons, Inc. Copyright

UNIT 18.1

Immunotoxicology


Figure 18.1.1 Changes in the onset, course, and outcome of infectious disease. Schematic shows factors which influence infectious disease susceptibility.

Associating Changes in the Immune System with Clinical Diseases

affect the etiology, progression, and/or severity of a much broader range of disorders, including certain cancers and autoimmune disease. Establishing the quantitative relationship between altered immune responses and frequency or severity of disease in human populations is challenging, as many factors may contribute (Morris and Potter, 1997). This is summarized schematically in Figure 18.1.1, where the appearance, progression, and outcome of infectious disease is viewed as an interrelationship between the virulence of the organism, infectious dose (number of organisms required to produce illness), integrity of the host’s anatomical and functional barriers, and overall immunocompetence of the individual. The latter is affected by genetics as well as age, gender, use of certain medications, drug/alcohol use, smoking history, stress, and nutritional status. These factors probably account for most of the variability reported in mean immune values, which can exceed two standard deviations. Another challenge when establishing quantitative associations between immune function and disease is to account for the functional overlap (i.e., redundancy) that exists between the various immune responses and disease. As multiple effector mechanisms are evoked in response to a disease, redundancy is often misinterpreted as reserve; however, as with other organ-systems, such as the liver or central nervous system, the existence of an immune reserve in the general population, where disease can occur in uncompromised individuals, is unlikely. In contrast, immune redundancy, as discussed by Halloran (1996), is

scientifically supported and can be empirically examined. The effect of redundancy on the interpretation of immunotoxicology studies was recently addressed by Keil et al. (2001) using factor analysis and multiple/logistic regression to quantitatively evaluate the contributions of different immune system parameters in host resistance.

Infectious Diseases While both infectious and neoplastic diseases are associated with secondary immunodeficiency, infectious diseases are often the focus of epidemiological studies as neoplastic diseases usually have such a long latency period. The particular microorganism responsible for an infection may assist in identifying the qualitative and quantitative nature of the immunodeficiency. For example, extracellular pathogens such as Streptococcus pneumoniae and Haemophilus influenza, multiply only outside phagocytic cells and, thus, produce disease only when they can resist phagocytosis. Facultative intracellular pathogens, (e.g., Mycobacterium tuberculosis), are generally phagocytized but resist intracellular killing. Thus, infections with extracellular or facultative intracellular organisms are more frequent in individuals with impaired phagocytic mechanisms, (e.g., neutropenia), or when humoral immune deficiencies are present. Obligate intracellular pathogens, which include all viruses, cannot multiply unless they are within a host cell and are more commonly observed in individuals with defects in cellular immunity.



Microbial agents associated with immunodeficiency disorders can also be classified into common, opportunistic, or latent pathogens. Common pathogens occur in the general population at frequencies associated with their infectious nature (e.g., virulence, ease of transmission), as represented by viruses that cause influenza infection and severe acute respiratory syndrome (SARS). The respiratory system is the most vulnerable target for common pathogens, as it is directly exposed to the external environment and has a large surface area: four times the combined total surface area of the gastrointestinal tract and skin (Gardner, 2001). Upper-respiratory infections occur in all age groups but produce the most severe effects in the very young and very old due to their lesser immunocompetency. Although influenza is responsible for more morbidity and mortality than any other infectious agent in recorded history (Patriarca, 1994), the low individual rates of infections in the general population (only one or two episodes in an individual per year), combined with underreporting, make it difficult to detect changes in infection rates in most epidemiological studies. While infections with common pathogens occur routinely in the healthy population, opportunistic infections are typically seen in individuals with severe immunosuppression, such as AIDS patients. Examples of microorganisms that can produce opportunistic infections are certain protozoans, including Toxoplasma gondii, which cause cerebral infections and intractable diarrhea, Candida albicans, Mycobacterium avium complex (MAC), and Pneumocystis carinii, a fungus causing severe lung diseases in AIDS patients (Morris and Potter, 1997). These organisms are commonly encountered in food, water, dust, or soil, but they cause disease in the general population at very low incidences. Certain pathogenic microorganisms are responsible for latent infections. In the case of members of the herpes virus family, including cytomegalovirus (CMV), herpes simplex virus (HSV), and Epstein-Barr virus (EBV), the virus remains in the tissue in a latent form following primary infection for the duration of the host’s life. In healthy individuals, the immune system maintains viral latency, with cellular immunity playing a major role. When the cellular immune response is compromised, viral replication can ensue and potentially cause severe complications or death. Preceding viral activation, a vigorous immune response to viral-specific antigens occurs in response to

replication. As will be discussed later, changes in virus-specific immune response or activation of latent viruses has been observed in individuals with secondary immunodeficiency disorders where mild-to-moderate immunosuppression may exist.

Virally Induced Tumors Immunodeficiency is also associated with an increased incidence of certain virally induced tumors, such as non-Hodgkin’s lymphomas (NHLs) and tumors of the skin (Penn, 2000). In contrast to cancers of internal organs, in particular those in the lung and liver, which are often induced by chemical carcinogens, virus-induced cancers are more immunogenic and, thus, more likely influenced by immunological factors. Suppression of cellmediated immunity has been associated with higher incidences of skin cancers, leukemias, and lymphoproliferative disorders in transplant patients, whereas Kaposi’s sarcoma and EBV-associated B cell lymphomas are associated with severe immunosuppression as seen in patients with AIDS. Natural killer (NK) cells are more likely to play a role in resisting the progression and metastatic spread of tumors once they develop, rather than preventing initiation (Herberman, 2001). Unexpectedly, studies of individuals with NK cell deficiency states, most of which are associated with single gene mutations, have helped identify a role for NK cells in defense against human infectious disease. A resounding theme of NK cell deficiencies is susceptibility to herpes viruses, suggesting that unexplained severe herpes viral infection should raise the possibility of an NK cell deficit (Orange, 2002).

CONSIDERATIONS IN THE USE OF EPIDEMIOLOGICAL DATA IN IMMUNOTOXICOLOGY RISK ASSESSMENT There are many advantages of using human data over experimental animal studies in quantitative risk assessment, especially as it avoids the difficulties in interspecies extrapolation and provides data on lower doses that are of interest to public health policy makers (Hertz-Picciotto, 1995). Human studies offer realistic exposure scenarios, including multiple routes of exposure, and include a much more diverse range of genetic backgrounds than experimental models, providing the potential to explore differences in susceptibility by genotype. The limitations and challenges of human studies, however, are considerable.

Immunotoxicology


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Here, a brief overview of issues surrounding the design and interpretation of human studies as it pertains to the assessment of risk due to immunotoxic exposures is provided.

Clinical Studies The design of human studies can range from controlled clinical trials to large, populationbased, observational studies. Clinical studies offer advantages in that exposure parameters of interest can often be controlled (e.g., chamber studies of inhaled toxicants, challenge studies of adenovirus infection), and outcomes can be prospectively monitored. There are also disadvantages, as ethical considerations provide little opportunity for exposure with toxic chemicals. Furthermore, studies with extensive biological monitoring and functional immune tests can be expensive, and exposures as well as outcomes of interest may be difficult to study in the available time frame as study participants are not typically available for longterm exposures or extended follow-up. For the purpose of obtaining data for immunotoxicological risk assessment, clinical studies are particularly useful as they can provide data on the frequency of infections or the level of immune response to vaccines. Variations on this type of study design might include follow-up of patient populations administered immunosuppressive therapy (i.e., transplant patients), that, as described below, may also have many of the characteristics of observational studies.

Epidemiological Studies

Associating Changes in the Immune System with Clinical Diseases

Other types of human studies that have been employed in immunotoxicology are typically classified as observational or epidemiological. Observational studies can be of varying size and they can be cross-sectional (one point in time), retrospective, or prospective in nature; each design has advantages and disadvantages. The initial means of control in observational studies is introduced through the study design and the quality and validity of results can be greatly affected by the methods used to select the study sample and the rigor with which exposures and outcomes are measured. In addition to high costs, observational studies are challenging for many reasons, including potential confounding by host (age, gender, and lifestyle) and environmental (frequency of exposure to chemicals and infectious agents) factors. A secondary measure of control in observational studies is through the use of multivariable analysis techniques (e.g., regression modeling), providing there is sufficient sample

size and information on potential confounders. Overall, well-designed epidemiological studies (e.g., absence of selection bias, exposure or outcome misclassification, control of confounding factors) can contribute valuable information to the assessment of risk due to immunotoxic exposures. Existing immunotoxicology studies in humans tend to be based upon either fairly small sample sizes, often in individuals with transient high-level occupational exposures, or large groups with chronic low-level exposures. Although in some instances body burdens of chemicals have been determined, drawing broadly applicable conclusions from some of these studies is difficult. This is because subjects may have been exposed to chemicals other than those specifically addressed by the study, and characterization of chemical exposure may rely on subject recall or rough estimates of the duration and intensity of exposure. Furthermore, in contrast to experimental animals, functional assessment is considerably more difficult in humans as it requires antigen challenge, which involves some risk to the individual. When such studies have been undertaken, subjects have been provided commercial vaccines, such as hepatitis antigen (Weisglas-Kuperus et al., 2000; van Loveren et al., 2001; Yucesoy et al., 2001; Sleijffers et al., 2003). The cellular and humoral immune response to vaccination is thought to be a sensitive indicator of immunosuppression (Glaser et al., 1993) and can reflect susceptibility to infectious disease (Deseda-Tous et al., 1978; van Loveren et al., 2001). In most epidemiological studies, testing in humans has been limited to blood collection where peripheral cell counts and differentials, immunoglobulin levels, or immunophenotyping are performed. While certainly of value, it is generally agreed these are not highly sensitive indicators of immunosuppression, making it difficult to detect low-to-moderate levels of immunosuppression (ITC, 1999).

IMMUNODEFICIENCY AND RELATIONSHIP TO INFECTIOUS DISEASE Environmental Chemicals PCBs The need to extend data obtained in experimental studies to humans has been recently reviewed (Tryphonas, 2001); however, epidemiologic data on the effects of chemical exposures on immune parameters and



infectious outcomes in human populations are limited. Some of the more complete immunotoxicology studies have focused on persistent organochlorine compounds formerly found in pesticides and industrial chemicals—e.g., polychlorinated biphenyls (PCBs)—in children following prenatal or perinatal exposure via maternal diet and breast milk. Accidental exposures of populations in Japan (Yusho) and China (Yu-Cheng) suggest an association of PCBs, their thermal breakdown products (quaterphenyls), and polychlorinated dibenzofurans, found in contaminated rice oil, with immune abnormalities and increased infections. Children born to exposed mothers between 1978 and 1987 in the Yu-Cheng population had lower levels of serum IgA and IgM, and a higher frequency of respiratory infections and otitis media compared to matched unexposed controls (Lu and Wu, 1985; Nakanishi et al., 1985; Yu et al., 1998). The association between PCBs and increased frequency of otitis media in children has also been described in other populations. A study of 343 children in the United States (Michigan) showed no general association between organochlorine levels and prevalence of infections, but there was a positive association between polychlorinated biphenyls (PCBs) and DDE (the primary metabolite of DDT), or PCBs and hexachlorobenzene with otitis media (Karmaus et al., 2001). In a study of Inuit infants in Arctic Quebec, Canada (Dewailly et al., 2000), the relative risk of recurrent episodes (at least three per year) of otitis media was higher in breast-fed infants in the second and third highest percentile of organochlorines exposure, compared to the lowest. At three months of age, breast-fed infants with higher exposure levels had lower numbers of white blood cells and lymphocytes, and lower serum IgA levels at ages 7 and 12 months, compared to bottle-fed infants. In Dutch preschool children (WeisglasKuperus et al., 2000), PCB levels in breast milk (nonortho and planar PCBs) were also associated with increased recurrent otitis media and other symptoms of respiratory infection. In this sample, the body burden of PCBs at age 42 months was associated with higher prevalence of recurrent otitis media and chicken pox. PCB body burden was not associated with differences in lymphocyte markers outside the normal range for age-matched children, although levels in breast milk and cord blood were positively correlated with lymphocyte counts and various T-cell subsets. While these findings linking otitis media with PCB exposure are

consistent across three studies, it was not possible to determine whether the changes in immune parameters mediated this association or simply represented parallel findings.

Pesticides The immunotoxicity of pesticides following human exposure has been reviewed by several authors (Thomas et al., 1995; Vial et al., 1996; Voccia et al., 1999; Luebke, 2002). Although some studies have described associations between pesticide exposure, altered immune function, and increased rates of infection, sample sizes were generally small and, in some cases, patients were self-selected based on symptoms rather than exposure. Furthermore, the frequency of infections was typically estimated by recall over several years and immune function data were scarce. Not all studies suffer from these shortcomings. For example, a relatively large (n = 1600) and well-defined population living in and around Aberdeen, North Carolina, near a pesticide dump site (a priority Superfund site containing organochlorine pesticides, volatile organic compounds, and metals), was evaluated for immune function and frequency of viral infections. Compared to a neighboring community, residents of Aberdeen, ages 18 to 40, were found to have higher incidence of herpes zoster (reactivated herpes infection causing shingles), but no difference in the frequency of other infectious diseases (Arndt et al., 1999). In a substudy of 302 individuals, those living in Aberdeen had significantly higher age-adjusted levels of plasma DDE than those living in neighboring communities. Furthermore, higher levels of plasma DDE were related to lower lymphocyte responses to mitogens but higher absolute lymphocyte counts and IgA levels (Vine et al., 2001). In a separate analysis, residents living nearer to the pesticide dump site had both a lower lymphocyte response to mitogen stimulation and a greater likelihood of having a lower percentage of CD16+ (NK) cells, (5%) non-specific lysis, as indicated by high counts per minute in spontaneous release (SR) wells

Insufficient washing of target cells Poor target cell viability

Add more washes or increase the volume of medium used in each wash. Poor target cell viability after labeling.

Very low counts per minute Poor 51 Cr uptake by target in MR wells cells Poor target cell viability

Use the freshest 51 Cr available, and do not use if decayed (even decaying by one half-life can adversely affect the sensitivity of these assays). Poor target cell viability after labeling.

Very low counts per minute Improper effector cell in samples containing (or sensitization. Poor cellular viability. expected to contain) effector cells

Validate the efficacy of effector cell sensitizaiton using an alternate method (e.g., flow cytometry, cytokine production, or proliferation). Use cells isolated from animals as quickly as possible after the animal is sacrificed. Check the pH of solutions and wash the cells more gently.

Poor target cell viability after 51 Cr labeling

Use only viable cells. Wash cells more gently and very carefully maintain solution pH. Do not let the labeled target cells sit unused for >30 min.

Poor cellular viability prior to labeling Cells were not used within 30 min of labeling

1. To test whether in vitro exposure to a chemical affects the clonal expansion and differentiation of T cells (see Basic Protocol 1). 2. To test whether in vivo exposure to a chemical affects the clonal expansion and differentiation of T cells (see Alternate Protocols 1 and 2). 3. To determine whether exposure to a particular substance affects the inherent cytolytic function of NK cells (see Basic Protocol 2). 4. To determine whether exposure to a substance has differential effects on the native and boosted cytolytic activity of NK cells (see Basic Protocol 2). 5. To determine whether exposure to an agent exhibits differential or selective effects on CTL and NK cells (a combination of Basic Protocol 1 or Alternate Protocols 1 or 2 and Basic Protocol 2 in separate but simultaneous assays).

Critical Parameters All animals used for these procedures should be housed in specific pathogen-free

conditions and all media should be free from bacterial endotoxin and other immunostimulatory contaminants. Cells isolated from animals should be used as quickly as possible after animal sacrifice.

Troubleshooting See Table 18.6.1 for troubleshooting tips.

Anticipated Results If the effector cells recognize and kill the target cells, then the counts per minute and calculated percent specific cytolytic activity should increase with increasing E:T ratios, as depicted in Figure 18.6.1. On the other hand, the amount of cytolytic activity in wells set up using control effector cells should not change with increasing E:T ratios. In fact, the counts per minute in wells containing the negative control cells cultured with target cells ought to be about the same as the spontaneous release of the target cells. The maximum achievable percent cytolytic activity will vary depending upon the type of

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Figure 18.6.1 This graph depicts the average cytolytic activity of spleen cells in an in vitro chromium release assay. Spleen cells were isolated from a C57B1/6 mouse 9 days after i.p. injection with P815 mastocytoma cells (closed circles). As a control, spleen cells from a na¨ıve mouse (open circles) were cultured with the labeled-P815 cells, which served as the target cells for this assay.

effector cells used and how they were sensitized. Lysis of 25% to 60% of the target cells at the highest E:T ratio is the typical range expected in chromium release assays for both NK cells and CTL. Lysis of 100% of the target cells by the effector cells should not be expected, and if observed, it likely indicates a technical problem with the assay. When measuring the activity of NK cells, treatment with poly I:C should lead to increased activity compared to native NK cell activity toward Yac-1 tumor cells, however, the magnitude of this change is not more than 2-fold (e.g., increasing the percent cytolytic activity from 35% to 50% at an E:T ratio of 100:1 is typical).

Time Considerations

Measuring the Activity of Cytotoxic Lymphocytes

A chromium release assay requires 1 full day. It takes ∼3 hr to collect the effector cells and label the target cells, and these steps need to be performed simultaneously. The total time required to set up the assay (i.e., combine the effector and target cells) depends upon the size of the assay, but generally takes ∼1 hr. The cells need to be incubated for 4 to 5 hr prior to harvesting supernatants. The duration of this step depends upon the size of the assay (number of 96-well plates), but generally takes ∼1 hr. While general steps and overall time commitment for the chromium release assays described here is similar among the protocols described, the total amount of time required to measure the cytolytic activity of CTL and NK cells is very different. CD8+ T cells require sensitization with antigen to develop into CTL.

Therefore, whether one uses an in vitro or in vivo approach, it takes 5 to 10 days to generate a population of CTL. The innate and in vitro– boosted cytolytic activity of NK cells can be measured in 1 day. However, if one wants to stimulate NK cells in vivo prior to measuring the cytolytic activity, then the assay will require 1 additional day for this step.

Literature Cited Barrett, T. and Inglis, S.C. 1985. Growth, purification and titration of influenza viruses. In Virology: A Practical Approach (B.W. J. Mahy, ed.), pp. 119-150. IRL Press, Washington, D.C. Cho, Y., Basta, S., Chen, W., Bennink, J.R., and Yewdell, W. 2003. Heat-aggregated noninfectious influenza virus induces a more balanced CD8+T lymphocyte immunodominance hierarchy than infectious virus. J. Virol. 77:4679-4684. Kagi, D., Ledermann, B., Burki, K., Zinkernagel, R.M., and Hengartner, H. 1996 Molecular mechanisms of lymphocyte-mediated cytotoxicity and their role in immunological protection and pathogenesis in vivo. Annu. Rev. Immunol. 14:207-232. Kos, F. and Engleman, E. 1996. Role of natural killer cells in the generation of influenza virus–specific cytotoxic T cells. Cell Immunol. 173:1-6. Mitchell, K.A. and Lawrence, B.P. 2003a. Exposure to 2,3,7,8-tetrachlorodibenzo-p-dioxin (TCDD) renders influenza virus–specific CD8+ T cells hyporesponsive to antigen. Toxicol. Sci. 74:7484. Mitchell, K.A. and Lawrence, B.P. 2003b. T cell receptor transgenic mice provide novel insights into understanding cellular targets of TCDD: Suppression of antibody production, but not the response of CD8+ T cells, during infection with influenza virus. Toxicol. Appl. Pharmacol. 192:275-286.



Nonacs, R., Humborg, C., Tam, F., and Steinman, R. 1992. Mechanisms of mouse spleen dendritic cell function in the generation of influenzaspecific cytolytic T lymphocytes. J. Exp. Med. 176:519-529. Russell, J.H. and Ley, T.J. 2002. Lymphocytemediated cytotoxicity Annu. Rev. Immunol. 20:323-370. Warren, T.K., Mitchell, K.A., and Lawrence, B.P. 2000. Exposure to 2,3,7,8-tetrachlorodibenzo-pdioxin (TCDD) suppresses the humoral and cellmediated immune responses to influenza A virus without affecting cytolytic activity in the lung. Toxicol. Sci. 56:114-123.

Internet Resources http://www.cdc.gov/od/ohs/biosfty/bmbl4/ bmbl4toc.htm A pdf manual of Biosafety in Microbiological and Biomedical Laboratories, 4th Edition (1999), U.S. Department of Health and Human Services, Centers for Disease Control and Prevention, and National Institutes of Health. US Government Printing Office Washington, D.C.

Contributed by B. Paige Lawrence Washington State University Pullman, Washington

Yokoyama, W.M. and Plougastel, B.F.M. 2003. Immune functions encoded by the natural killer gene complex. Nat. Rev. Immunol. 3:304-316.

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Solid-Phase Immunoassays

UNIT 18.7

Solid-phase quantitative immunoassays can be configured in a multitude of ways to detect the presence and concentration of a specific antigen or an antibody of a particular isotype and specificity. These assays can provide a wealth of information about cellular and molecular events that are altered by toxicant exposure. For example, these assays can produce information related to antibody or antigen concentrations, the affinity and avidity of the interactions that produce immune complexes, and the organ, tissue, cellular, and subcellular distribution of target molecules (Probst et al., 1995). In the context of toxicological assessment, these assays provide a window on the effects that chemical and physical agents can have on developmental processes and effector mechanisms in the body. Since the immune system itself is often a target of toxin effects, evaluation of immune function can be extremely informative, but these same assay formats can be used to probe other biological systems with equal ease.

STRATEGIC PLANNING The enzyme-linked immunosorbent assay (ELISA) is a multi-step process in which a target molecule (e.g., an antigen) is first immobilized onto an insoluble substrate—most often composed of polystyrene or another plastic in a 96-well microtiter plate configuration. Ordinarily, this immobilization is a non-specific binding event and must be followed with a blocking reagent to saturate any other non-specific binding sites on the substrate. A number of different kinds of reagents have been used to block subsequent non-specific binding, and examples of some of the common blocking agents are listed in Table 18.7.1. Bovine serum albumin (BSA) is most commonly used, but has some drawbacks under certain circumstances. BSA is a particularly sticky protein that can increase undesirable non-specific interactions. Moreover, polyvalent sera that have been raised against cultured cells may have secondary reactivity against BSA that was carried along with the immunizing cells from the fetal bovine serum that is often used to supplement culture media. Teleostean fish gelatin has the advantage of being antigenically distinct from the antigens present in samples associated with many common applications of the ELISA technology, and thus it will not often produce spurious cross-reactions. Since this protein derives from cold-water fish, it is soluble at room temperature, and will not gel in the microplate wells.

Table 18.7.1 Blocking Agents

Commonly used concentration range

Blocking agent

Suppliers

Bovine serum albumin (BSA)

Jackson ImmunoResearch Laboratories Sigma-Aldrich

1%–10%

Casein

Sigma-Aldrich

5%–10%

45% Teleostean fish gelatin in water

Sigma-Aldrich

1%–5% dilution of stock solution

Non-fat dried milk (NFDM)

Local grocery store, Sigma Chemical

2%–5%

Goat serum

Sigma-Aldrich

1%–10% Immunotoxicology

Contributed by Michael A. Lynes Current Protocols in Toxicology (2005) 18.7.1-18.7.19 C 2005 by John Wiley & Sons, Inc. Copyright


Table 18.7.2 Enzymes Used in Conjugates and Representative ELISA Substrates

Detection method for substrate

Enzyme

Complementary substrates

Enzyme limitations

Alkaline phosphatase (AP)

a. Para-nitrophenyl phosphate a. Measure absorption at (PNPP) 405 nm b. 4-Methyl-umbelliferyl phosphate b. Excitation at 360 nm and (4-MUP) emission at 440 nm

Samples that contain a high level of endogenous phosphatase are not compatible with this enzyme conjugate

ß-galactosidase (ß-gal)

a. o-Nitrophenyl-ß-Dgalactopyranosidase (ONPG) b. 4-Methyl-umbelliferyl-ß-Dgalactopyranosidase (MUM-Gal)

a. Measure absorption at 405 nm b. Excitation at 360 nm and emission at 440 nm

This enzyme is not as active as the alternatives and produces a lower signal over time

Horseradish peroxidase (HRP)

a. 3,3 ,5,5 -Tetramethylbenzidine (TMB) b. 2,2 -Azino-di-(3-ethylbenzthiazoline sulfonic acid (ABTS)

a. Measure endpoint at 370 or 655 nm, or acidify and measure at 450 nm b. Measure at 416 nm 405 nm will also work

Do not use this enzyme conjugate if the sample contains oxidizing or reducing agents, or some preservatives

Once the surface has been blocked, the complementary molecular partner (e.g., a cognate antibody conjugated to an enzyme label) can be added to the surface and allowed to selectively interact with the target molecule. Following an appropriate period, the surface can be washed to remove any unbound antibody-enzyme conjugate. The next step is to add an appropriate substrate for that enzyme. The more enzyme that is present, the more substrate will be converted to a product that can be measured in a microtiter plate reader. Measurements of the amount of substrate conversion to product can be interpreted to indicate the degree of antibody-enzyme conjugate that has bound to the surface. A list of the most common enzyme labels and their substrates is provided in Table 18.7.2. Substrates must be stable until they interact with enzyme, must produce a soluble product, and should have a large extinction coefficient to maximize the signal-to-noise ratio. In some cases, fluorescent substrates can improve the sensitivity of an assay by 100- to 1000-fold over an absorbance assay, but these assays do require fluorescent microtiter plate readers that are more expensive and less commonplace than standard absorbance microtiter plate spectrophotometers. In addition to the listed substrates, there are also commercially available chemiluminescent substrates (often used with horseradish peroxidase–labeled antibody conjugates) that can further enhance the detection limits of ELISA. The disadvantage of chemiluminescent systems is that the signal persists for only a brief time (for 5 to 30 min) and then begins to degrade. Moreover, readings must be taken in a microtiter plate luminometer.


Another substrate alternative that can enhance the sensitivity of the ELISA is an enzymatic recycling system (e.g., commercially available from Invitrogen). In this type of substrate system, immobilized alkaline phosphatase catalyzes the conversion of the reduced form of nicotinamide adenine dinucleotide phosphate (NADPH) to reduced nicotinamide adenine dinucleotide (NADH). The NADH can then participate in a cyclic reaction in which diaphorase reduces a tetrazolium salt (iodonitrotetrazolium violet or INT-violet) to colored formazan and produces oxidized NAD+ . To continue the cycle, NAD+ is then reduced back to NADH and ethanol is oxidized to acetaldehyde in the presence of alcohol dehydrogenase. The NADH can be recycled through this process and yields an approximately ten-fold signal amplification over that of standard ELISA protocols when absorption is read at 495 nm.



One of the initial issues to be considered for the design of an ELISA is the availability of antibodies specific for the target to be assessed. There are many sources of commercial antibodies that produce reagents of differing specificity, concentration, purity, quality, isotype, and species of origin. Identification of the appropriate reagent must take into account the singular or multiple applications for which the antibody will be used. For example, it may prove useful to employ a polyclonal antibody that cross-reacts with homologous antigens from different species to facilitate the assessment of cellular functions in cell cultures of different origins. Alternatively, a monoclonal antibody may enhance the specificity of the antibody/antigen interaction by minimizing the appearance of crossreactions that decrease the signal-to-noise ratio of the assay. Equally important, it may prove useful to employ antibodies that are biotinylated to facilitate the use of a spectrum of different labeling reagents that have each been coupled to avidin. In this way, the same biotinylated antibody reagents incubated with an avidin-enzyme conjugate that could be used for an ELISA would also be useful in fluorescent assays when incubated with an avidin-fluorescent tag. Once the appropriate antibodies have been identified, it is often useful to obtain a source of purified antigen for use as a standard in the reaction. While this is not always possible (or economical), it can serve to enrich the interpretations that are possible from data generated by the ELISA. Having obtained the central reagents, the next step is to select the specific kind of ELISA that will be used. Figure 18.7.1 illustrates the most commonly used ELISA configurations.

Figure 18.7.1

ELISA protocol formats.

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The simplest of these assays is the direct ELISA (Fig. 18.7.1A). This assay is usually configured with non-specifically adsorbed target antigen on the surface of the microtiter plate well. The amount of adsorbed antigen is detected using a pre-established amount of cognate antibody coupled to an enzyme label. The more antigen that is immobilized on the surface, the more antibody-enzyme conjugate will subsequently bind. Ultimately, the larger the amount of enzyme that is immobilized after binding to the surface, the more detection reagent will be converted to a signal that can be measured by light absorption, fluorescence, or luminescence. A common variation of this assay is used to quantitate the amount of antibody specific for a particular antigen that is present in a sample. In this version of the ELISA (Fig. 18.7.1B), known amounts of antigen are immobilized on the surface of the microtiter plate well. After blocking the surface to prevent further non-specific binding, the surface is incubated with the sample that may contain antibody specific for the target antigen. If there is antibody present in the sample, it will be retained on the surface after washing, and will serve as a target for a secondary antibody that has been conjugated to enzyme. The anti-immunoglobulin-enzyme conjugate is added in a standardized amount (usually in excess), and thus the amount of signal that develops is directly proportional to the amount of primary antibody that has bound to the surface. If the antigen to be detected is present in a heterogenous sample mixture that also contains other molecules, the actual detection of antigen will be subject to the influences of the competitive binding of those different molecules from the sample. For this reason, an ELISA that employs a capture antibody is more commonly used (Fig. 18.7.1C and D). Unconjugated antibodies against the target antigen are first immobilized on the substrate, and the surface is then blocked to prevent further irrelevant binding to the surface. At this point, antigen that is incubated on this surface will be captured by specific binding to the capture antibody. Irrelevant molecules will be left unbound. After washing away unbound molecules, antigen that remains bound to the surface can then be detected with antibody-enzyme conjugate in a direct ELISA with capture antibody, or in an indirect ELISA configuration. Finally, the ELISA can be configured as a competition assay (Fig. 18.7.1E and F). In these assays, soluble antibodies will bind to immobilized antigen unless additional free antigen is added to the solution. As the amount of free antigen in solution increases, the amount of antibody that will bind to the immobilized substrate decreases. Antibody binding is again determined by the amount of directly conjugated enzyme that is present in a direct ELISA, or by subsequent measures of the amount of anti-immunoglobulin antibody-enzyme conjugate binding that can be made in an indirect ELISA. Competition ELISAs are particularly useful for measurements of antigen concentration in complex mixtures when the unknown samples that may contain antigen are compared to similar samples that contain known amounts of purified antigen.


The ELISPOT (enzyme-linked immunoSPOT) assay is a useful variation of the basic ELISA format. While ELISA measures the concentrations of analyte in solution, the ELISPOT enumerates cells that are actively secreting a specific protein (e.g., cytokine or immunoglobulin) in a heterogeneous population of cells. This assay differs from the basic ELISA protocol in both the configuration of the microtiter plate, and in the chemical characteristics of the enzyme substrate that is used. The microtiter plate wells that are used have filter bottoms. The filters are first coated with sterile capture antibody or antigen. Cells are then added to the wells of the plate and settle to the filter surface. Cells that secrete molecules that can be bound by the capture antibody, or cells secreting antibodies specific for immobilized antigen will secrete those molecules to a region immediately surrounding the cell. The analyte will bind to the filter-immobilized cognate molecule in highest concentrations nearby the cell source of the analyte. As analyte diffuses away from the cell, its concentration diminishes. Following an appropriate incubation period,



a secondary antibody-enzyme conjugate that is specific for the analyte is added and allowed to bind. Finally, enzyme substrate that produces a precipitating colored product is added. As the substrate is converted to this precipitating product, color will accumulate in specific regions of the filter that correspond to the original locations of the cells that secreted the analyte. These spots of accumulated color are proportional to the amount of analyte bound to the filter, and the precipitate spots can be counted microscopically and compared to the number of cells originally added to each individual well. A new version of the solid-phase immunoassays is the grating-coupled surface plasmon resonance (GCSPR) assay. This assay has been used as a label-free variant of ELISA, and offers some interesting advantages over the more commonly used ELISA. GCSPR is based on the phenomenon of energy transfer that occurs at a metal-dielectric interface (e.g., the interface between gold and water) when light energy is coupled to the electrons within the metal. Under certain optical conditions, the energy of the light excites electron movement within the metal. This excitation reduces the intensity of the reflected light. SPR technologies match the wave vector of the illuminating beam of light with the plasmon wave vector using a prism (the Kretschman configuration) or with a grating embossed on the gold surface. There is a critical angle at which this vector matching is best, and at which the energy transfer into the metal is largest. The angle at which maximum energy transfer occurs is referred to as the SPR angle. Because the measurement of energy transfer is sensitive to the refractive index of the dielectric/metal interface, the addition of proteins to the surface (which have a higher index of refraction than water) will increase the angle at which coupling occurs. Antibodies or other capture molecules immobilized on the surface that subsequently capture analyte from the sample solution will cause an increase in the critical angle of reflection. GCSPR offers a significant advantage over the Kretschman configuration because the entire surface of the sensor chip can be illuminated, and images of the chip surface across a range of illuminating angles can be captured. Individual regions of interest (ROI) on the chip surface can be computationally assessed for protein capture by immobilized antibody according to the changes in SPR angle at each ROI. Thus, GCSPR offers a microarray format for the simultaneous analysis of multiple proteins in solution from a single small volume sample. This unit describes several prototypical variations of these three types of solid-phase immunoassays and considers the use of these assays in the evaluation of toxic chemical effects on biological systems. Several different configurations of ELISA are described in Basic Protocol 1 (direct ELISA), Alternate Protocol 1 (indirect ELISA), and Alternate Protocol 2 (competitive ELISA). Basic Protocol 2 describes the ELISPOT assay, and Basic Protocol 3 describes the GCSPR protein microarray application.

DIRECT ENZYME-LINKED IMMUNOSORBENT ASSAYS In this simplest of ELISA configurations (Fig. 18.7.2), known and potential sources of antigen are first incubated in wells of a microtiter plate. The known antigen solutions act as positive controls for those wells in which potential sources of antigen are added. It is advantageous to maximize antigen coupling to the surface, and so antigen is often diluted in a coupling buffer that will facilitate binding to the microtiter plate material. Incubation of the antigen solution in the well can be done for up to 1 hr at 37◦ C, or overnight at 4◦ C. Then the antigen is aspirated from the well, and the well is filled with a larger volume of blocking agent in phosphate-buffered saline. After blocking, the well is washed and then incubated with the antibody-enzyme conjugate, washed again, and then substrate is added. After incubating the substrate in the wells for a defined period, a single, endpoint reading can be taken to measure the amount of substrate conversion to product. Alternatively, a kinetic measurement can be made in which multiple sequential measurements of the optical density of the substrate are taken and then converted to a milli-optical density

BASIC PROTOCOL 1

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Figure 18.7.2


Sequential ELISA steps.

(mO.D.)/min reading, or a rate of substrate conversion. This second approach to quantifying the amount of enzyme in the well has the advantage of diminishing the differences that might result from the pipetting sequence. Wells initially loaded with substrate have more time to develop color than will wells loaded at a later time, but if the wells have equivalent amounts of enzyme, the rates of substrate conversion to colored product should be equal. Another advantage of the kinetic measurement of substrate conversion is that measurements can be started immediately after substrate addition. Usually 5 to 10 min of sequential measurements at 20- to 30-sec intervals from each well are sufficient to produce reliable estimates of substrate conversion rates. It is important to ensure that the conversion rate is linear and does not plateau, or the mathematical estimate of the absolute rate will be lower than the real rate. For that reason, the plate should be shaken in between readings to prevent substrate depletion at the surface of the microtiter plate well, and the assay should be set up to avoid approaching the maximum O.D. that the instrument can measure. For the same reason, substrate should be present in excess in each well. If the plate reader does not offer the capacity to do kinetic measurements, then the substrate should be incubated in each well for a fixed period of time (usually 30 min at 37◦ C), and the individual wells measured as an endpoint assay. If the rates of substrate conversion are rapid, the activity of the enzyme should be stopped before reading the plate. This is usually done by changing the pH of the substrate solution; for alkaline phosphatase conjugates conversion of p-nitrophenyl phosphate disodium hexahydrate (PNPP), the reaction can be stopped with the addition of 3 N NaOH.



Materials Antigen diluted in ELISA coupling buffer (see recipe for ELISA coupling buffer) Blocking reagent (Table 18.7.1) ELISA wash buffer (see recipe) Antibody-enzyme conjugate (e.g., Southern Biotech) Substrate (Table 18.7.2) dissolved in the appropriate substrate buffer (e.g., for alkaline phosphatase-based assays, the most common substrate is p-nitrophenyl phosphate, PNPP, Sigma; see recipe) 96-well flat-bottom microtiter plates (plates specifically treated to enhance binding of hydrophobic or hydrophyllic proteins, or DNA molecules; e.g., Immulon 2 HB plates, Thermo Electron) 37◦ C incubator Automated plate washer (optional) Plate sealer (adhesive-backed mylar sheets, Fisher) Microtiter plate reader with appropriate wavelength filters (e.g., Molecular Devices, Bio-Tek Instruments, Thermo Electron, and Tecan) Plate shaker Determine working dilutions of antigen and antibody-enzyme conjugate 1. Determine working dilutions of the antibody-enzyme conjugate and the immobilized target molecule in a 96-well microtiter plate. Set up the microtiter plate in a checkerboard fashion: with replicate wells containing 100 µl serial doubling dilutions of cognate antigen (approximate initial range: 10 µg/well to 0.01 µg/well) arranged across the plate from left to right. Use an underplate well indicator to guide the distribution of reagents. Incubate 60 min at 37◦ C. Include wells that lack one or both of the antigen and antibody-enzyme conjugate to determine the rate of spontaneous conversion of the chromogenic substrate to colored product. All measurements should be made in at least three replicate wells to allow the subsequent calculation of standard deviation, which should ideally be 20 years ago (Czerkinsky et al., 1983; Sedgwick and Holt, 1983) can provide information about the number of cells in a population that are secreting a specific protein (usually a cytokine or immunoglobulin). Cytokine secretion can be interpreted to indicate pathways of immune activation such as Th 1- or Th 2-dependent pathways (Lalvani et al., 2001), and immunoglobulin secretion



can indicate either antigen-specific responses to antigen challenge, or total amounts of specific isotype production indicative of overall immune capacity. There are some suppliers of complete ELISPOT kits (e.g., Sanquin; Mabtech; and Diaclone), but it is often far more economical to develop the specific ELISPOT assay for an antigen of specific interest in the laboratory. In the prototype assay described below, antigen coupled to the membrane surface serves as a target for the quantification of cells secreting an antibody specific for that antigen.

Materials Antigen ELISPOT coating buffer (see recipe) ELISPOT wash buffer (see recipe) RPMI-1640 supplemented with 10% FBS or ELISPOT blocking buffer (see recipe) Test cells Capture and detection antibodies for the target molecule of interest ELISPOT assay diluent ELISPOT culture medium ELISPOT substrate solution: BCIP (5-bromo,4-chloro,3-indolylphosphate)/NBT (Kirkegaard and Perry) 96-well filtration plates with sterile surfactant-free membrane (e.g., Millipore) Automated plate washer (optional) ELISPOT reader (ImmunoBiosys, Zeiss, Sanquin, or A. EL. VIS) or dissecting microscope and digital image analysis software (e.g., ImageJ available as freeware at http://rsb.info.nih.gov/ij/) Add antigen to plates 1. Incubate ELISPOT plate with 100 µl/well of filter-sterilized (0.2-µm filter) antigen at 100 µg/ml in ELISPOT coating buffer. Incubate specificity control wells with 100 µl/well of an irrelevant antigen at 100 µg/ml in ELISPOT coating buffer. Incubate overnight at 4◦ C. These 96-well filtration plates with sterile surfactant-free membranes have filters on the bottom, which have a pore size of 0.45µm and are comprised of hydrophobic PVDF.

2. Wash three times with sterile ELISPOT wash buffer in an automated plate washer, or wash plates manually. 3. Block plate with 200 µl/well sterile RPMI-1640 supplemented with 10% FBS for 1.5 to 2 hr at room temperature, or with ELISPOT blocking buffer.

Add cells 4. Add 100 µl of the test cell populations in sterile RPMI-1640 supplemented with 10% FBS to each well at serially diluted cell suspensions of 106 /well to 104 /well in triplicate and incubate in a 37◦ C, 5% CO2 humidified incubator for 24 hr. 5. Wash wells three times with ELISPOT wash buffer manually or in an automated plate washer.

Perform ELISPOT assay 6. Add 100 µl/well of ELISPOT detection antibody at 1 µg/ml in PBS (usually ∼1:500 dilution of antibody) and incubate 2 hr at 37◦ C. It is important that the specificities of the capture and detection antibodies are compatible. The pair must be capable of binding to distinct epitopes on the same target molecule simultaneously.

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Figure 18.7.4 Examples of colored precipitate spots generated in the ELISPOT assay. (A) Control well containing unstimulated cells. (B) Well containing stimulated cells.

7. Wash three times with ELISPOT wash buffer manually or in an automated plate washer. 8. Add 100 µl/well of ELISPOT substrate solution and develop for 10 to 30 min at room temperature until spots appear. 9. Wash plate three times with 200 µl/well distilled water. 10. Air-dry the plate and count spots by using a dissecting microscope or ELISPOT reader. Legitimate spots will be round and larger than a cell in size (usually 75 to 400 µm in diameter) when viewed with a dissecting microscope. An example of these spots can be seen in Figure 18.7.4. Artifacts will have eccentric shapes, and staining will not be uniform across the spot. It is often useful to look at a well that has not been seeded with cells before the staining protocol was done to identify known artifacts. While the trained eye is still best at identifying spots, it is advisable to have coded the wells so that enumeration can be done without bias. BASIC PROTOCOL 3

GRATING-COUPLED SURFACE PLASMON RESONANCE IMMUNOASSAY GCSPR is a new assay that offers the sensitivity of an ELISA in a microarray format (Fig. 18.7.5). This format enables the simultaneous evaluation of small volume samples to determine the presence of a large number of different antigenic determinants (Brockman and Fernández, 2001). Currently available chips have an active area that is ∼1 cm2 , and can accommodate ∼400 individual spots. With replicates, this allows for the assessment of >100 antibody/antigen interactions at the same time in a real-time assay that is label free. Currently, there is only one commercial supplier of GCSPR instrumentation (HTS Biosystems).

Materials Ethanol Capture antibody Antigen PBS (see recipe) GCSPR blocking reagent Analyte solution Solid-Phase Immunoassays



Figure 18.7.5

GCSPR optical diagram.

MicroCaster System, 8-pin manual arrayer (Schleicher and Schuell Bioscience), or another spotting system GCSPR sensor chips (Applied Biosystems) 96-well microtiter plates 37◦ C incubator and humid box Applied Biosystems 8500 affinity chip analyzer 1. Rinse pins of arrayer sequentially in ethanol and distilled water. 2. Rinse gold sensor chip surface sequentially in ethanol and distilled water. Allow to air dry. 3. Place chip in arrayer with the notch in the upper right hand corner. Determine which pins will spot inside the active area of the chip (the part of the chip that has the diffraction grating embossed on it). 4. Place captured antibodies in wells of a 96-well microtiter plate. Dip pins into antibody solutions. 5. Transfer samples to active area of chip on slide. Designate regions of interest (ROIs). 6. Move chip using MicroCaster indexing system and repeat for desired number of spots. Move chip in a horizontal direction. 7. Use a sharp needle to make a small scratch in the chip underneath the spots for orientation (make this mark at least 1 mm away from the ROIs).

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Figure 18.7.6 The GCSPR chip is illuminated following spotting of the chip surface with the capture antibody and reflected light intensity is measured at a range of angles (solid line). Following exposure of the chip to cognate antigen, the angle scan is repeated, and reflected light intensity is again measured (dashed line). The increase in the angle at which the maximum coupling occurs, which is the angle that has the least light reflected, is an indication of antigen binding to the capture antibody. The magnitude of this angle shift is a measure of the increase in molecular mass at the ROI.

8. Place spotted chips in 37◦ C incubator inside a humid box for 30 min to allow antibody to bind to the chip surface. Install chip in reader. 9. Initiate liquid flow at 200 µl/min and pass PBS over chip surface, take initial GCSPR data to assess the amount of protein that remains bound to the chip (compare to empty ROIs where no capture antibody has been placed) 10. Pass GCSPR blocking reagent over chip surface. 11. Pass analyte solution over surface and take GCSPR data to assess the change in the GCSPR angle (the illumination angle at which the greatest coupling and thus least reflection of light occurs). 12. Assess angle shift increases as an indicator of antigen capture (Fig. 18.7.6).

REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.

Alkaline phosphatase substrate buffer 9.7% (w/v) diethanolamine 0.02% (w/v) NaN3 0.01% (w/v) MgCl2 , pH 9.8 Store buffer for several months at 4◦ C. Add p-nitrophenyl phosphate (1 mg/ml PNPP; Sigma) 10 min before use to allow the PNPP to dissolve. Tablet or capsule forms of the substrate can simplify addition to the buffer.


ELISA blocking buffer (BSA-based) PBS (see recipe) 0.2% (w/v) NaN3 10% (w/v) BSA Store up to 1 week at 4◦ C.



ELISA coupling buffer 15 mM NaHCO3 35 mM Na2 CO3 0.2% (w/v) NaN3 , pH 9.6 Store up to 6 months at 4◦ C CAUTION: NaN3 is a toxic substance and should be handled with gloves and filter mask to avoid ingestion. Discarded solutions with azide should be substantially diluted with running water if poured down the drain to avoid the formation of lead azide in the pipes (a contact explosive).

ELISA wash buffer PBS 0.2% (w/v) NaN3 0.05% (v/v) Tween 20, pH 7.2 Store up to 6 months at 4◦ C ELISPOT blocking buffer 1% (w/v) Teleostean gelatin or 5% (w/v) nonfat dry milk in PBS (no azide). Store up to 1 week at 4◦ C. ELISPOT coating buffer 1.57% (w/v) Na2 CO3 2.93% (w/v) NaHCO3 0.2% (w/v) NaN3 , pH 9.7 Store up to 6 months at 4◦ C ELISPOT substrate solution Prepare stock solutions of nitroblue tetrazolium (100 mg NBT/2 ml of 70% dimethylformamide in water) and bromo-4-chloro-3-indolyl phosphate (100 mg BCIP in 100% DMF). Add 33 µl NBT stock plus 17 µl BCIP stock to buffer comprised of 100 mM Tris·Cl, pH 9.5, 100 mM NaCl, and 5 mM MgCl2 . Use working substrate within 1 hr. Store NBT and BCIP stock solutions at −18◦ C. ELISPOT wash buffer PBS (see recipe) 0.05% (w/v) NaN3 Store up to 6 months at 4◦ C Phosphate buffered saline (PBS) 8 g/liter NaCl 0.2 g/liter KCl 1.15 g/liter Na2 HPO4 0.2 g/liter KH2 PO4 Store up to 6 months at 4◦ C COMMENTARY Background Information ELISA has supplanted the radioimmunoassay (RIA) as the method of choice for measuring antibody/antigen interactions. Depending on the particular configuration

used, a wealth of information can be obtained with ELISA. For the most part, this assay can enable the characterization of cellular antigens, soluble antigens, and the antibodies that detect these targets. It is important to design

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Table 18.7.3 Troubleshooting Guide to ELISA/ELISPOT

Problem

Possible cause

Solution

No color development

Enzyme or substrate may be bad or left out of final step

Retain unused diluted antibody-enzyme conjugate and prepared substrate. Test for enzyme activity by mixing 10-µl aliquot of the diluted conjugate with 100 µl of substrate.

Antibody-enzyme conjugate denatured or aggregated

Do not freeze-thaw conjugate, store at 4◦ C

Excess color development; non-linear changes in optical density with changes in antigen or antibody concentration

Some plate readers produce linear measurements up to an O.D. of 2.0. Others are linear to an O.D. of 3.0

Dilute the antigen or the antibody to reduce the amount of enzyme that is bound in the well

False positives in ELISA/ELISPOT

Antibody-enzyme conjugates will aggregate under some conditions and become quite sticky

To reduce non-specific binding, centrifuge conjugate in microcentrifuge at top speed, or ultrafilter diluted conjugate just before use. Inappropriate cross-reactivity may exist in the antigen-specific antibody preparation, or contaminating antigens may be present.

the ELISA with potential cross-reactions in mind, and to include both positive and negative controls in the assay for quality control. It is equally important, especially in the context of toxicological assessments, to ensure that the toxic agent does not have an undue influence over the performance of the assay. For example, toxins may diminish viability of input cell populations in the ELISPOT assay, which could alter the interpretation of results. Metals can alter structural features of target molecules, making their detection less certain. Lead has been shown to interact with IFN-γ in this way (David Lawrence, pers. comm.). Thus, it may be useful to run preliminary assays in the presence of the toxicant itself to assess the impact of this reagent on the ability of the antigen to adsorb to the microtiter plate surface, on the ability of the antibody to bind antigen, and on the enzymatic conversion of substrate to product.

usually useful to test several dilutions of antigen or antibody to exclude artifactual results by showing that the reaction behaves as predicted (i.e., a ten-fold dilution will produce a ten-fold decrease in signal). For measurements of humoral immune responses, it is often of value to determine the titer of the solution—at what dilution the reactivity diminishes below a specific defined value. In assay configurations where the antigen must couple to a capture antibody and to an antibody-enzyme conjugate, care must be taken to match the antibody pairs; otherwise steric interference will impede the simultaneous binding of both antibodies to the antigen, thus making the assay unsuccessful. The ELISPOT assay is an outgrowth of the plaque assay, where cells in soft agar were assayed for the secretion of antibody that could lyse erythrocytes in an overlying layer of agarose. As cells secret relevant antibodies, they diffuse into the overlying layer containing the erythrocytes, and will bind to the erythrocyte membranes. After this incubation is complete, a solution containing complement proteins (often a dilution of guinea pig serum) is added to the top of the agarose layers and allowed to saturate the agarose. Where antibody has bound to erythrocyte membranes, the complement cascade will be activated, and cells will lyse. This can be seen as a clearing of the granular erythrocyte suspension (visualized as a clear plaque) and can further be identified by

Troubleshooting See Table 18.7.3 for a list of possible problems, possible causes, and solutions.

Anticipated Results


Typically, ELISA can distinguish proteins in complex solutions at concentrations in the micro- to nanogram range, but specifically selected antibodies can be sufficiently sensitive to detect antigen in the picogram range. It is



the lymophocyte that is located in the center of the plaque. The ELISPOT assay is a more universal assay because the antigens that can be assayed need not be naturally or artificially located on the erythrocyte membranes. An optimized ELISPOT can detect two to three secreting cells per 105 input cells. The GCSPR protein microarray platform is a further improvement on the basic assessment of antigen-antibody interactions. Sensitivity of the GCSPR assay approximates that of an ELISA, and may be further improved as the sensing technology is improved, and as the chemistry of antigen immobilization is enhanced. Moreover, current investigations are underway to explore the application of the GCSPR to ELISPOT-type investigations. Each of these assays can be used to assess the impact of toxins on the immune response by assessment of the degree of immunity that develops in the presence of different levels of toxin.

Time Considerations ELISA usually takes about a half day to complete, but this can be accelerated by increasing the antigen and antibody concentrations as the time per step is shortened. Alternatively, the assay can incorporate an overnight step to prepare the initial antigen-bound plate, or can employ plates that have been commercially prepared to reduce the time before experimental sample is applied. ELISPOT can take several hours to prepare, including the preparation of cells from harvested tissue or from cell culture. Once the cell culture incubation phase is complete (overnight incubation), the remainder of the assay can take an additional 1 to 2 hr to complete. Once stained, the spots can be enumerated under the microscope; this usually will take 1 or 2 hr to complete. GCSPR assays can take 1 or 2 hr to complete; on a per antigen basis, this assay can be the fastest of those discussed here since several hundred spots can be assessed on a single chip.

Literature Cited Amoura, Z., Chabre, H., Koutouzov, S., Lotton, C., Cabrespines, A., Bach, J.F., and Jacob, L. 1994. Nucleosome-restricted antibodies are detected before anti-dsDNA and/or antihistone antibodies in serum of MRL-Mp lpr/lpr and +/+ mice, and are present in kidney eluates of lupus mice with proteinuria. Arthritis Rheum. 37:1684-1688.

Brockman, J.M. and Fernández, S.M. 2001. Grating-coupled surface plasmon resonance for rapid, label-free, array-based sensing. American Laboratory 6:37-40. Czerkinsky, C., Nilsson, Langstrom., Nygren, H., Ouchterlony, O., and Tarkowski, A. 1983. A solid-phase enzyme-linked immunospot (ELISPOT) assay for enumeration of specific antibody-secreting cells. J. Immunol. Methods 65:109-121. Lalvani, A., Pathan, A.A., Durkan, H., Wilkinson, K.A., Whelan, A., Deeks, J.J., Reece, W.H., Latif, M., Pasvol, G., and Hill, A.V. 2001. Enhanced contact tracing and spatial tracking of Mycobacterium tuberculosis infection by enumeration of antigen-specific T cells. Lancet. 357:2017-2021. Probst, P., Kuntzlin, D., and Fleischer, B. 1995. Th2type infiltrating T cells in nickel-induced contact dermatitis. Cell. Immunol. 165:134-140. Sedgwick, J.D. and Holt, P.G. 1983. A solidphase immunoenzymatic technique for the enumeration of specific antibody-secreting cells. J. Immunol. Methods 57:301-309.

Internet Resources http://www.probes.com Molecular Probes is an excellent source for information and materials related to fluorescent immunoassays. http://www.southernbiotech.com http://www.rndsystems.com These companies provide a wide range of antibody/enzyme conjugates. http://www.abcam.com http://biocompare.com Online search engines for identifying sources of antibodies from a range of commercial vendors. Linscott’s directory of immunological and biological reagents is available as a CD, or in print and is also available online at http://www. linscottsdirectory.com/directoryonline.htm. http://www.piercenet.com http://www.sigmaaldrich.com Sources of antibodies, purified antigens, blocking reagents, and other useful reagents for these techniques. http://www.elispotresource.com/ A Website supported by many of the vendors of ELISPOT materials and instrumentation. http://www.htsbiosystems.com Website for GCSPR Flexchip Analyzer.

Contributed by Michael A. Lynes University of Connecticut Storrs, Connecticut

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Immune Cell Phenotyping Using Flow Cytometry

UNIT 18.8

Immunophenotyping is a general term used to describe the use of fluorescently labeled antibodies to identify and quantify distinct subpopulations of cells within a heterogeneous population of cells. Frequently, the term is used more specifically to refer to the characterization of subsets of cells associated with the immune system (i.e., immune cell phenotyping). In either case, the antibodies used for immunophenotyping are usually specific for cell surface proteins that are known to be differentially expressed on discrete subsets of cells. Immunophenotyping can be carried out using either flow cytometry or immunohistochemical analysis. However, flow cytometry is often the method of choice, because it permits the acquisition of nonsubjective data on thousands of cells within seconds. Furthermore, each cell suspension can be labeled with several different antibodies simultaneously, and the fluorescence associated with each antibody can be collected on a per-cell basis as correlated data. Modern flow cytometers can routinely measure five or more different fluorescence emissions simultaneously, providing the means to easily detect unique subsets within a heterogeneous population of cells. More recently, cell surface phenotyping with immunofluorescent probes has been performed to ascertain the functional status (e.g., DNA or cytokine content, mitochondrial membrane integrity, glutathione status) of cells, making multiparameter flow cytometry a very powerful tool for characterizing the activation state of a cell. The only drawback of flow cytometry is that cells must be stained and analyzed in a single-cell suspension, which is easier to prepare from some tissues than from others. Because cells derived from blood and lymphoid tissues are readily maintained in suspension, flow cytometry has been widely used to phenotype these cells. Immunophenotyping of lymphoid tissues, particularly spleen and thymus, has been used in the field of immunotoxicology for many years as part of a tiered screening approach for detecting immunotoxicity following chemical exposure. Since thymic involution frequently occurs after toxicant exposure, changes in thymocyte immunophenotypes are commonly evaluated. Similarly, just as white blood cell (WBC) counts and differentials have been used for many years to assess immune status in toxicant-exposed animals, newer screening approaches such as immunophenotyping of spleen and/or lymph nodes have been used to quantify frequencies of the major lymphocyte subtypes (B cells, T cells, and T cell subsets) following chemical exposure. In studies conducted by the National Toxicology Program, the utility of immune cell phenotyping has been validated using a large database of environmental toxins and certain pharmaceutical agents. In these studies, changes in lymphocyte phenotypes in the spleen or thymus following toxicant exposure was identified as one of the best single correlates with toxicant-induced changes in host resistance to pathogens or tumors (Luster et al., 1993). However, even though immunophenotyping of the spleen and thymus may correlate with toxic chemical exposure, it is not necessarily a robust predictor of immunotoxicity, particularly when naive animals are exposed to toxicants. Some highly immunotoxic chemicals, including cyclosporin A and 2,3,7,8-tetrachlorodibenzo-p-dioxin (TCDD), do not alter splenic immunophenotypes in the absence of antigen challenge (Kerkvliet and Brauner, 1990; Vandebriel et al., 1999). Likewise, changes in thymic phenotypes can occur indirectly, through stress-induced effects on the thymus that are independent of direct immunotoxicity. Thus, it is generally agreed that immunophenotypic analysis of B cells, T cells, and T cell subsets present in the thymus and/or spleen should be used in conjunction with conventional assessments of immune function when screening for the immunotoxic effects of drugs and chemicals (Immunotoxicology Technical Committee, 2001; Food and Drug Administration, 2002). Immunotoxicology Contributed by Julie A. Oughton and Nancy I. Kerkvliet Current Protocols in Toxicology (2005) 18.8.1-18.8.24 C 2005 by John Wiley & Sons, Inc. Copyright


Although immunotoxicology studies have focused primarily on the phenotypic analysis of B cells and T cell subsets in spleen and/or thymus from naive mice, several extended applications of immunophenotyping are emerging in the immunotoxicological literature. One important extension is the analysis of peripheral blood cells from mice and rats (Oughton et al., 1995; Nygaard and Lovik, 2002; Funatake et al., 2004). The use of peripheral blood samples from rodents allows more direct comparisons to be made with toxicity data that may be available from human studies, which nearly always rely on peripheral blood samples. Analysis of blood also allows evaluation of systemic changes that may result from toxicant effects in lymphoid tissues, as well as the testing of multiple samples obtained from the same animal over an extended time. Another important addition to standard immunophenotyping techniques is the analysis of activation markers. Most cells in the immune system respond to antigenic stimulation by altering their expression of a number of different proteins on the cell surface. Many of these proteins are known to be involved in the functional response of the cell (e.g., adhesion, migration, costimulation), while others may reflect the cell’s antigenic history (as in memory cells). Evaluation of activation markers is likely to provide an enhanced ability to detect immunotoxic effects. For example, when mice were exposed to low doses of TCDD for more than a year, no changes in major T- and B cell subsets were observed. However, analysis of the naive/memory phenotypes of CD4+ T cells revealed that TCDD exposure had significantly reduced the proportion of memory cells present (Oughton et al., 1995). Likewise, evaluation of activation markers in the context of an ongoing immune response may provide novel insight into the selective targets and mechanisms of action of immunotoxic chemicals. Several laboratories are now using immunophenotyping to track the effects of toxicants on the activation and fate of antigen-specific T cells following antigenic challenge, with very exciting results (Shepherd et al., 2000; Mitchell and Lawrence, 2003; Funatake et al., 2004). Since the specific antibodies that one might use for immunophenotyping will vary depending on the species tested, the flow cytometer used, and the experimental questions being asked, the following protocol describes a generic approach to the staining of cells with fluorescently labeled monoclonal antibodies (MAbs) to assess cell surface protein expression. Work in the authors’ laboratory has primarily involved mouse cells; however, all aspects of this unit easily apply to the analysis of cells from rats, dogs, and horses. The first protocol (Basic Protocol 1) describes the procedure by which cells obtained from lymphoid tissue are stained with fluorochrome-conjugated MAbs (direct staining method). A second protocol (Alternate Protocol) describes two indirect methods for the staining of cells with primary MAbs that are not conjugated to a fluorochrome. Finally, a third protocol (Basic Protocol 2) describes the staining of peripheral blood cells with MAbs. Support protocols are presented for the preparation of lymphoid cells (Support Protocol 1), the fixation of cells (Support Protocol 2), and discrimination of dead cells (Support Protocol 3). When freshly stained cells cannot be analyzed immediately, samples must be fixed so that the cells are stable for storage (Support Protocol 2). Furthermore, because the inclusion of dead cells in flow cytometric analysis can lead to the misinterpretation of data and possibly to the rendering of erroneous conclusions, it is desirable to assess cell viability using DNA dyes (Support Protocol 3). Finally, a procedure for the determination of optimal antibody concentrations is presented in Support Protocol 4. Immune Cell Phenotyping Using Flow Cytometry

NOTE: The investigator should consult the manufacturer’s instruction manual for specific information regarding the operation of his or her flow cytometer.



DIRECT STAINING OF CELLS PREPARED FROM LYMPHOID TISSUES This is the preferred method for staining cells obtained from lymphoid tissues (e.g., spleen, thymus, lymph nodes). Basic Protocol 1 is appropriate for immunophenotyping of cells from either rats or mice. Because this method uses MAbs that are directly conjugated to fluorochromes, staining is simple and quick and usually produces little background signal.

BASIC PROTOCOL 1

Toxicology studies usually involve the assessment of lymphoid tissues from several individual animals in a number of different treatment groups. For example, a 3-dose-pluscontrol experiment that examined spleen and thymus from 5 animals per treatment group would yield 40 individual cell samples that needed to be stained, usually in more than one staining configuration. Processing of large numbers of samples can be facilitated by staining cells in V-bottom 96-well microtiter plates, as opposed to 12 × 75–mm tubes. These plates can be centrifuged using a microtiter plate carrier, and the contents of the wells can be mixed easily using a vortex mixer (such as the Vortex-Genie 2, VWR Scientific Products) equipped with a platform head for a microtiter plate. Use of a multichannel pipettor (such as the 12-channel Pipet-Lite multichannel pipet from Rainin Instrument) facilitates rapid dispensing of media into the wells, as well as quick transfer of samples to Titertubes (Bio-Rad) for flow cytometric analysis. Titertubes are small tubes that can easily be inserted into 12 × 75–mm disposable tubes, which are commonly used to run samples on a flow cytometer. The abovementioned products greatly increase productivity and also simplify the processing of large numbers of samples. To help keep track of stained samples, a template of the microtiter plate can be prepared in which descriptions of the contents of each well (e.g., animal source, MAbs used) are recorded. In lieu of microtiter plates, cells can be stained in 12 × 75–mm plastic tubes or microcentrifuge tubes.

Materials Tissue of interest (e.g., lymphoid cells, Support Protocol 1) Fc receptor (FcR)-blocking immunoglobulin (Ig; e.g., normal rat IgG) solution, 200 µg/ml MAbs or isotype Igs directly conjugated to fluorochromes PBS supplemented with sodium azide and BSA (PAB; see recipe) Micropipettor or multichannel pipettor with tips 96-well V-bottom microtiter plates, disposable 12 × 75–mm polystyrene tubes, or microcentrifuge tubes Centrifuge, refrigerated and equipped with a microtiter plate carrier Vortex mixer equipped with a platform head for a microtiter plate Titertubes (Bio-Rad Laboratories; optional) Additional reagents and equipment for preparation of lymphocyte suspension to be analyzed (Support Protocol 1) NOTE: Several steps need to be performed to minimize antibody cross-linking and the subsequent internalization of cell surface receptors that may occur when these receptors are bound to MAbs. This process is often referred to as capping of the antigen-antibody complex. Some cell surface molecules are less likely to cap than others; however, surface immunoglobulin on B cells can cap almost instantaneously. The addition of sodium azide (final concentration, 0.1% w/v) to the staining medium will help to prevent capping. In addition, all cell preparations and staining procedures, including wash steps, should be performed at 4◦ C, a condition achieved by incubating the cells on ice. Staining samples on ice also reduces the off-rate constant associated with antibody binding by a factor of 10 compared with the off-rate constant at 25◦ C. Immunotoxicology


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Prepare lymphocyte samples for staining 1. Prepare a single-cell suspension of lymphocytes in 5% HBSS. Lyse red blood cells (RBCs) if present in large numbers (Support Protocol 1). 2. Using a micropipettor or microchannel pipettor, dispense 1–4 × 106 cells into each well of a V-bottom 96-well microtiter plate. Alternatively, dispense cells into 12 × 75–mm tubes or microcentrifuge tubes.

3. Pellet cells by centrifugation for 3 min at 200 to 1500 × g, 4◦ C. The range of centrifugation speeds quoted in this step reflects the fact that no one speed is accepted by the entire flow cytometry community. Although 1500 × g may sound excessive, this is the speed most commonly used by Stewart and Stewart (2001a) in order to minimize the loss of cells during washing.

4. Remove supernatant by holding the plate over a sink and inverting with a flick of the wrist or by decanting liquid from tubes. Blot the plate or the lip of each plastic tube on an absorbent towel to remove excess supernatant. 5. Vortex plate or tubes lightly to resuspend cell pellet.

Label with fluorochrome-conjugated MAbs 6. Add 50 µl FcR-blocking Ig (10 µg) to each cell suspension. Vortex cells lightly and incubate on ice for 10 min. A number of cells (especially myeloid cells) have receptors for the Fc portion of an immunoglobulin and will bind to any MAb. In order to assess the antigen-specific binding of a MAb, FcR must be blocked using normal IgG from the same animal species as the selected MAb. For example, for cells stained with rat anti–mouse CD4 MAb, normal rat IgG should be used to block FcR-mediated binding. As an alternative, a MAb specific for mouse FcR (CD16/CD32) can be used, although this involves a substantially higher cost. FcR should not be blocked when assessing FcR expression.

7. Without removing the FcR-blocking reagent, add the appropriate fluorochromelabeled MAbs (see Support Protocol 4 for determination of optimal antibody concentrations) or isotype Igs to each well or tube. Vortex plate or tubes and incubate cells on ice for 10 min, protected from light. Cells can be stained with a number of MAbs simultaneously as long as there are no interactions among these MAbs. For large studies, a cocktail of all the MAbs of interest can be used to stain each sample. For example, if cells are to be stained for CD3, CD4, and CD8, a single cocktail including all three MAbs can be prepared in a total volume of 10 µl. Cocktail staining greatly expedites sample preparation and ensures that all samples are stained uniformly. Over time, MAbs in solution tend to form aggregates. Cells that express Fc receptors can bind these antibody aggregates with far greater avidity than they can antibody monomers. Aggregates should be removed by microcentrifugation of the antibody solution for 5 min at 12,000 to 13,000 rpm, 4◦ C, prior to staining. Fluorochromes [especially the phycoerythrin (PE)–Cy5 and PE-Cy7 tandem conjugates] are particularly sensitive to photodegradation. Therefore, all staining procedures should be conducted in subdued light.

8. Add 150 µl PAB to each well or tube and centrifuge for 3 min at 200 to 1500 × g, 4◦ C. Remove supernatant as in step 4. Immune Cell Phenotyping Using Flow Cytometry



9. To remove unbound MAb, which can contribute to background fluorescence, wash cells once by adding PAB (total volume in microtiter plate not to exceed 200 µl) to each well or tube and centrifuging for 3 min at 200 to 1500 × g, 4◦ C. Remove supernatant as in step 4. 10. Resuspend cells in 500 µl PAB with gentle vortexing. For samples prepared in microtiter plates, resuspend each cell pellet in 200 µl PAB. Next, using a multichannel pipettor, transfer the contents of each well to a Titertube and add an additional 300 µl PAB, for a total volume of 500 µl. These Titertubes can then be inserted into 12 × 75–mm plastic tubes for flow cytometric analysis.

11. Keep samples on ice until analyzed using a flow cytometer. If stained cells cannot be analyzed on the day of preparation, they must be fixed for longer-term storage (Support Protocol 2).

INDIRECT STAINING OF CELLS PREPARED FROM LYMPHOID CELLS In general, it is best to stain lymphoid cells with a MAb that is directly conjugated to a fluorochrome (i.e., using the direct method). It is relatively easy to obtain MAbs specific for mouse or rat markers in a number of conjugated forms from commercial sources. However, there are very few mouse or rat MAbs that are conjugated to PE–Texas Red. This will impact studies requiring four or five colors for immunophenotypic analysis, since one MAb may require a second-step staining reagent. This concern is even more relevant when dealing with species other than mice, rats, or humans, as it is very difficult to find commercial sources of fluorochrome-conjugated MAbs for such species. In those situations, it may be necessary to resort to one of two indirect staining methods.

ALTERNATE PROTOCOL

In one method, cells are incubated first with an unconjugated primary MAb and then with a fluorochrome-conjugated antibody that is specific for the primary antibody. For instance, if cells are stained with an unconjugated rat IgG2a MAb that is specific for mouse CD4, the secondary antibody would be a fluorochrome-conjugated antibody that is specific for rat IgG. In most cases, the secondary antibody will be a polyclonal anti-Ig antibody, although Ig isotype–specific MAbs can sometimes be obtained from commercial sources. In this example, one can limit cross-reactivity of the secondary antibody with all other IgG isotypes by restricting the specificity of the secondary antibody to the rat IgG2a isotype, if available. To minimize FcR-mediated binding of the secondary anti-Ig antibody, it is preferable to use the F(ab )2 fragment rather than the whole immunoglobulin. An alternative indirect staining method relies on the use of a biotin-conjugated primary antibody, which is then detected with fluorochrome-conjugated streptavidin (SA). This biotin-SA method is the preferred indirect staining approach, because reagent interactions, which are often associated with the use of anti-Ig antibodies, are less of a concern. In addition, there are commercially available SA conjugates to a variety of fluorochromes, making SA useful in multicolor staining protocols. However, as with directly conjugated MAbs, biotin-conjugated MAbs are not always available. The use of secondary anti-Ig antibodies can be circumvented by labeling purified MAbs with specific fluorochromes in the laboratory. Detailed methods for MAb conjugation are available on the Internet (Roederer, 1997), and conjugation kits can be obtained from commercial vendors. The authors have had success using the Zenon Alexa Fluor 488 Mouse IgG Labeling Kit (Molecular Probes, 2004a) to label unconjugated MAbs. Using this antibody labeling kit, small quantities (≥0.4 µg) of primary MAbs can easily be conjugated to fluorescent labels. Currently, the Zenon system can only be used with mouse IgG1 , mouse IgG2a , mouse IgG2b , rabbit IgG, and human IgG antibodies.

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Additional Materials (see also Basic Protocol 1) Unconjugated primary MAb or biotin-labeled primary MAb Fluorochrome-conjugated anti-Ig F(ab )2 fragment (for use with unconjugated primary MAb) or fluorochrome-conjugated streptavidin (for use with biotin-labeled primary MAb) 1. Stain cells with primary MAbs according to steps 1 through 9 of Basic Protocol 1. If necessary, a combination of several fluorochrome-conjugated MAbs plus one nonfluorochrome-conjugated MAb (preferably a biotinylated MAb) can be used in a single MAb cocktail.

2. Resuspend cells in 50 µl PAB with gentle vortexing. 3. Add fluorochrome-labeled streptavidin or an appropriate secondary antibody. Vortex lightly and incubate on ice for 20 min. 4. Add 150 to 200 µl PAB and centrifuge cells for 3 min at 200 to 1500 × g, 4◦ C. Remove supernatant as in Basic Protocol 1, step 4. 5. Wash cells once in PAB as in Basic Protocol 1, step 9. 6. Resuspend cells in 500 µl PAB with gentle vortexing. Alternatively, for samples prepared in microtiter plates, resuspend each cell pellet in 200 µl PAB. Next, using a multichannel pipettor, transfer the contents of each well to a Titertube and add an additional 300 µl PAB, for a total volume of 500 µl. These Titertubes can then be inserted into 12 × 75–mm plastic tubes for flow cytometric analysis.

7. Keep samples on ice until analyzed using a flow cytometer. If stained cells cannot be analyzed on the day of preparation, they must be fixed for longer-term storage (Support Protocol 2). SUPPORT PROTOCOL 1

PREPARATION OF CELLS FROM LYMPHOID ORGANS The processing of mouse lymphoid organs into single-cell suspensions is a fairly simple procedure.

Materials Lymphoid organ of interest HBSS supplemented with 5% (v/v) FBS (5% HBSS; see recipe) Endotoxin-screened distilled water for cell culture (Gibco) 10× HBSS (Sigma) ACK lysis buffer (optional; see recipe) 60 × 15–mm untreated culture dish 25 × 75 × 1–mm frosted glass microscope slides 15-ml conical centrifuge tubes 12 × 75–mm disposable cell culture tubes Coulter counter or hemacytometer 1. Transfer a freshly dissected lymphoid organ to a 60 × 15–mm culture dish containing 4 ml of 5% HBSS.


2. When ready for processing, place the lymphoid organ between the frosted ends of two glass microscope slides. Disrupt the organ by gently pressing the frosted ends in a circular motion until only the empty capsule remains.



3. Transfer the resulting cell suspension to a 15-ml conical centrifuge tube. Rinse the culture dish and capsule with a combined total of 3 ml of 5% HBSS and then add this liquid to the cell suspension. 4. Centrifuge the cell suspension for 10 min at 200 × g, 4◦ C. Discard supernatant.

For spleen cell preparations 5a. Lyse RBCs by adding 4.5 ml cold distilled water (for cell culture) to the cell pellet and resuspending with gentle vortexing for 10 sec. After vortexing, immediately add 0.5 ml of 10× HBSS to restore isotonicity, followed by an additional 5 ml of 5% HBSS. Centrifuge cells as in step 4, discard supernatant, and resuspend the cell pellet in 10 ml of 5% HBSS with gentle vortexing. This procedure also lyses dead cells. If it is necessary to preserve dead cells (e.g., for apoptosis studies), RBCs can be removed by (1) resuspending the cell pellet in 5 ml of a standard ammonium chloride lysing solution (ACK lysing buffer; see recipe) and vortexing briefly; (2) incubating the resuspended pellet for 5 min at room temperature; (3) adding 5 ml of 5% HBSS (inverting the tube several times to mix); and then (4) centrifuging the cells for 10 min at 200 × g, 4◦ C, and discarding the supernatant. In general, the viability of spleen cell suspensions will be substantially higher when the hypotonic lysis method (>95%), as opposed to the ammonium chloride lysis method (85%), is used.

6a. Let cell debris settle for 10 min on ice, and then remove the cell-rich supernatant and transfer to a clean tube.

For thymus and lymph node cell preparations 5b. Resuspend the cell pellet in 10 ml of 5% HBSS with gentle vortexing. 6b. Let cell debris and clumps settle for 8 to 10 min on ice, and then remove the cell-rich supernatant and transfer to a clean tube. 7. Determine cell concentration using a Coulter counter or hemacytometer (APPENDIX 3B). Use 5% HBSS to adjust the final concentration of the cell suspension to 10–40 × 106 cells/ml.

CELL FIXATION If it is not possible to analyze freshly stained cells on the day of preparation, the cells must be fixed in electron microscopy (EM)–grade formaldehyde (such as Ultrapure formaldehyde, Polysciences) prior to storage at 4◦ C. Fixation not only cross-links proteins to make cells stable for storage but also inactivates the infectious activity of any viruses present. Hence, cell fixation is highly advisable when handling biological specimens capable of transmitting infection, and especially when handling human-derived cells.

SUPPORT PROTOCOL 2

Fixed samples should be analyzed within 5 days of fixation, since prolonged storage can result in increased autofluorescence. This is usually not a problem when antigens that are expressed at high levels are being analyzed. However, increased autofluorescence may make it more difficult to resolve negative cells from dimly stained cells. Cell fixation often leads to increased cell aggregation; therefore, it may be prudent to filter samples through a 40-µm nylon mesh prior to running them on the flow cytometer.

Additional Materials (see also Basic Protocol 1) 1% formaldehyde: 10% (v/v) Ultrapure EM-grade formaldehyde (Polysciences) diluted 1:10 in PAB (see recipe) 40-µm nylon mesh (Small Parts, Inc.; optional)

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1. Stain cells with MAbs according to steps 1 through 9 of Basic Protocol 1 or steps 1 through 5 of the Alternate Protocol. 2. Resuspend cell pellet in residual supernatant by gently vortexing. Cells must be thoroughly resuspended prior to fixation. If gentle vortexing does not completely resuspend the cell pellets in the microtiter plate, use a multichannel pipettor (equipped with plastic tips) to break up the pellets.

3. Add 200 µl 1% formaldehyde to each well and store cells on ice, protected from light, for 30 min. If necessary, cells can be stored overnight in 1% (v/v) formaldehyde at 4◦ C.

4. Prior to flow cytometric analysis, centrifuge cells 3 min at 1500 × g, 4◦ C. Remove supernatant as in Basic Protocol 1, step 4, and resuspend the cell pellet in 500 µl PAB. After resuspension in PAB, it may be desirable to filter samples through a nylon mesh to remove cell aggregates, as this will prevent clogging of the flow cell nozzle. BASIC PROTOCOL 2

STAINING OF PERIPHERAL BLOOD LEUKOCYTES This protocol details the immunophenotyping of peripheral blood leukocytes, a process often referred to as a “whole-blood” technique, as RBC lysis takes place after the sample has been stained with specific MAbs. It is appropriate for the immunophenotyping of blood cells from a number of animal species, including mouse, rat, horse, and dog.

Materials Animals of interest Sodium heparin, 250 IU per ml in PBS (APPENDIX 2A) PBS supplemented with sodium azide and BSA (PAB; see recipe) FcR-blocking Ig solution, 2 mg/ml (note that concentration differs from that used in Basic Protocol 1) Appropriate primary MAbs and (if necessary) secondary antibodies FACS Lysing Solution (BD Biosciences), diluted 1:10 in distilled water according to the manufacturer’s instructions 1-cc syringe equipped with 22-G needle Coulter counter Disposable 12 × 75–mm polystyrene tubes NOTE: Staining conditions are similar to those noted in Basic Protocol 1. However, all staining procedures should be performed in 12 × 75–mm polystyrene tubes, not microtiter plates. NOTE: Be sure to protect sample from light during incubation with fluorochromeconjugated MAbs.

Prepare peripheral blood leukocytes for staining 1. Using aseptic technique (APPENDIX 3B), collect blood (by cardiac puncture for mice) in a 1-cc syringe containing sodium heparin. A sufficient amount of heparin can be drawn into the syringe barrel by simply pulling the plunger back to the 0.05-cc mark. In order to reduce animal-to-animal variation, the same volume of blood should be collected from each animal (for example, ∼500 µl from each mouse). Immune Cell Phenotyping Using Flow Cytometry

2. Determine WBC concentration of blood sample using a Coulter counter. 3. Add 50 to 100 µl blood to a 12 × 75–mm polystyrene tube.



4. Wash cells twice by adding 2 to 3 ml PAB to tube, centrifuging for 5 min at 1500 × g, 4◦ C, and removing supernatant. This wash step minimizes artifactual staining of serum Igs with MAbs. Washing is performed by adding 2 ml PAB and centrifuging cells for 5 min at 1500 × g, 4◦ C. This centrifugation speed is required to produce a soft pellet of blood cells. The supernatant is removed by carefully decanting it and then touching the lip of the tube to an absorbent towel. Alternatively, the supernatant can be aspirated, taking care not to disturb the soft cell pellet.

5. Resuspend blood cells in residual medium (∼100 µl) by gently vortexing.

Label with fluorochrome-conjugated MAbs 6. To block FcR-mediated binding of Igs, add 5 µl FcR-blocking Ig (10 µg) to the cell suspension. Vortex cells lightly and incubate on ice for 10 min (Basic Protocol 1, step 6). 7. Without removing the FcR-blocking reagent, add the appropriate MAbs (see Support Protocol 4 for determination of optimal antibody concentrations) to the cell suspension, vortex gently, and incubate on ice for 10 min. All MAbs can be added concurrently in this step (Basic Protocol 1, step 7). It is highly advisable to include a MAb to CD45 to help define the leukocyte gate during data acquisition and analysis.

8. If all MAbs are fluorochrome-conjugated, proceed to step 11. 9. Add 2 ml PAB to sample and centrifuge for 5 min at 1500 × g, 4◦ C. Remove supernatant and wash cells once with PAB as in step 4. 10. Add secondary antibodies to sample, vortex gently, and incubate on ice for 20 min. 11. Without removing or washing away the MAbs, add 2.0 ml FACS Lysing Solution, 1:10 dilution. FACS Lysing Solution does not lyse nucleated erythrocytes, which are present in certain animal species.

12. Vortex cells thoroughly and incubate for 10 min at room temperature. 13. Centrifuge sample for 5 min at 1500 × g, 4◦ C. Remove supernatant and wash cells once with PAB as in step 4. 14. Resuspend cells in 500 µl PAB with gentle vortexing. 15. Keep samples on ice until analyzed using a flow cytometer. The FACS Lysing Solution contains formaldehyde, so no further cell fixation is required for longer-term storage.

ASSESSMENT OF CELL VIABILITY Cell viability should routinely be assessed in all flow cytometric analyses. Because the membrane integrity of dead cells has been compromised, most antibodies will freely pass through such membranes and stain dead cells nonspecifically. This nonspecific staining can make data difficult, if not impossible, to interpret and ultimately lead to erroneous conclusions. In general, cell viability is high (>95%) when lymphoid cells are taken directly from animal tissues, unless the mechanical process used to create single-cell suspensions has resulted in large numbers of dead cells. In contrast, the viability of cultured cells can be quite low, especially when these cells are chemically treated. In either case, to minimize

SUPPORT PROTOCOL 3

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cell death during sample processing, all cell preparations and staining procedures (including wash steps) should be performed at 4◦ C or on ice and in the presence of BSA or heat-inactivated FBS. The elimination of dead cells prior to staining is not recommended, since any manipulation of the cell sample could lead to the inadvertent loss of viable cells. Cells should be stained promptly after they are harvested and, if not fixed, analyzed in a flow cytometer as soon as possible. To assess cell viability, a sample of unstained cells is incubated with a DNA dye, such as propidium iodide (PI) or 7-aminoactinomycin D (7-AAD); dead cells will stain positively for either of these two nuclear dyes. Next, a data acquisition region is placed around the positively stained cells. By color-eventing or backgating on the PI+ or 7-AAD+ cells present, one can easily discern that most, but not all, dead cells exhibit lower forward scatter (FS) and higher side scatter (SS) than do viable cells. A gating region is then established around a cluster of viable cells (PI-negative) on the basis of their light scatter profile. This gating region can be used for all subsequent samples, even if these samples do not include a viability indicator. However, the best method for excluding dead cells from data analysis is to use a vital DNA dye in all samples. Some of the more common vital dyes used in multicolor analyses are PI, 7-AAD, TO-PRO-3 (Molecular Probes), and pyronin Y(G) [PY(G)].

Materials Nuclear staining compound dissolved in PBS (APPENDIX 2A): 200 µg/ml PI, 250 µg/ml 7-AAD, 250 µg /ml TO-PRO-3 (Molecular Probes), or 200 µg/ml PY(G) 12 × 75–mm polystyrene tubes Flow cytometer 1. Add 500 µl cells (1–2 × 106 ; unfixed and unstained) to a 12 × 75–mm polystyrene tube. 2. Add 5 µl PI to tube. Alternatively, add 4 µl 7-AAD, 4 µl TO-PRO-3, or 5 µl PY(G).

3. Incubate cells on ice for at least 5 min. 4. Analyze cells by flow cytometry. SUPPORT PROTOCOL 4

TITRATION TO DETERMINE OPTIMUM ANTIBODY CONCENTRATION The single most important factor in selecting an appropriate antibody is antibody quality. Good antibodies are characterized by their high specificities and high binding affinities, which allow specific staining to be distinguished from nonspecific staining. However, even with high-quality antibodies, it is important to determine the optimal staining concentration (or titer) for each MAb. This titer will ensure that cells are stained under saturating conditions. Under such conditions, specific fluorescence will not be readily influenced by cell number or incubation time (Kantor and Roederer, 1997). The method for titrating MAbs has previously been described by Stewart and Stewart (2001a).

Materials


Lymphoid cell suspension of interest (see Support Protocol 1) Fc receptor (FcR)-blocking immunoglobulin (Ig) solution, 200 µg/ml Fluorochrome-conjugated antibody solutions to be tested Centrifuge fitted with a microtiter plate adapter Vortex mixer fitted with a microtiter plate adapter Flow cytometer



1. Add 2 × 106 lymphoid cells to each of five wells in a 96-well microtiter plate. 2. Centrifuge cells for 3 min at 200 to 1500 × g, 4◦ C, in a centrifuge fitted with a microtiter plate adapter. 3. Block FcR-mediated binding by adding 50 µl FcR-blocking Ig to each well, vortexing gently using a vortex mixer fitted with a microtiter plate adapter, and then incubating samples for 10 min on ice. 4. Without removing the FcR-blocking Ig added in step 3, add 0.125, 0.25, 0.5, 1.0, and 2.0 µg of MAb, respectively, to the five wells containing lymphoid cells. Vortex cells gently and incubate for 10 min on ice. 5. Acquire immunofluorescence data using a flow cytometer 6. Create a two-parameter histogram of forward scatter versus side scatter and establish an appropriate gating region that encompasses the majority of viable cells. 7. Display fluorescence data from viable cells in single-parameter histograms. Establish one region that encompasses the negative population and a second region that brackets the positive population. Determine the median channel of fluorescence (MCF) for each region. 8. Compute the signal-to-noise (S/N) ratio, treating the positive population as the signal (S) and the negative population as the noise (N). Plot the calculated S/N ratios as a function of MAb concentration. 9. Identify the optimal MAb titer (i.e., the titer that produces the highest S/N ratio). Once the optimal titer is determined, it is not necessary to increase the amount of antibody used for staining unless the number of cells per sample exceeds 20 million. Even then, the amount of antibody needed for staining will only increase by a factor of two or three. However, it cannot be stressed enough that time and effort should be taken to optimize staining conditions (staining temperature, antibody concentration, staining time, and cell number) prior to any study. Doing so can also save money, as the manufacturer’s recommended staining concentration is often higher than necessary.

REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent for all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.

ACK lysing buffer 0.15 M NH4 Cl 10 mM KHCO3 0.1 mM disodium EDTA Adjust pH to between 7.2 and 7.4 using 1 M HCl Filter sterilize using a 0.2-µm filter Store at room temperature If the buffer is stored for an extended period, check its pH before use. HBSS, pH 7.2, supplemented with 5% (v/v) FBS 1× Hanks’ balanced salt solution (HBSS; Sigma) containing: 1 mM N-2-hydroxyethylpiperazine-N -2-ethanesulfonic acid (HEPES) 5% (v/v) fetal bovine serum (FBS), low-endotoxin, characterized (HyClone) 1 mM sodium pyruvate Adjust pH to 7.2 using 1 N NaOH Store up to 1 month at 4◦ C

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PBS, pH 7.2, supplemented with sodium azide and BSA Dulbecco’s PBS, calcium- and magnesium-free (e.g., Life Technologies), containing: 0.1% (w/v) sodium azide 1% (w/v) BSA Adjust pH to 7.2 using 1 N NaOH or 1 N HCl Filter solution through 0.22-µm filter paper Store up to 1 month at 4◦ C EDTA (1 mM) can be added to reduce cellular aggregation. CAUTION: Be extremely careful when handling sodium azide.



Secondary anti-Ig antibodies Although indirect staining with secondary fluorochrome-conjugated anti-Ig antibodies is the least desirable staining method, due to problems with nonspecific staining, there are certain precautions that can be exercised to minimize these problems. The secondary anti-Ig antibody is usually a polyclonal anti-Ig antibody that is specific for a large number of epitopes on the primary antibody. Since polyclonal secondary antibodies will be directed against all the Ig isotypes (e.g., IgG, IgA, IgM) found in the serum of the animal from which they were derived, these antibodies should be purified by affinity chromatography so that their binding is restricted to the heavy chain of the appropriate Ig isotype. The binding of the secondary anti-Ig antibody can be further restricted by selecting a reagent with no light-chain activity (i.e., one that is Fc fragment–specific). This will further ensure that the anti-Ig antibody is highly specific for a particular class of Ig (e.g., IgG) and thus minimize cross-reactions with other Ig isotypes. In addition, it is essential to use the F(ab) or F(ab )2 fragment of the affinity-purified secondary antibody to minimize FcR-mediated binding and, hence, reduce background staining. For example, a goat F(ab )2 anti–mouse IgG antibody (heavy and light chain–specific, purified by affinity chromatography) is specific for the heavy chain of mouse IgG but will also bind to other Igs that have the same light chain as IgG (e.g., IgA, IgM). Therefore, this reagent is not specific for mouse IgG at all. The authors have had great success using polyclonal secondary antibodies purchased from Jackson ImmunoResearch Laboratories. The binding of a secondary antibody can be further restricted to a specific Ig isotype and subclass. For example, if a rat IgG1 MAb specific for CD4 were being used as the primary

MAb, one could select a secondary MAb specific for rat IgG1 , if available. In all cases, it is necessary to determine the optimum antibody titer as well as assess the binding specificity of the secondary anti-Ig antibody prior to a study. First, the secondary antibody should not react with any cellular epitope. This can be assessed by simply incubating the cells of interest with the secondary antibody alone. The signal produced by this staining should be nearly identical to the signal yielded by unstained cells. If not, the reagent is inappropriate and should be replaced. Second, if this secondary antibody is used in multicolor analyses, it should not cross-react with any other MAb used for staining. To test for potential reagent interactions, incubate two samples of cells with the fluorochromeconjugated MAb, followed by washing. Next, add the secondary anti-Ig antibody to one sample and PAB to the other sample. The fluorescence pattern of the sample stained with the secondary anti-Ig reagent should be nearly identical to the pattern produced by the sample stained with PAB. If the fluorescence pattern is different, the secondary antibody in question should not be used in multicolor analyses. Fluorochrome choice Fluorochrome choice is highly contingent on the availability of the laser lines available on a particular flow cytometer. For instance, the Beckman Coulter XL flow cytometer is equipped with a single argon ion laser tuned to a wavelength of 488 nm, the most common wavelength used in flow cytometry. This instrument can handle up to four fluorescent emissions, all from dyes that can be excited by the 488-nm wavelength, but it cannot process the signal generated by a MAb conjugated to allophycocyanin (APC), which must be excited by a 633-nm laser line. However, APC signals can be processed on any benchtop analyzer, such as the FACScalibur



(BD Biosciences) or the FC500 (Beckman Coulter), that is equipped with a HeNe laser. For multicolor analyses, there are a number of factors to be considered when selecting a MAb conjugated to a particular fluorochrome. This selection is particularly important when using antigens that are expressed at low density, as some fluorochromes have higher absorption coefficients and/or higher quantum yields than others, making them more suitable for detecting weakly expressed antigens. In order to achieve the highest possible staining intensity, select antibodies conjugated to the brightest possible fluorochromes, e.g., in order, from brightest to dullest—PE, PE-Cy5, PE–Texas Red, APC, and FITC. In general, the fluorescence emitted by MAbs conjugated to PE, PE-Cy5, and APC is 5 to 10 times brighter than the fluorescence emitted by FITC conjugates. PE is often the fluorochrome of choice when examining the expression of an antigen that is expressed at low levels (e.g., CD25, CD69) or detecting the presence of intracellular cytokines. New fluorochromes are continually being developed, and they can be used in addition to or instead of the dyes mentioned above. Molecular Probes (2004b) has produced a new series of dyes, called the Alexa Fluor dyes, that exhibit more intense fluorescence than do other spectrally similar conjugates. For example, Alexa488, whose fluorescence spectrum is nearly identical to that of fluorescein isothiocyanate (FITC), is considered by many to be the best FITC-like reagent. A number of MAbs conjugated to Alexa488 can be obtained from commercial sources. If the experiment necessitates the use of a second-step reagent, the authors use fluorochrome-labeled SA, which is readily available from commercial sources, to stain biotin-labeled MAbs. For simple one- or two-color analyses, the second-step reagent of choice for the authors is PE-labeled SA, which will stain at the highest possible level. For multicolor studies, the authors routinely use a tandem conjugate PE–Texas Red–labeled SA. It is possible that PE-Alexa610-labeled SA, by virtue of its low background staining, may be a better option than the PE–Texas Red fluorochrome, which can exhibit the high background staining that is typical of Texas Red dyes. The authors rarely use FITC-labeled SA but have had great success with Alexa488-labeled SA, which is far superior to its FITC-labeled counterpart. All MAbs should be stored in the dark, as most fluorochromes are photosensitive. PE-Cy5 and PE-Cy7 tandem conjugates are

particularly sensitive to photodegradation. This phenomenon can easily be recognized by an increase in the PE signal and a concomitant decrease in the Cy5 or Cy7 signal. Use of cyanine tandem dyes can result in increased background staining, as these dyes can bind to monocytes and B cells. However, this binding is highly species-dependent; for instance, background binding can be high in mice with autoimmune disorders. Thus, again, it is prudent to assess the background staining of the reagents used prior to a large study. Multicolor staining combinations for routine immunophenotyping of murine cells Table 18.8.1 presents examples of possible staining strategies that can be used to phenotype various cell populations in the mouse. It is becoming increasingly clear that no single marker can be used to specifically identify a unique subset of cells. For instance, expression of CD4 and CD8a is not restricted to T cells, as these molecules are also expressed on subsets of dendritic cells (CD11c+ ). In addition, CD4 is expressed on a subset of natural killer (NK) cells. Thus, in order to ensure that only T cells are analyzed, a MAb specific for CD3e, a marker that is expressed on thymocytes and mature T cells, can be included in the MAb cocktail. The authors have also found that the correlated expression of two markers, CD11b and Gr-1, is needed to phenotype either macrophages or granulocytes in mouse spleen. Both of these cell types express CD11b and Gr-1; however, the level of Gr-1 is lower on macrophages than on granulocytes. The two populations are more clearly resolved by including additional markers that are expressed on the macrophage, such as F4/80, CD86, or CD54 (Choi et al., 2003). It is essential to recognize that many markers can be expressed at varying levels on different cell types.

Critical Parameters Autofluorescence Cellular autofluorescence is most often associated with myeloid cells, due to the presence of intracellular flavins in such cells. These flavins are easily excited by the 488-nm laser line, with peak emission occurring around 525 nm. Therefore, any signal generated by autofluorescent cells will be processed by the same photomultiplier tube (PMT) that processes FITC fluorescence. It is obvious that the presence of highly autofluorescent cells can contribute an unwanted addition to the FITC signal. Autofluorescence can

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Table 18.8.1 Examples of Possible Antibody Combinations for Staining Mouse Leukocytes Using Five Fluorochrome-Labeled Monoclonal Antibodies in a Staining Cocktail, with Analysis Performed on an FC500 Flow Cytometer (Beckman Coulter) with Dual Laser Excitation at 488 nm and 633 nma

Cell type(s)

Possible combination of stains

T cells, NK cells

FITC-anti-CD4 PE-anti-CD3e Biotin-anti-NK1.1b PE-Cy5–anti-CD8c PE-Cy7–anti-CD45d,e

Viable T cells

FITC-anti-CD4 PE-anti-CD3e Biotin-anti-NK1.1b 7-AADf PE-Cy7–anti-CD8

Granulocytes, dendritic cells, B cells, macrophages

FITC-anti-Gr-1 PE-anti-CD11c ECD-anti-B220g PE-Cy5–anti-CD11b PE-Cy7–anti-CD45

Activated T cells

FITC-anti-CD8 PE-anti-CD25 ECD-anti-CD4 PE-Cy5–anti-CD62Lh PE-Cy7–anti-CD8

a Abbreviations: 7-AAD, 7-aminoactinomycin D; APC, allophycocyanin; FITC, fluorescein isothiocyanate; MAb, monoclonal antibody; NK, natural killer; PE, phycoerythrin. b This biotinylated MAb necessitates the use of streptavidin labeled with PE–Texas Red or equivalent. c PE-Cy5-labeled MAb could be replaced by a MAb conjugated to APC or equivalent. d PE-Cy7-labeled MAb could be replaced by a MAb conjugated to APC-Cy7 or equivalent. e CD45 is a pan-leukocyte marker that is very useful in discriminating leukocytes from contaminating red blood cells, especially in the phenotyping of peripheral blood cells. A gating region can be established on the CD45+ leukocyte population as defined in a two-parameter histogram of side scatter versus CD45 expression. f 7-AAD is a DNA dye used as a viability indicator in unfixed cell samples. Its peak emission wavelength is 660 nm. g ECD is the trade name for the PE–Texas Red tandem conjugate produced by Beckman Coulter. Very few mouse marker–specific MAbs that are directly conjugated to this fluorochrome are available. h Because CD62L can easily be shed from the surface of cells, it is imperative to keep cells on ice throughout staining and data acquisition. The authors recommend that samples stained with anti-CD62L be kept on ice until loaded manually onto the flow cytometer for data collection. The use of a multisample carousel is not recommended when analyzing cells stained for CD62L, since cells cannot be kept cold while on such a device.


be particularly bothersome when measuring weak FITC fluorescence signals. If there are excessive numbers of myeloid cells in the experimental samples, the resulting high level of autofluorescence may make it difficult to interpret data. A sample of unstained cells can be used to determine the contribution of cellular autofluorescence to the background fluorescence signal as well as the minimum level of fluorescence exhibited by cells. There are several approaches that can be used to circumvent the problems associated with cellular autofluorescence. When studying myeloid cells, it may be prudent to use

MAbs that are conjugated to APC or APCCy7, as these dyes can be excited by a HeNe laser and emit at wavelengths greater than 600 nm, beyond the emission spectrum for cellular autofluorescence. When immunophenotyping lymphocytes in a sample containing elevated numbers of myeloid cells, one could include a MAb that specifically identifies the myeloid cells (such as one specific for Gr-1, a marker of myeloid/granulocyte lineage). The use of such an antibody allows one to gate out granulocytes and macrophages, thereby eliminating cellular autofluorescence from analysis.



Blocking FcR-mediated binding There are two ways that an antibody can specifically bind to cells. Antibodies can bind to a specific antigen (epitope-specific binding) through their antigen-binding sites, the F(ab ) fragment, or they can bind to myeloid cells that express receptors for the Fc portion of the Ig. Even though FcR-mediated binding is not antigen-specific, it is highly specific and needs to be eliminated in order to assess antigenspecific staining. FcR should be blocked with normal Ig prior to staining with epitope-specific MAbs. The authors typically incubate cells for 10 min at 4◦ C with purified normal Ig (Jackson ImmunoResearch Laboratories) taken from the same species that produced the antibodies used for staining. For instance, if a rat IgG2a anti-CD4 MAb is used, FcR-mediated binding should be blocked with an excess of purified rat IgG. The use of 10 µg of normal IgG in a staining volume of 50 µl (final concentration, 200 µg/ml) should be sufficient to block FcR. After 10 min of blocking, the MAbs are added directly to the samples without removing the FcR block, so that epitope-specific staining can proceed in the presence of FcR-block. Another approach to blocking FcR is to incubate cells with an unconjugated MAb that is specific for the FcR (CD16/CD32). It should be noted that this approach is more expensive than the former approach. In any case, do not block FcR if there is interest in assessing FcR expression. Isotype controls Isotype controls are also used to determine the minimum level of cellular fluorescence following staining and to assess the effectiveness of FcR blocking. An isotype control must match the primary MAb in terms of isotype and subclass, as well as fluorochrome label and concentration. For example, the appropriate isotype control for cells incubated with 0.5 µg FITC-labeled rat IgG2a MAb (specific for mouse CD4) is 0.5 µg FITC-labeled normal rat IgG2a . The appropriate isotype control for cells stained with biotin-labeled rat IgG1 MAb (specific for CD8) is treatment with biotin-labeled normal rat IgG1 , followed by incubation with fluorochrome-conjugated SA. For multicolor analyses, one isotype control sample can be set up to assess all the MAb isotypes in a single tube. It is not necessary to analyze each isotype control separately. For example, a sample of cells can be incubated, in the presence of an FcR block, with a cocktail of isotype-matched Igs for each specific MAb used in the staining procedure.

The fluorescence pattern of the isotype control should be nearly identical to that of the autofluorescence control. When this is not the case, it is indicative of an improperly titrated MAb, a poor-quality MAb, or both. This could have a profound effect on one’s ability to interpret positive fluorescence, especially when studying antigens that are expressed either transiently or at very low levels. In general, the authors have found that some isotype Igs (e.g., IgM, IgG2b ) exhibit more background staining than others, which can complicate the analysis of antigens expressed at very low levels. This is particularly relevant in analysis of intracellular antigens. In these cases, a cold block control is performed to verify antibody specificity. This can easily be accomplished by comparing the fluorescence of two cell samples, with one being pretreated with a cold blocking MAb (i.e., the same clone as the experimental MAb, but with no conjugate) for 15 min prior to the addition of the fluorochrome-conjugated MAb. The binding of the fluorochrome-conjugated MAb should be completely inhibited by the cold block. For example, to assess the expression of CD25, which can be present at low levels on activated T cells, cells are pretreated with unlabeled purified anti-CD25 MAb (at a 3-fold higher concentration compared with the conjugated MAb) for 15 min prior to the addition of the fluorochrome-conjugated antiCD25 MAb. This sample should represent a true negative control for CD25 staining. Background staining for cell surface expression of CD69, another T cell marker that is transiently up-regulated upon activation, can be assessed similarly. Due to the complexities involved in multicolor analyses, it is prudent to invest a little time to test these conditions prior to a large study. Background staining exhibited by cells stained with multiple reagents is very different from that seen in unstained cells or isotype controls, due to the broadening artifacts produced by compensation (see below) and/or reagentreagent interactions. Therefore, the best negative control for any given marker in a multicolor stain is a sample of cells stained with all but one reagent (commonly referred to as “fluorescence minus one,” or FMO). For example, when assessing CD25 expression on antigen-specific D011.10 T cells, the best negative control for CD25 expression is a cocktail containing MAbs to CD3, CD4, and KJ-126 (to identify the antigen-specific T cells in the D011.10 model) plus an isotype control for the CD25 MAb. In this way, the level of negative

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staining for CD25 expression can be assessed by gating on the antigen-specific T cells. Dead cells Cell viability must be assessed in each experimental study. Since most MAbs freely pass through the cell membranes of dead and damaged cells, dead cells will stain positively even if there is no specificity for a particular MAb. This nonspecific fluorescence can interfere with data analysis, and if ignored, it can result in erroneous interpretation of the data. For exclusion of dead cells, vital DNA dyes such as PI, 7-AAD, and TO-PRO-3 can be added to unfixed cell samples prior to analysis. These dyes cannot be used to assess the viability of cells that are permeabilized (as for intracellular staining) or fixed, since they will leak out of fixed cells. On the other hand, the vital dye ethidium monoazide (EMA; final concentration, 1 to 5 µg/ml) can be used to label dead cells prior to fixation (Reidy et al., 1991). Once inside a dead cell, EMA can be photochemically cross-linked to the DNA by exposure to visible light. This cross-linking allows EMA to be retained by fixed cells.


Compensation One of the most critical issues in performing multicolor phenotyping is dealing with overlaps in the spectral emissions of different fluorochromes. Once excited by the laser, a fluorochrome will emit a broad band of light. Even though a band-pass filter is often placed in front of a PMT to restrict the wavelengths of light that are transmitted to the tube, it is nearly impossible to exclude all of the light emitted by the other fluorochromes used in multicolor staining. Because of these spectral overlaps, each fluorochrome will contribute an unwanted light signal to several PMTs that are not assigned to detect that fluorochrome. For example, there is significant overlap in the spectral emissions of FITC and PE, so much so that some of the FITC emission spills over into the PE PMT and it must be eliminated from this PMT before analyzing PE fluorescence. By the same accord, PE contributes an unwanted signal to the FITC PMT, albeit at much lower levels, and this signal must be eliminated from the FITC PMT. If not eliminated, these spillovers will make a “false” contribution to the data. The process by which these false contributions are eliminated electronically is called compensation. Compensation is one of the least understood processes in multicolor analyses. It is important to consider compensation not only from the standpoint of applying it correctly but also

from the perspective of designing and analyzing multicolor experiments. Investigators need to be able to recognize data that has resulted from improper compensation, as failure to do so can easily result in data misinterpretation. Any spillover signal not completely compensated (i.e., undercompensated) out of a PMT can result in an overestimation of the positive population. Proper compensation is always essential when measuring antigen density by MCF. Compensation is often considered a process by which unwanted fluorescent signals are “subtracted” from the true fluorescent signal. However, compensation is not a subtraction process, but rather a straightforward application of linear algebra by which a proportion of the unwanted signal is eliminated from the true fluorescent signal. For some fluorochrome combinations, such as FITC and PE-Cy5 or APC, there is little, if any, overlap in spectral emissions; therefore, compensation is negligible. However, there is considerable overlap in the emission spectra of the PE, PE–Texas Red, and PE-Cy5 dyes, enough to make compensation quite challenging even for investigators who routinely perform multicolor analyses. In order to set up compensation properly, a sample of cells must be stained with a representative reagent for each fluorochrome used in the experiment. The number of compensation controls will be equivalent to the number of fluorochromes used in staining. For example, five-color staining requires five compensation controls, one for each fluorochrome. It is not necessary to use the same MAbs in the experiment as in the compensation controls. This is especially true when a particular experimental reagent is expected to stain just a small subset of cells. For example, the frequency of antigen-specific T cells (KJ1-26+ CD4+ ) adoptively transferred in the D011.10 model can be as low as 0.3% one day after injection with ovalbumin. Thus, it would be very difficult, if not impossible, to properly set compensation levels using the conjugated MAb that stains KJ1-26+ cells. What is important is using the brightest possible reagents for compensation controls. Therefore, the authors routinely set up compensation controls using CD45, CD8a, CD19, or CD45R MAbs conjugated to the appropriate fluorochromes. These markers are expressed at high levels on most immune cells, making it much easier to properly set compensation levels. In order to set up compensation properly, the compensation control sample must contain at least two populations— specifically, one bright population and one



negative or not-so-bright population. When examining spleen cells, there will be two distinct populations that stain for MAbs to CD8a and CD45R; one is positive, and the other is negative. Because CD45 stains all leukocytes, it is necessary to add an aliquot of unstained cells to the CD45 compensation control after the staining process but just prior to flow analysis to artificially create a negative population. There are two ways to perform compensation. Hardware compensation is performed during data acquisition, whereas software compensation is performed during data analysis. Because of the complexity of five-color analyses, the authors routinely collect flow cytometric data without performing hardware compensation in such analyses. Instead, prior to analyzing the experimental data using offline software, the data collected from compensation controls are used to perform software compensation using WinList software (Verity). The authors have found no disparity in the results obtained using software compensation as compared with hardware compensation. Compensation controls must be evaluated in every possible pairwise combination using two-parameter histograms. For example, in studies using FITC, PE, and PE-Cy5 fluorochromes in a single sample, the FITC compensation control must be evaluated using the following two-parameter histograms: FITC versus PE and FITC versus PE-Cy5. Twocolor samples require the assessment of just one pairwise combination, three-color samples require three combinations, four-color samples require six combinations; and five-color samples require ten combinations. Thus, it should be readily apparent that the complexity of compensation increases substantially with the addition of each fluorochrome. To properly set compensation levels, the center of the positive cell population (i.e., as determined by the median fluorescence intensity) is lined up with the center of the negative cell population. This is accomplished by adjusting compensation until the median channel for the positive population is equal to the median channel for the negative population. It is important to ensure that the fluorescence distribution for the negative population is not piled up on the baseline axis, as this will make it nearly impossible to determine the median channel and, thus, very difficult to set up compensation properly. An in-depth discussion on compensation can be found online (Roederer, 2000). Not all flow cytometers can handle all the possible pairwise combinations that may be necessary for hardware compensation. For

example, the Beckman Coulter FC500 and XL flow cytometers are capable of handling the pairwise compensation involving FITC and PE-Cy5, but the BD FACSCalibur is not. It can be argued that, in general, there is minimal, if any, FITC emission that spills over into the PECy5 PMT. However, when using carboxyfluorescein diacetate succinimidyl ester (CFSE), a low but measurable signal from this dye can spill over into the PE-Cy5 PMT. Thus, when using the BD FACSCalibur, in lieu of hardware compensation, one must rely on software compensation to eliminate unwanted CFSE signal from the PE-Cy5 detector. It is possible to minimize the need for compensation by carefully selecting reagents that are conjugated to fluorochromes whose emissions have little or no spectral overlap. For two-color analyses, the authors often select MAbs that are conjugated to FITC and either APC or PE-Cy5, since there is little, if any, overlap between the emission spectrum of FITC and the emission spectra of the latter two dyes. Beyond two-color analyses, fluorochrome choice gets more complicated with each additional fluorochrome. In general, PE– Texas Red–conjugated MAbs are used by the authors only when performing five-color immunophenotyping, as this fluorochrome has considerable spectral overlap with both PE and PE-Cy5 emissions, making compensation quite challenging. Of lesser importance, there are very few MAbs that are directly conjugated to PE–Texas Red, and two-step staining procedures are therefore often required. If a HeNe laser is available, the authors often select APC- or APC-Cy7-labeled reagents over PE-Cy5 conjugates. Control samples In order to ensure that data are collected and interpreted correctly, several control samples must be faithfully evaluated prior to running the experimental samples. These include controls to assess cellular autofluorescence, nonspecific binding, FcR-mediated binding, and compensation. Sometimes, the number of essential control samples exceeds the number of experimental samples. However, it is only through the proper use of all appropriate control samples that data can be collected and interpreted correctly. Controls can be performed for each animal, but this can easily lead to excessive numbers of control samples. For large studies, a pool of cells is created for each treatment group, and this pool is used to test for autofluorescence and isotype staining. In this way, a pool will

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contain representative cells from each animal in a given treatment group. For example, in an experimental study in which the effects of three different dose levels of a chemical are examined, there will be four pools of cells, each representing a particular treatment group (one for the vehicle control group and one for each of the three dose levels). Each treatment pool will have its own autofluorescence control and its own isotype control, resulting in a total of four autofluorescence controls and four isotype controls. To assess autofluorescence, a sample of unstained cells (with no MAbs added) is processed in a similar fashion as for all experimental samples. This sample should always be analyzed first in an experiment. To begin, the voltages for the photodetectors collecting the light-scatter signals (both forward scatter and side scatter) are adjusted to define the cells of interest. Second, the voltage of each PMT is adjusted to place the negative cell population in the first log decade of the histogram. It is important to apply enough voltage to ensure that the majority of cells are off of the baseline; this assures that any positive staining will fall beyond the first log decade. In essence, this control sample provides the baseline representing the minimum (or negative) fluorescence exhibited by cells. The next control to be analyzed is the isotype control. The fluorescence pattern of the isotype control should be similar to that of the unstained control if FcR-mediated binding has been blocked effectively. If not, one would need to evaluate the isotype controls used in the experiment. Like the autofluorescence control, the isotype control also determines the minimum level of fluorescence that is exhibited by cells. For multicolor experiments, compensation controls must be analyzed in order to account for any spectral overlap not removed by filters. The number of compensation controls will be equivalent to the number of fluorochromes used in staining. Compensation is greatly simplified in four- or five-color analyses when the PMTs are balanced, with their voltages set at nearly identical levels. However, it is crucial to remember that once compensation is set, any change in the PMT voltages will necessitate repetition of the entire compensation process.


Standardization of flow cytometric data There are three sources of error that can be introduced during immunophenotypic analysis: specifically, instrumental, technical, and biological. However, the amount of error

introduced by the instrument can easily be minimized. The instrument’s overall performance should be routinely evaluated prior to data acquisition using standardized beads, such as Beckman Coulter Flow-Check beads. These beads will assess the instrument’s fluidics and optics to ensure proper alignment of the laser with the sample stream. Once the fluidics and optics have been checked, the authors routinely run a sample of FlowSet beads (Beckman Coulter) to further standardize the instrument’s settings. These beads mimic the immunofluorescence exhibited by human and mouse cells and can be used at the voltages required for immunophenotypic analyses. The voltages and gain settings for each photodetector are adjusted to position the fluorescence emitted by the beads within a prescribed region, a process commonly referred to as channel positioning. In this way, any apparent change in marker expression over time can be expected to be biologically relevant, and not a result of changes in the instrument. Once it has been established that the instrument is performing under the expected experimental conditions, negative controls (autofluorescence and isotype) can be analyzed to ensure that the negative population falls within the first log decade for each channel being measured. Next, compensation controls should be performed to remove any unwanted fluorescence from the various PMTs due to overlaps in the spectral emissions of the fluorochromes used. It should be noted that improper compensation can result in false-positive or falsenegative results, so great care should be exercised when performing these controls. Once the instrument’s settings have been established, the authors routinely run a control sample, referred to as the verification control, in order to determine whether the system will provide accurate information. This control sample is stained with a set of MAbs that are conjugated to the same fluorochromes used in the experimental study but are specific for well-recognized markers. For example, in a five-color staining experiment (using MAbs conjugated to FITC, PE, PE–Texas Red, APC, and APC-Cy7), the verification tube would contain MAbs specific for CD4, CD8, CD3, CD45R, and Gr-1, with each of these MAbs being conjugated to one of the fluorochromes used in the study. In general, spleen cells from a naive mouse are used in these controls. Not only are the authors familiar with the staining profiles of these markers, but they have established a historical record that they can use for comparison with the observed relative



frequencies. In short, this verification control is expected to yield easily recognizable data that enable the authors to make any necessary final changes to compensation values. Once the verification control has produced the expected results, one can be confident that the flow cytometer will provide accurate results for the experimental samples. Data analysis The authors routinely acquire listmode data on 10,000 cells of interest when performing multiparametric analysis. For example, when immunophenotyping spleen cells for lineagespecific markers (such as markers of B cell, T cell, macrophage, or granulocyte lineage), they typically collect flow cytometric data on 10,000 viable spleen cells. If the authors are interested in analyzing cell surface markers gated on a discrete subset, such as antigenspecific CD4+ cells in the D011.1 model (which can represent as little as 0.5% to 4% of the total population), they collect data on 10,000 antigen-specific CD4+ cells. Following data acquisition, flow cytometric data can be displayed in either single-parameter or twoparameter histograms to monitor the cells of interest. In this way, gating and analysis strategies can be fine-tuned. Figure 18.8.1 illustrates one approach to the analysis of five-color flow cytometric data. The data represented in this figure are derived from studies monitoring the events of T cell activation in the graft-versus-host (GvH) response. T cells (both CD4+ and CD8+ ) were purified from the spleens of C57Bl/6 mice (H-2b ; referred to as donors) and injected into C57Bl/6 (H-2b ) × DBA/2 F1 mice (H-2d ; referred to as hosts). The donor T cells recognize the H-2d antigens contributed by the DBA/2 strain in the F1 host and become activated. Two days after the injection of these donor T cells, the F1 mice were sacrificed, and their spleens were processed into single-cell suspensions. Spleen cells were then stained simultaneously with a cocktail of five MAbs specific for CD4, H-2Dd , CD28, CD25, and CD62L. As shown in Figure 18.8.1A, a cluster of spleen cells was first identified by its light scatter profile (FS vs. SS), as is customary in flow cytometry. A rectilinear region was then established to exclude dead cells, debris, and cell aggregates from further analysis. This process is referred to as gating. The FS-versus-SS profile of dead cells, as determined by PI uptake, indicated that these cells were located below the gating region (data not shown). In general, the authors find that the viability of their

splenic suspensions often exceeds 97%. When immunophenotyping blood cells, the authors routinely include a MAb to the pan-leukocyte marker CD45, which allows them to set a gate around all the leukocytes present, thereby eliminating, or gating out, residual RBCs and platelets; this is most often performed by setting a gate around the appropriate cell clusters as defined by their CD45-versus-SS profiles. Two days after adoptive transfer into F1 mice, one can easily identify a small but distinct cluster of cells that represent the donor CD4+ cells (Fig. 18.8.1B). These cells are identified by their expression of CD4 and their lack of expression for H2Dd . As shown in this two-parameter histogram, the majority of cells, which are the host cells, express H2Dd . Only a small population of cells lacks H2Dd expression, representing the donor T cells. An elliptical region has been set to establish a gate for these CD4+ donor cells so that their expression of the T cell activation markers CD28, CD62L, and CD25 can be monitored throughout the course of the GvH response. Gating on the donor CD4+ cells (10,000) as shown in Figure 18.8.1B, one can generate single-parameter histograms that illustrate the expression of the T cell activation markers (Fig. 18.8.1C). The immunofluorescent profile for each marker is illustrated in the filled histograms, while the open histograms illustrate negative staining as determined by their appropriate isotype controls. Two populations can easily be distinguished on the basis of the differential expression of CD62L, CD62Lpos , and CD62Lneg . However, the staining profiles for CD25 or CD28, as seen in single-parameter histograms (Fig. 18.8.1C), are more homogeneous. Figure 18.8.1D illustrates the power of multiparametric analysis. Multiple subpopulations can be resolved by displaying the correlated expression of two markers in two-parameter histograms. While there is a hint of two populations in each of the single-parameter histograms for CD25 and CD28 expression, one could easily identify two cell clusters by correlating the expression of these two markers in a two-parameter histogram. It is interesting to note that three distinct cell clusters can be distinguished on the basis of the correlated expression of CD62L and CD25 expression. One strategy is to “color-event” the events of interest. As shown in Figure 18.8.1D, two regions have been set around the cells that have down-regulated their expression of CD62L, one population expresses CD25 (color-evented red), and another does not express CD25

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Figure 18.8.1

Legend at right.




(color-evented blue). The cells that have retained their CD62L expression have been color-evented gray. In this way, the expression of all their markers can be easily visualized for each population in all the histograms. For instance, the red population also expresses high levels of CD28 and elevated levels of CD4. In contrast, the blue population expresses intermediate levels of CD28 and CD4. By definition, both the red and blue cells are considered activated T cells since they have downregulated their expression of CD62L; however, the red cells are further along in the activation pathway since they have also up-regulated their expression of CD25, which occurs only after several cell divisions. The gray population represents CD4+ cells that have not become activated by day two of the GvH response. There is a new approach in visualizing flow cytometric data, i.e., changing the traditional four-decade logarithmic scale for immunofluorescent data displayed in two-parameter histograms to a new Logicle axis (Tung et al., 2004). The plot axes have been redefined to allow the log-transformed scale to have a zero as well as a negative region. There are several advantages to this approach. Once data have been properly compensated, a large number of negative cells pile up in the first channel, against the axis, as these cells exhibit little to no fluorescence. Using the new Logicle scale, these negative cells can be visualized as distinct cell

clusters. In addition, this new approach also makes it easier to assess compensation. Both FlowJo (Tree Star) and WinList (Verity Software House) offer the option to display flow cytometric data using the new Logicle axis. Statistics The data shown in Figure 18.8.1 represent the response of one individual animal. In most immunotoxicological studies, the authors use four to six animals per treatment group. In order to determine a treatment effect, data derived from the treatment group must be compared with data from the vehicle control group. The authors will often include a group of naive animals (N = 2 to 3) in order to monitor the antigen-induced response of the control animals. In general, the authors prefer to present flow cytometric data in the form of histograms and use statistics to state the level of confidence in the accuracy of each result. Flow cytometric data are usually presented as the frequency of any given population that expresses a particular marker and/or as a measure of the intensity of the fluorescent or light scatter signals. There are three measures commonly used to describe the central tendency when assessing fluorescence intensity: mean, median, and mode. The mean fluorescence can be significantly skewed higher or lower by the presence of a few outliers. Therefore, when possible, one should use the median fluorescence of the positives, since

Figure 18.8.1 (at left) Five-color analysis of T cell activation in a graft versus host (GvH) model. T cells purified from the spleens of C57Bl/6 mice (H-2b ; referred to as donors) were injected into C57Bl/6 (H-2b ) x DBA/2 (H-2d ) F1 mice. These donor T cells recognize the H-2d antigens contributed by the DBA/2 strain in the F1 host and become activated. Two days after the injection of donor T cells, the F1 mice were killed, and their spleens were processed into single-cell suspensions. The spleen cells were stained simultaneously with MAbs to CD4, H-2Dd , CD28, CD25, and CD62L. Cells were analyzed using an FC500 flow cytometer (Beckman Coulter). (A) Viable cells were identified on the basis of their forward scatter (FS)–versus–side scatter (SS) profiles. (B) CD4+ donor cells (elliptical region) were identified by their expression of CD4 and their lack of expression of H2Dd . In this sample, the donor CD4+ cells represent 1.9% of the total spleen cell population. (C) Expression of the T cell activation molecules CD28, CD25, and CD62L was assessed in single-parameter histograms by gating on 10,000 donor CD4+ cells. Cells with positive staining for the specified MAbs are represented by the gray-filled histograms. Open histograms represent negative staining associated with the isotype control antibodies. (D) Correlated expression of activation molecules on donor CD4+ cells as illustrated in two-parameter histograms. Donor CD4+ cells that have up-regulated expression of CD25 are color-evented as red events. Using color-eventing, a distinct cluster of donor CD4+ CD25+ cells that also express low levels of CD62L and high levels of CD28 is readily identifiable. When the data are displayed in a two-parameter histogram, the presence of at least three CD4+ subpopulations distinguished by their differential expression of CD25 and CD62L is evident (far left histogram). It is interesting to note that not all cells with low CD62L expression exhibit high CD25 expression. Also of interest is that CD4+ CD25+ cells have up-regulated expression of CD4, as shown in (B). This black and white facsimile of the figure is intended only as a placeholder; for full-color version of figure go to http://www.interscience.wiley.com/c p/colorfigures.htm

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the median is generally a more robust estimate of central tendency than the mean. The precision in cell counting, or the extent to which replicate samples agree with each other, is highly dependent on the number of events sampled. The statistic which is most often used to measure precision is the coefficient of variation (CV), which can be determined using the following simple equation: √ CV = [( N)/N] × 100% where N is the number of events sampled and the square root of N represents the expected standard deviation. As the difference between two populations becomes smaller, better and better precision (i.e., a lower CV) is needed. For example, the CV for 25 cells is 20%; for 100 cells, 10%; and for 10,000 cells, 1%. In the authors’ laboratory, in order to attain high precision, flow cytometric data are routinely collected on a minimum of 10,000 cells of interest, whether one is acquiring 10,000 viable cells or 10,000 antigen-specific T cells (such as the CD4+ donor cells shown in Fig. 18.8.1B).

Anticipated Results


The objective in flow cytometric analyses is to resolve epitope-positive cells from epitopenegative cells with a high degree of precision and accuracy. Sometimes, it is necessary to select a MAb that is labeled with a fluorochrome such as PE in order to maximize the resolution of positive cells from negative cells, especially when assessing dim immunofluorescence. However, there are several procedures that can be employed to ensure the collection of accurate results. Proper setup will help to eliminate instrumental error as a source of variation within an experiment. The systematic process by which an instrument’s performance is validated is known as quality control, and this process should be conducted for every experiment. There are several sequential quality control steps that should be followed (as discussed in Critical Parameters): (1) the use of standardized beads, such as Flow-Check fluorospheres (Beckman Coulter), to check laser alignment as well as the fluidics and optics of the system; (2) the use of standardized beads, such as Flow-Set fluorospheres (Beckman Coulter), for channel positioning (to ensure that changes observed in time-course studies are the result of biological effects and not of changes in the instrument); (3) the use of compensation controls to remove unwanted fluorescence from fluorochromes with overlapping spectral emis-

sions; and (4) the use of a verification control to establish the accuracy of the results yielded by the system. Once the instrument has been validated, there are several technical issues that must be addressed, such as cellular autofluorescence, FcR-mediated binding, and the presence of dead cells, which can result in the collection of inaccurate data and possibly lead to data misinterpretation or erroneous conclusions. It is easy to obtain high precision in flow cytometric data; this can be done by simply collecting data on a minimum of 10,000 cells of interest. As an example, when assessing activation markers on CD4+ T cells, data should be collected by gating on 10,000 of these cells. The precision of a given measurement is usually characterized by the statistic CV, and for 10,000 events, the expected CV would be 1%. Thus, the assessment of 10,000 cells of interest will help to ensure that small differences between two populations may still be biologically significant. It is only through these steps that one can be confident that the results obtained are accurate, and this can ensure the validity of data interpretation. This is of utmost importance when assessing treatment effects in toxicological studies. Treatment effects can be discerned by changes in the relative frequency of a particular lineage-specific population (e.g., CD4+ T cells, B cells, macrophages) or by changes in the relative frequency of a specific subset within these major leukocyte populations (e.g., activated CD4+ T cells, cytolytic T effector cells). Sometimes it is necessary to report changes in the MCF for a particular marker, especially when all the cells of interest homogeneously express that marker (e.g., CD11a, CD54). However, the bottom line in flow cytometric analysis is that each result is expected to accurately reflect the cell population(s) being sampled. In this way, one can be assured that the data can be correctly interpreted and valid conclusions derived. In-depth discussions on the issues presented can be found in Stewart and Stewart (2001b) and Shapiro (2003).

Time Considerations The time required to process, stain, and analyze samples is contingent on many factors, including the number of animals in the experiment and the number of staining combinations. The time needed to prepare single-cell suspensions from lymphoid tissues depends on the organ to be processed (spleen, thymus, and/or lymph nodes). Spleen cell suspensions will require an extra



step in sample processing to remove RBCs, whereas thymus and lymph node cell suspensions will not. The use of microtiter plates will greatly expedite the staining of large numbers of samples derived from lymphoid organs. Even though blood samples require no further processing once the blood is collected, the staining process requires the use of test tubes or microcentrifuge tubes, making sample manipulation more time-consuming. The time required for cell staining is dependent on the number of cell samples to be stained and the staining method employed (direct or indirect). In general, the direct staining method will take 10 min for FcR blocking and 10 min for incubation with fluorochrome-conjugated MAbs, followed by 10 to 20 min for wash steps. Indirect staining will require an additional 20 min to incubate cells with the secondstep reagent, followed by 10 to 20 min for wash steps. The time necessary to acquire flow cytometric data will vary depending on cell concentration and on the total number of events to be collected. For example, it should take less than 1 min per sample to collect 10,000 to 20,000 events when determining the various lineage-specific populations in a sample of spleen cells. However, it will definitely take longer to collect 10,000 events when gating on a smaller population. Instrumental setup will take much longer for multicolor analyses as compared with one-color analyses, as proper compensation will need to be established in multicolor studies. The time involved in data analysis is contingent on the number of fluorochromes used in each MAb cocktail as well as the number of samples processed. Data analysis can take as little as just a few minutes per sample for samples stained with one fluorochrome-conjugated MAb and as many as several hours for samples stained with five different colors.

Acknowledgement The authors would like to acknowledge Castle Funatake for her valuable editorial assistance. This unit was supported by the Cell and Tissue Analysis Facilities and Services Core of the Environmental Health Sciences Center, Oregon State University, grant number P30 ES00210, National Institute of Environmental Health Sciences, National Institutes of Health.

Literature Cited Choi, J.-Y., Oughton, J.A., and Kerkvliet, N.I. 2003. Functional alterations in CD11b+ Gr-1+ cells in mice injected with allogeneic tumor cells

and treated with 2,3,7,8-tetrachlorodibenzo-pdioxin. Int. Immunopharmacol. 3:553-570. Food and Drug Administration. 2002. Guidance for Industry. Immunotoxicology Evaluation of Investigational New Drugs. http://www.fda.gov/ cder/guidance/4945fnl.doc Funatake, C.J., Dearstyne, E.A., Steppan, L.B., Shepherd, D.M., Spanjaard, E.S., MarshakRothstein, A., and Kerkvliet, N.I. 2004. Early consequences of 2,3,7,8-tetrachlorodibenzo-pdioxin exposure on the activation and survival of antigen-specific T cells. Toxicol. Sci. 82:129142. Immunotoxicology Technical Committee, International Life Sciences Institute Health and Environmental Sciences Institute. 2001. Application of flow cytometry to immunotoxicity testing: Summary of a workshop. Toxicology 163:39-48. Kantor, A. and Roederer, M. 1997. FACS analysis of lymphocytes. In Handbook of Experimental Immunology, 5th ed. (L.A. Herzenberg, D.M. Weir, L.A. Herzenberg, and C. Blackwell, eds.) pp. 49.1–49.13. Blackwell Science, Cambridge, UK. Kerkvliet, N.I. and Brauner, J.A. 1990. Flow cytometric analysis of lymphocyte subpopulations in the spleen and thymus of mice exposed to an acute immunosuppressive dose of 2,3,7,8tetrachlorodibenzo-p-dioxin (TCDD). Environ. Res. 52:146-154. Luster, M.I., Portier, C., Pait, D.G., Rosenthal, G.J., Germolec, D.R., Corsini, E., Blaylock, B.L., Pollock, P., Kouchi, Y., Craig, W., White, D.L., Munson, A.E., and Comment, C.E. 1993. Risk assessment in immunotoxicology: II: Relationships between immune and host resistance tests. Fundam. Appl. Toxicol. 21:71-82. Mitchell, K.A. and Lawrence, B.P. 2003. Exposure to 2,3,7,8-tetrachlorodibenzo-p-dioxin (TCDD) renders influenza virus-specific CD8+ T cells hyporesponsive to antigen. Toxicol. Sci. 74:7484. Molecular Probes. 2004a. Zenon technology: Versatile reagents for immunolabeling. In Handbook of Fluorescent Probes and Research Products, 9th ed. http://www.probes.com/handbook/ sections/0703.html Molecular Probes. 2004b. Alexa Fluor dyes: Simply the best. In Handbook of Fluorescent Probes and Research Products, 9th ed. http://www. probes.com/handbook/sections/0103.html Nygaard, U.C. and Lovik, M. 2002. Blood and spleen lymphocytes as targets for immunotoxic effects in the rat: A comparison. Toxicology 174:153-161. Oughton, J.A., Pereira, C.B., DeKrey, G.K., Collier, J.M., Frank, A.A., and Kerkvliet, N.I. 1995. Phenotypic analysis of spleen, thymus, and peripheral blood cells in aged C57Bl6 mice following long-term exposure to 2,3,7,8tetrachlorodibenzo-p-dioxin. Fundam. Appl. Toxicol. 25:60-69. Riedy, M.C., Muirhead, K.A., Jensen, C.P., and Stewart, C.C. 1991. Use of a photolabeling

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technique to identify nonviable cells in fixed homologous or heterologous cell populations. Cytometry 12:133-139. Roederer, M. 1997. Conjugation of Monoclonal Antibodies. http://www.drmr.com/abcon/index. html Roederer, M. 2000. Compensation in Flow Cytometry: An Informal Perspective. http://www. drmr.com/compensation/index.html Shapiro, H.M. 2003. Practical Flow Cytometry, 4th ed. John Wiley & Sons, Hoboken, N.J. Shepherd, D.M., Dearstyne, E.A., and Kerkvliet, N.I. 2000. The effects of TCDD on the activation of ovalbumin (OVA)-specific DO11.10 transgenic CD4(+) T cells in adoptively transferred mice. Toxicol. Sci. 56:340-350. Stewart, C.C. and Stewart, S.J. 2001a. Cell preparation for the identification of leukocytes. In Methods in Cell Biology, Vol. 63 (Z. Darzynkiewicz, H.A. Crissman, and J.P. Robinson, eds.) pp. 217-251. Academic Press, San Diego.

Stewart, C.C. and Stewart, S.J. 2001b. Multiparameter data acquisition and analysis of leukocytes by flow cytometry. In Methods in Cell Biology, Vol. 63 (Z. Darzynkiewicz, H.A. Crissman, and J.P. Robinson, eds.) pp. 289-312. Academic Press, San Diego. Tung, J.W., Parks, D.R., Moore, W.A., and Herzenberg, L.A. 2004. New approaches to fluorescence compensation and visualization of FACS data. Clin. Immunol. 110:277-283. Vandebriel, R.J., Spiekstra, S.W., Hudspith, B.N., Meredith, C., and Van Loveren, H. 1999. In vitro exposure effects of cyclosporin A and bis(tri-nbutyltin)oxide on lymphocyte proliferation, cytokine (receptor) mRNA expression, and cell surface marker expression in rat thymocytes and splenocytes. Toxicology 135:49-66.

Contributed by Julie A. Oughton and Nancy I. Kerkvliet Oregon State University Corvallis, Oregon




In Vitro Model for Modulation of Helper T Cell Differentiation and Activation

UNIT 18.9

This unit provides an in vitro model for evaluating the effects of toxicants on murine helper T cell (TH ) reactivities. Protocols are included for studying the effects of such compounds on the development of precursor or naive CD4+ TH cells (TH0 cells) into type-1 (TH1 ) or type-2 (TH2 ) T helper effectors (see Basic Protocol 1) and on antigen-specific activation of cloned TH1 or TH2 cells (see Basic Protocol 2). The TH1 and TH2 subsets can be distinguished by patterns of cytokine secretion, and they exhibit different helper function for cell-mediated or humoral immunity (Mosmann and Coffman, 1989; Romagnani, 1996). In general, the TH subsets are best differentiated by their quantitative expression of select cytokines (TH0 , low levels of IL-2, IL-4, and IFN-γ; TH1 , high levels of IL-2 and IFN-γ; TH2 , IL-4, IL-5, IL-6, IL-10, and IL-13). For analysis of TH0 differentiation, use of T cells from clonotypic transgenic mice—i.e., DO11.10 ovalbumin-specific transgenic (OV-tg) mice—is described. TH1 and TH2 reactivities are most readily assayed with use of cloned T cell lines (Sanderson et al., 1985; Fiorentino et al., 1989). A helper T cell clone, namely the egg white conalbumin–specific D10.G4.1 TH2 clone from ATCC, is the only commercially available clone at the moment; therefore it is recommended that readers contact authors cited in this unit who have worked with specific helper T cell lines. An in vitro antigen-specific system is valuable for assessing the influence of toxins and toxicants on antigen-dependent interactions between antigen-presenting cells and T cells. The effect of a toxicant on TH development can be investigated by evaluating its modulation of the acquisition of IL-4- or IFN-γ-producing ability in vitro with antigenprimed naive CD4+ T cells. For example, spleens from DO11.10 OVA-tg mice suit the present investigation, because a large proportion (>50%) of the T cells are OVA-specific, naive (if raised in a pathogen-restricted environment), CD4+ T cells. Cloned T cell lines are also available and useful for evaluating toxicant-mediated alteration of antigen-dependent helper T cell activation resulting in cytokine secretion. The cloned TH1 (D1.6) and TH2 (CDC35) lines can be activated in vitro with rabbit IgG (RGG), if presented properly by I-Ad (major histocompatibility complex class II antigen, MHC II) antigen-presenting cells (APC), which include macrophages, dendritic cells, and B cells in the spleen or lymph nodes of H-2d mice (e.g., BALB/c mice). NOTE: All protocols using live animals must first be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) and must follow officially approved procedures for the care and use of laboratory animals.

ANTIGEN-DEPENDENT DEVELOPMENT OF NAIVE HELPER T CELLS TO MEMORY T CELLS

BASIC PROTOCOL 1

In this procedure, splenocytes from unimmunized OVA-tg mice are stimulated with ovalbumin (OVA) in the presence or absence of a test toxicant. T cells are expanded at 72 hr under the same culture conditions as were used for the primary stimulation, and harvested on Day 7. The cells are then washed and restimulated with OVA and antigen-presenting cells (APC) in the absence of the test toxicant. Culture supernatants collected at 48 hr are used for IL-4 and IFN-γ quantification by ELISA (UNIT 18.7).

Immunotoxicology Contributed by Yong Heo Current Protocols in Toxicology (2005) 18.9.1-18.9.10 C 2005 by John Wiley & Sons, Inc. Copyright


Materials OVA-tg mice (The Jackson Laboratory) RPMI 1640 medium (e.g., Invitrogen), ice-cold Red blood cell (RBC) lysing buffer (see recipe) Phosphate-buffered saline (PBS), pH 7.2 to 7.4 (APPENDIX 2A) Complete RPMI medium (see recipe) OVA antigen solution (see recipe) Recombinant human interleukin 2 (IL-2; e.g., Roche) Experimental toxicant(s) (e.g., PbCl2 ) Physiological saline: 0.9% (w/v) NaCl BALB/c mice (The Jackson Laboratory) 60 × 15–mm petri dishes Frosted-end microscope slides, sterilized by autoclaving 6- and 15-ml conical polypropylene centrifuge tubes Low-speed refrigerated centrifuge (e.g., Beckman GPKR or equivalent equipped with GH3.7 swinging-bucket rotor) Hemacytometer or automatic cell counter (e.g., Beckman Coulter) 24- and 12-well culture plates 137 Cs γ irradiator Sandwich ELISA kits for IL-4 and IFN-γ (e.g., BD Pharmingen; optional) Additional reagents and equipment for counting cells (APPENDIX 3B) and sandwich ELISA (UNIT 18.7; also see Heo et al., 1996) NOTE: All solutions and equipment coming into contact with living cells must be sterile and aseptic technique should be used accordingly. NOTE: All incubations are performed in a humidified 37◦ C, 5% CO2 incubator unless otherwise specified.

Prepare a single-cell suspension of spleen cells 1. Collect spleen from OVA-tg mouse and place it in a 60 × 15–mm petri dish containing 6 ml ice-cold RPMI. 2. Homogenize spleen as follows. Wet two frosted-end microscope slides with ice-cold RPMI. Place spleen between the frosted surfaces of the two slides and gently grind into small pieces to dissociate the spleen and free the cells. Rinse the slides with the medium to recover remaining cells. It is necessary to work under a laminar flow hood from this point on to maintain sterility.

3. Transfer the homogenate to a 15-ml centrifuge tube. Rinse the petri dish with 1 ml RPMI to collect remaining cells and transfer this to the tube containing the rest of the homogenate. Place tube on ice and allow cells to settle 3 min. This incubation on ice serves to remove dead cells and other tissue debris, which settle by gravity.

4. Collect supernatant from the settled homogenate and transfer to another 15-ml centrifuge tube. Centrifuge 10 min at 350 × g, 4◦ C. Aspirate and discard supernatant. Resuspend cell pellet in the residual liquid by tapping tube with fingers. 5. Add 2.5 ml RBC lysing buffer and incubate ∼3 min at room temperature. Centrifuge 10 min at 350 × g, room temperature, and discard supernatant. Helper T Cell Differentiation and Activation

It is recommended that RBC lysis not exceed 3 min, because viability of other spleen cells could be affected if the incubation lasts longer.



6. Wash cells by resuspending cell pellet in 10 ml cold PBS, centrifuging 10 min at 350 × g, 4◦ C, and discarding supernatant. Repeat the wash step. Resuspend the cell pellet in 2 ml ice-cold complete RPMI medium and count cells using a hemacytometer or automated cell counter (APPENDIX 3B). Anticipate 60 × 106 spleen cells from one 4- to 6-week old OVA-tg mouse.

Perform primary stimulation and expansion 7. Determine cell number required for the experiment and resuspend the appropriate number of cells in ice-cold complete RPMI. Adjust the final suspension to 2 × 106 cells/ml. 8. Place a 1-ml aliquot of cell suspension into each well of a 24-well culture plate (prepare sufficient wells to test each experimental condition in duplicate). To each of these cell-containing wells, add 0.5 mg OVA antigen and 10 to 12 U human recombinant IL-2. Immediately add experimental toxicant such as PbCl2 to appropriate wells. Incubate 72 hr. It is advisable to include a control well with medium only (no OVA and experimental additives). The final culture volumes of all wells should be made equal by substituting an appropriate amount of vehicle solution, such as physiological saline, for antigen or additives. The appropriate final concentrations of toxicant should be determined with consideration of cytotoxicity, determined by propidium iodide uptake assayed by flow cytometry. It is generally appropriate to use a nanomolar to micromolar range of concentrations for heavy metals or organic compounds.

9. At 72 hr, resuspend cells by vigorous pipetting with Pasteur pipet and transfer cells from each well of the 24-well culture plate to an individual 6 ml-polypropylene tube. Centrifuge 10 min at 350 × g, 4◦ C, and discard supernatants. Resuspend each cell pellet in 2 ml ice-cold complete RPMI and transfer cell suspension from each 6-ml tube to an individual well of a 12-well culture plate. 10. To the wells of the 12-well plate, add the same reagents that were added in step 8, but at 2-fold doses (e.g., 1 mg OVA and 20 to 24 U recombinant IL-2 per well, plus twice the dose of experimental toxicant). Incubate plates an additional 72 hr. 11. At end of expansion period (day 7) harvest cells by vigorous pipetting with 10 mlpipet and transfer to 15 ml-centrifuge tubes. Centrifuge 10 min at 350 × g, 4◦ C, and discard supernatants. Wash cells three times using the technique described in step 5. Resuspend each cell pellet in 1 ml ice-cold complete RPMI and count viable cells by trypan blue exclusion (APPENDIX 3B). Anticipate 12–18 × 106 cells from one well with OVA only, without experimental additives. In the author’s experience, it is not a problem to keep the cells from this step, which have undergone primary stimulation, on ice during the preparation of the APCs in steps 12 and 13, which will take ∼1 hr. However, it is also possible to prepare the APCs just before harvesting the cells that have undergone primary stimulation.

Set up cultures of restimulating memory cells (antigen-presenting cells, APC) 12. Prepare a single-cell suspension from BALB/c mouse spleen as described in steps 1 to 6. Anticipate 6–10 × 107 spleen cells from one 2- to 3-month old BALB/c mouse.

13. Using a 137 Cs γ irradiator, deliver 20 Gy of irradiation to the BALB/c mouse spleen cell suspension. These cells will function as antigen-presenting cells in the following restimulation steps. Immunotoxicology


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14. Prepare 24-well culture plates, adding to each well:

2.5 × 106 irradiated BALB/c mouse spleen cells in complete RPMI 0.5 mg OVA antigen 10 to 12 U recombinant human IL-2. 15. To each well, add the 2 × 105 of the cells obtained in step 11 (i.e., those that have undergone the primary stimulation), adjusting the final volume of each well to 1.3 ml with complete RPMI. Incubate 48 hr. The restimulation is perfomed in the absence of experimental additives to evaluate whether experimental additives present during the primary stimulation of naive helper T cells could direct skewing of helper T cell development toward the type-1 or type-2 helper T cell phenotype.

16. At the end of restimulation, transfer the culture suspension from each well to an individual 6 ml-centrifuge tube. Centrifuge 10 min at 350 × g, 4◦ C, and collect each supernatant in an individual 1.7-ml microcentrifuge tube. Keep the supernatants in freezer below −20◦ C prior to cytokine ELISA.

Determine skewing of naive helper T cells differentiation 17. Determine level of IL-4 and IFN-γ for each individual culture supernatant by a sandwich ELISA (UNIT 18.7). IL-4 and IFN-γ in culture supernatants can be quantitated by a sandwich ELISA method described in the literature (Heo et al. 1996) or supplied commercially (e.g., BD PharMingen). Briefly 96-well Immulon 4 plates (Dynex) are coated by overnight incubation at 4◦ C with capture MAb (100 µl/well; 2 µg/ml). The plates are washed three times and, after a 2-hr incubation with 200 µl per well of PBS containing 10% heat-inactivated FBS at room temperature, the supernatants or the cytokine standards are added to appropriate wells in duplicate. After overnight incubation at 4◦ C, the plates are washed six times and 100 µl per well (1 µg/ml) of biotinylated detection MAb is added. Plates are then incubated at room temperature for 2 hr, washed six times, and further incubated with 100 µl per well (2.5 µg/ml) of avidin-peroxidase for ∼1 hr before detection using the substrate, 2,2 -azino-bis (3-ethylbenzthiazoline 6-sulfonic acid). Absorbance is measured at 405 nm. The amounts of cytokines are calculated from the linear portion of the standard curves. The capture/detection MAb pairs are as follows: IL-4, BVD4-1D11/BVD6-24G2; IFN-γ , R4-6A2/XMG1.2. It is necessary to dilute the supernatants ≥ 10-fold for IFN-γ , but supernatants are used undiluted for IL-4 measurement.

18. Calculate IL-4 versus IFN-γ ratio, and compare the ratio with that for the OVA antigen control (OVA supplementation in the absence of experimental additives at primary stimulation described in step 8). The IL-4:IFN-γ ratio is calculated by dividing the amount of IL-4 by the amount of IFN-γ in supernatants from each individual well. BASIC PROTOCOL 2

ANTIGEN-DEPENDENT ACTIVATION OF CLONED TH CELLS In this procedure, resting D1.6 TH1 (Kurt-Jones et al., 1987) and CDC35 TH2 cells (Tony et al., 1985) are stimulated in the presence or absence of a test toxicant with irradiated BALB/c mouse splenocytes and antigen (RGG). Culture supernatants are collected at 48 hr and used for IL-4 and IFN-γ quantification by ELISA.

Materials

Helper T Cell Differentiation and Activation

BALB/c mice (The Jackson Laboratory) TH cells: D1.6 TH1 clone and CDC35 TH2 clone (available from Dr. David Lawrence, New York State Department of Health; david.lawrence@ wadsworth.org) Recombinant human interleukin 2 (IL-2; e.g., Roche)



RGG antigen stock solution (see recipe) Complete RPMI medium (see recipe) RPMI 1640 medium (e.g., Invitrogen) Ficoll-Paque (Amersham Biosciences) Phosphate-buffered saline (PBS), pH 7.2 to 7.4 (APPENDIX 2A) Experimental toxicant(s) (e.g., PbCl2 ) Physiological saline: 0.9% w/v NaCl 137 Cs γ irradiator 12- and 24-well culture plates 15- and 50-ml conical polypropylene centrifuge tubes 15-ml conical polystyrene centrifuge tubes Low-speed refrigerated centrifuge (e.g., Beckman GPKR or equivalent equipped with GH3.7 swinging-bucket rotor) Additional reagents and equipment for counting cells (APPENDIX 3B), preparing mouse spleen cells (see Basic Protocol 1, steps 1 to 6), and sandwich ELISA (UNIT 18.7; also see Heo et al., 1996) NOTE: All solutions and equipment coming into contact with living cells must be sterile and aseptic technique should be used accordingly. NOTE: All incubations are performed in a humidified 37◦ C, 5% CO2 incubator unless otherwise specified.

Prepare irradiated spleen cells 1. Prepare a single-cell suspension from BALB/c mouse spleen (see Basic Protocol 1, steps 1 to 6). Anticipate 6–10 × 107 spleen cells from one 2- to 3-month old BALB/c mouse.

2. Using a 137 Cs γ irradiator, deliver 20 Gy of irradiation to the BALB/c mouse spleen cell suspension. These cells will function as antigen-presenting cells in the following steps.

Prepare lymphocytes 3. Prepare resting TH cells by adding the following to each well of a 12-well culture plate: 5 × 106 irradiated BALB/c mouse spleen cells in complete RPMI 2 × 105 TH1 or TH2 cells 20 to 24 U recombinant human IL-2 0.2 mg RGG antigen Complete RPMI to 2 ml. Incubate plate 2 weeks. 4. Resuspend the resting TH cells by vigorous pipetting with a Pasteur pipet and transfer the contents of all wells containing TH1 cells to one 50-ml conical polypropylene centrifuge tube and the contents of all wells containing TH2 cells to a second 50-ml polypropylene centrifuge tube. Rinse each well with 1 ml plain RPMI 1640 medium to collect remaining cells, and transfer to the corresponding 50-ml tube containing the rest of the cells. Centrifuge the tubes 10 min at 350 × g, 4◦ C, and discard the supernatants. 5. Resuspend each cell pellet in the residual liquid by tapping the tube with the fingers, and add 2 ml complete RPMI to each of the pellets. Immunotoxicology


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6. For each tube prepared in steps 2 and 3, prepare a 15-ml conical polystyrene centrifuge tube containing 3 ml Ficoll-Paque. Carefully layer the 2-ml cell suspension (step 3) on top of the Ficoll-Paque. It is preferable to use polystyrene centrifuge tube for Ficoll-Paque density gradient isolation of viable cells because the tube is clear, making it easier to identify the lymphocyte layer.

7. Centrifuge the Ficoll-Paque gradients 15 min at 800 × g, room temperature. Using a sterile Pasteur pipet, transfer the lymphocyte layer to a clean 15-ml polypropylene centrifuge tube. The lymphocyte layer is generally found at the interface between the RPMI solution and the Ficoll-Paque.

8. Wash cells three times, each time by adding 10 ml ice-cold PBS, centrifuging 10 min at 350 × g, 4◦ C, and removing the supernatant. Resuspend cells in 1 to 2 ml complete RPMI. Take an aliquot and count viable cells (APPENDIX 3B).

Set up cultures for stimulation 9. Prepare irradiated BALB/c mouse spleen cells (in complete RPMI) as described in steps 1 and 2 of this protocol. Prepare 24-well culture plates, adding to each well: 2.5 × 106 irradiated BALB/c mouse spleen cells in complete RPMI 1 × 105 TH1 or TH2 cells (from step 8) 10 to 12 U recombinant human IL-2. Adjust the volume of each well to 1 ml with complete RPMI. 10. Add 0.1 mg RGG antigen to each well. Add experimental toxicants, e.g., PbCl2 , to appropriate wells. For each individual experimental condition, set up duplicate wells. It is desirable to include a control well with cells only (no RGG or experimental additives). The final culture volumes of all wells should be made equal by substituting appropriate amount of vehicle solution, such as physiological saline, for the RGG antigen or experimental toxicants. In the author’s experience, the final culture volume in each well will be equal to 1.3 ml.

11. Incubate plates 48 hr. Collect supernatant from each well in an individual 1.7-ml microcentrifuge tube. Keep the supernatants in freezer below −20◦ C prior to cytokine ELISA.

Evaluate cytokine expression 12. Determine level of IL-4 for TH2 supernatants or IFN-γ for TH1 supernatants by a sandwich ELISA (UNIT 18.7; also see Basic Protocol 1, step 17 annotation). Compare cytokine levels in experimental samples with those obtained for the RGG antigen control (RGG only without experimental additives; see step 10 annotation). REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.


Complete RPMI medium RPMI 1640 medium (e.g., Invitrogen) containing: 10% (v/v) fetal bovine serum (FBS), heat-inactivated 1 hr at 56◦ C 2 mM L-glutamine 50 µM 2-mercaptoethanol 1 mM sodium pyruvate 1× nonessential amino acids (Invitrogen) continued



1× penicillin/streptomycin/neomycin (PSN) mixture (Invitrogen) 0.075% (w/v) NaHCO3 Store up to 1 week at 4◦ C. OVA antigen stock solution Completely dissolve 2 g ovalbumin (OVA; Sigma, grade V; endotoxin < 10) in 100 ml physiological saline (0.9% w/v NaCl) and filter sterilize through a 0.45-µm low-protein-binding membrane filter (Corning). Determine OVA concentration by spectrophotometry at 280 nm, dividing the optical density value at that wavelength by 0.735 to obtain the concentration of OVA in mg/ml. Aliquot and store indefinitely in freezer below −20◦ C. RBC lysing buffer 4.415 g NH4 Cl (0.15 M) 0.5 g KHCO3 (10 mM) 18.6 mg Na2 EDTA (0.1 mM) 440 ml deionized, distilled H2 O Adjust pH to 7.2 to 7.4 with 1 N HCl. Add H2 O to 500 ml. Filter sterilize through a 0.22-µm membrane filter. Store indefinitely at room temperature. RGG antigen stock solution Completely dissolve 0.2 g rabbit IgG (RGG, Sigma cat. no. I-5006) in 100 ml physiological saline (0.9% w/v NaCl) and filter sterilize through a 0.45-µm low-proteinbinding membrane filter. Determine RGG concentration by spectrophotometry at 280 nm, dividing the optical density value at that wavelength by 1.5 to obtain the concentration of RGG in mg/ml. Aliquot and store indefinitely in freezer below −20◦ C. COMMENTARY Background Information Helper T cells have been subdivided into at least two distinct subsets based on the pattern of cytokine secretion; type-1 T cells produce IL-2 and IFN-γ (and other cytokines), but not IL-4, IL-5, or IL-10, while type-2 T cells produce IL-4, IL-5, and IL-10 (and other cytokines), but not IL-2 or IFN-γ (Cherwinski et al., 1987; Kurt-Jones et al., 1987). Among those cytokines, IL-4 and IFN-γ are known to be the cytokines most useful for defining the effector TH2 and TH1 phenotype subsets, respectively (Sedar and Paul, 1994). The production of distinct cytokines is believed to play a major role in the predominance of TH2 cells over TH1 cells or vice versa, which results in alteration of immune homeostasis between cell-mediated and humoral immunity, leading to various immunopathologic states such as lowered protection against microorganismal infection, allergic state, or autoimmune diseases. Concerning immune-modulatory ability of xenobiotics, differential influences of heavy metals (e.g., mercury and lead) or diesel exhaust particles on helper T cell activities have

been demonstrated, which can cause aberrant cell-mediated immunity or humoral immunity (Prigent et al., 1995; Heo et al., 1996, 2001). Murine T lymphocytes reactive with OVA have been primarily studied with the aim of obtaining specific T cell subsets in the presence of syngeneic BALB/c mouse spleen cells as a source of APC (Hsieh et al., 1992). When T cells from nonimmunized OVA-tg mice are stimulated in vitro with OVA and APC in the presence of immunotoxicant for 6 days, and subsequently restimulated in the absence of the immunotoxicant, these T cells acquire the capacity to produce substantial amounts of TH1 (e.g., IFN-γ) and TH2 (e.g., IL-4) cytokines, but they display an altered ratio of IL4 versus IFN-γ compared with T cells primed with OVA only. The in vitro differentiation model described in Basic Protocol 1 might be close to the in vivo environmental conditions in which differentiation occurs. Basic Protocol 1 can be further extended using naive CD4+ T cells from OVA-tg/RAG (recombinase activating gene)2−/− mice, in which endogenous T cell receptor TCR-α expression

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is completely depleted and CD44+ /LECAMlow memory T cells and B cells are not present (Rulifson et al., 1997). Owing to the complete lack of memory T cells and any TCR other than the DO11.10 clonotype in the OVAtg/RAG 2−/− mice, it is possible to exclude possible contamination with memory T cells in the in vitro differentiation model using OVA-tg mice. Mice bearing transgenic T cell receptor specific for hen egg lysozyme (Desbarats et al., 1999), pigeon cytochrome c (Vratsanos et al., 2001), or myelin basic protein (Olivares-Villagómez et al., 1998) could be used for antigen-dependent differentiation of naive helper T cells to memory T cells. Cell lines expressing I-Ad (e.g., A20, K46R, 2PK3, BCL1 B cell line, or PU5, WEHI265 monocytic cell line) can be employed as antigen-presenting cells (APC) for activating the CDC35 or D1.6 TH cells (Walker et al., 1982; Harris et al., 1984; Zuckerman et al., 1988; Spencer et al., 1993). Use of cell lines for the APC source provides two advantages: first, mice do not have to be sacrificed, and, second, homogeneous preparations of B cells versus monocytic lines can be compared. Furthermore, dendritic cells with high levels of MHC class II expression also can be used as APC; these are prepared by plastic adherence and rosetting with antibody-coated sheep erythrocytes following collagenase digestion of a BALB/c splenocyte suspension (Inaba et al., 1998). Resting cloned CDC35 or D1.6 TH cells can also be stimulated with polyclonal stimulators such as immobilized anti-mouse CD3 or phorbol 12-myristate 13-acetate (PMA) plus ionomycin (Basic Protocol 2). To investigate signaling pathways modulated by immunotoxicant, the author recommends parallel use of antigen/APC-dependent TCR-mediated stimulation with RGG and irradiated syngeneic APC, accessory cell–independent CD3/TCR complex–mediated stimulation with immobilized anti-CD3, and stimulation bypassing phospholipase C–initiated early signaling events with PMA for activation of protein kinase C, plus a calcium ionophore for upregulation of intracellular free calcium (Heo et al., 1997).

Critical Parameters and Troubleshooting Helper T Cell Differentiation and Activation

Experience has shown that cells should be completely harvested through vigorous pipetting at the time of expansion and restimulation of OVA-specific T cells (Basic Protocol 1). It will be necessary to rinse culture wells once

with RPMI and collect the rinsed-out suspension as well. It is recommended that exogenous recombinant IL-2 be added for in vitro differentiation of naive TH cells and activation of TH clones. Although cloned TH cells can proliferate and be activated in culture when stimulated with antigen in the absence of exogenous IL-2, better growth of TH cells is obtained when these are cultured with both IL-2 and antigen. IL-2 is known not to have direct effects on TH subset differentiation (Duncan and Lawrence, 1995). When an irradiation source is not available, treatment of APC with mitomycin C (a DNA cross-linking agent) can be used; like irradiation, this preserves the antigen-presenting function of syngeneic antigen-presenting cells. Mitomycin C should be completely washed out prior to mixing the APC with cloned TH cells, because mitomycin C exerts its cytotoxicity by blocking cell division. Since activated lymphocytes will tend to enlarge, microscopic examination of the cultures for blasts will give an indication of successful activation. Meanwhile, to obtain TH clonal cells at stationary phase, the cells should be passaged at least 14 days after initiating culture with soluble antigen (Kimoto and Fathman, 1982). With all assays, it is critical that cultures in each well be adjusted to equal final volumes in order to correctly compare the resulting levels of cytokines. Experience has shown that recovery of live lymphocytes from spleen or culture wells is significantly enhanced if the resuspensions after each centrifugation are done by first discarding the supernatant, then resuspending the cells by tapping the tube with the fingers, and finally adding the solution for resuspension, in that order.

Anticipated Results

Using Basic Protocol 1, ∼50 ng/ml of IFNγ and 1 ng/ml of IL-4 will be detected in a well in which 2 × 105 OVA-specific TH cells, differentiated in the absence of experimental additives, are stimulated with 2.5 × 106 APC. Skewing of TH cell development toward the TH2 phenotype has been reported in the presence of heavy metal (25 µM Pb) during the primary stimulation of naive TH cells with OVA. With this concentration of Pb, Heo et al. (1997) detected 1.2±0.2 ng/ml IL-4 and 32±7 ng/ml IFN-γ, while with no experimental additives, they detected 0.8±0.2 ng/ml IL-4 and 57±9 ng/ml IFN-γ. Using Basic Protocol 2, the levels of IL-4 and IFN-γ from CDC35 TH2 and D1.6 TH1 cells stimulated with RGG alone will be ∼2 to 4 ng/ml and 4 to 8 ng/ml, respectively.



Pb-induced differential effects on activities of TH2 and TH1 cells has been demonstrated using CDC35 TH2 and D1.6 TH1 clones, in that the levels of IL-4 from CDC35 and IFNγ from D1.6 clonal cells were 1.6±0.6 and 3.5±0.1 ng/ml, respectively. Addition of 25 µM Pb enhanced IL-4 production to 128% and inhibited IFN-γ production to 53% (Heo et al., 1997).

Time Considerations In Basic Protocol 1, 2 to 3 hr of work are required for the steps from removal of spleens up to the point where the cells are ready for incubation on culture plates. Following an additional 6 days of primary stimulation and expansion, preparation of antigen-presenting cells and mixed culture with resting OVA-specific TH cells takes ∼4 hr. In Basic Protocol 2, preparing APCs and setting up the mixed culture with resting D1.6 or CDC35 TH clonal cells takes ∼4 hr. The exact length of time will depend on the size of the experiment and the skill of investigator.

Literature Cited Cherwinski, H.M., Schumacher, J.H., Brown, K.D., and Mosmann, T.R. 1987. Two types of mouse helper T cell clone. III. Further differences in lymphokine synthesis between Th1 and Th2 clone revealed by RNA hybridization, functionally monospecific bioassays, and monoclonal antibodies. J. Exp. Med. 166:1229-1244. Desbarats, J., Wade, T., Wade, W.F., and Newell, M.K. 1999. Dichotomy between na¨ıve and memory CD4+ T cell responses to Fas engagement. Proc. Natl. Acad. Sci. U.S.A. 96:8104-8109. Duncan, D.D. and Lawrence, D.A. 1995. T cells and cloned and transformed T-cell lines to assess immune function. In Methods in Immunotoxicology Vol. 1 (G.R. Burleson, J.H. Dean, and A.E. Munson, eds.) pp. 483-505. Wiley-Liss, New York. Fiorentino, D.F., Bond, M.W., and Mosmann, T.R., 1989. Two types of mouse T helper cell. IV. Th2 clones secret a factor that inhibits cytokine production by Th1 clones. J. Exp. Med. 170:20812095. Harris, M.R., Kindle, C.S., Abruzzini, A.F., Pierce, C.W., and Cullen, S.E. 1984. Antigen presentation by the BCL1 murine B cell line: In vitro stimulation by LPS. J. Immunol. 133:12021208. Heo, Y., Parsons, P.J., and Lawrence, D.A. 1996. Lead differentially modifies cytokine production in vitro and in vivo. Toxicol. Appl. Pharmacol. 138:149-157. Heo, Y., Lee, W.T., and Lawrence, D.A. 1997. Differential effects of lead and cAMP on development and activities of Th1- and Th2lymphocytes. Toxicol. Sci. 43:172-185.

Heo, Y., Saxon, A., and Hankinson, O. 2001. Effect of diesel exhaust particles and their components on the allergen-specific IgE and IgG1 response in mice. Toxicology 159:3143-3158. Hsieh, C.-S., Heimberger, A.B., Gold, J.S., O’Garra, A., and Murphy, K.M., 1992. Differential regulation of T helper phenotype development by interleukin 4 and 10 in an αβT-cellreceptor transgenic system. Proc. Natl. Acad. Sci. U.S.A. 89:6065-6069. Inaba, K., Swiggard, W.J., Steinman, R.M., Romani, N., and Schuler, G. 1998. Isolation of dendritic cells. In Current Protocols in Immunology (J.E. Coligan, A.M. Kruisbeek, D.H. Margulies, E.M. Shevach, and W. Strober, eds.) pp. 3.7.1-3.7.15. John Wiley & Sons, Hoboken, N.J. Kimoto, M. and Fathman, C.G., 1982. Immunization and long-term culture of murine immune lymph node cells. In Isolation, Characterization, and Utilization of T Lymphocyte Clones. (C.G. Fathman and F.W. Fitch, eds.) pp. 525-526. Academic Press, San Diego, Calif. Kurt-Jones, E.A., Hamberg, S., Ohara, J., Paul, W.E., and Abbas, A.K. 1987. Heterogeneity of helper/inducer T lymphocytes I. Lymphokine production and lymphokine responsiveness. J. Exp. Med. 166:1774-1787. Mosmann, T.R. and Coffman, R.L. 1989. TH1 and TH2 cells: Different patterns of lymphokine secretion lead to different functional properties. Annu. Rev. Immunol. 7:145-173. Olivares-Villagómez, D., Wang, Y., and Lafaille, J.J. 1998. Regulatory CD4+ T cells expressing endogenous T cell receptor chains protect myelin basic protein-specific transgenic mice from spontaneous autoimmune encephalomyelitis. J. Exp. Med. 188:1883-1894. Prigent, P., Saoudi, A., Pannetier, C., Graber, P., Bonnefoy, J.Y., Druet, P., and Hirsch, F. 1995. Mercuric chloride, a chemical responsible for T helper cell (Th2)-mediated autoimmunity in Brown Norway rats, directly triggers T cells to produce interleukin-4. J. Clin. Invest. 96:14841489. Romagnani, S. 1996. Th1 and Th2 Cells in Health and Disease, Karger, Basel, Switzerland. Rulifson, I.C., Sperling, A.I., Field, P.E., Fitch, F.W., and Bluestone, J.A. 1997. CD28 costimulation promotes the production of Th2 cytokines. J. Immunol. 158:658-665. Sanderson, C.J., Strath, M., Warren, D.J., O’Garra, A., and Kirkwood, T.B. 1985. The production of lymphokines by primary alloreactive T-cell clones: A co-ordinate analysis of 233 clones in seven lymphokine assays. Immunology 56:575584. Sedar, R.A. and Paul, W.E. 1994. Acquisition of lymphokine-producing phenotype by CD4+ T cells. Annu. Rev. Immunol. 12:635-673. Spencer, J.S., Freed, J.H., and Kubo, R.T. 1993. Expression and function of mixed iostype MHC class II molecules in normal mice. J. Immunol. 151:6822-6832.

Immunotoxicology


Supplement 24

Tony, H.P., Phillips, N.E., and Parker, D.C. 1985. Role of membrane immunoglobulin (Ig) crosslinking in membrane Ig-mediated, major histocompatibility complex-restricted T cell–B cell cooperations. J. Exp. Med. 162:1695-1708. Vratsanos, G.S., Jung, S.-S., Park, Y.-M., and Craft, J. 2001. CD4+ T cells from lupus-prone mice are hyperresponsive to T cell receptor engagement with low and high affinity peptide antigens: A model to explain spontaneous T cell activation in lupus. J. Exp. Med. 193:329-337. Walker, E., Warner, N.L., Chestnut, R., Kappler, J., and Marrack, P. 1982. Antigen-specific, Iregion-restricted interactions in vitro between tumor cell lines and T cell hybridomas. J. Immunol. 128:2164-2169. Zuckerman, S.H., Schreiber, R.D., and Marder, P. 1988. Reactivation of class II antigen

expression in murine macrophage hybrids. J. Immunol. 140:978-983.

Key References Heo et al., 1996. See above. Shows how metal toxicants differently modify helper T cell subset activation in vitro. Heo et al., 1998. See above. Contains the first description of lead(Pb)-mediated helper T cell development skewing toward type-2 effectors in vitro.

Contributed by Yong Heo Catholic University of Daegu Kyongbuk Province Republic of Korea




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APPENDIX 1

Safe Use of Radioisotopes

APPENDIX 1A

The use of radioisotopes to label specific molecules in a defined way has greatly furthered the discovery and dissection of biochemical pathways. The development of methods to inexpensively synthesize such tagged biological compounds on an industrial scale has enabled them to be used routinely in laboratory protocols, including many detailed in this manual. Although most of these protocols involve the use of only microcurie (µCi) amounts of radioactivity, some (particularly those describing the metabolic labeling of proteins or nucleic acids within cells) require amounts on the order of millicuries (mCi). In all cases where radioisotopes are used, depending on the quantity and nature of the isotope, certain precautions must be taken to ensure the safety of the scientist. This appendix outlines a few such considerations relevant to the isotopes most frequently used in biological research. In designing safe protocols for the use of radioactivity, the importance of common sense, based on an understanding of the general principles of isotopic decay and the importance of continuous monitoring with a hand-held radioactivity monitor (e.g., Geiger counter), cannot be overemphasized. In addition, it is also critical to take into account the rules, regulations, and limitations imposed by each specific institution. These are usually not optional considerations: an institution’s license to use radioactivity normally depends on strict adherence to such rules. Many of the protocols described have evolved (and are evolving still) over the years. The authors are indebted to those who have trained them in the safe use of radioactivity, in particular to the members of the Salk Institute Radiation Safety Department. Most of the designs for the shields and other safety equipment shown in Figures A.1A.2A, A.1A.3, and A.1A.4 were created at the Salk Institute in collaboration with Dave Clarkin and Mario Tengco. Safety equipment of similar design is available from several commercial vendors, including CBS Scientific and Research Products International.

BACKGROUND INFORMATION The Decay Process As anyone who has taken a basic chemistry course will remember, each element is characterized by its atomic number, defined as the number of orbital electrons or the number of protons in the nucleus of that atom. Isotopes of a given element exist because some atoms of each element, while by definition having the same number of protons, have a different number of neutrons and therefore a different nuclear weight. It should be noted that the number of electrons outside the nucleus remains the same for all isotopes of a given element, and so all isotopes of a given element are equivalent with respect to their chemical reactivity. Radioactive decay occurs when subatomic particles are released from the nucleus of an atom of a heavy isotope. This often results in the conversion of an atom of one isotope to an isotope of a different element, because the original isotope’s atomic number changes after decay. The subatomic particles released from naturally occurring radioisotopes are of three basic types: α and β particles and γ rays (see Table A.1A.1). An α particle is essentially the nucleus of a helium atom, or two protons plus two neutrons. It is a relatively large, heavy particle that moves slowly and usually only across short distances before it encounters some other atom with which it interacts. These particles are released from isotopes with large nuclei (atomic number >82; e.g., plutonium or uranium); such isotopes are not commonly used in biological research. In contrast to α particles, β particles are light, high-speed charged particles. Negatively charged β particles are essentially electrons of nuclear origin that are released when a neutron is converted to a proton. Release of a β particle thus changes the atomic number and elemental status of the isotope. γ radiation has both particle and wave properties; its wavelength falls within the range of X-ray wavelengths. The distinction between γ rays and X rays was made when primitive X-ray machines produced X rays with a wavelength longer than those of the γ rays produced naturally by radioisotopes. Modern X-ray machines

Contributed by Jill Meisenhelder and Kentaro Semba Current Protocols in Toxicology (1999) A.1A.1-A.1A.13 Copyright © 1999 by John Wiley & Sons, Inc.

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A.1A.1

Table A.1A.1

Nuclide 3

Physical Characteristics of Commonly Used Radionuclidesa

Half-life

H 14 C 32 b P

12.43 years 5370 years 14.3 days

33 b

25.4 days 87.4 days 60 days 8.04 days

P S 125 c I 131 c I 35

Emission β β β

β β γ β γ

Approx. specific Energy, max Range of Atom resulting activity at 100% Target organ (MeV) emission, max from decay enrichment (Ci/mg) 0.0186 0.156 1.71

0.42 cm (air) 21.8 cm (air) 610 cm (air) 0.8 cm (water) 0.76 cm (Plexiglas) 0.249 49 cm 0.167 24.4 cm (air) 0.27–0.035 0.2 mm (lead) 0.606 165 cm (air) 0.364 2.4 cm (lead)

9.6 4.4 mCi/mg 285

3 2

156 43 14.2 123

33 16 35 17 125 52 130 54

He

14N 7 33 16

S

S Cl Te Xe

Whole body Bone, fat Bone

Bone Testes Thyroid Thyroid

aTable compiled based on information in Lederer et al. (1967) and Shleien (1987). bRecommended shielding is Plexiglas; half-value layer measurement is 1 cm. cRecommended shielding is lead; half-value layer measurement is 0.02 mm.


produce a much broader spectrum of wavelengths, including γ radiation; currently this sort of X-ray radiation is termed γ when it is of nuclear origin. Unlike β-particle release, the release of γ radiation by itself produces an isotopic change rather than an elemental one; however, the resultant nuclei are unstable and often decay further, releasing β particles. The energy of all α particles and γ rays (measured in electron volts) is fixed, because they are of specific composition or wavelength. The energy of β particles, however, varies depending on the atom they originate from (and with concomitant release of neutrinos or antineutrinos that serve to balance the conservation of energy aspect of the decay equation). Thus there are (relatively) high-energy β particles released during the decay of 32P and low-energy β particles released when tritium (3H) decays. Isotopic decay usually involves a chain or sequence of events rather than a single loss of a particle, because the resultant, equally unstable atoms try to achieve equilibrium. During this course of decay, secondary forms of radiation can be generated that may also pose a hazard to workers. For example, when high-energy β particles released during the decay of 32P encounter the nuclei of atoms with a large atomic number, a strong interaction occurs. The β particle loses some energy in the form of a photon. Such photons are called Bremsstrahlung radiation; they are detectable using a monitor suitable for the detection of γ or X rays. Following their release, α, β, and γ emissions (as well as secondary forms of radiation) travel

varying average distances at varying average speeds, depending on their energy and the density of the material through which they are moving. The distance they actually travel before encountering either the electrons or nucleus of another atom is termed their degree of penetrance. This value is expressed as an average for each type of particle. The energy of the particles released (and therefore their potential penetrance) thus dictates what type of shielding, if any, is necessary for protection against the radiation generated by the decay of a given isotope. α, β, and γ emissions all have the potential, upon encountering an atom, to knock out its electrons, thereby creating ions. Thus, these three types of emissions are called ionizing radiation. The formation of such ions may result in the perturbation of biological processes: therein lies the danger associated with radioactivity!

Measuring Radioactivity and Individual Exposure to It The radioactivity of a given substance is measured in terms of its ionizing activity. A curie (Ci) by definition is the amount of radioactive material that will produce 3.7 × 1010 disintegrations per second (see Table A.1A.2). This, not coincidentally, happens to be the number of disintegrations that occur during the decay of one gram of radium and its decay products. Exposure to such radiation is measured as the amount of energy absorbed by the recipient, which, of course, is directly related to the potential damage such radiation may cause. One rad is the dose of radiation that will

A.1A.2 Current Protocols in Toxicology

Table A.1A.2

Conversion Factors for Radioactivity

Measurement of Radioactivity The SI unit for measurement of radioactivity is the becquerel: 1 becquerel (Bq) = 1 disintegration per second The more commonly encountered unit is the curie (Ci): 1 Ci = 3.7 × 1010 Bq = 2.22 × 1012 disintegrations per minute (dpm) 1 millicurie (mCi) = 3.7 × 107 Bq = 2.22 × 109 dpm 1 microcurie (µCi) = 3.7 × 104 Bq = 2.22 × 106 dpm Conversion factors: 1 day = 1.44 × 103 min = 8.64 × 104 sec 1 year = 5.26 × 105 min = 3.16 × 107sec counts per minute (cpm) = dpm × (counting efficiency) Measurement of Dose The SI unit for energy absorbed from radiation is the gray (Gy): 1 Gy = 1 joule/kg Older units of absorbed energy are the rad (r) and roentgen (R): 1 r = 100 ergs/g = 10−2 Gy 1 R = 0.877 r in air = 0.93 − 0.98 r in water and tissue The SI unit for radiation dosage is the sievert (Sv), which takes into account the empirically determined relative biological effectiveness (RBE) of a given form of radiation: dosage [Sv] = RBE × dosage [Gy] (biological effect of a dose of standard radiation [Gy]) RBE = (biological effect of a dose of other radiation [Gy]) RBE = 1 for commonly encountered radionuclides The older unit for dosage is the rem (roentgen-equivalent-man): 1 rem = 0.01 Sv cause 100 ergs of energy to be absorbed per gram of irradiated material. Another unit commonly used to measure radiation doses is the rem; this is related to the rad but takes into account a “quality factor” based on the type of ionizing radiation being received. For β particles and γ or X rays this factor is 1; therefore, rems β equal rads β. In contrast, the quality factor associated with α particles is 20, so an exposure of one rad due to α particles would be recorded as 20 rem. The amount or dose of radiation received by materials (cells, scientists, etc.) near the source depends not only on the specific type and energy (penetrance) of the radiation being produced, but also on the subject’s distance from the source, the existence of any intervening layers of attenuating material (shielding), and the length of time spent in the vicinity of the radiation source. To best measure such doses, every person working with or around radioactivity should wear an appropriate type of radiation detection badge (in addition to carrying a portable radiation monitor that can give an immediate, approximate reading). This is nor-

mally a requirement for compliance with an institution’s license to use radioisotopes. Such badges are usually furnished by the safety department and collected at regular intervals for reading by a contracted company. At most institutions, the old-style film badges have been replaced with the more accurate TLDs (thermoluminescent dosimeters). These take advantage of chemicals such as calcium or lithium fluorides which, following exposure to ionizing radiation, will luminesce at temperatures below their normal thermal luminescence threshold. Different types of badges are sensitive to different types of radiation: always be sure to wear one that is appropriate for detecting exposure to the isotope being used! In most places pregnant women are asked to wear a more sensitive (and more expensive) dosimeter to better monitor their (and the developing fetus’) exposure. Most often workers will be asked to wear a radiation detection badge on the labcoat lapel in order to measure whole-body radiation. When working with 32P or 125I it is also advisable to wear a ring badge to measure exposure to the unshielded extremities (fingers). The

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limit set for “acceptable” exposure to wholebody radiation is several-fold less than the limit set for extremities. Nevertheless, the authors have found that the exposure recorded on ring badges is often significant with respect to the limit for extremities set by our institution. What is known about the effects on humans of exposure to low levels of radiation (i.e., levels which would be received when briefly handling millicurie or microcurie amounts of radioactivity)? Not much, for the obvious reason that direct studies have not been undertaken. Accordingly, guidelines for exposure levels are set using extrapolations—either by extrapolating down from population statistics obtained following accidents or disasters (the Chernobyl meltdown, atomic bombings) or by extrapolating up from numbers obtained from animal experiments. Each form of extrapolation is subject to caveats, and given that predictions based on such extrapolations cannot be perfect, most health and safety personnel aim for radiation exposure levels said to be ALARA or “as low as reasonably achievable.” Further discussion of exposure limits and the statistics on which they are based may be found in B. Shleien’s health physics text (Shleien, 1987). Limiting exposure to radiation can be accomplished by adjusting several parameters of the exposure: the duration of exposure, distance from the source, and the density of the material (air, water, shielding) between the individual and the source. Time is of the essence When designing any experiment using radioactivity, every effort should be made to limit the time spent directly handling the vials or tubes containing the radioactive material. Speed should be encouraged in all manipulations, though not to the point of recklessness! Have everything needed for the experiment ready at hand before the radioactivity is introduced into the work area.


Distance helps to determine dose When possible, experiments involving radioactivity should be performed in an area separate from the rest of the lab. Many institutions require that such work be performed in a designated “hot lab”; however, if many people in the laboratory routinely use radioisotopes, it is less than feasible to move them all into what is usually a smaller space. No matter where an individual is working, it is his or her responsibility to monitor the work area and ensure his or her own safety and the safety of those work-

ing nearby by using adequate shielding. Obviously, when handling the radioactive samples, it is necessary to work rapidly behind any required shielding. To protect bystanders, remember that the intensity of radiation from a source (moving through air) falls off in proportion to the square of the distance. Thus, if standing 1 foot away from a source for 5 min would result in an exposure of 45 units, standing 3 feet away for the same amount of time would result in an exposure (1/3)2 of 45 units, or 5 units. This factor is also relevant when considering the storage of large amounts of radioactivity, particularly 125I or 32P, as no amount of shielding can completely eliminate radiation. Shielding is the key to safety As mentioned above, the energy of the particle(s) released during the decay of an isotope determines what, if any, type of shielding is appropriate (see Table A.1A.3). β particles released during the decay of 14C and 35S possess roughly ten times the energy of those released when 3H decays. All three β particles are of relatively low energy, do not travel very far in air, and cannot penetrate solid surfaces. No barriers are necessary for shielding against this type of β radiation. The major health threat from these isotopes occurs through their accidental ingestion, inhalation, or injection. β particles released during the decay of 32P have 10-fold higher energy than those released from 14C and pose a significant threat to workers. (One reported hazard is the potential for induction of cataracts in the unshielded eye.) The fact that these high-energy beta particles can potentially generate significant amounts of Bremsstrahlung radiation is the reason that low-density materials are used as the primary layer of shielding for 32P β radiation. Water, glass, and plastic are suitable low-density materials (as opposed to lead). Obviously water is unsuitable as a shielding layer for work on the bench, although it does a reasonable job when samples are incubating in a water bath. Shields made from a thickness of glass sufficient to stop these particles would be extremely heavy and cumbersome (as well as dangerous if dropped). Fortunately, plastic or acrylic materials—variously called Plexiglas, Perspex, or Lucite—are available for shielding against 32P β radiation. Shields as well as storage boxes constructed of various thicknesses of Plexiglas are necessary equipment in laboratories where 32P is used. When millicurie amounts of 32P are used at one time it is necessary to also block the Bremsstra-


Table A.1A.3

Shielding Radioactive Emissiona

β emitters Energy Mass (mg)/cm2 to reduce (MeV) intensity by 50% 0.1 1.0 2.0 5.0

Thickness (mm) to reduce intensity by 50% Water Glass Lead Plexiglas

1.3 48 130 400

0.013 0.48 1.3 4.0

0.005 0.192 0.52 1.6

0.0011 0.042 0.115 0.35

0.0125 0.38 1.1 4.2

γ emitters Energy (MeV) 0.5 1.0 2.0 3.0

Thickness of material (cm) to attenuate a broad beam of γ-rays by a factor of 10 Water Aluminum Iron Lead 54.6 70.0 76.0 89.0

20.3 24.4 32.0 37.0

6.1 8.2 11.0 12.0

1.8 3.8 5.9 6.4

aFrom Dawson et al. (1986). Reprinted with permission.

hlung radiation by adding a layer of high-density material (such as 4 to 6 mm lead) to the outside of the Plexiglas shield (covering the side farthest from the radioactive source). γ rays released during the decay of 125I have much higher penetrance than the β particles from 32P decay; this radiation must be stopped by very-high-density material, such as lead. Lead foil of varying thicknesses (2 to 6 mm) can be purchased in rolls and can be cut and molded to cover any container, or taped to a Plexiglas shield (used in this instance for support). Obviously this latter arrangement has the disadvantage that it is impossible to see what one is doing through the shield. For routine shielding of manipulations involving 125I, it is useful to purchase a lead-impregnated Plexiglas shield that is transparent, albeit inevitably very heavy (as well as relatively expensive). Although it seems logical that the use of more radioactivity necessitates the use of thicker layers of shielding, it is also true that no shielding material is capable of completely stopping all radiation. When deciding how thick is “thick enough,” consult the half-value layer measurement for each type of shielding material. This number gives the thickness of a given material necessary to stop half the radiation from a source; Table A.1A.3 lists half-value layer measurements for the β particles released from isotopes commonly used in the biology laboratory. In general, 1 to 2 cm of Plexiglas and/or 0.02 mm of lead are sufficient to shield the amounts of radioactivity used in most experiments.

GENERAL PRECAUTIONS Before going on to a discussion of specific precautions to be taken with individual isotopes, a short list of general precautions to be taken with all isotopes seems pertinent: 1. Know the rules. Be sure that each individual is authorized to use each particular isotope and uses it in an authorized work area. 2. Don the appropriate apparel. Whenever working at the lab bench, it is good safety practice to wear a labcoat for protection. Disposable paper/synthetic coats of various styles are commercially available: at $4 each these may be conveniently thrown out if contaminated with radioactivity during an experiment, rather than held for decay as might be preferable with cloth coats costing about $30 each. As an alternative, disposable sleeves can be purchased and worn over those of the usual cloth coat. Other necessary accessories include radiation safety badges, gloves, and protective eyewear. It’s handy to wear two pairs of gloves at once when using radioactivity; when the outer pair becomes contaminated, it is possible to strip it off and continue working without interruption. 3. Protect the work area as well as the workers. Lab benches and the bases of any shields used should be covered with some sort of disposable, preferably absorbent, paper sheet. Underpads or diapers (the kind normally used in hospitals) are convenient for this purpose. 4. Use appropriate designated equipment. It is very convenient, where use justifies

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the expense, to have a few adjustable pipettors dedicated for use with each particular isotope. Likewise, it is good practice to use only certain labeled centrifuges and microcentrifuge rotors for radioactive samples so that all the lab’s rotors do not become contaminated. Although such equipment should be cleaned after each use, complete decontamination is often not possible. A few pipettors or a single microcentrifuge can easily be stored (and used) behind appropriate shielding. Actually, contamination of the insides and tip ends of pipettors can be greatly reduced by using tips supplied with internal aerosol barriers; these have recently become very popular for setting up PCR reactions and are available to fit a variety of pipettors. To prevent contamination of the outside of the pipettor’s barrel, simply wrap the hand-grip in Parafilm, which can be discarded later. Several manufacturers sell disposable paper inserts for their microcentrifuges that protect the wall of the rotor chamber from contamination that might spin off the outside of sample tubes. Trying to fashion homemade liners of this sort is not recommended, as we have had disastrous experiences using laminated adhesive paper that “unstuck” during microcentrifuge spins. These liners would get caught by the rotor, shattering sample tubes and creating an even bigger mess! 5. Know where to dispose of radioactive waste, both liquid and solid. Most institutions require that radioactive waste be segregated by isotope. This is done not only so that appropriate shielding can be placed around waste containers, but so that some waste can be allowed to decay prior to disposal through normal trash channels (see Table A.1A.4 and Fig. A.1A.1). With a decreasing number of radioactive waste disposal facilities able or willing to accept radioactive waste for burial (and a concomitant increase in dumping charges from those that still do), this practice of on-site decay can save an institution thousands of dollars a year in disposal charges. 6. Label your label! It is only common courtesy (as well as common sense) to alert coworkers to the existence of anything and everything radioactive that is left where they may come in contact with it! A simple piece of tape affixed to the sample box—with the investigator’s name, the amount and type of isotope, and the date written on it—should do the trick. Yellow hazard tape printed with the international symbol for radioactivity is commercially available in a variety of widths.

7. Monitor radioactivity early and often. Portable radiation detection monitors are essential equipment for every laboratory using radioactivity. No matter how much or how little radioactivity is being used, the investigator should keep a hand-held monitor nearby—and it should be on! Turn it on before touching any radioactivity to avoid contaminating the monitor’s switch. Use a monitor with the appropriate detection capacity (β for 35S and 32P; γ for 125I) before, during, and after all procedures. The more frequently fingers and relevant equipment are monitored, the more quickly a spill or glove contamination will be detected. Such timely detection will keep both the potential mess and the cleanup time to a minimum. Because lowenergy β emitters such as 3H cannot be detected using such monitors, wipe tests of the bench and equipment used are necessary to ensure that contamination of the work area did not occur.

SPECIFIC PRECAUTIONS The following sections describe precautions to be taken with individual isotopes in specific forms. Although the sections dealing with 35Sor 32P-labeling of proteins in intact cells are presented in terms of mammalian cells, most of the instructions are also pertinent (with minimal and obvious modifications) to the labeling of proteins in other cells (bacterial, insect, etc.).

Working with 35S Using 35S to label cellular proteins and proteins translated in vitro As discussed above, the β radiation generated during 35S decay is not strong enough to make barrier forms of shielding necessary. The risk associated with 35S comes primarily through its ingestion and subsequent concentration in various target organs, particularly the testes. Although willful ingestion of 35S seems unlikely, accidental or unknowing ingestion may be more common. As reported several years ago (Meisenhelder and Hunter, 1988), 35S-labeled methionine and cysteine, which are routinely used to label proteins in intact cells and by in vitro translation, break down chemically to generate a volatile radioactive component. The breakdown occurs independent of cellular metabolism. Thus the radioactive component is generated to the same extent in stock vials as in cell culture dishes. The process seems to be promoted by freezing and thawing 35S-labeled materials. The exact identity of this component is not known, although it is probably SO2 or CH3SH. What is known is that it dissolves read-


Table A.1A.4 Decay factors for calculating the amount of radioactivity present at a given time after a reference date. For example, a vial containing 1.85 MBq (50 µCi) of a 35S-labeled compound on the reference date will have the following activity 33 days later: 1.85 × 0.770 = 1.42 MBq; 50 × 0.770 = 38.5 µCi.

0 20 40 60 80 100 120 140 160 180 200 220 240

131I

Days 0 1.000 0.794 0.630 0.500 0.397 0.315 0.250 0.198 0.157 0.125 0.099 0.079 0.063

2 0.977 0.776 0.616 0.489 0.388 0.308 0.244 0.194 0.154 0.122 0.097 0.077 0.061

4 0.955 0.758 0.602 0.477 0.379 0.301 0.239 0.189 0.150 0.119 0.095 0.075 0.060

6 0.933 0.741 0.588 0.467 0.370 0.294 0.233 0.185 0.147 0.117 0.093 0.073 0.058

8 0.912 0.724 0.574 0.456 0.362 0.287 0.228 0.181 0.144 0.114 0.090 0.072 0.057

10 0.891 0.707 0.561 0.445 0.354 0.281 0.223 0.177 0.140 0.111 0.088 0.070 0.056

12 0.871 0.691 0.548 0.435 0.345 0.274 0.218 0.173 0.137 0.109 0.086 0.069 0.054

14 0.851 0.675 0.536 0.425 0.338 0.268 0.213 0.169 0.134 0.106 0.084 0.067 0.053

16 0.831 0.660 0.524 0.416 0.330 0.262 0.208 0.165 0.131 0.104 0.082 0.065 0.052

18 0.812 0.645 0.512 0.406 0.322 0.256 0.203 0.161 0.128 0.102 0.081 0.064 0.051

0 4 8 12 16 20 24 28 32 36 40 44 48 52

Half-life: 14.3 days Hours 0 1.000 0.824 0.679 0.559 0.460 0.379 0.312 0.257 0.212 0.175 0.144 0.119 0.098 0.080

0 3 6 9 12 15 18 21 24 27 30 33 36

Hours 0 1.000 0.772 0.596 0.460 0.355 0.274 0.212 0.164 0.126 0.098 0.075 0.058 0.045

6 0.979 0.756 0.583 0.450 0.348 0.269 0.207 0.160 0.124 0.095 0.074 0.057 0.044

12 0.958 0.740 0.571 0.441 0.340 0.263 0.203 0.157 0.121 0.093 0.072 0.056 0.043

18 0.937 0.724 0.559 0.431 0.333 0.257 0.199 0.153 0.118 0.091 0.071 0.054 0.042

24 0.917 0.708 0.547 0.422 0.326 0.252 0.194 0.150 0.116 0.089 0.069 0.053 0.041

Days

33P

12 0.976 0.804 0.662 0.546 0.449 0.370 0.305 0.251 0.207 0.170 0.140 0.116 0.095 0.078

35S

Half-life: 8.04 days 30 0.898 0.693 0.533 0.413 0.319 0.246 0.190 0.147 0.113 0.088 0.068 0.052 0.040

36 0.879 0.678 0.524 0.405 0.312 0.241 0.186 0.144 0.111 0.086 0.066 0.051 0.039

42 0.860 0.664 0.513 0.396 0.306 0.236 0.182 0.141 0.109 0.084 0.065 0.050 0.039

48 0.842 0.650 0.502 0.387 0.299 0.231 0.178 0.138 0.106 0.082 0.064 0.049 0.038

54 0.824 0.636 0.491 0.379 0.293 0.226 0.175 0.135 0.104 0.080 0.063 0.048 0.037

60 0.806 0.622 0.481 0.371 0.286 0.221 0.171 0.132 0.102 0.079 0.061 0.047 0.036

66 0.789 0.609 0.470 0.363 0.280 0.216 0.167 0.129 0.100 0.077 0.059 0.046 0.035

24 0.953 0.785 0.646 0.533 0.439 0.361 0.298 0.245 0.202 0.166 0.137 0.113 0.093 0.077

36 0.930 0.766 0.631 0.520 0.428 0.353 0.291 0.239 0.197 0.162 0.134 0.110 0.091 0.075

48 0.908 0.748 0.616 0.507 0.418 0.344 0.284 0.234 0.192 0.159 0.131 0.108 0.089 0.073

60 0.886 0.730 0.601 0.495 0.408 0.336 0.277 0.228 0.188 0.155 0.127 0.105 0.086 0.071

72 0.865 0.712 0.587 0.483 0.398 0.328 0.270 0.223 0.183 0.151 0.124 0.102 0.084 0.070

84 0.844 0.695 0.573 0.472 0.389 0.320 0.264 0.217 0.179 0.147 0.121 0.100 0.082 0.068

Half-life: 87.4 days

0 1 2 3 4 5 6 7 8 9 10 11 12

Days 0 1.000 0.946 0.895 0.847 0.801 0.758 0.717 0.678 0.641 0.607 0.574 0.543 0.514

6 0.849 0.646 0.492 0.374 0.285 0.217 0.165 0.126 0.096 0.073 0.055 0.042 0.032

7 0.826 0.629 0.479 0.364 0.277 0.211 0.161 0.122 0.093 0.071 0.054 0.041 0.031

Weeks

Days

32P

Half-life: 60.0 days

Days

Days

125I

1 0.992 0.939 0.888 0.840 0.795 0.752 0.711 0.673 0.636 0.602 0.569 0.539 0.510

2 0.984 0.931 0.881 0.833 0.788 0.746 0.705 0.667 0.631 0.597 0.565 0.534 0.506

3 0.976 0.924 0.874 0.827 0.782 0.740 0.700 0.662 0.626 0.592 0.560 0.530 0.502

4 0.969 0.916 0.867 0.820 0.776 0.734 0.694 0.657 0.621 0.588 0.556 0.526 0.498

5 0.961 0.909 0.860 0.814 0.770 0.728 0.689 0.652 0.616 0.583 0.552 0.522 0.494

6 0.954 0.902 0.853 0.807 0.764 0.722 0.683 0.646 0.612 0.579 0.547 0.518 0.490

Half-life: 25.4 days 0 10 20 30 40 50 60 70 80 90 100 110 120

Days 0 1.000 0.761 0.579 0.441 0.336 0.256 0.195 0.148 0.113 0.086 0.065 0.050 0.038

1 0.973 0.741 0.564 0.429 0.327 0.249 0.189 0.144 0.110 0.084 0.064 0.048 0.037

2 0.947 0.721 0.549 0.418 0.318 0.242 0.184 0.140 0.107 0.081 0.062 0.047 0.036

3 0.921 0.701 0.534 0.406 0.309 0.236 0.179 0.136 0.104 0.079 0.060 0.046 0.035

4 0.897 0.683 0.520 0.395 0.301 0.229 0.174 0.133 0.101 0.077 0.059 0.045 0.034

5 0.872 0.664 0.506 0.385 0.293 0.223 0.170 0.129 0.098 0.075 0.057 0.043 0.033

8 0.804 0.612 0.466 0.355 0.270 0.205 0.156 0.119 0.091 0.069 0.053 0.040 0.030

9 0.782 0.595 0.453 0.345 0.263 0.200 0.152 0.116 0.088 0.067 0.051 0.039 0.030

Number of half-lives elapsed 6

1

5

2

3

3

4

4

5

6

7

8 9 10

1

2

15

20

25

30

40

50

60 70 80 90 100

Percent of radioactivity remaining Figure A.1A.1 Correlation of loss of radioactivity with elapsing half-lives of an isotope. Useful Information



ily in water and is absorbed by activated charcoal or copper. The amount of this volatile radioactive component released, despite stabilizers added by the manufacturers, is about 1/8000 of the total radioactivity present. The amount of this radioactivity that a scientist is likely to inhale while using these compounds is presumably even smaller. Nevertheless, such a component can potentially contaminate a wide area because of its volatility, and also tends to concentrate in target organs. Thus, it is advisable to thaw vials of 35S-labeled amino acids in a controlled area such as a hood equipped with a charcoal filter. This charcoal filter will become quite contaminated and should be changed every few months. If such an area is not available, the stock vial should be thawed using a needle attached to a charcoal-packed syringe to vent and trap the volatile compound. Anyone who has ever added 35S-labeled amino acids to dishes of cells for even short periods knows that the incubator(s) used for such labelings quickly becomes highly contaminated with 35S. Such contamination is not limited to the dish itself, nor to the shelf on which the dish was placed. Rather, the radioactive component’s solubility in water allows it to circulate throughout the moist atmosphere of the incubator and contaminate all the inside surfaces of the incubator. For this reason, in laboratories where such metabolic labelings are routine, it is highly convenient to designate one incubator to be used solely for working with 35S-labeled samples. Such an incubator can be fitted with a large honeycomb-style filter, the size of an incubator shelf, made of pressed, activated charcoal. These filters are available from local air-quality-control companies. Such a filter will quickly become quite contaminated with radioactivity and should therefore be monitored and changed as necessary (usually about every three months if the incubator is used several times a week). The water used to humidify the incubator will also become quite “hot” (contaminated with radioactivity); keeping the water in a shallow glass pan on the bottom of the incubator makes it easy to change it after every use, thus preventing contamination from accumulating. Even with the charcoal filter and water as absorbents, the shelves, fan, and inner glass door of the incubator will become contaminated, as will the tray on which the cells are carried and incubated. Routine wipe tests and cleaning when necessary will help to minimize potential spread of this contamination.

If such labelings are done infrequently or there is no “spare” incubator, dishes of cells can be placed in a box during incubation. This box should be made of plastic, which is generally more easily decontaminated than metal. Along with the dishes of cells, a small sachet made of activated charcoal wrapped loosely in tissue (Kimwipes work well) should be placed in the box. If the box is sealed, it will obviously need to be gassed with the correct mixture of CO2; otherwise small holes can be incorporated into the box design to allow equilibration with the incubator’s atmosphere. In either case, the incubator used for the labeling should be carefully monitored for radioactivity after each experiment.

Working with 32P Microcurie amounts of 32P The amount of 32P-labeled nucleotide used to label nucleic acid probes for northern or Southern blotting is typically under 250 µCi, and the amount of [γ-32P]ATP used for in vitro phosphorylation of proteins does not usually exceed 50 µCi for a single kinase reaction (or several hundred microcuries per experiment). However, handling even these small amounts, given the time spent on such experiments, can result in an unacceptable level of exposure if proper shielding is not employed. With no intervening shielding, the dose rate 1 cm away from 1 mCi 32P is 200,000 mrads/hr; the local dose rate to basal cells resulting from a skin contamination of 1 µCi/cm2 is 9200 mrads/hr (Shleien, 1987). Such a skin contamination could be easily attained though careless pipetting and the resultant creation of an aerosol of radioactive microdroplets, because the concentration of a typical stock solution of labeled nucleotide may be 10 µCi/µl. For proper protection during this sort of experiment, besides the usual personal attire (glasses, coat, safety badges, and gloves) it is necessary to use some form of Plexiglas screen between the body and the samples (see Fig. A.1A.2A). Check the level of radiation coming through the outside of the shield with a portable monitor to be sure the thickness of the Plexiglas is adequate. Hands can be shielded from some exposure by placing the sample tubes in a solid Plexiglas rack, which is also useful for transporting samples from the bench to a centrifuge or water bath (see Fig. A.1A.2B). Experiments of these types often include an incubation step performed at a specific temperature, usually in a water bath. Although the water


surrounding the tubes or hybridization bags will effectively stop β radiation, shielding should be added over the top of the tubes (where there is no water)—e.g., a simple flat piece of Plexiglas. If the frequency of usage justifies the expense, an entire lid for the water bath can be constructed from Plexiglas. When hybridization reactions are performed in bags, care should be taken to monitor (and shield) the apparatus used to heat-seal the bags. The waste generated during the experiments should also be shielded. It is convenient to have a temporary waste container right on the bench. Discard pipet tips and other solid waste into a beaker lined with a plastic bag and placed behind the shield. This bag can then be emptied into the appropriate shielded laboratory waste

A

container when the experiment is done. Liquid waste can be pipetted into a disposable tube set in a stable rack behind the shield (see Fig. A.1A.2C). When radiolabeled probes or proteins must be gel-purified, it may be necessary to shield the gel apparatus during electrophoresis if the samples are particularly hot. Be advised that the electrophoresis buffer is likely to become very radioactive if the unincorporated label is allowed to run off the bottom of the gel; check with a radiation safety officer for instructions on how to dispose of such buffer. It is also prudent to check the gel plates with a portable detection monitor after the electrophoresis is completed, because they sometimes become contaminated as well.

12 in. (300 mm)

12-14 in. (300-350 mm)

18-24 in. (450-600 mm)

14 in. (350 mm)

12-15 in. (300-375 mm) L shield

T shield

B

3 in. (75 mm)

12 in. (300 mm) rack for microcentrifuge tubes

C 3.5 in. (90.5 mm)

tube holder for liquid waste

Figure A.1A.2 Plexiglas shielding for 32P. (A) Two portable shields (L and T design) made of 0.5-in. Plexiglas. Either can be used to directly shield the scientist from the radioactivity he or she is using. Turned on its side, the L-shaped shield can be used to construct two sides of a cage around a temporary work area, providing shielding for other workers directly across or to the sides of the person working with 32P. (B) Tube rack for samples in microcentrifuge tubes. (C) Tube holder for liquid waste collection.

Useful Information


Millicurie amounts of 32P In order to study protein phosphorylation in intact mammalian cells, cells in tissue culture dishes are incubated in phosphate-free medium with 32P-labeled orthophosphate for a period of several hours or overnight to label the proteins. The amount of 32P used in such labelings can be substantial. Cells are normally incubated in 1 to 2 mCi of 32P/ml labeling medium; for each 6-cm dish of cells, 2.5 to 5 mCi 32P may be used. When this figure is multiplied by the number of dishes necessary per sample, and the number of different samples in each experiment, it is clear that the amount of 32P used in one experiment can easily reach 25 mCi or more. Because so much radioactivity is used in the initial labeling phase of such experiments, it is necessary for a researcher to take extra precautions in order to adequately shield him or herself and coworkers.

A

When adding label to dishes of cells, it is important to work as rapidly as possible. An important contribution to the speed of these manipulations is to have everything that will be needed at hand before even introducing the label into the work area. Prepare the work area, arranging shielding and covering the bench with diapers, in advance. Set out all necessary items, including any pipettors and tips needed, a portable detection monitor, extra gloves, and a cell house (see Fig. A.1A.3A). Work involving this much radioactivity should be done behind a Plexiglas shield at least 3⁄ in. (2 cm) thick; the addition of a layer of lead 4 to the outside lower section of this shield to stop Bremsstrahlung radiation is also needed. If one shield can be dedicated to this purpose at a specific location, a sheet of lead several centimeters thick can be permanently screwed to the

door 2 in. (5 cm)

3.5 in. (9 cm) 7.5 in. (19 cm) cell house

B

Plexiglas 0.75-1 in. (20-25 mm) thick

lead plate 0.25 in. (4-6 mm) thick

Plexiglas 0.5 in. (12.5 mm) thick

leaded shield

C ID = 5.75 in. (140 mm) 1.25 in. (45 mm)

ID = 2.5 in. (65mm)

5.5 in. (135 mm) square block

box made of 0.5 in. (12.5 mm) thick Plexiglas rack and storage box


Figure A.1A.3 (A) Box for cell incubation (a “cell house”). (B) Stationary leaded shield. (C) Sample storage rack and box made of 0.5-in. Plexiglas. Abbreviations: ID, interior dimension.


Plexiglas (as shown in Fig. A.1A.3B); however, this lead makes the shield extremely heavy and therefore less than portable. If space constraints do not permit the existence of such a permanent labeling station, a layer or two of lead foil can be taped temporarily to the outside of the Plexiglas shield. Again, each worker should take care to shield not only him or herself, but bystanders on all sides. Handling of label should be done away from the central laboratory if possible to take maximum advantage of distance as an additional form of shielding. It is also advisable not to perform such experiments in a tissue culture room or any other room that is designed for a purpose vital to the whole laboratory. An accident involving this much 32P would seriously inconvenience future work in the area, if not make it altogether uninhabitable! If care is taken to minimize the amount of time the dish of cells is open when adding the label, use of a controlled air hood to prevent fungal or bacterial contamination of the cells should not be necessary. Once the label has been added to the dishes of cells, they will also need to be shielded for transport to and from the incubator and other work areas. Plexiglas boxes that are open at one end (for insertion of the dishes) and have a handle on top (for safe carrying) make ideal “cell houses” (see Figure A.1A.3A). A Plexiglas door that slides into grooves at the open end is important to prevent dishes from sliding out if the box is tilted at all during transport. If this door is only two-thirds the height of the house wall, the open slot thus created will allow equilibration of the CO2 level within the house with that in the incubator. Obviously, this slot will also allow a substantial stream of radiation to pass

out of the cell house, so the house should be carried and placed in the incubator with its door facing away from the worker (and others)! Following incubation with label and any treatments or other experimental manipulations, the cells are usually lysed in some type of detergent buffer. It is during this lysis procedure that a worker’s hands receive their greatest exposure to radiation, because it is necessary to directly handle the dishes over a period of several minutes. It is therefore very important to streamline this procedure and use any shielding whenever possible. If the cell lysates must be made at 4°C, as required by most protocols, working on a bench in a cold room is preferable to placing the dishes on a slippery bed of ice. In either case, make the lysate using the same sort of shielding (with lead if necessary) that was used when initially adding the label. Pipet the labeling medium and any solution used to rinse unincorporated radioactivity from the cells into a small tube held in a solid Plexiglas holder (shown in Fig. A.1A.2C). The contents of this tube can later be poured into the appropriate liquid waste receptacle. If possible, it is a good practice to keep this high-specific-activity 32P liquid waste separate from the lower-activity waste generated in other procedures so that it can be removed from the laboratory as soon as possible following the experiment. If it is necessary to store it in the laboratory for any time, the shielding for the waste container should also include a layer of lead. The solid waste generated in the lysis part of these experiments (pipet tips, disposable pipets, cell scrapers, and dishes) is very hot and should be placed immediately into some sort of shielded container to avoid further exposure of

ID = 5 in. (125 mm)

ID = 6.75 in. (175 mm)

ID = 10.75 in. (275 mm) box for solid waste

Figure A.1A.4 Box for solid waste collection made of 0.5-in. Plexiglas. Abbreviation: ID, interior dimension.

Useful Information



the hands. A Plexiglas box similar in design to that in Figure A.1A.4 is convenient; placed to the side of the shield and lined with a plastic bag, it will safely hold all radioactive waste during the experiment and is light enough to be easily carried to the main laboratory waste container where the plastic bag (and its contents) can be dumped after the experiment is done. If the lid of the box protrudes an inch or so over the front wall, it can be lifted using the back of a hand, thus decreasing the possibility of contaminating it with hot gloves. When scraping the cell lysates from the dishes, it is good practice to add them to microcentrifuge tubes that are shielded in a solid Plexiglas rack; this will help to further reduce the exposure to which the hands are subjected. At this point, the lysates are usually centrifuged at high speed (>10,000 × g) to clear them of unsolubilized cell material. Use screw-cap tubes for this clarification step, as these will contain the labeled lysate more securely than flip-top tubes, which may open during centrifugation. No matter what type of tube is used, the rotor of the centrifuge often becomes contaminated, most probably due to tiny drops of lysate (aerosol) initially present on the rim of the tubes that are spun off during centrifugation. Monitor the rotor and wipe it out after each use. The amount of 32P taken up by cells during the incubation period varies considerably, depending on the growth state of the culture as well as on the cell type and its sensitivity to radiation. This makes it difficult to predict the percentage of the radioactivity initially added to the cells that is incorporated into the cell lysate; however, this figure probably does not exceed 10%. Thus, the amount of radioactivity being handled decreases dramatically after lysis, making effective shielding much simpler. However, at least ten times more radioactivity than is usual in other sorts of experiments is still involved! It is easy to determine if the shielding is adequate— just use both β and γ portable monitors to measure the radiation coming through it. Again, be sure to check that people working nearby (including those across the bench) are also adequately shielded. It is sometimes necessary to construct a sort of cage of Plexiglas shields around the ice bucket that contains the lysates. At the end of the day or the experiment, it may be necessary to store radioactive samples; in some experiments, it may be desirable to save the cell lysates. These very hot samples are best stored in tubes placed in solid Plexiglas racks that can then be put into Plexiglas boxes (see Fig. A.1A.3C). Such boxes may be of similar

construction to the cell houses described above; however, they should have a door that completely covers the opening. Be sure to check the γ radiation coming through these layers and add lead outside if necessary.

Working with 33P Using 33P-labeled nucleotides to label nucleic acid probes or proteins Several of the major companies that manufacture radiolabeled biological molecules have recently introduced nucleotides labeled with 33P (both α- and γ-labeled forms). 33P offers a clear advantage over 32P with respect to ease of handling, because the energy of the β particles it releases lies between that of 35S and 32P and thus its use does not require as many layers of Plexiglas and lead shielding as for 32P. In fact, the β radiation emitted can barely penetrate through gloves and the surface layer of skin, so the hazard associated with exposure to even millicurie amounts of 33P is thought to be insignificant (as reported in the DuPont NEN product brochure). Gel bands visualized on autoradiographs of 33P-labeled compounds are sharper than bands labeled with 32P because the lower-energy β radiation does not have the scatter associated with that from 32P. The halflife of 33P is also longer (25 days compared to 14 days for 32P). Despite its higher cost, these features have led many researchers to choose 33P-labeled nucleotides for use in experiments such as band/gel shift assays where discrimination of closely-spaced gel bands is important. The best way to determine what degree of shielding is needed when using 33P is to monitor the source using a portable β monitor and add layers of Plexiglas as necessary.

Working with 125I Using 125I to detect immune complexes (immunoblots) 125I that is covalently attached to a molecule such as staphylococcal protein A is not volatile and therefore is much less hazardous than the unbound or free form. Most institutions do not insist that work with bound 125I be performed in a hood, but shielding of the γ radiation is still necessary. Lead is a good high-density material for stopping these γ rays; its drawbacks are its weight and opacity. Commercially available shields for 125I are made of lead-impregnated Plexiglas—though heavy, these are at least seethrough. Alternatively, a piece of lead foil may be taped to a structural support, although this


arrangement does not provide shielding for the head as a worker peers over the lead! Incubations of the membrane or blot with the [125I] protein A solution and subsequent washes are usually done on a shaker. For shielding during these steps, a piece of lead foil may simply be wrapped around the container. Solutions of 125I can be conveniently stored for repeated use in a rack placed in a lead box. Using 125I to label proteins or peptides in vitro Any experiments that call for the use of free, unbound 125I should be done behind a shield in a hood that contains a charcoal filter to absorb the volatile iodine. Most institutions require that such experiments be done in a special hot lab to which access is limited. Ingested or inhaled iodine is concentrated in the thyroid; a portable γ monitor should therefore be used to scan the neck and throat before beginning and after completing each experiment. Similar scans should routinely be performed on all members of any laboratory in which unbound iodine is used.

DEALING WITH ACCIDENTS Despite the best intentions and utmost caution, accidents happen! Accidents involving spills of radioactivity are particularly insidious because they can be virtually undetectable yet pose a significant threat to laboratory workers. For this reason it is best to foster a community spirit in any laboratory where radioisotopes are routinely used—a sense of cooperativity that extends from shielding each other properly to helping each other clean up when such accidents occur. The specific measures to be taken following an accident involving radioactivity naturally depend on the type and amount of the isotope involved, the chemical or biological hazards of the material it is associated with, and the physical parameters of the spill (i.e., where and onto what the isotope was “misplaced”). However, following any accident there are several immediate steps that should be taken: 1. Alert coworkers as well as Radiation Safety personnel to the fact that there has been an accident. This will give them the opportunity to shield themselves if necessary—and to help clean up as well! 2. Restrict access to and away from the site of the accident to ensure that any uncontained

radioactive material is not spread around the laboratory. When leaving the site be sure to monitor the bottoms of the shoes as well as the rest of the body. 3. Take care of all contaminated personnel first, evacuating others if necessary. If anyone’s skin is contaminated, first use a portable monitor to identify specific areas of contamination. Then wipe these areas with a damp tissue to remove as much surface radioactivity as possible. Try to scrub only small areas at a time to keep the contamination localized. If the contamination is not easily removed with paper tissues, try a sponge or an abrasive pad, but be careful not to break the skin! Sometimes soaking is required: do this only after all easily removed contamination is gone and keep the soaked area to a minimum. Contaminated strands of hair can be washed (or perhaps a new hairstyle may be in order). 4. When attempting to clean any contaminated equipment, floors, benches, etc., begin by soaking up any visible radioactive liquid with an absorbent material. Use a small amount of soap and water to clean the contaminated area, keeping the area wiped each time to a minimum to avoid smearing the contamination over an even greater surface. Many surfaces prove resistant to even Herculean cleaning efforts; in these instances the best that can be done is to remove all contamination possible and then shield whatever is left until the radioactivity decays sufficiently for safety.

LITERATURE CITED Dawson, R.M.C., Elliot, D.C., Elliott, W.H., and Jones, K.M. (eds.) 1986. Data for Biochemical Research. Alden Press, London. Lederer, C.M., Hollander, J.M., and Perlman, I. (eds.) 1967. Table of Radioisotopes, 6th edition. John Wiley and Sons, New York. Meisenhelder, J. and Hunter, T. 1988. Radioactive protein-labelling techniques. Nature 335:120. Shleien, B. (ed.) 1987. Radiation Safety Manual for Users of Radioisotopes in Research and Academic Institutions. Nucleon Lectern Associates, Olney, Md.

Contributed by Jill Meisenhelder and Kentaro Semba The Salk Institute La Jolla, California

Useful Information


Transgenic and Gene-Targeted Mouse Lines for Toxicology Studies The isolation of mammalian genes is of utmost importance to toxicology because of the contributions these studies can make to the understanding of normal physiology and development as well as chemical-induced perturbations of normal biochemical processes. The development of methods that allow functional expression of specific genes, including human genes, in experimental animal models provides exciting and novel avenues for investigation into gene-environment interactions. Likewise, the ability to delete the function of a normal gene in living animals (e.g., “knockout” mice) also provides a unique approach to studying gene function. Methods for the direct microinjection of DNA into pronuclei of fertilized embryos are well established (see UNIT 1.3). Foreign genes can be incorporated into somatic germ-line tissues, with expression of these elements in the progeny of founder mice. An additional gene-transfer technology in mice involves the use of stem cells from the early embryo, so-called embryonic stem (ES) cells. The capacity of ES cells to undergo differentiation makes them useful for investigating the effects of genetic modifications of either gain or loss of function. These pluripotent genetically modified ES cells can then be targeted with specific genes and vectors aimed at creating point mutations or null mutations. During the past several years, a large number of genetically engineered mouse models have been described and utilized for toxicology stud-

APPENDIX 1B

ies, including carcinogenicity testing, mutation analysis, chemical bioassays and risk assessment, gene action and the environment, and environmental stress response mechanisms. Table A.1B.1 briefly describes a selected number of transgenic and gene-targeted mouse models that have been used, or may be of interest, in investigations and testing relevant to the field of toxicology. The table is designed as an introductory guide and not an exhaustive and conclusive reference source. A number of the models are available from the Induced Mutant Mouse Resource at The Jackson Laboratory (http://www.jax.org), while others are available commercially from various animal vendors. Many, however, are only available from the individuals who originally described them. Attempts to increase the number and capacity of mutant mouse resource centers may provide the means of making more of these genetically altered mouse lines available to the toxicological community (Battey et al., 1999).

LITERATURE CITED Battey, J., Jordon, E., Cox, D., and Dove, W. 1999. An action plan for mouse genomics. Nature Genet. 21:73-75.

Contributed by George Sanders, Carol Ware, and Warren Ladiges University of Washington Seattle, Washington

Useful Information Contributed by George Sanders, Carol Ware, and Warren Ladiges Current Protocols in Toxicology (1999) A.1B.1-A.1B.11 Copyright © 1999 by John Wiley & Sons, Inc.

A.1B.1 Supplement 1

A.1B.2

Supplement 1


Tm

Tm

Tg

Apc716 Adenoma polyposis coli

Apc1638 Adenoma polyposis coli

Atase Alkylguanine-DNA alkyltransferase Atase Alkylguanine-DNA alkyltransferase CD2 promoter Tg

Chemical mutant

Tm

Tm

Tg

Mutant typea

Apc Adenoma polyposis coli

Alox Arachidonate 5-lipoxygenase

Ahr Aryl-hydrocarbon receptor AhR

AGT Angiotensinogen

Gene symbol and name

Description

More severe phenotype than apc or apc 1638. Multiple colonic polyps and gastrointestinal neoplasia. Model for evaluating the role of gene in intestinal tumorgenesis. Less severe phenotype than apc or apc716. Multiple colonic polyps and gastrointestinal neoplasia. Model for evaluating role of gene in intestinal tumorgenesis. Hepatic expression. Model for evaluating role of gene in adverse biological effects of alkylating agents. Hepatic expression prevents N-methyl-N-nitrosourea–induced thymic lymphomas. Model for evaluating mechanisms involved in carcinogenesis.

Causes multiple intestinal neoplasia. Model for studying carcinogen-induced neoplasia in transgenic mice.

Exhibits selective opposition to certain inflammatory insults. Model for studying importance of leukotrienes.

Hepatic fibrosis and immune system impairment. Model for studying development of liver, immune system, and expression of dioxin-induced genes, as well as AhR activation in genital tubercle, palate, and other embryonic tissues. Model for evaluating endogenous role of AhR in proliferation or differentiation and developmental targets of dioxin-like compounds.

Expression in liver, kidney, heart, adrenal gland, ovary, brain, and white and brown fat. Exogenous renin administration yields transient increase in blood pressure. Potential human hypertension model.

Table A.1B.1 Transgenic and Gene-Targeted Mice for Toxicology Studies

— —

Dumenco et al., 1993. Science 259:219

—

—

JL

JL

JL

JL

Available fromb

Fan et al., 1990. NAR 18:5723

Fodde et al., 1994. PNAS 91:8969

Moser et al., 1990. Science 247:322; Su et al., 1992. Science 256:668 [Erratum, Science 256:1114] Oshima et al., 1996. Cell 87:803

Chen et al., 1994. Nature 372:179

Fernandez-Salguero et al., 1995. Science 268:722; Schmidt et al., 1996. PNAS 93:6731; Willey et al., 1998. Toxicol. Appl. Pharmacol. 151:33

Yang et al., 1994. JBC 269:32497

Reference(s)

A.1B.3


Supplement 1

cat Catalase

brca2 Breast cancer susceptibility gene 2

Causes embryonic developmental arrest at day 6.5 and hypersensitivity to γ irradiation. Model for evaluation of role of gene in DNA repair and tumor suppression. Increased catalase activity attenuates adriamycin-induced cardiotoxicity.

Tm

Tg

Tm

brca1 Breast cancer susceptibility gene 1

Embryonic lethality prior to gastrulation on day 7.5. Evaluation of role of gene in cellular proliferation during embryogenesis, cell cycle progression regulation, and function during oncogenesis.

Tg

Tg

bcl-2 HHK1 Human keratin 1 (Hk-1)

Tg

bcl-2 B cell leukemia lymphoma bcl-2-Ig NL bcl-2-Ig SM bcl-2 H2K-BCL-2 Tg

Overexpression in cells of hematolymphoid system. Model for increasing resistance of animals to apoptosis-inducing challenges like irradiation. Overexpression in cells of the nervous system, protects neuronal cells from ischemia-induced death and causes neuronal hypertrophy. Model for evaluating role of gene in developing nervous system and neurodegenerative disorders. Epidermal expression yields multifocal hyperplasia with keratinocytes more resistant to cell death from radiation and chemical carcinogen exposure. Model for evaluating pathogenesis of nonmelanoma skin cancers.

Tg

bcl-2 2-bcl-2

bcl-2 NSE-Hu-BCL-2

Causes growth retardation, neurologic dysfunction, infertility, immunologic abnormalities, lymphoreticular malignancies, chromosomal instability, and extreme sensitivity to ionizing radiation. Model for evaluating role of gene in normal cellular function and as a model for ataxia-telangiectasia. Expression of high levels of protein in thymocytes and peripheral T cells. Model for evaluating variable resistance to chemotherapy regimens. Extended B cell survival with follicular lymphoproliferation. Model for studying pathogenesis of B lymphoid neoplasms.

Tm

Atm Ataxia telangiectasia, mutated

Kang et al., 1996. JBC 271:12610

continued

—

—

—

Hakem et al., 1996. Cell 85:1009

Sharan et al., 1997. Nature 386:804

—

Rodriguez-Villanueva et al., 1998. Oncogene 16:853

—

Martinou et al., 1994. Neuron 13:1017

—

McDonnell et al., 1989. Cell 57:79

—

—

Siegel et al., 1992. PNAS 89:7003

Domen et al., 1998. Blood 91:2272

JL

Barlow et al., 1996. Cell 86:159; Xu et al., 1996. Genes Dev. 10:2411

A.1B.4

Supplement 1


Tm

Tg

COX-2 Cyclooxygenase-2

CRP C-reactive protein cyD1 Cyclin D1 EBVED-L2 cyD1

Cyp Cytochrome P-450 CYP2B2-19 CYP2B2-39

Tm

Tm

Tg

Negative and positive regulation of transgene by phenobarbital administration. Phenobarbital induction mechanism system model.

Exposure to an aromatic hydrocarbon increases hepatic expression of transgene, which may be gender-related. Model for evaluating the gender-related expression of the gene. Neonatal lethality via respiratory distress. Potential neonatal respiratory distress syndrome models.

Null mutation yields a decreased incidence of indomethacin-induced gastric ulceration, reduced platelet aggregation, and decreased response to arachidonic acid. Model for the evaluation of physiological roles of COX-1 and COX-2. Targeted disruption yields severe nephropathy and exhibits increased susceptibility to peritonitis. Models for COX isoform evaluation and improvement of NSAIDs, mitigation of endotoxin-induced hepatocellular cytotoxicity, and evaluation of COX-2 in early pregnancy. Inflammation-induced liver-specific expression. Model to study liver development and regeneration. Epithelial dysplasia. Model for evaluating associated malignancies.

Tm

COX-1 Cyclooxygenase-1 Ptgs1

cyp Cytochrome P-450 1a-1 Cyp 1a-1-CAT Cyp Cytochrome P-450 1a-1, 1a-2

Overexpression in large intestine, cecum, and stomach. Model for antigen-specific vaccine studies, antibody-targeting studies, and other forms of immunotherapy.

Tg

CEA TgN(CEAGe)18FJP

Tg

Epithelial spatiotemporal expression pattern similar to humans. Model for tumor immunotherapy.

Description

Tg

Mutant typea

CEA Carcinoembryonic antigen gene cosCEA1


Table A.1B.1 Transgenic and Gene-Targeted Mice for Toxicology Studies, continued

—

Dinchuk et al., 1995. Nature 378:406; Lim et al., 1997. Cell 91:197

JL

—

Ramsden et al., 1993. JBC 268:21722

—

—

Pineau et al., 1995. PNAS 92:5134; Diliberto et al., 1997. Biochem. Biophys. Res. Commun. 236:431

Jones et al., 1991. NAR 19:6547

Mueller et al., 1997. Cancer Res. 57:5542

—

—

Langenbach et al., 1995. Cell 83:483

Ciliberto et al., 1987. EMBO J. 6:4017

—

—

Available fromb

Clarke et al., 1998. Cancer Res. 58:1469

Eades-Perner et al., 1994. Cancer Res. 54:4169

Reference(s)

A.1B.5


Supplement 1

Epidermal hyperplasia and decreased hair growth. Role in epithelial growth and tumorgenesis. Model for evaluating role in apoptosis and cell proliferation suppression. Higher numbers of T cells in the thymus, lymph node, and spleen with increased numbers of T cells resistant to Fas-mediated apoptosis. Model for evaluating Fas-mediated apoptosis. Expression yields early abnormal bone development. Model for studying pathways responsible for specificity of induced tumorgenesis. Gene deficiency yields growth retardation, lethal diffuse colitis, alteration in thymocyte maturation and function, plus colonic adenocarcinoma. Model for ulcerative colitis and colonic carcinogenesis. Increased activity confers increased tolerance to ischemia-reperfusion injury, H2O2-induced DNA-strand breakage in lens epithelial cells, and paraquat toxicity. Model for brain hydroperoxide metabolism and oxidative stress homeostasis. Model for evaluating thermoregulation and processes involving actions of hydroxy and lipid peroxidases.

Tg Tm

E2F1 K5-E2F1

E2F1 Transcriptional activator gene fasL fas ligand fasL/TCR-β Fas ligand

Null mutant mice are more susceptible to acute paraquat toxicity and myocardial ischemia-reperfusion injury. Renal convoluted tubular epithelium expression. Model for in vivo study of GST modulation in carcinogenesis and drug toxicity.

Tg

Tm Tg

GPx-1 Glutathione peroxidase

GST mu Mu-class glutathione transferase

Tm

Tg

GPx-1 Glutathione peroxidase

fos MT-c-fos-209-4 MT-c-fos-211-5 GαI2 G-protein subunit that functions as tumor-suppressor gene

Liver, kidney, lung, spleen, testis, small intestine, thymus, brain, skin, and heart expression. Model for studying the toxicological significance of P-450s during embryonic development.

Tm

Cyp Cytochrome P-450 MT-1-CYP3A7

Tg

Modulation of tissue expression post male castration and post testosterone/rat growth hormone treatment in female. Model for evaluating transcriptional regulation of gene.

Tm

Cyp Cytochrome P-450 Cyp2C2-luc

—

Connelly et al., 1993. Pathobiology 61:7

continued

—

Cheng et al., 1998. J. Nutr. 128:1070

—

— Rudolph et al., 1995. Nature Genet. 10:143

Mirault et al., 1994. Ann. N.Y. Acad. Sci. 738:104; Mirochnitchenko et al., 1995. PNAS 92:8120; Yoshida et al., 1996. J. Mol. Cell. Cardiol. 28:1759; Reddy et al., 1997. Ophthalmologica 211:192; Cheng et al., 1998. J. Nutr. 128:1070

—

—

JL

—

—

—

Ruther et al., 1987. Nature 325:412

Field et al., 1996. Cell 85:549; Yamasaki et al., 1996. Cell 85:537 Cheng et al., 1997. J. Immunol. 159:674

Pierce et al., 1998. Oncogene 16:1267

Li et al., 1996b. Arch. Biochem. Biophys. 329:235

Li et al., 1996a. Mol. Cell Endocrinol. 120:77

A.1B.6

Supplement 1


Increased incidence of epithelial tumorgenesis after carcinogen exposure. Role as a determinant of carcinogen exposure and cancer susceptibility. Protein overexpression in neurons of various brain regions. Model for evaluating the function of carbon monoxide as a neuromodulator, iron as a gene regulator, and bile pigments as in vivo antioxidants. Model for evaluation of role of carbon monoxide as retrograde neuronal messenger. Increased myocardial expression demonstrates cardioprotective effect against ischemia. Model for evaluating cardiac post-ischemic recovery. Impairment of immune and acute-phase inflammatory responses. Model for studying response to injury or infection. Polyclonal plasmacytosis of thymus, lymph node, spleen, lung, liver, and kidney. Useful in determining role of gene in polyclonal and monoclonal plasma cell abnormalities. Tongue, esophagus, and forestomach epithelial dysplasia. Model of upper-digestive tract tumor initiation.

Tm

Tg

Tm Tg

Tm

Tg

GST pi Pi-class glutathione transferase

HO-1 Heme oxygenase

HO-2 Heme oxygenase

hsp-70 Heat-shock protein

IL-6 Interleukin 6

IL-6 Interleukin 6 Eµ-IL-6 L2hD

JL

Suematsu et al., 1989. PNAS 86:7547

Increased total cholesterol and HDL cholesterol levels. Model of hyperalphalipoprotenemia.

Specific expression in endothelial cells of vessel walls and Harats et al., 1995. J. Clin. Invest. 95:1335 pulmonary epithelium. System for direct expression of molecules by vascular wall to evaluate select proteins.

Tm

lipox Lipoxygenase

Vaisman et al., 1995. JBC 270:12269

Gossen et al., 1989. PNAS 86:7971

Kohler et al., 1991. PNAS 88:7958

Tg

Plasmid rescue from genome. Used in induced mutation assays.

Tm

LacZ β-galactosidase λ shuttle vector

—

—

—

JL

—

—

—

Marber et al., 1995. J. Clin. Invest. 95:1446; Radford et al., 1996. PNAS 93:2339 Kopf et al., 1994. Nature 368:339

Nakagawa et al., 1997. Oncogene 14:1185

—

—

—

Available fromb

Poss et al., 1995. Neuron 15:867

Maines et al., 1998. J. Neurochem. 70:2057

Henderson et al., 1998. PNAS 95:5275

Reference(s)

LCAT Lecithin cholesterol acyltransferase

Plasmid rescue from genome. Used in induced mutation assays.

Tm

LacI β-galactosidase λ shuttle vector

Tg

Description

Mutant typea



A.1B.7


Supplement 1

Overexpression in bone marrow confers resistance to cytotoxic drug–induced leukopenia. Model for evaluating MDR1-dependent multidrug resistance and high- dose chemotherapy regimens.

Tg

mdr-1a Multidrug resistance 1 β-actin/MDR1

JL, CR

Overexpression yields B cell lymphomas or no increase in Adams et al., 1985. Nature 318:533 tumor incidence. Models for evaluating lymphoma development, B cell ontogeny, and immunoglobulin regulation. Tg

myc (c-myc) Proto-oncogene B cell promoters

continued

—

Deficiency yields increased incidence of lymphoid tumors Reitmair et al., 1995. Nature Genet. 11:64 with microsatellite instabilities in 2-month-old mice. Model for evaluating progression of tumors and screening carcinogenic and anticancer agents.

Tm

MSH2 DNA mismatch repair gene

—

Baker et al., 1995. Cell 82:309

Null mutation yields microsatellite instability in several tissues, increased incidence of sarcomas and lymphomas, and male infertility. Model for evaluation of role in DNA repair, genetic recombination, and meiotic chromosome synapsis.

JL

Becker et al., 1996. Cancer Res. 56:3244

Tm

—

TF

—

—

—

Walter et al., 1993. Carcinogenesis 14:1537

Galski et al., 1989. Mol. Cell Bio. 9:4357

Schinkel et al., 1994. Cell 77:491

Andersen et al., 1992. Ann. N.Y. Acad. Sci. 648:241

Cases et al., 1995. Science 268:1763

MS2 DNA mismatch repair gene

Tm

Overexpressed in liver and brain. Possible use in evaluation of cellular resistance to DNA-alkylation toxicity and tumor induction. Significant reduction in skin tumor incidence post topical carcinogen application. Model for studying role in prevention of tumor initiation.

Deficiency of the blood-brain barrier with subsequent sensitivity to drugs. Model for compartmental drug accumulation.

Tm

Mdr-1a Multidrug resistance 1

Tg

Neuronal-specific overexpression. Model for evaluating effects of high MAO-B levels on development and possible role in neurologic disease.

Tg

MAO-B Monamine oxidase B pNSE

mgmt Methylguanine-DNA methyltransferase DNA repair protein Mgmt Methylguanine-DNA methyltransferase DNA repair protein

Increased CNS serotonin and norepinephrine levels. Model for MAO-A deficiency.

Tm

MAO-A Monamine oxidase A Tg(H2-INF-β)8

A.1B.8

Supplement 1


Overexpression of NOS2 in pancreatic islets. Model for the evaluation of NOS2 in pancreatic β-cell degeneration. Neuronal NOS deficiency yields gastric distention, pyloric stenosis, and decreased NOS catalytic activity. Model of infantile pyloric stenosis. Targeted deletion results in multiorgan hyperplasia, gigantism, tumorgenesis, and female sterility. Model for the evaluation of p27Kip1 function. Development of spontaneous tumors by 6 months of age. Model for studying role in tumorigenesis and testing suspected carcinogens.

Tg

NOS2 Nitric oxide synthase gene NOS2 Nitric oxide synthase gene neuronal NOS p27Kip1 Cyclin-dependent kinase inhibitor

Tg

Tg

p53 Tumor suppressor gene p53 172R-H

paf Platelet activating factor PARF

Tm

p53 Tumor suppressor gene

Tm

Decrease in spontaneous tumors with concurrent acceleration of carcinogen and oncogene-mediated tumorgenesis. Model for evaluation of early events in mammary tumorgenesis. Preclinical model for new chemotherapeutic testing. Bronchial hyperreactivity, increased endotoxin lethality, and melanocytic tumorgenesis. Model for study of bronchial asthma and endotoxin-induced death.

One-step mammary tumor induction, used to study major determinants of malignant progression of tumors.

Tg

neu (c-neu) Proto-oncogene MMTV-c-neu

Tm

All mice fed a specifically formulated diet developed Rao et al., 1997. Breast Cancer Res. palpable mammary tumors by 28 weeks of age. Model for Treat. 45:149 evaluation of dietary intervention strategies for breast cancer.

Tg

neu (c-neu) c-neu/erbB2 TG.NK

Ishii et al., 1997. EMBO J. 16:133

Li et al., 1998. Oncogene 16:997

Donehower et al., 1992. Nature 356:215

Fero et al., 1996. Cell 85:733

Huang et al., 1993. Cell 75:1273

Takamura et al., 1998. JBC 273:2493

Muller et al., 1988. Cell 54:105

Increased incidence of spontaneous T cell tumors and Stewart et al., 1993. Int. J. Cancer inducible leukemia. System for screening novel myc genes 53:1023 involved in T cell lymphoma.

Tg

myc (c-myc) Proto-oncogene T cell promoter

Weiss et al., 1997. EMBO J. 16:2985

Reference(s)

Neuroectodermal expression yields neuroblastoma development. Model for evaluating pathogenesis and therapy of neuroblastoma.

Description

Tg

Mutant typea

myc (c-myc) Proto-oncogene Pth-mycn



—

—

JL, TF

—

BK, JL

—

CR

—

—

—

Available fromb

A.1B.9


Supplement 1

—

Geraci et al., 1998. Chest 114:99S

Tg

Tg

ras c-Ha-ras ras H2

Neonatal lethality due to loss of epidermal barrier function and transepidermal water. Model for evaluating lipid metabolism modulation in relation to epidermal formation and functional maintenance. Multiple skin papilloma induction with further development to neoplasia. Model for screening tumor promoters and assessing antitumor and antiproliferative agents. Increased incidence of tumorgenesis in vascular endothelium, lymphocytes, skin, lung, and Harderian gland epithelium. Model for evaluation of somatic point mutations associated with transgene. Bioassay model for rapid carcinogenicity testing.

Saitoh et al.,1990. Oncogene 5:1195; Yamamoto et al., 1998. Environ. Health Perspect. 106(suppl. 1):57

Leder et al., 1990. PNAS 87:9178

Imakado et al., 1995. Genes Dev. 9:317

Tg

Tm

—

Bonventre et al., 1997. Nature 390:622

continued

—

TF, CR

—

—

—

—

Allen et al., 1997. Oncogene 15:1133

Expression induces craniofacial abnormalities. Model for Damm et al., 1993. PNAS 90:2989 evaluating mechanism of gene regulation by retinoic acid. Specific epidermal expression. Model for determination of Tsou et al., 1994. Exp. Cell Res. 214:27 sites of transcriptional activation by retinoic acid.

Increased incidence of T cell lymphomas. Model for evaluating the tumorigenic properties of pim-2. Null mutation yielded reduced fertility and decreased the ischemic brain injury associated with cerebral arterial occlusion. Model for evaluation of regulation of eicosanoid and platelet activation factor in inflammation process.

TF

—

Lim et al., 1990. Cancer Res. 50:1701

Overexpression promotes development of lymphomas van Lohuizen et al., 1989. Cell 56:673 before 7 months of age. Model for evaluating the cooperation between pim-1 and myc in lymphomagenesis.

Expression in liver and kidney regulated by dietary manipulation. Model for evaluating role of DNA repair in protection from carcinogenesis induced by N-nitro compounds. Selective pulmonary prostacyclin expression. Lung displays less hypoxic vasoconstriction. Model for evaluation of role in vasoconstriction.

Tg

ras Activated v-Ha-ras oncogene TG.AC

RAR Retinoic acid receptor RAR Retinoic acid receptor βRARE-LacZ RAR Retinoic acid receptor HK1-RARα403

Tg

pim-2 pim-2 proto-oncogene PLA2 Phospholipase A2 Tm

Tg

Tg

Tg

pim-1 pim-1 proto-oncogene Eµ-pim-1

pepck-ada Phosphoenolpyruvate carboxykinase Bacterial O6 alkylguanine-DNA alkyltransferase pgis Prostacyclin synthase hSPC-rPGIS

A.1B.10

Supplement 1


β-gal promoter trap selection for identification of active promoters/genes. Mouse model of colorectal adenocarcinoma with lymph node metastasis.

Tm

SOD-1 Copper-zinc superoxide dismutase

Tm

Tg

Increased Cu-Zn SOD activity in CNS. Animal model of Down’s syndrome. Increased Cu-Zn SOD expression in nervous tissue confers protection against various forms of brain trauma and focal cerebral ischemia. Model for evaluation of role of oxygen in ischemic and traumatic injury. Null mutant mice show a pronounced sensitivity to paraquat and myocardial ischemia reperfusion injury.

Progressive thyroid tumors. Model for evaluating neoplastic progression of human thyroid carcinomas.

Tm

Ret Proto-oncogene ret/ptc1 SAβ-gal, SAβ-geo, and ROSAβ-geo Tm

Friedrich and Soriano, 1991. Genes Dev. 5:1513 Zhu et al., 1998. Cell 94:703

Aberrant melanogenesis with melanocytic tumor development. Model for systematic development of melanocytic tumors.

Tg

Ret Proto-oncogene MT/ret

Smad3 Transforming growth factor β-receptor gene SOD-1 Copper-zinc superoxide dismutase

Santoro et al., 1996. Oncogene 12:1821

Increased incidence of mammary, salivary, and male reproductive tract tumors. Model for evaluating oncogenicity of activated ret gene.

Tg

ret Proto-oncogene MMTV/ret

Ho et al., 1998. Environ. Health Perspect. 106:1219

Epstein et al., 1987. PNAS 84:8044; Chan et al., 1994. Ann. N.Y. Acad. Sci. 738:93

Iwamoto et al., 1991. EMBO J. 10:3167

Iwamoto et al., 1990. Oncogene 5:535

Neuronal cell death and defective erythropoiesis in 14-15 Jacks et al., 1992. Nature 359:295; Hu et al., 1994. Oncogene 9:1021 day old homozygous pups and development of pituitary tumors in heterozygous adult mice. Model for evaluating normal function in cells and for human familial retinoblastoma. Heterozygous mice exhibit high incidence of spontaneous pituitary tumors between 2 and 11 months of age. Model for addressing tissue-specific tumor predisposition by inactivation of ubiquitously expressed tumor-suppressor gene.

Tm

Rb Retinoblastoma tumor-suppressor gene

Andres et al., 1987. PNAS 84:1299

Reference(s)

Increased incidence of mammary and salivary gland tumors. Model for in vivo evaluation of molecular mechanisms of proto-oncogene activation.

Description

Tg

Mutant typea

ras Ha-ras oncogene Whey acidic protein (wap)-ras



—

JL

—

—

—

—

—

JL

JL

Available fromb

A.1B.11


Supplement 1

Reduced incidence of diabetes. Used to assess oxidative stress in pancreatic β cell destruction in autoimmune diabetes. Deletion yields increased susceptibility to ultraviolet-B and carcinogen–induced skin and eye tumors and papillomas. Model for human XPA. Gene deficiency yields increased susceptibility to ultraviolet-induced carcinogenesis. Animal model for human XPC.

Tm

Tm

Tm

trx Thioredoxin, a reduction/oxidation active protein Ins-TRX

XPA Xeroderma pigmentosum, group A

XPC Xeroderma pigmentosum, group C

—

—

TF

Hotta et al., 1998. J. Exp. Med. 188:1445

deVries et al., 1995. Nature 377:169; O’Neill, 1996. 24:642 Sands et al., 1995. Nature 377:162

—

JL, TF

—

the author of the cited paper.

bAbbreviations: BK, B & K Universal; JL, The Jackson Laboratory; CR, Charles River Labs; TF, Taconic Farms. See SUPPLIERS APPENDIX for contact information. If no commercial supplier is listed, consult

aAbbreviations: Tg, transgenic; Tm, targeted mutation.

Expression of truncated protein in thymocytes predisposes Ceci et al., 1997. Genes Dev. 11:688 to development of T cell lymphomas. Model for studying involvement in oncogenesis.

Tg

tpl-2 Tumor progression loci 2, an oncogenic kinase

Probert et al., 1993. J. Immunol. 151:1894

Diffuse vascular thrombosis and tissue necrosis neutralizable by human TNF mAb administration. Model for evaluating the different contributions of the two TNF receptors in thymus development and signaling.

Tm

Keffer et al., 1991. EMBO J. 10:4025

TNF-α Tumor necrosis factor CD2-TNF

Development of chronic inflammatory polyarthritis with prevention possible by administration of anti-hTNF monoclonal antibody. Model for human arthritis.

JL

—

Osman et al., 1998. Int. Immunol. 10:1613 Factor et al., 1998. JBC 273:15846

—

Wispe et al., 1992. JBC 267:23937

Tg

Model of oxidation DNA damage and acceleration of hepatocarcinogenesis.

Increased expression confers pulmonary epithelial cell protection against oxygen injury. Used to study the primary role of lung epithelial cell gene expression in oxygen-injury protection. Expression of type II collagen–specific T cell receptor transgenes accelerates onset of arthritis in DBA/1 inbred mice.

TNF-α Tumor necrosis factor

Tg double

Tg

Tcr-α/β T cell receptor α/β chain genes

TGF-α/c-myc Transforming growth factor α and the proto-oncogene c-myc

Tg

SOD-2 Manganese superoxide dismutase SP-C-Mn-SOD-hGH

LABORATORY STOCK SOLUTIONS AND EQUIPMENT

APPENDIX 2

Common Stock Solutions and Buffers

APPENDIX 2A

This section describes the preparation of buffers and reagents used in this manual. When preparing solutions, use Milli-Q-purified water (or equivalent) and reagents of the highest available grade. Sterilization—by filtration through a 0.22-µm filter or by autoclaving—is recommended for most solutions stored at room temperature. Where storage conditions are not specified, store up to 6 months at room temperature. Discard any solution that shows evidence of contamination, precipitation, or discoloration. Standardized and reference reagents are sometimes required—e.g., for quality control, positive and negative controls, equipment calibration, and assay validation. Environmental and chemical standards are available from a number of suppliers, including the National Institute of Standards and Technology (NIST), AccuStandard Inc., and Sigma Chemical Co. Consult the LabGuide, published annually by the American Chemical Society (ACS) for other suppliers (e.g., ACS, 1999). Acid, concentrated stock solutions See Table A.2A.1. Ammonium acetate, 10 M Dissolve 385.4 g ammonium acetate in 150 ml H2O Add H2O to 500 ml Ammonium hydroxide, concentrated stock solution See Table A.2A.1. Ammonium sulfate, saturated 76 g ammonium sulfate 100 ml H2O Heat with stirring to just below boiling point Let stand overnight at room temperature

Table A.2A.1

Molarities and Specific Gravities of Concentrated Acids and Basesa

Acid/base Acetic acid (glacial) Ammonium hydroxide Formic acid Hydrochloric acid Nitric acid Perchloric acid Phosphoric acid Sulfuric acid

Molecular weight

% by weight

Molarity (approx.)

Specific gravity

1M solution (ml/liter)

99.6 28 90 98 36 70 60 72 85 98

17.4 14.8 23.6 25.9 11.6 15.7 9.2 12.2 14.7 18.3

1.05 0.90 1.205 1.22 1.18 1.42 1.54 1.70 1.70 1.835

57.5 67.6 42.4 38.5 85.9 63.7 108.8 82.1 67.8 54.5

60.05 35.0 46.03 36.46 63.01 100.46 98.00 98.07

aCAUTION: Handle strong acids and bases carefully.

Current Protocols in Toxicology (1999) A.2A.1-A.2A.6 Copyright © 1999 by John Wiley & Sons, Inc.

Laboratory Stock Solutions and Equipment

A.2A.1

ATP, 100 mM 1 g ATP (adenosine triphosphate) 12 ml H2O Adjust pH to 7.0 with 4 M NaOH Adjust volume to 16.7 ml with H2O Store in aliquots indefinitely at −20°C Base, concentrated stock solutions See Table A.2A.1. CaCl2 , 1 M 147 g CaCl2⋅2H2O H2O to 1 liter Carbonate buffer 1.6 g Na2CO3 (15 mM final) 2.9 g NaHCO3 (35 mM final) 0.2 g NaN3 (3.1 mM final) H2O to 1 liter Adjust to pH 9.5 CAUTION: Sodium azide is poisonous; follow appropriate precautions for handling, storage, and disposal.

CMF-DPBS (calcium- and magnesium-free Dulbecco’s phosphate-buffered saline) 8.00 g NaCl (0.137 M) 0.20 g KCl (2.7 mM) 2.16 g Na2HPO4⋅7H2O (8.1 mM) 0.20 g KH2PO4 (1.1 mM) H2O to 1 liter DPBS (Dulbecco’s phosphate-buffered saline) 8.00 g NaCl (0.137 M) 0.20 g KCl (2.7 mM) 0.20 g KH2PO4 (1.1 mM) 0.10 g MgCl2⋅6H2O (0.5 mM) 2.16 g Na2HPO4⋅7H2O (8.1 mM) 0.10 g anhydrous CaCl2 (0.9 mM) H2O to 1 liter DTT (dithiothreitol), 1 M Dissolve 1.55 g DTT in 10 ml water and filter sterilize. Store in aliquots at −20°C. EDTA (ethylenediaminetetraacetic acid), 0.5 M (pH 8.0) Dissolve 186.1 g disodium EDTA dihydrate in 700 ml water. Adjust pH to 8.0 with 10 M NaOH (∼50 ml; add slowly). Add water to 1 liter and filter sterilize. Begin titrating before the sample is completely dissolved. EDTA, even the disodium salt, is difficult to dissolve at this concentration unless the pH is increased to between 7 and 8.


HBSS (Hanks’ balanced salt solution) 0.40 g KCl (5.4 mM final) 0.09 g Na2HPO4⋅7H2O (0.3 mM final) 0.06 g KH2PO4 (0.4 mM final) 0.35 g NaHCO3 (4.2 mM final) 0.14 g CaCl2 (1.3 mM final) 0.10 g MgCl2⋅6H2O (0.5 mM final) continued


0.10 g MgSO4⋅7H2O (0.6 mM final) 8.0 g NaCl (137 mM final) 1.0 g D-glucose (5.6 mM final) 0.2 g phenol red (0.02%; optional) Add H2O to l liter and adjust pH to 7.4 with 1 M HCl or 1 M NaOH Filter sterilize and store up to 1 month at 4°C HBSS may be made or purchased without Ca2+ and Mg2+ (CMF-HBSS). These components are optional and usually have no effect on an experiment; in a few cases, however, their presence may be detrimental. Consult individual protocols to see if the presence or absence of these components is recommended. Bottles should be kept tightly closed to prevent CO2 loss and subsequent alkalinization.

HCl, 1 M Mix in the following order: 913.8 ml H2O 86.2 ml concentrated HCl KCl, 1 M 74.6 g KCl H2O to 1 liter MgCl2 , 1 M 20.3 g MgCl2⋅6H2O H2O to 100 ml MgSO4 , 1 M 24.6 g MgSO4⋅7H2O H2O to 100 ml NaCl, 5 M 292 g NaCl H2O to 1 liter NaCl (saline), 0.9% (w/v) 9 g NaCl (154 mM final) H2O to 1 liter NaOH, 10 M Dissolve 400 g NaOH in 450 ml H2O H2O to 1 liter PBS (phosphate-buffered saline) 8.00 g NaCl (0.137 M) 0.20 g KCl (2.7 mM) 0.24 g KH2PO4 (1.4 mM) 1.44 g Na2HPO4 (0.01 M) H2O to 1 liter Potassium acetate buffer, 0.1 M Solution A: 11.55 ml glacial acetic acid per liter (0.2 M) in water. Solution B: 19.6 g potassium acetate (KC2H3O2) per liter (0.2 M) in water. Referring to Table A.2A.2 for desired pH, mix the indicated volumes of solutions A and B, then dilute with water to 100 ml. Filter sterilize if necessary. Store up to 3 months at room temperature. continued



This may be made as a 5- or 10-fold concentrate by scaling up the amount of sodium acetate in the same volume. Acetate buffers show concentration-dependent pH changes, so check the pH by diluting an aliquot of concentrate to the final concentration. To prepare buffers with pH intermediate between the points listed in Table A.2A.2, prepare closest higher pH, then titrate with solution A.

Potassium phosphate buffer, 0.1 M Solution A: 27.2 g KH2PO4 per liter (0.2 M final) in water. Solution B: 34.8 g K2HPO4 per liter (0.2 M final) in water. Referring to Table A.2A.3 for desired pH, mix the indicated volumes of solutions A and B, then dilute with water to 200 ml. Filter sterilize if necessary. Store up to 3 months at room temperature. This buffer may be made as a 5- or 10-fold concentrate simply by scaling up the amount of potassium phosphate in the same final volume. Phosphate buffers show concentration-dependent changes in pH, so check the pH of the concentrate by diluting an aliquot to the final concentration. To prepare buffers with pH intermediate between the points listed in Table A.2A.3, prepare closest higher pH, then titrate with solution A. Table A.2A.2 Preparation of 0.1 M Sodium and Potassium Acetate Buffersa

Desired pH

Solution A (ml)

Solution B (ml)

3.6 3.8 4.0 4.2 4.4 4.6 4.8 5.0 5.2 5.4 5.6

46.3 44.0 41.0 36.8 30.5 25.5 20.0 14.8 10.5 8.8 4.8

3.7 6.0 9.0 13.2 19.5 24.5 30.0 35.2 39.5 41.2 45.2

aAdapted by permission from CRC (1975).

Table A.2A.3 Preparation of 0.1 M Sodium and Potassium Phosphate Buffersa


Desired pH

Solution A (ml)

Solution B (ml)

Desired pH

Solution A (ml)

Solution B (ml)

5.7 5.8 5.9 6.0 6.1 6.2 6.3 6.4 6.5 6.6 6.7 6.8

93.5 92.0 90.0 87.7 85.0 81.5 77.5 73.5 68.5 62.5 56.5 51.0

6.5 8.0 10.0 12.3 15.0 18.5 22.5 26.5 31.5 37.5 43.5 49.0

6.9 7.0 7.1 7.2 7.3 7.4 7.5 7.6 7.7 7.8 7.9 8.0

45.0 39.0 33.0 28.0 23.0 19.0 16.0 13.0 10.5 8.5 7.0 5.3

55.0 61.0 67.0 72.0 77.0 81.0 84.0 87.0 90.5 91.5 93.0 94.7

aAdapted by permission from CRC (1975).


Table A.2A.4

Preparation of SDS Sample Buffer

Ingredient

2×

4×

Final conc. in 1× buffer

0.5 M Tris⋅Cl, pH 6.8a SDS Glycerol Bromphenol blue 2-Mercaptoethanolb,c H2O

2.5 ml 0.4 g 2.0 ml 20 mg 400 µl to 10 ml

5.0 ml 0.8 g 4.0 ml 40 mg 800 µl to 10 ml

62.5 mM 2% (w/v) 10% (v/v) 0.1% (w/v) ∼300 mM —

aSee recipe below. bAlternatively, dithiothreitol (DTT), at a final concentration of 100 mM, can be substituted

for 2-mercaptoethanol. cAdd just before use.

SDS, 20% (w/v) Dissolve 20 g SDS (sodium dodecyl sulfate or sodium lauryl sulfate) in H2O to 100 ml total volume with stirring. Filter sterilize using a 0.45-µm filter. It may be necessary to heat the solution slightly to fully dissolve the powder.

SDS electrophoresis buffer, 5× 15.1 g Tris base 72.0 g glycine 5.0 g SDS Distilled, deionized H2O to 1 liter Store up to 1 month at 0° to 4°C Dilute to 1× before use Do not adjust the pH of the stock solution; the pH is 8.3 when diluted to 1×. Use purified SDS if appropriate.

SDS sample buffer See Table A.2A.4. Sodium acetate, 3 M Dissolve 408 g sodium acetate trihydrate (NaC2H3O2⋅3H2O) in 800 ml H2O Adjust pH to 4.8, 5.0, or 5.2 (as desired) with 3 M acetic acid (see Table A.2A.1) Add H2O to 1 liter Filter sterilize Sodium acetate buffer, 0.1 M Solution A: 11.55 ml glacial acetic acid per liter (0.2 M) in water. Solution B: 27.2 g sodium acetate (NaC2H3O2⋅3H2O) per liter (0.2 M) in water. Referring to Table A.2A.2 for desired pH, mix the indicated volumes of solutions A and B, then dilute with water to 100 ml. Filter sterilize if necessary. Store up to 3 months at room temperature. This may be made as a 5- or 10-fold concentrate by scaling up the amount of sodium acetate in the same volume. Acetate buffers show concentration-dependent pH changes, so check the pH by diluting an aliquot of concentrate to the final concentration. To prepare buffers with pH intermediate between the points listed in Table A.2A.2, prepare closest higher pH, then titrate with solution A. Laboratory Stock Solutions and Equipment


Sodium phosphate buffer, 0.1 M Solution A: 27.6 g NaH2PO4⋅H2O per liter (0.2 M final) in water. Solution B: 53.65 g Na2HPO4⋅7H2O per liter (0.2 M) in water. Referring to Table A.2A.3 for desired pH, mix the indicated volumes of solutions A and B, then dilute with water to 200 ml. Filter sterilize if necessary. Store up to 3 months at room temperature. This buffer may be made as a 5- or 10-fold concentrate by scaling up the amount of sodium phosphate in the same final volume. Phosphate buffers show concentration-dependent changes in pH, so check the pH by diluting an aliquot of the concentrate to the final concentration. To prepare buffers with pH intermediate between the points listed in Table A.2A.3, prepare closest higher pH, then titrate with solution A.

TBS (Tris-buffered saline) 100 mM Tris⋅Cl, pH 7.5 (see recipe below) 0.9% (w/v) NaCl Store up to several months at 4°C Tris⋅Cl, 1 M Dissolve 121 g Tris base in 800 ml H2O Adjust to desired pH with concentrated HCl Adjust volume to 1 liter with H2O Filter sterilize if necessary Store up to 6 months at 4°C or room temperature Approximately 70 ml HCl is needed to achieve a pH 7.4 solution, and ∼42 ml for a solution that is pH 8.0. IMPORTANT NOTE: The pH of Tris buffers changes significantly with temperature, decreasing approximately 0.028 pH units per 1°C. Tris-buffered solutions should be adjusted to the desired pH at the temperature at which they will be used. Because the pKa of Tris is 8.08, Tris should not be used as a buffer below pH ∼7.2 or above pH ∼9.0.

LITERATURE CITED American Chemical Society (ACS). 1999. LabGuide 98-99. ACS, Washington, D.C. [also available online at http://pubs.acs.org./labguide]. Chemical Rubber Company (CRC). 1975. CRC Handbook of Biochemistry and Molecular Biology, Physical and Chemical Data, 3d ed., Vol. 1. CRC Press, Boca Raton, Fla.



Standard Laboratory Equipment

APPENDIX 2B

Listed below are pieces of equipment that are standard in the modern toxicology laboratory—i.e., items used extensively in this manual and thus not usually included in the individual materials lists. No attempt has been made to list all items required for each procedure in the Materials list of each protocol; rather, those lists note those items that might not be readily available in the laboratory or that require special preparation. See SUPPLIERS APPENDIX for contact information for commercial vendors of laboratory equipment. Applicator, cotton-tipped and wooden

Gloves, disposable plastic and heat-resistant

Autoclave

Graduated cylinders

Bag sealer Balances, analytical and preparative

Heating blocks, thermostatically controlled for test tubes and microcentrifuge tubes

Beakers

Homogenizer

Bench protectors, plastic-backed (including “blue” pads)

Ice bucket

Biohazard disposal containers and bags Biosafety cabinet, tissue culture or laminar flow hood; filters air and maintains air flow pattern to protect cultured cells from investigator and vice versa Bottles, glass, plastic, and squirt Bunsen burners Centrifuges, low-speed (to 20,000 rpm) refrigerated, ultracentrifuge (20,000 to 80,000 rpm), large-capacity low-speed, tabletop, with appropriate rotors and adapters Centrifuge tubes and bottles, plastic and glass, various sizes Clamps Conical centrifuge tubes, plastic and glass Containers, assortment of glass and plastic, for gel and membrane washes Coplin jars, glass, for 25 × 75–mm slides Cuvettes, glass and quartz

Humidified CO2 incubator Ice maker Immersion oil for microscopy Lab coats Laboratory glassware Light box Liquid nitrogen Lyophilizer Magnetic stirrer, with and without heater, and stir bars Markers, including indelible markers, chinamarking pens, and luminescent markers Microcentrifuge, Eppendorf-type with 12,000 to 14,000 rpm maximum speed Microcentrifuge tubes, 0.2-, 0.5-, 1.5-, 2-ml Mortar and pestle Ovens, drying and microwave Paper cutter, large Paper towels

Desiccator and desiccant

Parafilm

Dry ice

Pasteur pipets and bulbs

Electrophoresis equipment, agarose and acrylamide, full-size and mini, with power supplies

pH meter

Film developing system and darkroom

pH paper Pipets, graduated

Filtration apparatus

Pipettors, adjustable delivery, 0.5- to 10-µl, 10- to 200-µl, and 200- to 1000-µl

Forceps

Plastic wrap (e.g., Saran Wrap)

Freezers, −20°C, −70°C, and liquid nitrogen Fume hood Geiger counter Gel dryer

Current Protocols in Toxicology (1999) A.2B.1-A.2B.2 Copyright © 1999 by John Wiley & Sons, Inc.

Polaroid camera or video documentation system Power supplies, 300-V for polyacrylamide gels, 2000- to 3000-V for other applications


A.2B.1

Radiation shield, Lucite or Plexiglas

Tape, masking, electrician’s black, autoclave, and Time tape

Radioactive waste containers for liquid and solid wastes

Test tubes, glass and plastic, various sizes, with and without caps

Refrigerator, 4°C

Timer

Ring stand and rings

Toolbox with common tools

Rubber policemen or plastic scrapers

Trays, plastic and glass, various sizes

Rubber stoppers

Tubing, rubber and Tygon

Safety glasses

UV transparent plastic wrap (e.g., Saran Wrap)

Racks, test tube and microcentrifuge tubes

Scalpels and blades Scintillation counter, β Scissors Shakers, orbital and platform, room temperature or 37°C

Vacuum desiccator Vacuum oven Vacuum supply Vortex mixers

Spectrophotometer, visible and UV range

Waring Blendor or equivalent blender

Speedvac evaporator

Water bath with adjustable temperature

Syringes and needles

Water purification system X-ray film cassettes and intensifying screens

Standard Laboratory Equipment

A.2B.2 Current Protocols in Toxicology

USEFUL TECHNIQUES

APPENDIX 3

Molecular Biology Techniques

APPENDIX 3A

Protocols presented in Current Protocols in Toxicology (CPTX) may include molecular biological, biochemical, and other biological techniques that may not be fully described in this manual. Although it may be reasonable to assume that readers have at least a basic understanding of these techniques, there are times when a full step-by-step description of a procedure is helpful. For that reason we have included the following table (Table A.3A.1), which lists common molecular biology techniques, some of which are described in this manual and provides references to specific units in its sister publications Current Protocols in Molecular Biology (CPMB; Ausubel et al., 1999), Current Protocols in Immunology (CPI; Coligan et al., 1999), and Current Protocols in Cell Biology (CPCB; Bonifacino et al., 1999), which describe the methods in more detail. Protocols for some of these techniques will be added as units or as sections of this appendix in future supplements. Alternatively, the reader may wish to consult any of the many other texts and manuals specifically devoted to recombinant DNA technology and biochemical analysis. Table A.3A.1 Molecular Biology Techniques

Technique

Unit(s)

Animal care and handling Antibody affinity purification monoclonal polyclonal purification Antigen-hapten conjugation Autoradiography Bacterial cell culture extracts Blotting northern Southern Cells, preparation from tissues Chemiluminescent detection Chromatography cation-exchange gel-filtration immunoaffinity ion-exchange size-exclusion Cloning PCR products subcloning DNA fragments Colorimetry, for protein quantitation

CPI

Chapter 1

CPTX 2.3, CPI 2.7 & 2.9 CPMB 11.4-11.11, CPI 2.5 CPMB 11.12, CPI 2.4 CPMB 11.11 & 11.13, CPI 2.7 CPTX 2.3, CPI 9.4 CPMB APPENDIX 3A, CPI APPENDIX 3J, CPCB 6.3 CPMB 1.2 & 1.3 CPTX 9.3 CPMB 4.9, CPI 10.12 CPMB 2.9, CPI 10.6A CPTX 6.3 CPMB 10.8, CPI 8.10 CPTX 8.2 CPTX 7.1, CPMB 10.9, CPI 9.2 CPTX 3.2, CPMB 10.11A, CPI 8.2 CPTX 4.2, CPMB 10.10 CPMB 10.9, CPI APPENDIX 3I CPMB 15.2 CPMB 3.16 CPMB 10.1A

continued Useful Techniques Current Protocols in Toxicology (1999) A.3A.1-A.3A.3 Copyright © 1999 by John Wiley & Sons, Inc.

A.3A.1

Table A.3A.1 Molecular Biology Techniques, continued

Technique

Molecular Biology Techniques

Dialysis DNA extraction from mammalian tissue ligation preparation, miniprep purification and concentration quantification, spectrophotometric recovery from agarose gels sequencing DNase I digestion E. coli growth in liquid medium growth on solid medium lysate preparation membrane preparation transformation ELISA Ethanol precipitation of DNA Fixation, embedding, and sectioning Gel electrophoresis agarose, basic agarose, low gelling/melting temperature agarose agarose, minigels denaturing polyacrylamide (oligonucletide) nondenaturing polyacrylamide (DNA) SDS-PAGE, basic SDS-PAGE, minigels SDS-PAGE, two-dimensional staining gels with Coomassie blue Genetic analysis of bacteria HPLC analytical purification Hybridization of Southern blots in situ Immunoblotting Immunohistochemistry Immunoprecipitation Interaction trap/two-hybrid system Isoelectric focusing Kinetic assay methods Mammalian cell tissue culture Mass spectrometry Media, preparation and use E. coli mammalian cell yeast

Unit(s) CPMB APPENDIX 3C, CPI APPENDIX 3H CPTX 2.2 & 3.2 CPMB 2.2, CPI 10.2 CPMB 3.14 CPMB 1.6, CPI 10.3 CPMB 2.14, CPI 10.1 CPMB APPENDIX 3D, CPI APPENDIX 3I CPMB 2.6, CPI 10.1 CPMB

Chapter 7, CPI 10.25

CPTX 3.2 CPMB 1.2, CPI 10.3 CPMB 1.3 CPMB 1.7, CPI 10.3 CPTX 8.5 CPMB 1.8, CPI APPENDIX 3N CPTX 2.3, CPMB 11.2, CPI 2.2 CPMB 2.1A, CPI 10.1 CPMB 14.1, CPI 5.8 CPTX 2.2, CPMB 2.5A, CPI 10.4 CPMB 2.6, CPI 10.5 CPMB 2.5A, CPI 10.4 CPMB 2.12, CPI 10.4 CPMB 2.7, CPI 10.7 CPMB 10.2, CPI 8.4 CPTX 2.2 CPMB 10.3 & 10.4, CPI 8.5 CPMB 10.6, CPI 10.9 CPTX 3.1 CPTX 6.2 & 6.3 CPTX 10.2 CPMB 2.10, CPI 10.6A CPMB

Chapter 14, CPI 12.8

CPTX 2.3, CPMB 10.8, CPI 8.10, CPCB 6.2, CPTX 2.3 CPMB 14.5, CPI 5.8, CPMB 4.6 CPTX 7.1, CPMB 10.16, CPI 8.3 CPMB 20.1 CPTX 9.4 CPMB APPENDIX 3H CPMB APPENDIX 3F CPTX 3.2 CPMB 1.1-1.3 CPMB APPENDIX 3F, CPI APPENDIX 2, CPCB 1.2 CPMB 13.1 & 13.2

continued


Table A.3A.1 Molecular Biology Techniques, continued

Technique Microsome preparation Mitochondrial preparation PCR anchored end-labeling primers for general primer design primer synthesis using end-labeled primers Phenol/chloroform extraction DNA RNA Plasmid preparation Protein preparation from bacteria quantitation, spectrophotometric and colormetric Pulsed-field gel electrophoresis Quantitation of DNA and RNA Radioimmunoassay Radiometric enzyme assay Random primer labeling Restriction endonuclease digestion RNA preparation and purification extraction with guanidinium isothiocyanate poly(A)+ total Silanization of glassware Spectrophotometry for quantitation of DNA and RNA for quantitation of protein T4 DNA ligase Thin-layer chromatography Tissue fractionation Transfection calcium-mediated DEAE-dextran-mediated electroporation liposome-mediated TUNEL assay for DNA fragmentation

Unit(s) CPTX 4.1, 4.3 & 9.3 CPTX 4.1 & 6.3 CPMB 15.6, CPI 10.24 CPMB 3.10, CPI 10.10 CPMB 15.1, CPI 10.20 CPMB 15.1, CPI 10.20 CPMB 2.11 CPMB 7.4 CPMB 2.1, CPI 10.4 CPMB 4.1, CPI 10.11 CPMB 1.6, 1.7 & 2.1B, CPI 17.1 CPTX 9.4 CPMB 10.1A CPTX 2.2 CPMB APPENDIX 3D, CPI APPENDIX 3L CPTX 10.4, CPMB 9.7A, CPI 18.3 CPTX 8.2 CPMB 3.5, CPI 10.22 CPMB 3.1-3.3, CPI 10.8 CPMB 4.2, CPI 10.11 CPMB 4.5, CPI 10.11 CPMB 4.1 & 4.2, CPI 10.11 CPMB APPENDIX 3B, CPI APPENDIX 3K CPMB APPENDIX 3D, CPI APPENDIX 3L CPMB APPENDIX 10.1A CPMB 3.14 CPTX 4.3, CPI 11.1 CPTX 4.1 CPMB 9.1, CPI 10.13 CPMB 9.2, CPI 10.14 CPMB 1.8, CPI 10.15 CPMB 9.4, CPI 10.16 CPTX 2.2, CPI 3.17

LITERATURE CITED Ausubel, F.A., Brent, R., Kingston, R.E., Moore, D.D., Seidman, J.G., Smith, J.A., and Struhl, K. (eds.). 1999. Current Protocols in Molecular Biology. John Wiley & Sons, New York. Bonifacino, J.S., Dasso, M., Harford, J.B., Lippincott-Schwartz, J., and Yamada, K.M. (eds.). 1999. Current Protocols in Cell Biology. John Wiley & Sons, New York. Coligan, J.E., Kruisbeek, A.M., Margulies, D.H., Shevach, E.M., and Strober, W. 1999. Current Protocols in Immunology. John Wiley & Sons, New York. Useful Techniques


Techniques for Mammalian Cell Tissue Culture

APPENDIX 3B

Tissue culture technology has found wide application in biological research. Monolayer cell cultures and suspension are utilized in cytogenetic, biochemical, and molecular laboratories for diagnostic as well as research studies. In most cases, cells or tissues must be grown in culture for days or weeks to obtain sufficient numbers of cells for analysis. Maintenance of cells in long-term culture requires strict adherence to aseptic technique to avoid contamination and potential loss of valuable cell lines. The first section of this appendix discusses basic principles of aseptic technique. An important factor influencing the growth of cells in culture is the choice of tissue culture medium. Many different recipes for tissue culture media are available and each laboratory must determine which medium best suits their needs. Individual laboratories may elect to use commercially prepared medium or prepare their own. Commercially available medium can be obtained as a sterile and ready-to-use liquid, in a concentrated liquid form, or in a powdered form. Besides providing nutrients for growing cells, medium is generally supplemented with antibiotics, fungicides, or both to inhibit contamination. The second section of this appendix discusses medium preparation. As cells reach confluency, they must be subcultured or passaged. Failure to subculture confluent cells results in reduced mitotic index and eventually in cell death. The first step in subculturing monolayers is to detach cells from the surface of the primary culture vessel by trypsinization or mechanical means (see Basic Protocol). The resultant cell suspension is then subdivided, or reseeded, into fresh cultures. Suspension cells are more easily passaged (see Alternate Protocol). Secondary cultures are checked for growth, fed periodically, and may be subsequently subcultured to produce tertiary cultures, etc. The time between passaging cells varies with the cell line and depends on the growth rate. Support protocols describe freezing of monolayer (see Support Protocol 1) and suspension cells (see Support Protocol 2), thawing and recovery of cells (see Support Protocol 3), cell counting using a hemacytometer (see Support Protocol 4), and preparation of cells for transport (see Support Protocol 5). ASEPTIC TECHNIQUE It is essential that aseptic technique be maintained when working with cell cultures. Aseptic technique involves a number of precautions to protect both the cultured cells and the laboratory worker from infection. Laboratory workers must realize that cells handled in the lab are potentially infectious and should be handled with caution. Protective apparel such as gloves, lab coats or aprons, and eyewear should be worn when appropriate (Knutsen, 1997). Care should be taken when handling sharp objects such as needles, scissors, scalpel blades, and glass that could puncture the skin. Sterile disposable plastic supplies may be used to avoid the risk of broken or splintered glass (Rooney and Czepulkowski, 1992). Frequently, specimens received in the laboratory are not sterile, and cultures prepared from these specimens may become contaminated with bacteria, fungus, or yeast. The presence of microorganisms can inhibit growth, kill cell cultures, or lead to inconsistencies in test results. The contaminants deplete nutrients in the medium and may produce substances that are toxic to cells. Antibiotics (penicillin, streptomycin, kanamycin, or gentamycin) and fungicides (amphotericin B or mycostatin) may be added to tissue culture medium to combat potential contaminants (see Table A.3B.1). An antibiotic/anContributed by Mary C. Phelan Current Protocols in Toxicology (1999) A.3B.1-A.3B.15 Copyright © 1999 by John Wiley & Sons, Inc.

Useful Techniques

A.3B.1 Supplement 2

timycotic solution or lyophilized powder that contains penicillin, streptomycin, and amphotericin B is available from Sigma. The solution can be used to wash specimens prior to culture and can be added to medium used for tissue culture. Similar preparations are available from other suppliers. All materials that come into direct contact with cultures must be sterile. Sterile disposable dishes, flasks, pipets, etc., can be obtained directly from manufacturers. Reusable glassware must be washed, rinsed thoroughly, then sterilized by autoclaving or by dry heat before reusing. With dry heat, glassware should be heated 90 min to 2 hr at 160°C to ensure sterility. Materials that may be damaged by very high temperatures can be autoclaved 20 min at 120°C and 15 psi. All media, reagents, and other solutions that come into contact with the cultures must also be sterile; medium may be obtained as a sterile liquid from the manufacturer, autoclaved if not heat-sensitive, or filter sterilized. Supplements can be added to media prior to filtration, or they can be added aseptically after filtration. Filters with 0.20- to 0.22-µm pore size should be used to remove small gram-negative bacteria from culture media and solutions. Contamination can occur at any step in handling cultured cells. Care should be taken when pipetting media or other solutions for tissue culture. The necks of bottles and flasks, as well as the tips of the pipets, should be flamed before the pipet is introduced into the bottle. If the pipet tip comes into contact with the benchtop or any other nonsterile surface, it should be discarded and a fresh pipet obtained. Forceps and scissors used in tissue culture can be rapidly sterilized by dipping in 70% alcohol and flaming. Although tissue culture work can be done on an open bench if aseptic methods are strictly enforced, many labs prefer to perform tissue culture work in a room or low-traffic area reserved specifically for that purpose. At the very least, biological safety cabinets are recommended to protect the cultures as well as the laboratory worker. In a laminar-flow hood, the flow of air protects the work area from dust and contamination and acts as a barrier between the work surface and the worker. Many different styles of safety hoods are available and the laboratory should consider the types of samples being processed and the types of potential pathogenic exposure in making a selection. Manufacturer recommendations should be followed regarding routine maintenance checks on air flow and filters. For day-to-day use, the cabinet should be turned on for at least 5 min prior to beginning work. All work surfaces both inside and outside of the hood should be kept clean and disinfected daily and after each use.


Some safety cabinets are equipped with ultraviolet (UV) lights for decontamination of work surfaces. However, use of UV lamps is no longer recommended, as it is generally ineffective (Knutsen, 1997). UV lamps may actually produce a false sense of security as they maintain a visible blue glow long after their germicidal effectiveness is lost. Effectiveness diminishes over time as the glass tube gradually loses its ability to transmit short UV wavelengths, and it may also be reduced by dust on the glass tube, distance from the work surface, temperature, and air movement. Even when the UV output is adequate, the rays must directly strike a microorganism in order to kill it; bacteria or mold spores hidden below the surface of a material or outside the direct path of the rays will not be destroyed. Another rule of thumb is that anything that can be seen cannot be killed by UV. UV lamps will only destroy microorganisms such as bacteria, virus, and mold spores; they will not destroy insects or other large organisms (Westinghouse Electric Company, 1976). The current recommendation is that work surfaces be wiped down with ethanol instead of relying on UV lamps, although some labs use the lamps in addition to ethanol wipes to decontaminate work areas. A special metering device is available to measure the output of UV lamps, and the lamps should be replaced when they fall below the minimum requirements for protection (Westinghouse Electric Company, 1976).

A.3B.2 Supplement 2


Cultures should be checked routinely for contamination. Indicators in the tissue culture medium change color when contamination is present: for example, medium that contains phenol red changes to yellow because of increased acidity. Cloudiness and turbidity are also observed in contaminated cultures. Once contamination is confirmed with a microscope, infected cultures are generally discarded. Keeping contaminated cultures increases the risk of contaminating other cultures. Sometimes a contaminated cell line can be salvaged by treating it with various combinations of antibiotics and antimycotics in an attempt to eradicate the infection (e.g., see Fitch et al., 1997). However, such treatment may adversely affect cell growth and is often unsuccessful in any case. In addition to microscopically detectable bacteria and fungi, cultures can be contaminated by mycoplasma, which require more elaborate procedures to detect (Coté, 1999). PREPARING CULTURE MEDIUM Choice of tissue culture medium comes from experience. An individual laboratory must select the medium that best suits the type of cells being cultured. Chemically defined media are available in liquid or powdered form from a number of suppliers. Sterile, ready-to-use medium has the advantage of being convenient, although it is more costly than other forms. Powdered medium must be reconstituted with tissue culture–grade water according to manufacturer’s directions. Distilled or deionized water is not of sufficiently high quality for medium preparation; double- or triple-distilled water or commercially available tissue culture water should be used. The medium should be filter sterilized and transferred to sterile bottles. Prepared medium can generally be stored ≤1 month in a 4°C refrigerator. Laboratories using large volumes of medium may choose to prepare their own medium from standard recipes. This is the most economical approach, but it is time-consuming. Basic media such as Eagle’s minimal essential medium (MEM), Dulbecco’s modified Eagle medium (DMEM), Glasgow modified Eagle’s medium (GMEM), and RPMI 1640 and Ham’s F10 nutrient mixtures (e.g., Life Technologies) are composed of amino acids, glucose, salts, vitamins, and other nutrients. A basic medium is supplemented by addition of L-glutamine, antibiotics (typically penicillin and streptomycin sulfate), and usually serum to formulate a “complete medium.” Where serum is added, the amount is indicated as a percentage of fetal bovine serum (FBS) or other serum. Some media are also supplemented with antimycotics, nonessential amino acids, various growth factors, and/or drugs that provide selective growth conditions. Supplements should be added to medium prior to sterilization or filtration, or added aseptically just before use. The optimum pH for most mammalian cell cultures is 7.2 to 7.4. Adjust the medium’s pH as necessary after all supplements are added. Buffers such as bicarbonate and HEPES are routinely used in tissue culture medium to prevent fluctuations in pH that might adversely affect cell growth. HEPES is especially useful in solutions used for procedures that do not take place in a controlled-CO2 environment. Most cultured cells will tolerate a wide range of osmotic pressure and an osmolarity between 260 and 320 mOsm/kg is acceptable for most cells. The osmolarity of human plasma is ∼290 mOsm/kg, and this is probably the optimum for human cells in culture as well (Freshney, 1993). Fetal bovine serum (FBS) is the most frequently used serum supplement. Calf serum, horse serum, and human serum are also used; some cell lines are maintained in serum-free medium (Freshney, 1993). Complete medium is supplemented with 5% to 30% (v/v) serum, depending on the requirements of the particular cell type being cultured. Serum Useful Techniques


Supplement 2

Table A.3B.1 Working Concentrations of Antibiotics and Fungicides for Mammalian Cell Culture

Additive

Final concentration

Penicillin Streptomycin sulfate Kanamycin Gentamycin Mycostatin Amphotericin B

50–100 U/ml 50–100 µg/ml 100 µg/ml 50 µg/ml 20 µg/ml 0.25 µg/ml

that has been heat inactivated (30 min to 1 hr at 56°C; see recipe) is generally preferred, because this inactivates complement and is thought to reduce the number of contaminants. Serum is obtained frozen, then is thawed, divided into smaller portions, and refrozen until needed. There is considerable lot-to-lot variation in FBS. Most suppliers will provide a sample of a specific lot and reserve a supply of that lot while the serum is tested for its suitability. The suitability of a serum lot depends upon the use. Frequently the ability of serum to promote cell growth equivalent to a laboratory standard is used to evaluate a serum lot. Once an acceptable lot is identified, enough of that lot should be purchased to meet the culture needs of the laboratory for an extended period of time. Commercially prepared media containing L-glutamine are available, but many laboratories choose to obtain medium without L-glutamine and then add it to a final concentration of 2 mM just before use. L-glutamine is an unstable amino acid that, upon storage, converts to a form cells cannot use. Breakdown of L-glutamine is temperature and pH dependent. At 4°C, 80% of the L-glutamine remains after 3 weeks, but at incubator temperature (35°C) the amount diminishes more rapidly (Brown and Lawce, 1997). To prevent degradation, 100× L-glutamine (see recipe) should be stored frozen in aliquots until needed. As well as practicing good aseptic technique, most laboratories add antimicrobial agents to medium to further reduce the risk of contamination. A combination of penicillin and streptomycin is the most commonly used antibiotic additive; kanamycin and gentamycin are used alone. Mycostatin and amphotericin B are the most commonly used fungicides (Rooney and Czepulkowski, 1992). Table A.3B.1 lists suggested working concentrations for the most commonly used antibiotics and antimycotics. Combining antibiotics in tissue culture medium can be tricky, as some antibiotics are not compatible, and one may inhibit the action of another. Furthermore, combined antibiotics may be cytotoxic at lower concentrations than is true for the individual antibiotics. In addition, prolonged use of antibiotics may cause cell lines to develop antibiotic resistance. For this reason, some laboratories add antibiotics and/or fungicides to medium when initially establishing a culture but eliminate them from medium used in later subcultures.


All tissue culture medium, whether prepared commercially or within the laboratory, should be tested for sterility prior to use. A small aliquot from each lot of medium is incubated 48 hr at 37°C and monitored for evidence of contamination such as turbidity (infected medium will be cloudy) and color change (if phenol red is the indicator, infected medium will turn yellow). Any contaminated medium should be discarded.

A.3B.4 Supplement 2


TRYPSINIZING AND SUBCULTURING CELLS FROM A MONOLAYER 2

A primary culture is grown to confluency in a 60-mm petri dish or 25-cm tissue culture flask containing 5 ml tissue culture medium. Cells are dispersed by trypsin treatment and then reseeded into secondary cultures. The process of removing cells from the primary culture and transferring them to secondary cultures constitutes a passage or subculture.

BASIC PROTOCOL

Materials Cells cultures HBSS without Ca2+ and Mg2+ (APPENDIX 2A), 37°C 0.25% (w/v) trypsin/0.2% EDTA solution (see recipe), 37°C Complete medium with serum: e.g., DMEM supplemented with 10% to 15% (v/v) FBS (complete DMEM-10; see recipe), 37°C Sterile Pasteur pipets 37°C warming tray or incubator Tissue culture plasticware or glassware including pipets and 25-cm2 flasks or 60-mm petri dishes, sterile NOTE: All incubations should be performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. 1. Remove all medium from culture with a sterile Pasteur pipet. Wash adhering cell monolayer once or twice with a small volume of 37°C HBSS without Ca2+ and Mg2+ to remove any residual FBS, which may inhibit the action of trypsin. Use a buffered salt solution that is Ca2+- and Mg2+-free to wash cells. Ca2+ and Mg2+ in the salt solution can cause cells to stick together. If this is the first medium change from primary culture, rather than discarding medium that is removed from primary culture, transfer it to a fresh dish or flask. The medium contains unattached cells that may attach and grow, thereby providing a backup culture.

2. Add enough 37°C trypsin/EDTA solution to culture to cover adhering cell layer. 3. Place plate on a 37°C warming tray 1 to 2 min. Tap bottom of plate on the countertop to dislodge cells. Check culture with an inverted microscope to be sure that cells are rounded up and detached from the surface. If cells are not sufficiently detached, return plate to warming tray for an additional minute or two.

4. Add 2 ml of 37°C complete medium. Draw cell suspension into a Pasteur pipet and rinse cell layer two or three times to dissociate cells and to dislodge any remaining adherent cells. As soon as cells are detached, add serum or medium containing serum to inhibit further trypsin activity that might damage cells. If cultures are to be split 1/3 or 1/4 rather than 1/2, add sufficient medium such that 1 ml of cell suspension can be transferred into each fresh culture vessel.

5. Add an equal volume of cell suspension to fresh dishes or flasks that have been appropriately labeled. Alternatively, cells can be counted using a hemacytometer (see Support Protocol 4) or Coulter counter and diluted to the desired density so a specific number of cells can be added to each culture vessel. A final concentration of ∼5 × 104 cells/ml is appropriate for most subcultures. For primary cultures and early subcultures, 60-mm petri dishes or 25-cm2 flasks are generally used; larger petri dishes or flasks (e.g., 150-mm dishes or 75-cm2 flasks) may be used for later subcultures. Cultures should be labeled with cell line identification, lab number, date of subculture, and passage number. Useful Techniques


Supplement 2

6. Add 4 ml fresh medium to each new culture. Incubate in a humidified 37°C, 5% CO2 incubator. If using 75-cm2 culture flasks, add 9 ml medium per flask. Some labs now use incubators with 5% CO2 and 4% O2. The low oxygen concentration is thought to simulate the in vivo environment of cells and to enhance cell growth. However, some media (e.g., DMEM) may require altered levels of CO2 to maintain pH.

7. If necessary, feed subconfluent cultures after 3 or 4 days by removing old medium and adding fresh 37°C medium. 8. Passage secondary culture when it becomes confluent by repeating steps 1 to 7, and continue to passage as necessary. ALTERNATE PROTOCOL

PASSAGING CELLS IN SUSPENSION CULTURE A suspension culture is grown in culture flasks in a humidified 37°C, 5% CO2 incubator. Passaging of suspension cultures is somewhat less complicated than passaging of monolayer cultures. Because the cells are suspended in medium rather than attached to a surface, it is not necessary to disperse them enzymatically before passaging. However, before passaging, cells must be maintained in culture by feeding every 2 to 3 days until they reach confluency (i.e., until the cells clump together in the suspension and the medium appears turbid when the flask is swirled). NOTE: All culture incubations should be performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. 1. Feed cells as follows every 2 to 3 days until the cultures are confluent: a. Remove flask of suspension cells from incubator, taking care not to disturb those that have settled to the flask bottom. b. Aseptically remove and discard about one-third of the medium from flask and replace with an equal volume of prewarmed (37°C) medium. If the cells are growing rapidly, add an additional 10% medium by volume in order to maintain optimum concentration of 1 × 106 cells/ml. Gently swirl flask to resuspend cells. c. Return flask to incubator. If there is 1 kb, and as little as a 2-sec pause for targets 40) can reduce the polymerase specific activity, increase nonspecific amplification, and deplete substrate (nucleotides). Many investigators lengthen the time for the last extension step—to 7 min, for example—to try to ensure that all the PCR products are full length. These guidelines are appropriate for most commercially available thermal cyclers. For rapid cyclers, consult the manufacturers’ protocols.

Analyze the product 5. Electrophorese 10 µl from each reaction on an agarose, nondenaturing polyacrylamide, or sieving agarose gel appropriate for the PCR product size expected. Stain with ethidium bromide. For resolution of PCR products between 100 and 1000 bp, an alternative to nondenaturing polyacrylamide gels or sieving agarose is a composite 3% (w/v) NuSieve (FMC Bioproducts) agarose/1% (w/v) SeaKem (FMC Bioproducts) agarose gel. SeaKem increases the mechanical strength of the gel without decreasing resolution. An alternative to ethidium bromide, SYBR Gold Nucleic Acid Gel Stain (Molecular Probes), is 25 to 100 times more sensitive than ethidium bromide, is more convenient to use, and permits optimization of 10- to 100-fold lower starting template copy number.

6. Examine the stained gel to determine which condition resulted in the greatest amount of product. Minor, nonspecific products may be present even under optimal conditions.

Useful Techniques

A.3C.3 Current Protocols in Toxicology

Supplement 3

Table A.3C.2

Master Mixes for Optimizing First-Cycle Reactions

Components 10× PCR buffer MgCl2 (L, M, or H) Primer 1 Primer 2 Additive Template DNA 25 mM 4dNTP mixb Taq polymerase Taq pol + TaqStart H 2O Preparation temperature

Final concentration 1× Optimal 0.5 µM 0.5 µM Optimal —b 0.2 mM 2.5 U 2.5 U To 100 µl

Master mix (µl) A 10 10 1.0 1.0 Va Va 0.8 0.5 — Va Room temperature

B 10 10 1.0 1.0 Va Va 0.8 0.5 — Va Ice slurry

C 10 10 1.0 1.0 Va Va 0.8 — — Va Room temperature

D 10 10 1.0 1.0 Va Va 0.8 — 1.0 Va Room temperature

aV, variable amount (total volume should be 100 µl). bUse undiluted or diluted template DNA based on results obtained in step 6.

7. To ensure that the major product is the correct one, digest an aliquot of the reaction with a restriction endonuclease known to cut within the PCR product. Check buffer compatibility for the restriction endonuclease of choice. If necessary, add Na+ or precipitate in ethanol (APPENDIX 3), then resuspend in the appropriate buffer. Electrophorese the digestion product on a gel to verify that the resulting fragments have the expected sizes. Alternatively, transfer the PCR products to a nitrocellulose or nylon filter and hybridize with an oligonucleotide derived from the sequence internal to the primers. With appropriately stringent hybridization and washing conditions, only the correct product (and possibly some minor related products) should hybridize.

Optimize the first cycle These optional steps optimize initial hybridization and may improve efficiency and yield. They are used when primer-dimers and other nonspecific products are detected, when there is only a very small amount of starting template, or when a rare sequence is to be amplified from a complex mixture. For an optimal reaction, polymerization during the initial denaturation and annealing steps should be prevented. Taq DNA polymerase activity can be inhibited by temperature (reaction B), physical separation (reaction C), or reversible antibody binding (reaction D). PCR without hot start is performed for comparison (reaction A). 8. Prepare four reaction mixtures using the optimal MgCl2 concentration and additive requirement determined in step 6. Prepare the mixes according to the recipes in Table A.3C.2. Use the following variations for addition of Taq polymerase.

Enzymatic Amplification of DNA by PCR

a. Prepare reactions A and C at room temperature. b. Chill all components of reaction B in an ice slurry before they are combined. c. For reaction D, combine 1.0 µl TaqStart antibody with 4.0 µl of the dilution buffer provided with the antibody, add 1.0 µl Taq DNA polymerase (for 1:4:1 mixture of these components), mix, and incubate 5 to 10 min at room temperature before adding to reaction mixture D (glycerol and PMPE are compatible with TaqStart antibody but DMSO will interfere with antibody binding).

A.3C.4 Supplement 3


To ensure that the reaction does not plateau and thereby obfuscate the results, use the smallest amount of template DNA necessary for visualization of the PCR product by ethidium bromide staining. Use the results from step 6 to decide how much template to use. If the desired product stains intensely, dilute the starting material as much as 1/100. If only a faint signal is apparent, use undiluted sample.

9. Overlay each reaction mixture with 50 to 100 µl mineral oil. 10. Heat all reactions 5 min at 94°C. It is most convenient to use the automated thermal cycler for this step and then initiate the cycling program directly.

11. Cool the reactions to the appropriate annealing temperature as determined in step 4. Add 0.5 µl Taq DNA polymerase to reaction C, making sure the pipet tip is inserted through the layer of mineral oil into the reaction mix. Time is also an important factor in this step. If the temperature drops below the annealing temperature and is allowed to remain low, nonspecific annealing will occur. Taq DNA polymerase retains some activity even at room temperature.

12. Begin amplification of all four reactions at once, using the same cycling parameters as before. 13. Analyze the PCR products on an agarose gel and evaluate the results as in steps 5 and 6. 14. Prepare a batch of the optimized reaction mixture, but omit Taq DNA polymerase, TaqStart antibody, PMPE, and 4dNTP mix—these ingredients should be added fresh just prior to use. If desired, add Ficoll 400 to a final concentration of 0.5% to 1% (v/v) and tartrazine to a final concentration of 1 mM. Adding Ficoll 400 and tartrazine dye to the reaction mix precludes the need for a gel loading buffer and permits direct application of PCR products to agarose or acrylamide gels. At these concentrations, Ficoll 400 and tartrazine do not decrease PCR efficiency and do not interfere with PMPE or TaqStart antibodies. Other dyes, such as bromphenol blue and xylene cyanol, do inhibit PCR. Tartrazine is a yellow dye and is not as easily visualized as other dyes; this may make gel loading more difficult. Ficoll 400 and tartrazine dye may be prepared as 10× stocks and stored indefinitely at room temperature.

REAGENTS AND SOLUTIONS Use deionized, distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.

Enhancer agents For a discussion of how to select enhancer agents, see Commentary. 5× stocks: 25% acetamide (20 µl/reaction; 5% final) 5 M N,N,N-trimethylglycine (betaine; 20 µl/reaction; 1 M final) 40% polyethylene glycol (PEG) 8000 (20 µl/reaction; 8% final) 10× stocks: Glycerol (concentrated; 10 µl/reaction; 10% final) 20× stocks: Dimethylsulfoxide (DMSO; concentrated 5 µl/reaction; 5% final) Formamide (concentrated; 5 µl/reaction; 5% final) Useful Techniques


Supplement 3

100× stocks: 1 U/µl Perfect Match Polymerase Enhancer [Strategene; 1 µl (1 U) per reaction, final] 10 mg/ml acetylated bovine serum albumin (BSA) or gelatin (1 µl/reaction; 10 µg/ml final) 1 to 5 U/µl thermostable pyrophosphatase [PPase; Boehringer Mannheim; 1 µl (1 to 5 U) per reaction, final] 5 M tetramethylammonium chloride (TMAC; betaine hydrochloride; 1 µl/reaction; 50 mM final) 0.5 mg/ml E. coli single-stranded DNA-binding protein (SSB; Sigma; 1 µl/reaction; 5 µg/ml final) 0.5 mg/ml Gene 32 protein (Amersham Pharmacia Biotech; 1 µl/reaction; 5 µg/ml final) 10% Tween 20, Triton X-100, or Nonidet P-40 (1 µl/reaction; 0.1% final) 1 M (NH4)2SO4 (1 µl/reaction; 10 mM final; use with thermostable DNA polymerases other than Taq) MgCl2-free PCR amplification buffer, 10× 500 mM KCl 100 mM Tris⋅Cl, pH 9.0 (at 25°C) 0.1% Triton X-100 Store indefinitely at −20°C This buffer can be obtained from Promega; it is supplied with Taq DNA polymerase.

4dNTP mix For 2 mM 4dNTP mix: Prepare 2 mM each dNTP in TE buffer, pH 7.5 (see recipe). Store up to 1 year at −20°C in 1-ml aliquots. For 25 mM 4dNTP mix: Combine equal volumes of 100 mM dNTPs (Promega). Store indefinitely at −20°C in 1-ml aliquots. TE (Tris/EDTA) buffer, pH 7.5 10 mM Tris⋅Cl, pH 7.5 (APPENDIX 2A) 1 mM EDTA, pH 8.0 (APPENDIX 2A) Store up to 6 months at room temperature COMMENTARY Background Information


The theoretical basis of the polymerase chain reaction (PCR) was probably first described in a paper by Kleppe et al. (1971). However, this technique did not excite general interest until the mid-1980s, when Kary Mullis and co-workers at Cetus developed PCR into a technique that could be used to generate large amounts of single-copy genes from genomic DNA (Saiki et al., 1985, 1986; Mullis et al., 1986; Embury et al., 1987). The initial procedure entailed adding a fresh aliquot of the Klenow fragment of E. coli DNA polymerase I during each cycle because this enzyme was inactivated during the subsequent denaturation step. The introduction of thermostable Taq DNA polymerase from Thermus aquaticus (Saiki et al., 1988) alleviated this

tedium and facilitated automation of the thermal cycling portion of the procedure. Taq DNA polymerase also permitted the use of higher temperatures for annealing and extension, which improved the stringency of primer–template hybridization and thus the specificity of the products. This also served to increase the yield of the desired product. All applications for PCR depend upon an optimized PCR. The basic protocol in this unit optimizes PCR for several variables, including MgCl2 concentration, enhancing additives— dimethyl sulfoxide (DMSO), glycerol, or Perfect Match Polymerase Enchancer (PMPE)— and prevention of pre-PCR mispriming. These and other parameters can be extremely important, as every element of PCR can affect the outcome; see Critical Parameters and Trou-

A.3C.6 Supplement 3


bleshooting for discussion of individual parameters. There are several PCR optimization kits and proprietary enhancers on the market (Table A.3C.3). Optimization kits generally provide a panel of buffers in which the pH, buffer, nonionic detergents, and addition of (NH4)2SO4 are varied, MgCl2 may be added at several concentrations, and enhancers (e.g., DMSO, glycerol, formamide, betaine, and/or proprietary compounds) may be chosen. The protocol presented here is aimed at keeping the costs low and the options broad.

Critical Parameters and Troubleshooting MgCl2 concentration Determining the optimum MgCl2 concentration, which can vary even for different primers from the same region of a given template (Saiki, 1989), can have an enormous influence on PCR success. In this protocol three test concentrations are suggested—1.5 mM (L), 3.0 mM (M), and 4.5 mM (H). If further optimization is necessary, the MgCl2 range can be extended or narrowed around the most successful concentration. A 10× buffer optimized for a given enzyme and a separate vial of MgCl2 are typically provided with the polymerase, so that the user may titrate the MgCl2 concentration for their unique primer-template set. Note that some enhancers may broaden the MgCl2 optimal range. Reagent purity For applications that amplify rare templates, reagent purity is the most important parameter, and avoiding contamination at every step is critical. To maintain purity, store multiple small volumes of each reagent in screw-cap tubes. For many applications, simply using highquality reagents and avoiding nuclease contamination is sufficient. However, avoid one common reagent used to inactivate nucleases— diethylpyrocarbonate (DEPC). Even the tiny amounts of chemical left after treatment of water and autoclaving are enough to ruin a PCR. Primer selection This is the factor that is least predictable and most difficult to troubleshoot. Simply put, some primers just do not work. To maximize the probability that a given primer pair will work, pay attention to the following parameters.

General considerations. An optimal primer set should hybridize efficiently to the sequence of interest with negligible hybridization to other sequences present in the sample. If there are reasonable amounts of template available, hybridization specificity can be tested by performing oligonucleotide hybridization. The distance between the primers is rather flexible, ranging up to 10 kb. There is, however, a considerable drop-off in synthesis efficiency with distances >3 kb (Jeffreys et al., 1988). Small distances between primers, however, lessen the ability to obtain much sequence information or to reamplify with nested internal oligonucleotides, should that be necessary. Design primers to allow demonstration of the specificity of the PCR product. Be sure that there are diagnostic restriction endonuclease sites between the primers or that an oligonucleotide can detect the PCR product specifically by hybridization. Several computer programs can assist in primer design. These are most useful for avoiding primer sets with intra- and intermolecular complementarity, which can dramatically raise the effective melting temperature (Tm). Given the abundance of primers relative to template, this can preclude template priming. Computer primer design is not foolproof. If possible, start with a primer or primer set known to efficiently prime extensions. In addition, manufacturers’ Web sites offer technical help with primer design. Complementarity to template. For many applications, primers are designed to be exactly complementary to the template. For others, however, such as engineering of mutations or new restriction endonuclease sites (UNIT 8.5), or for efforts to clone or detect gene homologs where sequence information is lacking, basepair mismatches will be intentionally or unavoidably created. It is best to have mismatches (e.g., in a restriction endonuclease linker) at the 5′ end of the primer. The closer a mismatch is to the 3′ end of the primer, the more likely it is to prevent extension. If cloned template is available, primers can be checked for suitability by using them in a sequencing reaction with Taq DNA polymerase. The use of degenerate oligonucleotide primers to clone genes where only protein sequence is available, or to fish out gene homologs in other species, has sometimes been successful— but it has also failed an untold (and unpublished) number of times. When the reaction works it can be extremely valuable, but it can Useful Techniques


Supplement 6

Table A.3C.3

PCR Optimization Products

Optimization goal

Supplier

Product

Optimization support Optimization support

PE Biosystems Promega

Optimization kits

Boehringer Mannheim, Invitrogen, Stratagene, Sigma, Epicentre Technologies, Life Technologies Amersham Pharmacia Biotech

Technical information in appendix to catalog PCR troubleshooting program on the Internet: http://www.promega.com/amplification/assistant Several buffers, Mg2+, and enhancers which may include DMSO, glycerol, formamide, (NH4)2SO4, and other unspecified or proprietary agents

Quick startup

Quick startup

Fisher

Quick startup

Life Technologies

Quick startup

Marsh Biomedical

Hot-start/physical barrier Fisher, Life Technologies

Hot-start/separate MgCl2 Invitrogen Hot-start/separate MgCl2 Stratagene Hot Start/separate polymerase

Promega

Hot-start/reversible inactivation of polymerase by antibody binding Hot-start/antibody binding Hot-start/antibody binding Hot-start/reversible chemical modification Hot-start/reversible chemical modification Enhancer

Clontech

Ready-To-Go Beads “optimized for standard PCR” and Ready-To-Go RAPD Analysis Beads (buffer, nucleotides, Taq DNA polymerase) EasyStart PCR Mix-in-a-Tube—tubes prepackaged with wax beads containing buffer, MgCl2, nucleotides, Taq DNA polymerase PCR SuperMix—1.1× conc.—premix containing buffer, MgCl2, nucleotides, Taq DNA polymerase Advanced Biochemicals Red Hot DNA Polymerase—a new rival for Taq polymerase with convenience features Molecular Bio-Products HotStart Storage and Reaction Tubes—preadhered wax bead in each tube; requires manual addition of one component at high temperature HotWax Mg2+ beads—wax beads contain preformulated MgCl2 which is released at first elevated-temperature step StrataSphere Magnesium Wax Beads—wax beads containing preformulated Mg2+ TaqBead Hot Start Polymerase—wax beads encapsulating Taq DNA polymerase which is released at first elevated-temperature step TaqStart Antibody, TthStart Antibody—reversibly inactivate Taq and Tth DNA polymerases until first denaturation at 95°C

Life Technologies

PlatinumTaq—contains PlatinumTaq antibody

Sigma

JumpStart Taq—contains TaqStart antibody

PE Biosystems

AmpliTaq Gold—activated at high temperature

Qiagen

HotStarTaq DNA Polymerase—activated at high temperature Tth pyrophosphatase, thermostable

Enhancer Enhancer

Boehringer Mannheim, New England Biolabs Clontech CPG

Enhancer

Fisher

Enhancer Enhancer Enhancer Enhancer Enhancer

Life Technologies Promega Qiagen Stratagene Stratagene

GC-Melt (in Advantage-GC Kits)—proprietary Taq-FORCE Amplification System and MIGHTY Buffer—proprietary Eppendorf MasterTaq Kit with TaqMaster Enhancer—proprietary PCRx Enhancer System—proprietary E.coli Single Stranded Binding Protein (SSB) Q-Solution—proprietary Perfect Match Polymerase Enhancer—proprietary TaqExtender PCR Additive—proprietary

A.3C.8 Supplement 6


also generate seemingly specific products that require much labor to identify and yield no useful information. The less degenerate the oligonucleotides, especially at the 3′ end, the better. Caveat emptor. Primer length. A primer should be 20 to 30 bases in length. It is unlikely that longer primers will help increase specificity significantly. Primer sequence. Design primers with a GC content similar to that of the template. Avoid primers with unusual sequence distributions, such as stretches of polypurines or polypyrimidines as their secondary structure can be disastrous. It is worthwhile to check for potential secondary structure using one of the appropriate computer programs that are available. “Primer-dimers.” Primer-dimers are a common artifact most frequently observed when small amounts of template are taken through many amplification cycles. They form when the 3′ end of one primer anneals to the 3′ end of the other primer, and polymerase then extends each primer to the end of the other. The ensuing product can compete very effectively against the PCR product of interest. Primer-dimers can best be avoided by using primers without complementarity, especially in their 3′ ends. Should they occur, optimizing the MgCl2 concentration may minimize their abundance relative to that of the product of interest. Template Aside from standard methods for preparing DNA (APPENDIX 3), a number of simple and rapid procedures have been developed for particular tissues (Higuchi, 1989). Even relatively degraded DNA preparations can serve as useful templates for generation of moderate-sized PCR products. The two main concerns regarding template are purity and amount. A number of contaminants found in DNA preparations can decrease the efficiency of PCR. These include urea, the detergent SDS (whose inhibitory action can be reversed by nonionic detergents), sodium acetate, and, sometimes, components carried over in purifying DNA from agarose gels (Gelfand, 1989; Gyllensten, 1989; K. Hicks and D. Coen, unpub. observ.). Additional organic extractions, ethanol precipitation from 2.5 M ammonium acetate, and/or gel purification on polyacrylamide rather than agarose, can all be beneficial in minimizing such contamination if the simplest method (precipitating the sample with ethanol and repeatedly washing the pellet with 70% ethanol) is not sufficient.

Clearly the amount of template must be sufficient to be able to visualize PCR products using ethidium bromide. Usually 100 ng of genomic DNA is sufficient to detect a PCR product from a single-copy mammalian gene. Using too much template is not advisable when optimizing for MgCl2 or other parameters, as it may obscure differences in amplification efficiency. Moreover, too much template may decrease efficiency due to contaminants in the DNA preparation. Amount of template, especially in terms of the amount of target sequence versus nonspecific sequences, can have a major effect on the yield of nonspecific products. With less target sequence, it is more likely that nonspecific products will be seen. For some applications, such as certain DNA sequencing protocols where it is important to have a single product, gel purification of the specific PCR product and reamplification are advisable. Taq and other thermostable DNA polymerases Among the advantages conferred by the thermostability of Taq DNA polymerase is its ability to withstand the repeated heating and cooling inherent in PCR and to synthesize DNA at high temperatures that melt out mismatched primers and regions of local secondary structure. The enzyme, however, is not infinitely resistant to heat, and for greatest efficiency it should not be put through unnecessary denaturation steps. Indeed, some protocols (e.g., the “hot start” method described here) recommend adding it after the first denaturation step. Increasing the amount of Taq DNA polymerase beyond 2.5 U/reaction can sometimes increase PCR efficiency, but only up to a point. Adding more enzyme can sometimes increase the yield of nonspecific PCR products at the expense of the product of interest. Moreover, Taq DNA polymerase is not inexpensive. A very important property of Taq DNA polymerase is its error rate, which was initially estimated at 2 × 10−4 nucleotides/cycle (Saiki et al., 1988). The purified enzyme supplied by manufacturers lacks a proofreading 3′→5′ exonuclease activity, which lowers error rates of other polymerases such as the Klenow fragment of E. coli DNA polymerase I. For many applications, this does not present any difficulties. However, for sequencing clones derived from PCR, or when starting with very few templates, this can lead to major problems. Direct sequencing of PCR products, sequencing numerous PCR-generated clones, and/or

Useful Techniques


Supplement 3

the use of appropriate negative controls can help overcome these problems. Alternatively, changing reaction conditions (Eckert and Kunkel, 1990) or changing to a non–Taq DNA polymerase (with greater fidelity) may be useful. Another important property of Taq DNA polymerase is its propensity for adding nontemplated nucleotides to the 3′ ends of DNA chains. This can be especially problematic in cloning PCR products. It is frequently necessary to “polish” PCR products with enzymes such as other DNA polymerases before adding linkers or proceeding to blunt-end cloning. Conversely, addition of a nontemplated A by Taq DNA polymerase can be advantageous in cloning. Table A.3C.4 lists currently available thermostable DNA polymerases by generic and trade names, the original source of native and recombinant enzymes, the supplier, the end generated (3′A addition versus blunt), and associated exonuclease activities. A 3′ to 5′ exonuclease activity is proofreading. Removal of the 5′ to 3′ exonuclease activity of Taq DNA polymerase (N-terminal deletion) is reported to produce a higher yield. A 5′ to 3′ exonuclease activity may degrade the primers somewhat. Proofreading enzymes synthesize DNA with higher fidelity and can generate longer products than Taq, but tend to generate low yields. Enzyme blends (Table A.3C.5) have been optimized for increased fidelity and length along with sensitivity and yield.


Hot start What happens prior to thermal cycling is critical to the success of PCR. Taq DNA polymerase retains some activity even at room temperature. Therefore, under nonstringent annealing conditions, such as at room temperature, products can be generated from annealing of primers to target DNA at locations of low complementarity or having complementarity of just a few nucleotides at the 3′ ends. The latter would in effect create new templates “tagged” with the primer sequences. Subsequent cycles amplify these tagged sequences in abundance, both generating nonspecific products and possibly reducing amplification efficiency of specific products by competition for substrates or polymerase. Thus conditions preventing polymerization prior to the first temperature-controlled steps are desirable. In this protocol, three methods of inhibiting polymerization prior to the temperature-controlled step are compared. These include physical separation of an essen-

tial reaction component prior to the first denaturation step, cooling reagents to 0°C, and reversibly blocking enzymatic activity with an antibody. Denaturation of the template before Taq polymerase or MgCl2 is added to the reaction provides a dramatic improvement in specificity and sensitivity in many cases (Chou et al., 1992). The main drawback of this method is that it requires opening the reaction tubes a second time to add the essential missing component. This creates both an inconvenience and an increase in the risk of contamination, an important consideration when testing for the presence of a given sequence in experimental or clinical samples. Cooling all components of the reaction mixture to 0°C prior to mixing is more convenient and the least expensive method but is also the least reliable. Transferring the PCR reaction tubes from the ice slurry to a 95°C preheated thermocycler block may improve the chance of success. Reversible inhibition of Taq DNA polymerase by TaqStart antibody (Clontech) is the most convenient and is very effective (Kellogg et al., 1994). Complete reactions can be set up, overlaid with oil, and stored at 4°C for up to several hours prior to thermal cycling with no loss of sensitivity or specificity compared to the other hot-start methods (M.F.K. and D.M.C., unpub. observ.). Cycling is initiated immediately following 5-min denaturation of the antibody at 94°C. DMSO inhibits antibody binding and should not be used with TaqStart. Several hot-start products are now commercially available (Table A.3C.3). Success with each may depend on strict adherence to the manufacturer’s protocols, even on a specific thermocycler. Wax barrier and reversible antibody binding methods are more forgiving, while chemical modifications have more stringent activation temperature requirements. Deoxyribonucleoside triphosphates In an effort to increase efficiency of PCR, it may be tempting to increase the concentration of dNTPs. Don’t. When each dNTP is 200 µM, there is enough to synthesize 12.5 µg of DNA when half the dNTPs are incorporated. dNTPs chelate magnesium and thereby change the effective optimal magnesium concentration. Moreover, dNTP concentrations >200 µM each increase the error rate of the polymerase. Millimolar concentrations of dNTPs actually inhibit Taq DNA polymerase (Gelfand, 1989).

A.3C.10 Supplement 3


Table A.3C.4

Thermostable DNA Polymerases

DNA polymerase Generic name

Trade name

Pfu Pfu (exo-) Psp Psp (exo-)

— — Deep Vent Deep Vent (exo-)

Pwo Taq (native and/or recombinant)

—

Taq, N-terminal Stoffel deletion fragment Klen-Taq Tbr DyNAzyme Tfl Tli

Vent

Tli (exo-)

Vent (exo-)

Tma Tth

UlTma —

Biological source

Supplier

Product ends

Exonuclease activity

Pyrococcus furiosus Pyrococcus furiosus Pyrococcus sp.GB-D Pyrococcus sp.GB-D

Stratagene, Promega Stratagene New England Biolabs New England Biolabs

Blunt Blunt Blunt Blunt

3′-5′ (proofreading) No 3′-5′ (proofreading) No

Pyrococcus woesei Thermus aquaticus

Boehringer Mannheim Blunt 3′A Ambion, Amersham Pharmacia Biotech, Boehringer Mannheim, Clontech, Fisher, Life Technologies, Marsh Biomedical, PE Biosystems, Promega, Qiagen, Sigma, Stratagene PE Biosystems, Sigma 3′A

3′-5′ (proofreading) 5′-3′

Thermus aquaticus

Thermus brocianus Thermus flavus

MJ Research Promega, Epicentre Technologies New England Biolabs (Vent), Promega New England Biolabs

Thermococcus litoralis Thermococcus litoralis Thermotoga maritima PE Biosystems Thermus thermophilus Amersham Pharmacia Biotech, Boehringer Mannheim, Epicentre Technologies, PE Biosystems, Promega

No

—a Blunt

5′-3′ —a

Blunt

3′-5′ (proofreading)

Blunt

No

Blunt 3′ A

3′-5′ (proofreading) 5′-3′

aNo information at this time.

The protocol in this unit calls for preparing 4dNTPs in 10 mM Tris⋅Cl/1 mM EDTA (TE buffer), pH 7.4 to 7.5. This is easier and less prone to disaster than neutralization with sodium hydroxide. However, EDTA also chelates magnesium, and this should be taken into account if stocks of dNTPs are changed. Alternatively, to lower the risk of contamination, a 4dNTP mix can be made by combining equal volumes of commercially prepared stocks. Enhancers Enhancers are used to increase yield and specificity and to overcome difficulties encountered with high GC content or long templates. Nonionic detergents (Triton X-100, Tween 20, or Nonidet P-40) neutralize charges

of ionic detergents often used in template preparation, and should be used in the basic reaction mixture, rather than as optional enhancers. Higher yields can be achieved by stabilizing/enhancing the polymerase activity with enzyme-stabilizing proteins (BSA or gelatin), enzyme-stabilizing solutes such as betaine or betaine⋅HCl (TMAC), enzyme-stabilizing solvents (glycerol), solubility-enhancing solvents (DMSO or acetamide), molecular crowding solvents (PEG), and polymerase salt preferences [(NH4)SO4 is recommended for polymerases other than Taq]. Greater specificity can be achieved by lowering the TM of dsDNA (using formamide), destabilizing mismatchedprimer annealing (using PMPE or hot-start strategies), and stabilizing ssDNA (using E.

Useful Techniques


Supplement 3

Table A.3C.5

Thermostable DNA Polymerase Blends

Product (trade name)

Supplier

Thermostable DNA polymerases and other components

Expand High Fidelity, Expand Long Template, Boehringer Mannheim Taq + Pwo and Expand 20kb PCR Systems KlenTaq LA Polymerase Mix

Clontech, Sigma

Advantage-HF PCR Kit

Clontech

Advantage-cDNA and Advantage-GC cDNA Polymerase Mixes and Kits

Clontech

Advantage Genomic and Advantage-GC Genomic Polymerase Mixes and Kits

Clontech

Tth + unspecified proofreading polymerase + TthStart Antibody; GC Kit contains GC Melt

eLONGase Enzyme Mix

Life Technologies

Platinum Taq DNA Polymerase

Life Technologies

Taq + Psp + unspecified proofreading polymerase(s) + eLONGase Buffer Taq + Psp + Platinum Taq Antibody

Platinum High Fidelity DNA Polymerase DyNAzyme EXT Polymerase

Life Technologies MJ Research

Taq + Psp + Taq Antibody Tbr with unspecified enhancer

GeneAmp XL PCR and XL RNA PCR Kits OmniBase Sequencing Enzyme Mix

PE Biosystems Promega

Tth + Tli Unspecified proofreading polymerase(s) with thermostable pyrophosphatase

AccuTaq LA DNA Polymerase Mix Sigma TaqPlus Long and TaqPlus Precision PCR Stratagene Systems Accurase Fidelity PCR Enzyme Mix; Calypso Tetralink High Fidelity Single Tube RT-PCR System


coli SSB or Gene 32 Protein). Amplification of high-GC-content templates can be improved by decreasing the base pair composition dependence of the TM of dsDNA (with betaine; Rees et al., 1993). Betaine is an osmolyte widely distributed in plants and animals and is nontoxic, a feature that recommends it for convenience in handling, storage, and disposal. Betaine may be the proprietary ingredient in various commercial formulations. For long templates, a higher pH is recommended (pH 9.0). The pH of Tris buffer decreases at high temperatures, long-template PCR requires more time at high temperatures, and increased time at lower pH may cause some depurination of the template, resulting in reduced yield of specific product. Inorganic phosphate (PPi), a product of DNA synthesis, may accumulate with amplification of long products to levels that may favor reversal of polymerization. Accumulation of PPi may be prevented by addi-

KlenTaq-1 (5′-exonuclease deficient Taq) + unspecified proofreading polymerase KlenTaq-1 + unspecified proofreading polymerase + TaqStart Antibody KlenTaq-1 + unspecified proofreading polymerase + TaqStart Antibody; GC Kit contains GC Melt

Taq + unspecified proofreading polymerase Pfu + Taq; TaqPlus Precision Reaction Buffer (proprietary) Thermus sp. + Thermococcus sp.; Calypso also contains AMV-RT

tion of thermostable PPase. When large numbers of samples are being analyzed, the convenience of adding PCR products directly to a gel represents a significant time savings. Some companies combine their thermostable polymerase with a red dye and a high density component to facilitate loading of reaction products onto gels without further addition of loading buffer. Thermal cycling parameters Each step in the cycle requires a minimal amount of time to be effective, while too much time can be both wasteful and deleterious to the DNA polymerase. If the amount of time in each step can be reduced, so much the better. Denaturation. It is critical that complete strand separation occur during the denaturation step. This is a unimolecular reaction which, in itself, is very fast. The suggested 30-sec denaturation used in the protocol ensures that the tube

A.3C.12 Supplement 3


contents reach 94°C. If PCR is not working, it is well worth checking the temperature inside a control tube containing 100 µl water. If GC content is extremely high, higher denaturation temperatures may be necessary; however, Taq DNA polymerase activity falls off quickly at higher temperatures (Gelfand, 1989). To amplify a long sequence (>3 kb), minimize the denaturation time to protect the target DNA from possible effects, such as depurination, of lowered pH of the Tris buffer at elevated temperatures. Annealing. It is critical that the primers anneal stably to the template. Primers with relatively low GC content (1.5 mm thick, cracking can be reduced by using an alternative fixing solution containing glycerol (3% glycerol/10% glacial acetic acid/20% methanol; see Support Protocol 1). Among the biggest problems encountered in autoradiography are images that are either too weak or too intense. Such problems can be solved by varying the exposure time. Estimating initial exposure time is difficult, since the

Table A.3D.2

Film Choice and Exposure Temperature for Autoradiography

Isotope

Enhancement method

Film

Exposure temperature

3H

Fluorography None Fluorography CaWO4 intensifying screens

Double-coated Single-coated Double-coated Double-coated

−70°C Room temperature −70°C −70°C

35S, 14C, 32P 35S, 14C, 32P 32P, 125I

Useful Techniques

A.3D.9 Current Protocols in Toxicology

Supplement 7

amount of radioactivity in the sample is often unknown. A Geiger counter can offer some guidance with certain isotopes. For highly exposed film, the length of time in developer can be reduced to produce a lighter image. It is particularly important to remember that if accurate quantification of the film image is desired, film must be preflashed so that there is a linear relationship between the amount of radioactivity in the sample and the image intensity. Artifacts, such as black spots and stripes, can be avoided during developing by making sure that no moisture comes in contact with the film and that films exposed at −70°C are brought to room temperature before developing. Also, it must be noted that β particles from weak isotopes such as 3H cannot penetrate plastic wrap, and plastic wraps can attenuate signals from 35S and 14C up to two-fold.

for fixing and ∼30 additional minutes will be required for drying. The length of exposure for films in autoradiography can range from a few minutes to a few weeks, depending on the strength of the radioactivity in the sample. Most exposures last from several hours to a few days. Exposure time can be reduced more than 10-fold with a phosphor imager.

Anticipated Results

Laskey, R.A. and Mills, A.D. 1975. Quantitative film detection of 3H and 14C in polyacrylamide gels by fluorography. Eur. J. Biochem. 56:335-341.

The protocols described here should yield a film image of a gel that can be quantified, stored, and photographed for publication.

Time Considerations

Fixing a gel will require ∼45 min. Drying will take an additional 2 hr for a gel 1 mm in thickness. Incorporation of a fluor will add ∼45 min to the processing time. For gels >1.5 mm thick or with >15% acrylamide, an additional 30 min will be required

Literature Cited Chamberlain, J.P. 1979. Fluorographic detection of radioactivity in polyacrylamide gels with the water-soluble fluor, sodium salicylate. Anal. Biochem. 98:132-135. Johnston, R.F., Pickett, S.C., and Barker, D.L. 1990. Autoradiography using storage phosphor technology. Electrophoresis 11:355-360. Laskey, R.A. 1980. The use of intensifying screens or organic scintillators for visualizing radioactive molecules resolved by gel electrophoresis. Methods Enzymol. 65:363-371.

Laskey, R.A. and Mills, A.D. 1977. Enhanced autoradiographic detection of 32P and 125I using intensifying screens and hypersensitized film. FEBS Lett. 82:314-316.

Contributed by Daniel Voytas and Ning Ke Iowa State University Ames, Iowa

Detection and Quantitation of Radiolabeled Proteins in Gels and Blots

A.3D.10 Supplement 7


Northern Blot Analysis of RNA

APPENDIX 3E

Northern blot hybridization is used to determine quantitatively the presence of specific mRNA transcripts in cell or tissue lysates and to evaluate the specificity of a select oligonucleotide probe. First the RNA is extracted from a tissue or cell culture (see Support Protocol 1). Following agarose gel separation, the RNA is transferred by capillary action to a nylon membrane (see Basic Protocol). The RNA is then UV-crosslinked to the membrane and stained with methylene blue to verify RNA integrity and equal RNA loading and transfer and then used for hybridization. After prehybridization, which blocks the nonspecific binding sites, the membrane is probed with a specific cDNA oligonucleotide, which is labeled with 32P using terminal deoxynucleotidyl transferase (TdT; see Support Protocol 2). Following successive washes of increasing stringency, the membrane is exposed to an autoradiographic film. The presence of a band on the film emulsion indicates RNA hybridization with the radioactive probe. The probe can then be removed from the membrane, and subsequent probes rehybridized for further analysis (see Support Protocol 3). In general, it is preferable to isolate mRNA instead of total RNA, as mRNA represents only 1% to 5% of the total cellular RNA; therefore, mRNA samples give a substantially higher signal than total RNA. For mRNA isolation, use commercially available kits (e.g., Invitrogen’s Fastrack 2), which allow mRNA isolation directly from tissues or cells without prior total RNA extraction. Although extraction of total RNA is simpler and faster, and verification of RNA integrity is easier (the prominent bands corresponding to the 18 S and 28 S ribosomal subunits, which are 1.8 to 2 kb and 4 to 5 kb, respectively, can be used as an indication that the RNA has not been degraded during the extraction protocol), total RNA should be used for northern blots only if there is preliminary information of abundant expression of the transcript. If total RNA has to be used because of a limited amount of cells or tissues and no signal can be detected by northern blot analysis, reverse transcriptase amplification (RT-PCR) may be used. The major problem in the analysis of RNA is contamination with ribonucleases (RNase), which are very stable enzymes that do not require any cofactors for activity. Avoid RNase contamination by wearing clean powder-free gloves for all procedures and by using RNase-free reagents, plasticware, and glassware. Autoclaving solutions, glassware, and plasticware is usually not enough to inactivate all RNases. Instead, all heat-resistant glassware and plasticware should be baked at 140° to 150°C overnight. Equipment such as the electrophoresis tank should be rinsed in RNase-free water for 30 min prior to use. All solutions (except Tris-containing solutions) can be made from water treated with the RNase inhibitor diethylpyrocarbonate (DEPC; see UNIT 2.9). The use of RNase-free solutions (made up with DEPC-treated water, unless otherwise stated) is recommended for the preparation of all solutions before and during the hybridization steps. Because DEPC-treated water from commercial sources is quite expensive, the use of low-cost, DEPC-treated water made in the lab is recommended. The use of RNase-free solutions is not required in post-hybridization procedures because of the resistance of RNA:RNA hybrids to RNase degradation. Because even the smallest contaminants of DEPC may affect many subsequent enzymatic steps such as labeling or amplification, purified RNA should be solubilized in water from a known RNase/DNase-free source (not DEPCtreated and pyrogen-free). RNA samples should be kept on ice when they are not protected by RNase inhibitors. NOTE: For commentary concerning northern blot analysis, see UNIT 2.9. Useful Techniques Contributed by Marcelle Bergeron, Jari Honkaniemi, and Frank R. Sharp Current Protocols in Toxicology (2001) A.3E.1-A.3E.13 Copyright © 2001 by John Wiley & Sons, Inc.

A.3E.1 Supplement 7

NORTHERN BLOT HYBRIDIZATION

BASIC PROTOCOL

Isolated RNA (see Support Protocol 1) is size fractionated by gel electrophoresis using a 1% (w/v) agarose gel to separate RNA in the 1- to 10-kb range prior to transfer. If ethidium bromide is incorporated into the gel, bands can be visualized using a transilluminator. After size fractionation, the RNA is transferred to a membrane. Capillary flow is the most common and inexpensive method to transfer nucleic acids (RNA and DNA) from an agarose gel to a membrane. The nucleic acid is washed out of the gel onto the membrane using a stack of dry paper towels, which draws the transfer solution from the reservoir through a wick of Whatman 3 MM blotting paper, then through the gel, the membrane, and into the dry paper towels. Two variants of the capillary flow transfer method currently exist: upward and downward transfer (Figs. A.3E.1A and A.3E.1B, respectively). In both systems, the gel is closest to the reservoir. The membrane is in direct contact with the surface of the gel followed by some blotter paper and a stack of dry paper towels. Although both setups work very well, downward transfer (e.g., using the TurboBlotter, Schleicher and Shuell) is recommended since it results in somewhat sharper bands as no extra weight is applied on the gel. Once the materials are prepared, the nylon membrane bearing size-fractionated RNA is incubated with the radiolabeled oligonucleotide probe to allow hybridization between the single-stranded DNA and the single-stranded RNA. After hybridization the membrane is washed with a series of solutions of increasing stringency and then exposed to autoradiographic film to visualize the hybridized bands. This protocol is adapted for the use of short oligonucleotides (30- to 45-mer) in northern blot hybridization. Any variations in temperature, salts or denaturing conditions may greatly affect the probe hybridization. Materials Ultrapure agarose, low-melting temperature (e.g., Life Technologies, Bio-Rad, Sigma, or Fisher) DEPC-treated H2O (UNIT 2.9) 10× and 1× MOPS (see recipe) 37% formaldehyde (Sigma) Purified RNA (20 to 40 µg total or 4 to 10 µg mRNA; see Support Protocol 1) ≥99.5% formamide, deionized (Sigma) RNA loading dye (see recipe) 10 mg/ml ethidium bromide (optional; see recipe) Nylon membrane, positively charged (Hybond from Amersham Pharmacia Biotech or Nytran SuPerCharge from Schleicher & Schuell )

A

B weight

cover

paper towels blotting paper membrane gel buffer bridge support

Figure A.3E.1 Capillary flow transfer of RNA from agarose gels to a membrane. (A) Upward capillary transfer. (B) Downward capillary transfer.

A.3E.2 Supplement 7


20× and 2× SSC (UNIT 2.9) 0.02% (w/v) methylene blue RNA staining solution (UNIT 2.9) 20× SSPE (UNIT 2.9) 20% (w/v) SDS (APPENDIX 2A) 100× Denhardt’s solution (UNIT 2.9) 10 mg/ml heat-denatured salmon sperm DNA (Life Technologies) 50% (w/v) dextran sulfate (UNIT 2.9) 100 cpm/µg [α-32P]dATP-labeled oligonucleotide probe (see Support Protocol 2) 1× and 2× SSPE/0.1% SDS 1× STE buffer (see recipe) Horizontal gel electrophoresis system with casting box and Teflon combs with 3to 4-mm teeth (e.g., Bio-Rad, Amersham Pharmacia Biotech, Owl, Shadel) and appropriate power supply Microcentrifuge tubes, RNase-free UV transilluminator, ruler, and camera (optional) Elevated support (e.g., empty pipet tip box) Whatman 3 MM papers 10- to 15-cm stack of paper towels Small weight (~500 g) or light-weight cover UV Cross-linker (Stratagene) Pencil or fluorescent pen (optional) Hybridization oven with glass hybridization bottle (Hybaid) or 42°C water bath and sealable bag Geiger counter Autoradiographic cassette, light-tight with intensifying screen X-ray film (e.g., X-Omat AR5 or BioMax MS from Kodak) CAUTION: When working with radioactivity, take appropriate precautions to avoid contamination of the experimenter and the surroundings. Carry out the experiment and dispose of wastes in an appropriately designated area, following the guidelines provided by the local radiation safety officer (also see APPENDIX 1A). CAUTION: Radiolabeled probes are hazardous; see APPENDIX 1A for guidelines on handling, storage, and disposal. To protect from radioactivity and RNase contamination as well as finger smudges, always wear clean, powder-free gloves when working with RNA and radioactive substances. Do not reuse gloves. Prepare agarose gel 1. Set up the casting box (mold) following the manufacturer’s instructions. 2. In a flask, mix 1.0 g ultrapure low-melting-temperature agarose with 85 ml DEPCtreated water. Heat on a heating/stirring plate (or in a microwave oven) until all crystals are dissolved. NOTE: Do not boil the agarose solution. The volume of agarose solution needed depends on the size of the electrophoresis chamber used. Keep the gel thickness between 4 and 6 mm to obtain sharper bands and to allow efficient RNA transfer. Transcripts of 1 to 10 kb are easily separated by 1% agarose gels, whereas transcripts shorter than 0.5 to 4 kb are better separated by 1.6% to 1.8% agarose gels. Gels with

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