Immunotoxicity Testing: Methods and Protocols (Methods in Molecular Biology Vol598)

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Methods

in

Molecular Biology™

Series Editor John M. Walker School of Life Sciences University of Hertfordshire Hatfield, Hertfordshire, AL10 9AB, UK

For other titles published in this series, go to www.springer.com/series/7651

Immunotoxicity Testing Methods and Protocols

Edited by

Rodney R. Dietert Department of Microbiology and Immunology, College of Veterinary Medicine, Cornell University, Ithaca, NY 14853, USA

Editor Rodney R. Dietert Department of Microbiology and Immunology College of Veterinary Medicine Cornell University Ithaca, NY 14853 USA [email protected]

ISSN 1064-3745 e-ISSN 1940-6029 ISBN 978-1-60761-400-5 e-ISBN 978-1-60761-401-2 DOI 10.1007/978-1-60761-401-2 Springer New York Dordrecht Heidelberg London Library of Congress Control Number: 2009936794 © Humana Press, a part of Springer Science+Business Media, LLC 2010 All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Humana Press, c/o Springer Science+Business Media, LLC, 233 Spring Street, New York, NY 10013, USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. While the advice and information in this book are believed to be true and accurate at the date of going to press, neither the authors nor the editors nor the publisher can accept any legal responsibility for any errors or omissions that may be made. The publisher makes no warranty, express or implied, with respect to the material contained herein. Printed on acid-free paper Humana Press is part of Springer Science+Business Media (www.springer.com)

Preface Immunotoxicology as an interdisciplinary area of research, assessment, and instruction has been formally recognized since at least the 1970s. The science supporting immunotoxicology has driven both mechanistic research defining the interactions between xenobiotics and the immune system as well as safety testing for chemicals, drugs, and medical devices. During that time, there have emerged societies, specialty sections, and entire journals devoted solely to Immunotoxicology. While several important books have been prepared on this and related topics, Immunotoxicity Testing is among the first to meld consideration of immunotoxicity testing approaches and strategies with a comprehensive presentation of detailed laboratory protocols. The goal of the book is to utilize the expertise of scientists actually engaged in immunotoxocity testing to provide the reader with lab-ready procedures and the background information needed to identify effective testing approaches. The book includes an introduction to the topic with a description of the evolution of immunotoxicity testing and ideas concerning its future direction. Additionally, the importance of immunotoxicity testing for health risk reduction is presented by categories of disease. Given this scope, the book is appropriate for a broad audience reaching beyond immunotoxicology itself. Chapters are designed to be accessible by students, technicians, lab and safety office personnel as well as biology- and chemistry-oriented researchers. Above all, the book provides a one-stop reference resource for the most important and commonly used laboratory protocols in immunotoxicology. As an editor, I thank the expert authors for the time and effort they devoted to each chapter and hope that this novel reference work will aid the continued evolution and the application of immunotoxicity testing. Dr. Rodney R. Dietert Ithaca, NY March 2009

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Contents Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Contributors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Part I Introduction 1 Immunotoxicology Testing: Past and Future . . . . . . . . . . . . . . . . . . . . . . . . . . . . Michael I. Luster and G. Frank Gerberick

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Part II Overview and Health-Risk Considerations 2 Developmental Immunotoxicity (DIT): The Why, When, and How of DIT Testing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Rodney R. Dietert and Jamie DeWitt 3 An In Vivo Tiered Approach to Test Immunosensitization by Low Molecular Weight Compounds . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Irene S. Ludwig, Lydia M. Kwast, Daniëlle Fiechter, and Raymond H.H. Pieters 4 Risk of Autoimmune Disease: Challenges for Immunotoxicity Testing . . . . . . . . . Rodney R. Dietert, Janice M. Dietert, and Jerrie Gavalchin 5 Markers of Inflammation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Dori R. Germolec, Rachel P. Frawley, and Ellen Evans 6 Evaluating Macrophages in Immunotoxicity Testing . . . . . . . . . . . . . . . . . . . . . . John B. Barnett and Kathleen M. Brundage

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39 53 75

Part III Immunotoxicity and Host Resistance Models 7 Host Resistance Assays Including Bacterial Challenge Models . . . . . . . . . . . . . . . 97 Florence G. Burleson and Gary R. Burleson 8 Viral Host Resistance Studies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 109 Wendy Jo Freebern 9 Parasite Challenge as Host Resistance Models for Immunotoxicity Testing . . . . . . 119 Robert W. Luebke 10 Tumor Challenges in Immunotoxicity Testing . . . . . . . . . . . . . . . . . . . . . . . . . . . 143 Sheung Ng, Kotaro Yoshida, and Judith T. Zelikoff

Part IV Testing Protocols in Rodents and Other Laboratory Animals 11 The T-Dependent Antibody Response to Keyhole Limpet Hemocyanin in Rodents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 159 Lisa M. Plitnick and Danuta J. Herzyk 12 The Sheep Erythrocyte T-Dependent Antibody Response (TDAR) . . . . . . . . . . . 173 Kimber L. White, Deborah L. Musgrove, and Ronnetta D. Brown

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13 The Delayed Type Hypersensitivity Assay Using Protein and Xenogeneic Cell Antigens . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Rodney R. Dietert, Terry L. Bunn, and Ji-Eun Lee 14 The Cytotoxic T Lymphocyte Assay for Evaluating Cell-Mediated Immune Function . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Gary R. Burleson, Florence G. Burleson, and Rodney R. Dietert 15 NK Cell Assays in Immunotoxicity Testing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Qing Li 16 The Local Lymph Node Assay and Skin Sensitization Testing . . . . . . . . . . . . . . . Ian Kimber and Rebecca J. Dearman 17 Use of Contact Hypersensitivity in Immunotoxicity Testing . . . . . . . . . . . . . . . . . Jacques Descotes 18 Evaluation of Apoptosis in Immunotoxicity Testing . . . . . . . . . . . . . . . . . . . . . . . Mitzi Nagarkatti, Sadiye Amcaoglu Rieder, Dilip Vakharia, and Prakash S. Nagarkatti 19 Dendritic Cells in Immunotoxicity Testing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Donghong Gao and David A. Lawrence 20 Evaluating Cytokines in Immunotoxicity Testing . . . . . . . . . . . . . . . . . . . . . . . . . Emanuela Corsini and Robert V. House 21 Flow Cytometry in Preclinical Drug Development . . . . . . . . . . . . . . . . . . . . . . . . Patrick B. Lappin 22 Enhanced Histopathology Evaluation of Lymphoid Organs . . . . . . . . . . . . . . . . . Susan A. Elmore 23 Immunotoxicity Testing in Nonhuman Primates . . . . . . . . . . . . . . . . . . . . . . . . . Stephanie Grote-Wessels, Werner Frings, Clifford A. Smith, and Gerhard F. Weinbauer

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259 283 303 323 341

Part V Evaluation in Humans 24 Fundamentals of Clinical Immunotoxicology . . . . . . . . . . . . . . . . . . . . . . . . . . . . 363 Robert V. House

Part VI Wildlife Testing 25 In Vivo Functional Tests for Assessing Immunotoxicity in Birds . . . . . . . . . . . . . . 387 Keith A. Grasman

Part VII In Vitro Alternatives 26 In Vitro Testing for Direct Immunotoxicity: State of the Art . . . . . . . . . . . . . . . . 401 D.P.K. Lankveld, H. Van Loveren, K.A. Baken, and R.J. Vandebriel Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 425

Contributors K. A. Baken • Laboratory for Health Protection Research, National Institute for Public Health & the Environment, Bilthoven, The Netherlands John B Barnett • Department of Microbiology, Immunology and Cell Biology, West Virginia University, Morgantown, WV, USA Ronnetta D. Brown • Department of Pharmacology and Toxicology, School of Medicine, Virginia Commonwealth University, Richmond, VA, USA Kathleen M. Brundage • Department of Microbiology, Immunology and Cell Biology West Virginia University, Morgantown, WV, USA Terry L. Bunn • Department of Preventive Medicine and Environmental Health, University of Kentucky, Lexington, KY, USA Florence G. Burleson • Burleson Research Technologies, Morrisville, NC, USA Gary R. Burleson • Burleson Research Techologies, Morrisville, NC, USA Emanuela Corsini • Department of Pharmacological Sciences, University of Milan, Milan, Italy Rebecca Dearman • Faculty of Life Sciences, University of Manchester, Manchester, UK Jacques Descotes • Poison Center and Claude Bernard University, Lyon, France Jamie DeWitt • Department of Phamacology and Toxicology, East Carolina University, Greenville, NC, USA Janice M. Dietert • Performance Plus Consulting, Lansing, NY, USA Rodney R. Dietert • Department of Microbiology and Immunology, College of Veterinary Medicine, Cornell University, Ithaca, NY, USA Susan A. Elmore • Cellular and Molecular Pathology Branch, National Institute of Environmental Health Sciences, Research Triangle Park, NC, USA Ellen Evans • Clinical Pathology and Immunotoxicology, Schering Plough Research Institute, Lafayette, NJ, USA Rachel P. Frawley • Toxicology Branch, National Toxicology Program, National Institute of Environmental Health Sciences, Research Triangle Park, NC, USA Wendy J. Freebern • Department of Immunotoxicology, Drug Safety Evaluation, Research and Development, Bristol-Myers Squibb Co., Syracuse, NY, USA Danielle Fiechter • Institute for Risk Assessment Sciences, Utrecht University, Utrecht, The Netherlands Werner Frings • Covance Laboratories, Muenster, Germany Donghong Gao • Wadsworth Center, Albany, NY, USA Jerri Gavalchin • Department of Animal Science, Cornell University, Ithaca, NY Department of Medicine, SUNY Upstate Medical University, Syracuse, NY G. Frank Gerberick • Miami Valley Innovation Center, Proctor & Gamble, Cincinnati, OH, USA Dori R. Germolec • Toxicology Branch, National Toxicology Program, National Institute of Environmental Health Sciences, Research Triangle Park, NC, USA Keith A. Grasman • Department of Biology, Calvin College, Grand Rapids, MI, USA Stephanie Grote-Wessels • Covance Laboratories, Muenster, Germany ix

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Danuta J. Herzyk • Department of Safety Assessment, Merck Research Laboratories, West Point, PA, USA Robert V. House • DynPort Vaccine Company LLC, Frederick, MD, USA Ian Kimber • Faculty of Life Sciences, University of Manchester, Manchester, UK Lydia M. Kwast • Institute for Risk Assessment Sciences, Utrecht University, Utrecht, The Netherlands D. P. K. Lankveld • Laboratory for Health Protection Research, National Institute for Public Health & the Environment, Bilthoven, The Netherlands Patrick B. Lappin • Investigative Pathology, Pfizer, Inc. San Diego, CA, USA David A. Lawrence • Wadsworth Center, Albany, NY, USA Ji-Eun Lee • Product Safety, Colgate-Palmolive Company, Piscataway, NJ, USA Qing Li • Department of Hygiene and Public Health, Nippon Medical School, Japan Irene S. Ludwig • Institute for Risk Assessment Sciences, Utrecht University, Utrecht, The Netherlands Robert W. Luebke • Immunotoxicology Branch, Division of National Health and Environmental Effects, United State Environmental Protection Agency, Research Triangle Park, NC, USA Michael I. Luster • Luster Associates, Morgantown, WV, USA Deborah L. Musgrove • Department of Pharmacology and Toxicology School of Medicine,Virginia Commonwealth University, Richmond, VA, USA Mitzi Nagarkatti • Department of Pathology, Microbiology and Immunology, University of South Carolina School of Medicine, Columbia, SC, USA Prakash S. Nagarkatti • Department of Pathology, Microbiology and Immunology, University of South Carolina School of Medicine, Columbia, SC, USA Sheung Ng • Nelson Institute of Environmental Medicine, New York University School of Medicine, Tuxedo, NY, USA Raymond H. H. Pieters • Institute for Risk Assessment Sciences, Utrecht University, Utrecht, Netherlands Research Centre for Innovative Testing, Institute for Life Sciences and Chemistry, Utrecht University of Applied Sciences, Utrecht, The Netherlands Lisa M. Plinick • Department of Safety Assessment, Merck Research Laboratories, West Point, PA, USA Sadiye Amcaoglu Rieder • Department of Pathology, Microbiology and Immunology, University of South Carolina School of Medicine, Columbia, SC, USA Clifford A. Smith • Covance, Harrogate, UK Dilip Vakharia • Department of Pathology, Microbiology and Immunology, University of South Carolina School of Medicine, Columbia, SC, USA Rob J. Vandebriel • Laboratory for Health Protection Research, National Institute for Public Health & the Environment, Bilthoven, The Netherlands Henk van Loveren • Laboratory for Health Protection Research, National Institute for Public Health & the Environment, Bilthoven, The Netherlands Gerhard F. Weinbauer • Covance Laboratories, Muenster, Germany Kimber L. White Jr. • Department of Pharmacology and Toxicology, School of Medicine, Virginia Commonwealth University, Richmond, VA, USA Kotaro Yoshida • Nelson Institute of Environmental Medicine, New York University School of Medicine, Tuxedo, NY, USA Judith T. Zelikoff • Nelson Institute of Environmental Medicine, New York University School of Medicine, Tuxedo, NY, USA

Part I Introduction

Chapter 1 Immunotoxicology Testing: Past and Future Michael I. Luster and G. Frank Gerberick Abstract A brief historical perspective of immunotoxicology is presented describing the early development of predictive screening tests to identify xenobiotics that may cause immunosuppression or skin sensitization. This includes a discussion of the evolution of the discipline to support a better understanding of basic science and improvement of human risk assessment. The last section describes the need for additional validated screening tests and recent efforts to address this gap in the other areas of immunotoxicology including food and respiratory allergy, autoimmunity and immunostimulation. Key words: Testing, Guidelines, Immunosuppression, Hypersensitivity, Risk assessment

1. Introduction The identification and regulation of xenobiotic agents that inadvertently alter the immune system and affect human health have been of concern to the chemical/agricultural, pharmaceutical and consumer product industries, as well as to the federal regulatory agencies for over 40 years. Initial interest originated in the area of sensitization from the observations made by Landsteiner and Jacobs (1) that low molecular weight chemicals or drugs can be antigenic and capable of producing organ-specific (i.e., skin, lung or gastrointestinal tract) allergic responses. Subsequently, other studies reported that certain xenobiotics, such as halogenated aromatic hydrocarbons, could suppress, or in rare instances stimulate, the immune system resulting in an increased risk of infectious or neoplastic diseases, or in the latter case, exacerbate autoimmune disease. Of particular concern have been the xenobiotic effects in the neonate as increasing evidence suggests that the developing immune system is particularly sensitive to damage. Other materials, particularly certain pharmaceuticals, cause autoimmune-like syndromes R.R. Dietert (ed.), Immunotoxicity Testing: Methods and Protocols, Methods in Molecular Biology, vol. 598 DOI 10.1007/978-1-60761-401-2_1, © Humana Press, a part of Springer Science + Business Media, LLC 2010

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in which the disease dissipates following cessation of exposure while other chemicals appear to exacerbate existing autoimmune disease. The development and adoption of appropriate experimental methods to assess the influence of xenobiotics to cause these various toxicities were for many years the major focus of immunotoxicology, and for some effects such as autoimmunity, respiratory allergy and the so-called systemic allergies still remain an issue. The following provides a brief historical review on the development of immunotoxicity testing and a perspective of what testing strategies are needed in the future.

2. Immunosuppression As it is relatively difficult to determine the contribution of chronic low-level immunosuppression or the cumulative effects of modest changes in immune function to the background incidence of disease in the human population, efforts have been made to examine the relationships between laboratory measures of immune response and disease resistance in experimental animal models. Although the experimental methods initially adopted by immunotoxicologists to assess immune function were those common to immunology laboratories, the tests that were commonly performed and the experimental design by which they were conducted were performed ad hoc. Even the experimental species that have been selected varied with the earliest studies using rabbits and guinea pigs and later studies conducted using rats and mice. While rodents became the test species of choice, debate occurred on species selection with those trained in toxicology usually preferring the rat to allow comparison with other toxicology studies, and those trained in immunology preferring the mouse as the mouse immune system was well studied. The lack of standardized testing made it difficult to compare the chemical-specific effects and led Dean et al. (2), to suggest a “Tier” approach with the idea that each subsequent tier provided identification of a more defined effect on the immune system. Subsequently, the National Toxicology Program (NTP) organized a series of workshops composed of experts in immunotoxicology, basic immunology, toxicology, risk assessment, epidemiology and clinical medicine to help identify the most appropriate tests for immunotoxicology testing (3). Two major points were agreed upon from these workshops: First, since the immune system is not fully operational until it is challenged, the most appropriate tests would be those that incorporate an antigen challenge. Second, since it may be construed that an inadequate response to antigenic challenge does not represent an “adverse effect,” tests should also be added that could be readily identified with disease.

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The former recommendation highlighted several common assays including measurement of an antibody response following antigen challenge as a measure of humoral immunity and quantification of delayed hypersensitive response (DHR) or cytotoxic T lymphocyte response (CTL) as a measure for cell-mediated immunity. These assays were based upon the measurement of a primary immune response rather than secondary since it is generally thought that memory responses are less sensitive to inhibition than primary responses. To address the need to identify a clear adverse effect, a set of tests, usually referred to as “host resistance assays,” was suggested. These tests would also be used to validate the usefulness of other methods and extrapolate the potential for environmental agents to alter host susceptibility in the human population. In these assays, groups of experimental animals are challenged with either an infectious agent or transplantable tumor at a challenge level sufficient to produce disease in control animals and increased incidence is examined in the treated groups. As the endpoints in these tests have evolved from relatively non-specific (e.g., animal morbidity and mortality) to quantitative, such as tumor numbers, viral titers or bacterial cell counts, the sensitivity of these models has significantly increased. However, they are still somewhat limited by the need to use relatively large numbers of animals. Eventually, a three tier approach emerged in which Tier 1 included screening assays that would likely detect immunotoxic xenobiotics, Tier 2 allowed for defining the immune component(s) effected as well as establish effects on host resistance and Tier 3 provided, in very general terms, approaches that could be used to identify the mode of action. An interlaboratory validation effort involving four laboratories1 and sponsored by the NTP was conducted using Tier 1 and 2 tests (4). In addition to the demonstration of interlaboratory reproducibility, this effort helped identifying the relative sensitivity of the various immune tests and the degree to which they agreed with the commonly employed host susceptibility tests. This effort was followed several years later in which the concordance between various histological, hematological and immune function tests to identify immunotoxicity and host susceptibility changes were determined in a large dataset (5, 6). These latter studies were important, not only as a validation exercise for tier testing, but for providing a basis for moving immunotoxicology assessment forward. The analyses indicated that inclusion of a functional test, most notably the T-dependent antibody response (TDAR) to

The four participating laboratories were the National Institute of Environmental Health Sciences (Research Triangle Park, NC), Chemical Industry Institute of Toxicology (Research Triangle Park, NC), Virginia Commonwealth University (Richmond, VA) and IIT Research Institute (Chicago, IL).

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sheep red blood cells, along with a non-functional test, such as thymus weights, allowed achieving concordance, with respect to identifying potential immunotoxic agents, of well over 90%, although a number of other immune test groupings provided excellent levels of accuracy. These studies also provided evidence of a linear relationship between many of the immune tests and host resistance assays. In the Unites States, the preferred species for testing was the mouse, but subsequent validation studies were conducted in rats (7–9) and results from either rodent species are now equally accepted. Tiered screening panels have been the basis for several risk assessment guidelines and most regulatory agencies in the United States, European Union and Japan have established or are developing requirements or guidelines (reviewed by (10)). However, the Office of Prevention, Pesticides and Toxic Substances (OPPTS), US Environmental Protection Agency (EPA) was responsible for developing the first immunotoxicology test guideline (11) and over the years has taken the lead in their development. It should be noted that the configurations of these testing panels vary depending upon the agency/ organization/program under which they are conducted. The most notable difference is whether a functional immune test (i.e., incorporates antigen challenge) is included in Tier 1 or Tier 2. Although, as indicated earlier, it is generally agreed that functional testing provides the greatest sensitivity for identifying immunosuppression, it has been argued that a careful histological and hematological evaluation, particularly inclusion of extended histopathology endpoints, would identify a large proportion of potential immunotoxic agents (12–14). This is reflected in published and proposed immunotoxicity testing guidelines by the Committee for Proprietary Medicinal Products (CPMP), Organization of Economic Cooperation and Development (OECD) and International Conference on Harmonization of Technical Requirements for Registration of Pharmaceuticals for Humans Use (ICH) (reviewed by (10)). While testing for potential immunotoxicty in experimental animals has gained increased acceptance, few systematic epidemiological immunotoxicological studies had been undertaken. This is due to a number of difficulties in working with human populations (15): (a) Lack of validated immunological assays of sufficient sensitivity (b) Difficulty in accurately determining infectious disease incidence (c) Large cost and difficulty of sample acquisition at sites geographically distant from the investigator

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Table 1.1 Tier 1 immunotoxicty testing recommendations for human studies NRC recommendations

WHO recommendations

Serum IgG, IgM, IgA and IgE levels

Hematological profile

Natural antibody levels to ubiquitous antigens

Antibody mediated immunity

Secondary antibody response to proteins and polysaccharides

Immunophenotyping

Immunophenotyping

Delayed hypersensitivity response

Secondary DTH response

C-Reactive protein

Autoantibody titers (DNA, mitochondria)

Autoantibody titers (DNA, mitochondria) IgE to common allergens NK cells activity or numbers Phagocytosis Clinical chemistry

The US National Academy of Sciences (NAS) in understanding these challenges, proposed a three tier testing scheme to be used for study of populations known or suspected to have been exposed to an immunotoxicant (16). As with experimental animals, it was proposed that tests be conducted from the first Tier and the results used to consider proceeding to the next Tier. The tests included in the first Tier are shown in Table 1.1. The International Programme on Chemical Safety of the World Health Organization (IPCS/ WHO) issued a report on principles and methods for assessing direct immunotoxicity associated with exposure to chemicals (17). Although many assays overlapped, that report recommended a larger number of assays that can be used to evaluate possible immunotoxicity than the NRC tier system (Table 1.1). A symposium on Epidemiology of Immunotoxicity remarked on the need for welldesigned studies of immunotoxicity in humans and supported the application of the NRC three Tier approach (18). Over the years, a number of immunotoxicology population studies were conducted based on a selection of immune biomarkers from both the WHO and NRC Tier 1 recommendations (e.g., 19–21).

3. Skin Sensitization Testing

For many decades, the guinea pig has been the animal of choice for predictive studies of skin sensitization potential. This arose largely as

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a result of the use of the guinea pig in the pioneering investigations into mechanisms of skin sensitization to chemicals (1, 22). The first definition of a real predictive test came from the work of Draize more than 60 years ago (23). Since that time, numerous protocols have been described whose aims have been, in one way or another, to make improvements to the sensitivity and predictivity of the guinea pig as a surrogate for man. In essence, all the test protocols follow similar principles. Typically, a combination of intradermal and/or epicutaneous treatments is administered to 10–20 guinea pigs, with or without adjuvant, over a 2–3 week period in an attempt to induce skin sensitization, then a 1–2 week rest period to allow any immune response to mature, followed finally by a topical challenge to assess the extent to which the skin sensitization might have been induced. A set of 5–10 sham treated controls is also challenged. Evaluation of the skin reactions is usually by subjective visual assessments 24–48 h after the challenge application, the main reaction element being erythema. The protocol of Magnusson and Kligman (24) and that of Buehler (25, 26) are the two most studied and accepted guinea pig methods used for regulatory purposes worldwide (27). The Local Lymph Node Assay (LLNA) is a validated alternative approach to the traditional guinea pig test methods for skin sensitization testing that provides important animal welfare benefits (28, 29). In this method, skin sensitizing potential is measured as a function of lymph node cell proliferative responses induced in mice following repeated topical exposure to the test chemical (30, 31). Not least due to the improved animal welfare benefits, the LLNA has become the preferred method for assessing skin sensitization hazard by various regulatory authorities (32, 33). The OECD test guideline 429 for the LLNA indicates that a minimum of 3 test concentrations and a vehicle control group with a minimum of 4 animals per group are needed (27). A chemical is classified as a skin sensitizer if, at one or more test concentrations, it induces a three-fold or greater increase in draining lymph node cell proliferation compared with concurrent vehicletreated controls (Stimulation Index [SI] ≥3). In the standard LLNA, lymph nodes are pooled and processed on an experimental group basis using 4 mice per group. Alternatively, using 5 mice per group, lymph nodes are pooled on an individual animal basis providing the opportunity to employ statistical analyses and appropriate power (34). The LLNA has been evaluated extensively in both national and international inter-laboratory collaborative trials and has been the subject of comparisons with guinea pig predictive test methods and human sensitization data. An important point is that the LLNA was subjected to rigorous independent scrutiny and validated by the International Coordinating Committee on the Validation of Alternative Methods (ICCVAM) (29). There soon

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followed a similar endorsement by the European Centre for the Validation of Alternative Methods (ECVAM) (28). Thus, the tests traditionally used for the identification of chemicals possessing the intrinsic ability to cause skin sensitization are the guinea pig maximization test, the Buehler occluded patch test and the LLNA. The capacity of these methods to identify skin sensitization hazard has only been formally validated for the LLNA. However, both within this validation and via the publication of other datasets, the guinea pig methods are also recognized to be of sufficient sensitivity and specificity.

4. Needs in Immunotoxicity Screening Testing

As described above, the most attention to development of validated methods for immuntoxicity testing thus far has been devoted to identify xenobiotics that have the potential to produce skin sensitization or immunosuppression. Although not validated, opportunities exist to improve the current testing schemes in terms of improved sensitivity, less reliance on experimental animals and cost reduction, such as cytokine and gene expression profiling (35–37). These procedures offer the additional opportunity to make direct comparisons with humans using samples from serum or isolated leuokocytes. The use of in vitro systems, such as dendritic cell activation, peptide reactivity and T-cell activation are also being applied to hypersensitivty testing (38, 39). Unfortunately, these traditional testing paradigms are inadequate for many issues relevant to immunotoxicity testing that are of signi ficant current importance. For example, food allergies, which are often life-threatening, are common and affect 6–8% of children under the age of 4 and 1–4% of adults (40). While considerable attention has been given to the types of food products that can produce an allergic response, few studies have addressed development of appropriate test methods for identifying sensitizers in food. Rodent models employed for the evaluation of food allergy have utilized strains inherently skewed towards a Th2 allergic phenotype and high IgE production such as the Brown Norway rat or BALB/c and C3H/ HeJ mouse (41). Rodent models tend to replicate the IgE response to food allergens seen in humans but often fail to present similar clinical symptoms. Other animal models, including dogs and pigs, demonstrate many clinical characteristics of human food allergy including respiratory involvement, digestive problems and even anaphylaxis (41). However, lack of standardization due to variable use of adjuvant, varying routes of exposure, not to mention most appropriate test model, make assessing chemical-induced modulation of responses to protein allergens difficult. Importantly, however, dialog on standardized testing protocols for food allergens is continuing (42).

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Respiratory sensitizers can be identified in inhalation studies using the guinea pig (43). However, the technical difficulty, particularly as it relates to conducting inhalation, does not lend this procedure to a routine screening test. Most of the issues related to screening tests for food sensitizers, particularly as it relates to the Th2 phenotype, also apply for respiratory sensitizers. A number of studies have and still continue to address screening tests for respiratory sensitizers. These include among others, monitoring IgE or IgG1 levels in various test species and employing bronchial associated lymphoid tissues for immunophenotyping, cytokine profiling, gene expression and a modified LLNA (43). The observation that many drugs produce autoimmune-like syndromes and environmental chemicals induce onset and modulate autoimmune disease severity, has led the efforts to identify reliable screening tests for xenobiotic-induced autoimmunity. Animal models of autoimmunity have been used to explore both molecular mechanisms and therapeutic interventions for a variety of autoimmune diseases (44). However, while a number of syndromes that are similar to those clinically observed in humans can be mimicked in animal models, the diversity of autoimmune diseases limits the utility of any single model as a screening tool. The popliteal lymph node assay, which measures non-specific stimulation and proliferation in the lymph nodes draining chemically exposed tissues, has been used in conjunction with reporter antigens as a tool to screen for immunostimulating compounds (45). However, this assay falls short of measuring the potential to produce disease. Finally, an ongoing need in immunotoxicology testing is to develop screening tests to identify adverse health consequences from xenobiotics that produce immunostimulation or modulate inflammatory responses. Although these may include some industrial chemicals, they primarily represent therapeutics designed to treat immune-mediated diseases, such as asthma, autoimmunity, or chronic inflammatory diseases. Some general examples include Toll-like receptor (TLR) agonists, cytokine agonists or antagonists, modulators of adhesion molecules, angiogenic therapies, novel vaccine adjuvants and arachidonic acid modulators. Since the immune system represents a vast network of regulatory loops, altering the production or expression of one regulatory immune mediator to treat a disease would likely influence other mediators, the consequences of which may have adverse effects that outweigh the benefits of its intended use.

5. Conclusion In this survey, we have provided the reader with a brief historical perspective of immunotoxicology conveying that its foundation was in the development of predictive screening tests to identify

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xenobiotics that may cause immunosuppression or skin sensitization. The discipline, however, continues to evolve to include contributions to basic science as well as in improving human risk assessment. It is essential, however, that we understand the fact that immunotoxicology represents the study of a number of distinct diseases associated with perturbances of the immune system, and that there is a critical need to develop standardized and validated screening tests for all these immunotoxicities.

References 1. Landsteiner K, Jacobs J (1935) Studies on the sensitization of animals with simple chemical compounds I. J Exp Med 61:643–657 2. Dean JH, Padarathsingh ML, Jerrells TR (1979) Assessment of immunobiological effects induced by chemicals, drugs or food additives. I Tier testing and screening approach. Drug Chem Toxicol 2:5–16 3. Dean JH, Luster MI, Boorman GA, Lauer LD (1982) Procedures available to examine the immunotoxicity of chemicals and drugs. Pharmacol Rev 34:137–148 4. Luster MI, Munson AE, Thomas P, Holsapple MP, Fenters J, White K, Lauer LD, Dean JH (1988) Development of a testing battery to assess chemical-induced immunotoxicity. Fund Appl Toxicol 10:2–19 5. Luster MI, Portier C, Pait DG, White KL, Gennings C, Munson AE, Rosenthal GJ (1992) Risk assessment in immunotoxicology. I. Sensitivity and predictability of immune tests. Fund Apppl Toxicol 18:200–210 6. Luster MI, Portier C, Pait DG, Rosenthal GJ, Germolec DR, Corsini E, Blaylock BL, Pollock P, Kouchi Y, Craig W, White KL, Munson AE, Comment CC (1993) Risk assessment in immunotoxicology. II. Relationship between immune and host resistance tests. Fund Appl Toxicol 21:71–82 7. van Loveren H, Vos J (1989) Immunotoxicological considerations: a practical approach to immunotoxicity testing in the rat. In: Dayan A, Paine A (eds) Advances in applied toxicology. Taylor & Francis, New York, NY, pp 143–164 8. White K, Jennings P, Murray P, Dean J (1994) International validation study carried out in 9 laboratories on the immunological assessment of cyclosporin A in the Fisher 344 rat. Toxicol In Vitro 8:957–962 9. Ladics GS, Smith CE, Elliott GS, Slone TW, Loveless SE (1998) Further evaluation of the incorporation of an immunotoxicological functional asay for assessing humoral

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18. van Loveren H, Germolec DR, Koren HS, Luster MI, Nolan C, Repetto R, Smith E, Vos JG, Vogt RF (1999) Report of the Bilthoven Symposium: advancement of epidemiological studies in assessing the human health effects of immunotoxic agents in the environment and the workplace. Biomarkers 4:135–157 19. Weisglas-Kuperus N, Patandin S, Berbers GAM, Sas TCJ, Mulder PGH, Sauer PJJ, Hooijkaas H (2000) Immunologic effects of background exposure to polychlorinated biphenyls and dioxins in Dutch preschool children. Environ Health Perspect 108:1203–1207 20. Leonardi GS, Houthuijs D, Steerenberg PA, Fletcher T, Armstrong B, Antova T, Lochman I, Lochmanova A, Rudnai P, Erdei E, Musial J, Jazwiec-Kanyion B, Niciu EM, Durbaca S, Fabianova E, Koppova K, Lebret E, Brunekreef B, van Loveren H (2000) Immune biomarkers in relation to exposure to particulate matter: a cross-sectional survey in 17 cities of Central Europe. Inhalation Toxicol 12(Supp 4):1–14 21. Pinkerton L, Biagini R, Ward EM, Hull RD, Deddens JA, Boeniger MF, Schnoor TM, Luster MI (1998) Immunologic findings among lead-exposed workers. Am J Indus Med 33:400–408 22. Landsteiner K, Jacobs J (1936) Studies on the sensitization of animals with simple chemical compounds II. J Exp Med 64:625–639 23. Draize JH, Woodard G, Calvery HO (1944) Methods for the study of irritation and toxicity of substances applied topically to the skin and mucous membranes. J Pharmacol Exp Ther 8:377–390 24. Magnusson B, Kligman AM (1970) Allergic contact dermatitis in the guinea pig. Identification of contact allergens. Charles C. Thomas, Springfield IL 25. Buehler EV (1965) Delayed contact hypersensitivity in the guinea pig. Arch Dermatol 91:171–177 26. Robinson MK, Nusair TL, Fletcher ER, Ritz HL (1990) A review of the Buehler guinea pig skin sensitization test and its use in a risk assessment process for human skin sensitization. Toxicology 61:91–107 27. Organisation for Economic Cooperation and Development (2002) Test Guideline 429: The local Lymph Node Assay. OECD, Paris 28. Balls M, Hellsten E (2000) Statement on the validity of the local lymph node assay for skin sensitization testing. ECVAM Joint Research Centre, European Commission, Ispra. Altern Lab Anim 28:366–367 29. Dean JH, Twerdock LE, Tice RR, Sailstad DM, Hattan DG, Stokes WS (2001) ICCVAM evaluation of the murine local lymph node

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assay. II. Conclusions and recommendations of an independent scientific review panel. Regul Toxicol Pharmacol 34:258–273 Kimber I, Mitchell JA, Griffin AC (1986) Development of a murine local lymph node assay for the determination of sensitizing potential. Fd Chem Toxic 24:585–596 Kimber I, Hilton J, Weisenberger C (1989) The murine local lymph node assay for identification of contact allergens: a preliminary evaluation of in situ measurement of lymphocyte proliferation. Contact Derm 21:215–220 United States Environmental Protection Agency Health Effects Test Guidelines (2003) OPPTS 870.2600 Skin sensitization. US/ EPA, Washington, DC Cockshott A, Evans P, Ryan CA, Gerberick GF, Betts CJ, Dearman RJ, Kimber I, Basketter DA (2006) The local lymph node assay in practice: a current regulatory perspective. Hum Exp Toxicol 25:387–394 Gerberick GF, Ryan CA, Dearman RJ, Kimber I (2007) Local lymph node assay (LLNA) for detection of sensitizing capacity of chemicals. Methods 41:54–60 Loveless SE, Ladics GS, Smith C, Holsapple MP, Woolhiser MR, White KL, Musgrove DL, Smialowicz RJ, Williams W (2007) Interlaboratory study of the primary antibody response to sheep red blood cells in outbred rodents following exposure to cyclophosphamide or dexamethasone. J Immunotoxicol 4:233–238 Luebke R, Holsapple M, Ladics G, Luster MI, Selgrade M-J, Woolhiser M, Germolec DR (2006) Immunotoxicogenomics: the potential of genomics technology in the immunotoxicity risk assessment process. Toxicol Sci 94:22–27 Boverhof DR, Gollapudi BB, Hotchkiss JA, Osterioh-Quiroz M, Woolhiser MR (2008) Evaluation of a toxicogenomic approach to the local lymph node assay (LLNA). Toxicol Sci 107(2):427–439 Ryan CA, Hulette BC, Gerberick GF (2001) Review: approaches for the development of cell based in vitro methods for contact sensitization. Toxicol In Vitro 15:43–55 Ryan CA, Gerberick GF, Gildea LA, Hulette BC, Betts CJ, Cumberbatch M, Dearman RJ (2005) Interactions of chemicals with dendritic cells – a novel approach for identification of potential allergens. Toxicol Sci 88:4–11 Teufel M, Biedermann T, Rapps N, Hausteiner C, Henningsen P, Enck P, Zipfel S (2007) Psychological burden of food allergy. World J Gastroenterol 3:3456–3465 McClain S, Bannon GA (2006) Animal models of food allergy: opportunities and barriers. Curr Allergy Asthma Rep 2:141–144

Immunotoxicology Testing: Past and Future 42. Ladics GS, Selgrade MK (2008) Identifying food proteins with allergenic potential: evolution of approaches to safety assessment and research to provide additional tools. Regul Toxicol Pharmacol 54(3 Suppl): S2–S6 43. Holsapple MP, Jones D, Kawabata TT, Kimber I, Sarlo K, Selgrade MK, Shah J, Woolhiser MR

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(2006) Assessing the potential to induce respiratory hypersensitivity. Toxicol Sci 91:4–13 44. Germolec DR (2005) Autoimmune diseases, animal models. In: Vohr H-W (ed) Encyclopedic reference of immunotoxicology. Springer, Berlin, pp 75–79 45. Pieters R (2007) Detection of autoimmunity by pharmaceuticals. Methods 41:112–117

Part II Overview and Health-Risk Considerations

Chapter 2 Developmental Immunotoxicity (DIT): The Why, When, and How of DIT Testing Rodney R. Dietert and Jamie DeWitt Abstract Developmental immunotoxicity (DIT) has emerged as a serious health consideration given the increases in the prevalence of many immune-based childhood diseases and conditions, including allergic diseases and asthma, recurrent otitis media, pediatric celiac disease, and type 1 diabetes. As a result, the use of DIT testing to identify potential environmental risk factors contributing to these and other diseases has become a higher priority. This introductory chapter considers: (1) the basis for an increased and earlier use of DIT testing in safety evaluations and (2) the general features of DIT testing strategies designed to reduce health risks. Key words: Developmental immunotoxicity, DIT, Developmental immunotoxicology, Pediatric health risks, Safety testing, Autoimmunity, Allergic hypersensitivity, Inflammation, Immuno suppression, Host resistance

1. Introduction Developmental immunotoxicity (DIT) testing is a significant consideration under the larger umbrella of immunotoxicity testing covered in this book. DIT received only occasional research consideration before the mid-1990s (1–3). However, it has grown sufficiently in scope and impact to be the subject of a stand-alone book by Holladay (4) and has been an integral part of virtually every book on immunotoxicology to appear since that time (5–10). Luster et al. (10) recently defined developmental immunotoxicology as “the effects on the immune system resulting from pre- and/or postnatal exposure to physical factors (e.g., ionizing and ultraviolet radiation), chemicals (including drugs), biological materials, medical devices, and in certain instances, physiological factors, collectively referred to as agents.” DIT increases the risk R.R. Dietert (ed.), Immunotoxicity Testing: Methods and Protocols, Methods in Molecular Biology, vol. 598 DOI 10.1007/978-1-60761-401-2_2, © Humana Press, a part of Springer Science + Business Media, LLC 2010

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of autoimmunity, allergic hypersensitivity, susceptibility to infectious diseases and cancer, and inflammatory diseases in humans as well as in wildlife. The increased risk exists because the immune system is central not only to host defense but also to physiological homeostasis. Since environmentally induced immune dysfunction encompasses both suppression and inappropriate enhancement of immune responses, DIT testing should be capable of detecting both types of changes. This introductory chapter on DIT considers three key topics that significantly impact DIT testing. These are: (1) the “why” of DIT testing or the scientific basis for early-life vulnerability that has led DIT testing to become a central issue within safety testing, (2) the “when” of DIT testing or the circumstances that would be expected to result in DIT testing, and (3) the “how” of DIT testing or the key considerations that can guide an effective testing strategy for health-risk reduction.

2. The “Why” of DIT Testing Epidemiological studies and animal studies have consistently demonstrated the adverse effects of exogenous agents on the developing immune system that last longer or that occur at lower doses than effects of the same agents on adults. Therefore, assessing immunotoxicity in adult animals may not adequately reflect the severity or the persistence of the adverse effects following developmental exposure. From a risk assessment perspective, if early-life exposure to toxicants poses the greatest environmental risk for the immune system, then it poses the greatest effect on human health (11). Epidemiological studies of humans environmentally exposed to exogenous agents provide concrete examples of how developmental exposure to toxicants alters immunocompetence and subsequent susceptibility to infections (12–17). Populations in Canada, China, the Netherlands, and Japan accidentally exposed to polychlorinated biphenyls (PCBs) and their associated breakdown products are well-documented examples of DIT. In each of these populations, rates of recurrent otitis media (inflammation of the inner ear), recurrent respiratory infections, and other types of immune dysfunction were higher in developmentally exposed children than in matched controls. Myriad animal studies with PCBs and other chlorinated compounds corroborate these epidemiological data that developmental exposure increases the risk of infections later in life. Numerous other exogenous agents have been implicated as developmental immunotoxicants in animal models, exposed human populations, or both. These include therapeutic agents

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(diazepam, diethylstilbestrol, and dexamethazone), additional environmental chemicals such as pesticides (chlordane, heptachlor, and hexa-chlorobenzene), and metals (lead, mercury, and organotin compounds). Luebke et al. (18) compared the immunotoxicity of five different compounds (diethylstilbestrol, diazepam, lead, TCDD, and tributyltin oxide) following adult or developmental exposure and concluded that until information to the contrary is available, the developing immune system is more sensitive to toxicant exposure than the adult immune system. One important consideration is that the timing of exposure determines the immuno-toxicological outcome, which means that DIT produces a myriad of effects. For example, if lead exposure occurs throughout gestation, the juvenile and adult delayed-type hypersensitivity (DTH) response (a functional measure of cell-mediated immunity) is decreased; if exposure is restricted to the first half of gestation, macrophage function is impaired but later-life DTH response is unaffected (19). Therefore, the evaluation of DIT requires: (1) the understanding of immune system development, (2) the utilization of relevant age-based exposure regimes, and (3) the selection of appropriate immunological outcomes for assessment. These considerations are discussed in the subsequent section.

3. The “When” of DIT Testing As discussed in the prior section, the developing immune system is generally accepted as a more sensitive toxicological target compared with the immune system of an adult. In fact, even the nature of adverse immune outcomes resulting from early-life exposures is not reliably predictable, based on adult-exposure immunotoxicity results (11). This disconnection between adult-exposure immunotoxicity data and early-life immunotoxic risk raises two key questions in immunotoxicity testing: (1) Are age-relevant immunotoxicity data needed to ensure adequate protection of the nonadult from exposure to a chemical or drug? and (2) When should DIT studies be conducted in safety testing? In recent years, there has been an increased concern over the protection of children’s health that has been reflected in new government and international agency-sponsored activities (20–25). Not surprisingly, this has extended to an increased interest in DIT. Since many of the significant chronic diseases of childhood feature immune dysfunction (26), this increased interest appears warranted. A comparison of DIT publications from the years 1982– 1994 vs.1996–2008 makes it clear that the number of DIT studies, workshops, conference symposia, and reviews has

Additional host resistance testing may be requested

Developmental immune testing after most adultexposure testing

DIT testing may be triggered with adverse adult outcome

Secondary tier immune testing of adult exposed animals: may or may not include secondary immunizations

Pediatric-relevant immune data: are highly drug-specific and usually entail histopathology on unchallenged animals

Adult exposure immune data requested on challenged animals

Cause of concern requirement before proceeding to functional testing

Histopathology on unchallenged animals: routinely adult-exposure

Drugs

Adult exposure immune functional testing

DIT functional immune testing on primary and secondarily challenged animals exposed across all non-adult windows

Any additional narrow age-based or gender-specific testing

Chemicals and Drugs*

Histopathology and associated immune and/or host resistance parameters evaluated on adult-exposed animals

DIT histopathology and associated immune and/or host resistance parameters evaluated

*DIT testing would not be expected for a drug never consumed by a pregnant woman or a child

Sample Immunotoxicity Testing Flow Chart Based on Disease Risk Potential

Fig. 2.1. The flow diagrams on the left half of the figure illustrate the placement of DIT safety testing for chemicals and drugs based on recent regulatory expectations. In contrast, the flow chart on the right half of the figure illustrates a potential placement of DIT testing driven by its potential for disease risk reduction.

LEVEL 3

LEVEL 2

LEVEL 1

First tier immune testing of adult exposed animals: usually primary immunization only

Chemicals

Placement of DIT Testing in Recent Regulatory Requirements

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increased significantly in recent years (10, 11, 18, 26–36). As a result, more information is now available on age-based immune safety for selected chemicals and drugs. However, this increase appears to be largely a result of increased government- and industry-funded research into DIT as well as recent National Toxicology Program (NTP) contracts for early-life exposure studies. In contrast, regulatory expectations for DIT testing have remained relatively unchanged across the years with limited exceptions (e.g., pesticide safety). As illustrated in Fig. 2.1, current immunotoxicity testing of drugs and chemicals expected by regulatory agencies is focused on adult-exposure assessment. Additionally, the collection of specific immune data from adult exposures is often predicated on a “cause for concern” triggered by initial, more general data. DIT exposure assessment is not routinely expected and would usually be triggered only by evidence of clear immunotoxicity from the adult-exposure data. The problem with this priority of testing is that adverse immune outcomes from an insensitive toxicological response system (the fully matured adult immune system) are used as a prerequisite for pursuing DIT testing. The strategy of evaluating risk to the developing immune system only if and when the fully matured immune system (i.e., the adult trigger) is altered should produce two types of errors. The first is essentially a quantitative error. In this case, the adultexposure immunotoxicity data would not provide an indication of the dose–response sensitivity, persistence of adverse effects, or range of adverse effects that the same chemical or drug might produce with early-life exposure (11, 18). If direct DIT testing is pursued, this information is then available. But in the absence of additional DIT testing, addition of safety factors may be applied to reduce age-based risk for some immunotoxicants. Because age-based sensitivities vary widely (18), these may or may not be sufficient. The second type of error is more qualitative and of greater concern. If a chemical or drug is not identified as an immunotoxicant (based on adult-exposure results), then it may never be tested for DIT. Yet, the chemical or drug may be capable of producing an adverse immunotoxic outcome in early life at relevant exposures. While it is not clear if examples of this second type of error do exist, it is also likely that adult-defined “non-immunotoxicants” will never be tested for early-life risk of facilitating allergy, autoimmunity, or targeted immunosuppression. Therefore, the data may not exist to address the likelihood of overlooking developmental immunotoxicants. One advantage of using DIT testing earlier in the safety testing regime (shown in Fig. 2.1) is the benefit of having evaluated the most sensitive age-group for potential immunotoxicity. A negative finding in a comprehensive DIT assessment should

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provide safety information that extends to the adult immune system as well. However, the reverse is not true in that negative adult immune exposure results provide little assurance for earlylife immune safety.

4. The “How” of DIT Testing The specific protocols described in the assay-specific chapters within this book are directly relevant not only to adult-exposure assessment but also to DIT testing. Therefore, they should be considered as having direct relevance across age groups. Hence, there is no need to repeat details of these same assays in this background chapter on DIT. Most of the scientific discussions surrounding the use of specific assays and immune parameters in DIT testing concern: (1) the most informative version of each assay to use, (2) the optimum (or minimum) combination of assays needed for an informative DIT assessment, and (3) the timing of applying these assays in DIT assessment. There is general agreement that a routine screen of DIT would normally include exposure to the environmental agent over the entire developmental period of the nonadult (prenatal, neonatal, juvenile) with immune assessment occurring during the juvenile and/or young adult periods (8). More restricted or specialized exposure regimes could be applied as needed to address specific age-based concerns or the most relevant human exposure (e.g., proposed use of a drug in the pediatric population). DIT assessment should be performed on a challenged immune system (immunized or exposed to an infectious agent) to provide the opportunity to detect potential immune dysfunction (32). Beyond those basic suggestions, the optimum combination of assays and parameters to be employed in the assessment is the subject of considerable discussion. However, given that DIT testing is unlikely to be structured into a multiple tier approach, as are some adult immunotoxicity testing regimes, the probable one-time assessment needs to include a sufficient range of immune measures to address health concerns. Like adult-exposure immunotoxicity assessment, DIT testing is designed to identify potentially problematic exposures and to reduce immune-associated health risks. To accomplish the latter, it is useful to identify target diseases that could be impacted by a reduction of childhood and adult adverse immune outcomes and would serve as the justification for DIT testing. Dietert (26) started with eight of the most significant diseases or conditions of children and young adults having environmental risk factors and featuring immune dysfunction. Most of these diseases are chronic in nature, have increased in prevalence in recent decades and

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include: allergies (including asthma), autism, childhood leukemia, late-onset sepsis, multiple sclerosis, otitis media, pediatric myalgic encephalomyelitis, and type 1 diabetes. A reverse engineering approach was taken toward the goal of optimizing DIT testing for human health-risk reduction. The author asked how one might identify the earliest signs of the immune problems found with the eight childhood diseases and then progressed backwards to the immune parameters that were most useful for this identification. The results from this exercise led to a series of generalized priorities that may prove helpful in guiding effective DIT testing. These are presented here as a series of nine questions that should be useful in a consideration of specific DIT testing regimes and the desired combination of immune parameters to be included in DIT assessments. 1. Was the immune system adequately challenged to permit detection of immune dysfunction, including those parameters prominent during the secondary immune responses? 2. Did the measures permit a sensitive detection of changes in T helper (Th) balance (Th1, Th2, Th17)? 3. Was there an adequate assessment of cell-mediated immunity? 4. Was the potential risk of autoimmunity determined (involving changes in T regulatory cells and/or T cell receptor diversity)? 5. Was innate immune maturation adequately evaluated? 6. Was the status of marginal zone B lymphocytes and the potential responsiveness to T-independent antigens determined? 7. Was the status of resident macrophage populations such as microglia evaluated? 8. Did the assessment parameters evaluate immune cell recruitment and trafficking? 9. Was mucosal immune status determined [including the status of the bronchus-associated lymphoid tissue (BALT) and the gastrointestinal-associated lymphoid tissue (GALT)]?

5. Summary Numerous immunotoxicity testing protocols are detailed in the subsequent chapters of this book and virtually all of these are directly applicable to use in DIT testing. A broader issue concerns the conditions under which these protocols would be applied to DIT testing within a safety testing regime. In the past, DIT testing has been a relatively rare regulatory requirement. However,

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the increasing prevalence of immune-based childhood diseases coupled with the uncertainties inherent in applying adult-exposure immunotoxicity data to predict early-life health risks should increase the role of DIT testing in safety evaluation. With this in mind, it is important that DIT testing designs optimize detection of those adverse immune outcomes that contribute to the most significant health risks of the at-risk population (prenatal, neonatal, juvenile). This introductory chapter has provided disease-pertinent immune information that should prove useful in designing DIT testing strategies.

Acknowledgments The authors thank Janice Dietert for her editorial assistance.

References 1. Luster MI, Faith RE, Kimmel CA (1978) Depression of humoral immunity in rats following chronic developmental lead exposure. J Toxicol Environ Pathol 1:397–402 2. Barnett JB, Holcomb D, Menna JH, Soderberg LS (1985) The effect of prenatal chlordane exposure on specific anti-influenza cell-mediated immunity. Toxicol Lett 25:229–238 3. Dietert RR, Qureshi MA, Nanna UC, Bloom SE (1985) Embryonic exposure to aflatoxin-B1: mutagenicity and influence on development and immunity. Environ Mutagen 7:715–725 4. Holladay SD (2005) Developmental immunotoxicology. CRC Press, Boca Raton, FL 5. Barnett JB (1996) Developmental immunotoxicology. In: Smialowicz RJ, Holsapple MP (eds) Experimental Immunotoxicology. CRC Press, Boca Raton, FL, pp 47–62 6. Smialowicz RJ, Brundage KB, Barnett JB (2007) Immune system ontogeny and developmental immunotoxicology. In: Luebke R, House R, Kimber I (eds) Immunotoxicology and Immunopharmacology, 3rd edn. CRC Press, Boca Raton, FL, pp 327–346 7. Dietert RR (2009) Developmental immunotoxicology. In: Ballantyne B, Marrs T, Syversen T (eds) General and applied toxicology, 3rd edn. Wiley, Chichester, UK, pp 1977–1991 8. Dietert RR, Burns-Naas LA (2008) Develop mental immunotoxicity in rodents. In: Herzyk DJ, Bussiere JL (eds) Immunotoxicology strategies for pharmaceutical safety assessment. Wiley, Hoboken, NJ, pp 273–297

9. Holsapple MP, van der Laan JW, van Loveren H (2008) Development of a framework for developmental immunotoxicity (DIT) testing. In: Luebke R, House R, Kimber I (eds) Immunotoxicology and Immunopharma cology, 3rd edn. CRC Press, Boca Raton, FL, pp 327–346 10. Luster MI, Dietert RR, Germolec DR, Luebke RW, Makris SL (2008) Developmental immunotoxicology. In: Sonawane B, Brown R (eds) Encyclopedia of environmental health. Elsevier, Oxford 11. Dietert RR, Piepenbrink MS (2006) Perinatal immunotoxicity: why adult exposure assessment fails to predict risk. Environ Health Perspect 114:477–483 12. Lu YC, Wu YC (1985) Clinical findings and immunological abnormalities in Yu-Cheng patients. Environ Health Perspect 59:17–29 13. Nakanishi Y, Shigematsu N, Kurita Y, Matsuba K, Kanegae H, Ishimaru S, Kawazoe Y (1985) Respiratory involvement and immune status in Yusho patients. Environ Health Perspect 59:31–36 14. Yu ML, Hsin JW, Hsu CC, Chan WC, Guo YL (1998) The immunologic evaluation of the Yucheng children. Chemosphere 37: 1855–1865 15. Dewailly E, Ayotte P, Bruneau S, Gingras S, Belles-Isles M, Roy R (2001) Susceptibility to infections and immune status in Inuit infants exposed to organochlorines. Environ Health Perspect 108:205–211

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associated with exposure to chemicals. WHO Publications, Geneva 26. Dietert RR (2008) Developmental immunotoxicity (DIT) in drug safety testing: matching DIT testing to adverse outcomes and childhood disease risk. Curr Drug Saf 3:216–226 27. Holladay SD, Smialowicz RJ (2000) Develop ment of the murine and human immune system: differential effects of immunotoxicants depend on time of exposure. Environ Health Perspect 108(Suppl. 3):463–473 28. Dietert RR, Etzel RA, Chen D, Halonen M, Holladay SD, Jarabek AM, Landreth K, Peden DB, Pinkerton K, Smialowicz RJ, Zoetis T (2000) Workshop to identify critical windows of exposure for children’s health: immune and respiratory systems work group summary. Environ Health Perspect 108(Suppl. 3): 483–490 29. Luster MI, Dean JH, Germolec DR (2003) Consensus workshop on methods to evaluate developmental immunotoxicity. Environ Health Perspect 111:579–583 30. Holsapple MP, Paustenbach DJ, Charnley G, West LJ, Luster MI, Dietert RR, Burns-Naas L (2004) Symposium summary: children’s health risk-What’s so special about the developing immune system? Toxicol Appl Pharmacol 199:61–70 31. Luster MI, Johnson VJ, Yucesoy B, Simeonova PP (2005) Biomarkers to assess potential developmental immunotoxicity in children. Toxicol Appl Pharmacol 206:229–236 32. Dietert RR, Holsapple M (2007) Methodo logies for developmental immunotoxicity (DIT) testing. Methods. 41:123–131 33. Selgrade MK (2007) Immunotoxicity: the risk is real. Toxicol Sci 100:328–332 34. Burns-Naas LA, Hastings KL, Ladics GS, Makris SL, Parker GA, Holsapple MP (2008) What’s so special about the developing immune system? Int J Toxicol 27:223–254 35. Dietert RR (2009) Developmental immunotoxicology: focus on health risks. Chem Res Toxicol 22(1):17–23 36. Dietert RR, Zelikoff JT (2008) Early-life environment, developmental immunotoxicology, and the risk of pediatric allergic disease including asthma. Birth Defects Res B Develop Reprod Toxicol 83(6):547–560

Chapter 3 An In Vivo Tiered Approach to Test Immunosensitization by Low Molecular Weight Compounds Irene S. Ludwig, Lydia M. Kwast, Daniëlle Fiechter, and Raymond H.H. Pieters Abstract New chemical entities are tested in general toxicity assays during development before entering clinical trials. However, immunosensitization of these entities is not tested on a standard basis. There are no in vitro or in vivo standardized methods available for testing immunosensitization or immunostimulation. In this chapter, we describe a tiered strategy oral exposure model for assessing immunosensitization or immunostimulation capacity of low molecular weight compounds. The strategy starts from a set of data that may provide information on bioactivation, conjugation (hapten–protein conjugate formation), cytotoxicity and signs of inflammation in any of the animals in a 28 day-toxicity study. In case of concern, a reporter antigen–popliteal lymph node assay (RA–PLNA) and, subsequently, an oral exposure experiment with the reporter antigen can be performed. Based on the presence of RA-specific immune responses an indication for immunosensitization can be found. Key words: Drug-induced hypersensitivity, Reporter antigen, Mouse model, Oral, Relevant route, Antibodies

1. Introduction Many drugs are known to induce immune-mediated adverse effects in susceptible patients. The incidence of these reactions is usually low for a certain drug; however, the impact on the affected individuals can be very high. In some cases, a hypersensitivity response to a drug can lead to toxic epidermal necrosis with lethal effects. The background of susceptibility is not fully understood, yet. New drugs are tested in general toxicity assays during the development process before entering clinical trial. Toxicity based on pharmacological properties is assessed in these assays. However,

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idiosyncratic drug reactions, e.g., immunosensitization, cannot be predicted with these tests. There are also no standardized in vitro or in vivo methods available for testing immunosensitization or immunostimulation. The reporter antigen popliteal lymph node assay (RA–PLNA) is an in vivo assay that can be indicative of immuno-irritation or immunosensitization of low molecular weight compounds (1). Two well-defined antigens, Trinitrophenyl– Ovalbumin (TNP–OVA), which is a “regular” T cell-dependent antigen, and TNP–Ficoll, which is a so called T cell-independent type 2 antigen, are used in the RA-approach in the PLNA. In this assay, the immuno-enhancing capacity of low molecular weight compounds is measured by increased antibody responses to the reporter antigen that is co-injected in a low non-sensitizing dose. Although this assay is indicative of possible effects of the low molecular weight compound on the immune system, it also has its disadvantages. First, the effect of biotransformation of the low molecular weight compound is neglected since it is injected s.c.. Second, only local responses are measured. Since most drugs are administered orally in humans, there is the need for an oral exposure model. In this chapter, we describe a tiered approach to test for immunosensitization by a low molecular weight compound (2) with the combination of RA–PLNA and a mouse model with repeated oral exposure to a low molecular weight drug which offers the possibility to measure systemic responses. In the oral exposure model, similar to the RA–PLNA assay, a reporter antigen is used for detection of a response. Mice are exposed to drugs once daily for several consecutive days, and on the first day co-exposed to the RA (3). Systemic responses as delayed type hypersensitivity (DTH) and serum antibody levels can be determined as well as local secondary responses in the draining lymph node. A DTH response to the RA is indicative of a T-cell-mediated response. RA-specific antibody secreting cells (ASC) and RA-antibodies are indicative of B-cell responses. Furthermore, a distinction between Th1 and Th2 responses can be made based on the type of antibody induced by the low molecular weight compound; in the mouse, Th1 responses are characterized by IgG2a and Th2 responses by IgG1 induction. The cytokine profile of in vitro restimulated lymphocytes can give an indication of T cell skewing to Th1 or Th2 response with the induction of IFNg and IL-4 respectively. Altogether, these series of experiments can give an indication of induction of an immune response and the type of response by low molecular weight drugs.

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2. Materials 2.1. Subcutaneous Exposure

1. Female mice (weighing ~20 g), 5–6 weeks old at the time of arrival. 2. Test compound(s) and vehicle. 3. TNP–OVA (Biosearch Technology, Inc. Novato CA, USA). Prepare stock solution of 20 mg/mL TNP–OVA in NaCl and store in aliquots at −20°C until use. 4. 1 mL syringe and 25 G 16 mm needle. 5. Forceps and scissors. 6. Petri dish. 7. RPMI supplemented with penicillin/streptomycin and FCS (2 and 10%).

2.2. Oral Exposure

1. Female mice (weighing ~20 g), 5–6 weeks old at the time of arrival. 2. Test compound(s) and vehicle. 3. Gavage needle (21 G, 34 mm) with a rounded bulb at the end and 1 mL syringes. 4. TNP–OVA (Biosearch Technology, Inc. Novato CA, USA). Prepare stock solution of 20 mg/mL TNP–OVA in NaCl and store in aliquots at −20°C until use. 5. 18 G needle and minicollect tubes (Greiner) for blood collection. 6. Euthanasate (Pentobarbital or CO2). 7. Forceps and scissors. 8. Petri dish. 9. RPMI supplemented with penicillin/streptomycin and FCS (2 and 10%).

2.3. Delayed Type Hypersensitivity Assay

1. TNP–OVA 10 mg/20 mL NaCl and sterile NaCl. 2. Insulin needle with syringe (29 G, 12 mm, Terumo Europe N.V., Leuven, Belgium). 3. Digital microcalliper. 4. Anesthetic (e.g., ketamine/xylamzine mixture or isoflurane).

2.4. Preparation of Single Cell Suspension of Lymph Nodes

1. RPMI supplemented with penicillin/streptomycin and FCS (2 and 10%). 2. Glass slides with frosted ends. 3. 25 G 16 mm needles and 1 mL syringes. 4. Conic tubes (4 mL). 5. 25 G needles and 1 mL syringes.

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2.5. Serum Ig ELISA

1. Carbonate buffer 0.05 M pH 9.6: 318 mg Na2CO3, 586 mg NaHCO3. (a) Dissolve in 200 mL bidest. (b) Adjust pH to 9.6 with 6 N NaOH. (c) Store at 4°C. 2. Diethanol amine buffer pH 9.8: 97 mL diethanol amine, 0.2 g MgCl2⋅6H2O, (a) Dissolve in ±800 mL bidest. (b) Adjust pH with 1 N HCl to 9.8. (c) Complete up to a liter. (d) Store at 4°C. (e) Protect from light. 3. Substrate solution: 1 mg/mL 4-nitrophenylphosphate in diethanol amine buffer. Prepare just before use. 4. PBS 10×: 80 g NaCl, 2 g KCl, 17.4 g Na2HPO4⋅7H2O, 2 g KH2PO4. (a) Dissolve ±800 mL bidest. (b) Complete up to a liter with bidest. (c) Store at room temperature. (d) Dilute PBS 10× before use 10 times with bidest, and adjust pH (7.2–7.4) to get PBS 1×. 5. PBS/Tween 10×: 80 g NaCl, 2 g KCl, 17.4 g Na2HPO4·7H2O, 2 g KH2PO4. (a) Dissolve ± 800 mL bidest. (b) Add 5 mL Tween-20. (c) Complete up to a liter with bidest. (d) Autoclave and store at room temperature. (e) Dilute PBS/Tween 10× before use 10 times with bidest, and adjust pH (7.2–7.4) to get PBS/Tween 1×. 6. PBS/Tween/BSA (1%): add 1 g BSA (Sigma A-4503) to 100 mL PBS/Tween. 7. Automated reader ELX800 (Bio-Tek Instruments, Winooski, VT).

2.6. ELISpot

1. PBS/Tween/1%BSA: add 1 g BSA (Sigma A-4503) to 100 mL PBS/Tween. 2. Alkaline–phosphatase (AP) conjugated anti mouse-IgG1 or IgG2a (goat-anti-mouse IgG1-AP (SBA 1070-04) and goatanti-mouse IgG2a-AP (SBA 1080-04) from Southern Biotechnology Associates (SBA), Inc., Birmingham, AL). 3. AP-buffer (500 mL): 6.05 g Trizma base (100 mM) (Sigma T1503), 2.92 g NaCl (100 mM), 5.08 mg MgCl2⋅6H2O (5 mM).

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(a) Dissolve in 400 mL MiliQ water. (b) Adjust pH to 9.5 with HCl. (c) Add MiliQ water until a volume of 500 mL. (d) Store at room temperature. 4. Substrate solution; prepare this solution just before use. (a) Dissolve 1 tablet of Nitro Blue Tetrazolium (NBT, Sigma N5514) in 30 mL of AP-buffer. (b) Filter solution over filter paper when tablet is dissolved. (c) Prepare BCIP (5-Bromo-4-chloro-3-indolyl phosphate p-toluidine salt, Sigma B8503) solution just before use (35 mg/mL in dimethylformamide). (d) Add 142 mL of BCIP solution to 30 mL AP buffer with NBT. Be careful: compounds are toxic: wear gloves, and prepare solutions in fume cab. 5. ELISpot blocks (made in house) and protein blot membranes (Immobilon PVDF Transfer, Millipore, Etten-Leur, Netherlands) or commercial ELISpot plates with a well surface of approximately 2 cm2.

3. Methods 3.1. Subcutane Exposure and Experimental Setup RA–PLNA

1. Acclimate female mice (5–6 weeks old at arrival) for 1 week before starting the experiment. 2. Randomize the mice into 6–8 animals per treatment group. 3. House the animals in solid-bottom cages with woodchip bedding. Provide drinking water and standard laboratory food pellets ad libitum. House the animals according to the guidelines of The Association for Assessment of Laboratory Animal Care. 4. Administer the low molecular weight compound in combination with 10 mg reporter antigen in a total of 50 mL vehicle (see Note 1). 5. Prepare at least three different doses and one vehicle as control (see Note 2). 6. Inject the compound and reporter antigen solution subcutaneously in the footpad toe-to-heel with a 1 mL syringe with a 25 G 16 mm needle. 7. Check animals daily for irritation of the paw and general health state. 8. Sacrifice the mice 7 days after the s.c. injection by cervical dislocation. 9. Wet the hind leg with 70% ethanol.

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10. Make an incision in the skin from the heel without cutting the tendon. 11. Tear the skin slowly from the leg until the knee joint is exposed. 12. Stretch the leg and use a forceps to isolate the PLN. 13. Place lymph nodes in a small Petri dish with RPMI supplemented with 2% fetal calf serum (FCS) on ice until further use. The PLNs can be used for immunohistology, or for the preparation of single cell suspensions to be used in functional assays. Several assays can be performed with the single cell suspensions, e.g., an Enzyme Linked Immunosorbent Spot (ELISpot) Assay (see Subheading 3.3.5) to determine the formation of RA-specific ASC, ex vivo restimulation (see Subheading 3.3.7) to detect cytokine production profile, and flow cytometry (see Subheading 3.3.6) to detect changes in cell populations or activation status of cells (4). 3.2. Oral Exposure and Experimental Setup

1. Acclimate female C3H/HeOuJ mice (5–6 weeks old upon arrival) for a week before starting the experiment.

2. Randomize the mice in 6–8 animals per treatment group and house the animals in solid-bottom cages with woodchip bedding. 3. Provide drinking water and standard laboratory food pellets ad libitum. 4. House the animals according to the guidelines of The Association for Assessment of Laboratory Animal Care. 5. Give the mice a daily dose of low molecular weight drugs for several days orally using a gavage needle (21 G, 3.4 cm long) (see Notes 1 and 3) 6. Preferably, dilute the test compound in 200 mL sterile H2O or PBS. 7. On the first day, give a low dose of reporter antigen (TNP– OVA) (10 mg/mouse) i.p. in 100 mL sterile NaCl simultaneously with the first oral dose of drugs. 8. Check the mice daily for general health state. 3.2.1. Collecting Blood

1. Collect blood one day before the first drug administration, 10 days after the first drug administration and at the final day of the experiment, before euthanizing the animals. 2. Collect blood in minicollect tubes (Greiner Bio-One, Alphen a/d Rijn, Netherlands) with serum separation gel by a puncture with an 18 G needle in the submandibular vein. This vein is localized in the cheek of the mice.

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3. Centrifuge the blood at 2,000 × g for 7 min at 4°C to obtain serum. 4. Store serum samples at −20°C until analysis. 5. Determine reporter antigen specific antibodies in the serum using a sandwich ELISA (see Subheading 3.3.4). 3.2.2. Delayed Type Hypersensitivity Response

1. On day 15 after the start of the experiment, determine the DTH response to the RA. 2. Measure ear thickness of both left and right ear of the mice with a digital microcalliper before injection. 3. Inject one ear of each mouse with a low dose of RA (10 mg TNP–OVA in 20 mL sterile NaCl). To control the effect of injection liquid in the ear, inject one ear per mouse with 20 mL sterile NaCl only as a reference. (a) Use an insulin syringe (29 G, 12 mm) because of the fine needle and small scale on the syringe. (b) Apply the needle from the top of the ear to the base. (c) Inject the 20 mL very carefully avoiding disruption of the ear. (d) All the operations are performed under isoflurane anesthetics. 4. After 24 h, determine the thickness of both ears again with the digital caliper. 5. Calculate the DTH response as the difference in ear thickness of TNP–OVA injected ear minus the difference in ear thickness of the vehicle injected ear. 6. The DTH response can also be performed in the hind foot pad (see Note 4).

3.2.3. Isolation of the Auricular Lymph Node

1. At day 21, 6 days after DTH measurement, take blood from the submandibular vein. 2. Euthanize the mice. Cervical dislocation is not preferable since this method could disrupt the structure around the auricular lymph node (ALN). 3. Moisten the neck and facial area with 70% ethanol. 4. Isolate the ALN at the side of the TNP–OVA injected ear by making a cut from the ear down the cheek. 5. Carefully remove the skin from the underlying tissue by gently tearing the skin with two forceps. The ALN is localized at the bifurcation of the jugular vein posterior of the masseter muscle. 6. Carefully remove the ALN with a pair of forceps. 7. Place lymph nodes in a small Petri dish with RPMI supplemented with 2% fetal calf serum (FCS) on ice until further isolation of cells.

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3.3. Preparation of Cell Suspensions

1. For the preparation of a single cell suspension, place the individual lymph nodes between two glass slides at the frosted sides and carefully crush the lymph nodes. Keep the glass slides wet with RPMI all the time. 2. Take the suspension from the dish with a syringe with an orange needle and place in a conic 4 mL tube. 3. Spin down the cell suspension at 230 × g for 10 min and resuspend in RPMI 10% FCS. 4. Count the cell numbers. 5. Dilute the cell suspensions to the desired concentration. A concentration of 10e6 cells/mL is sufficient for ELISpot.

3.4. RA-IgG1 and IgG2a ELISA for Serum Samples

1. Coat hibond 96 wells plates (costar 3590) with TNP–BSA in carbonate buffer (20 mg/mL) 100 mL/well overnight at 4°C. 2. Wash wells three times with PBS/Tween (200 mL/well). 3. Block the plates with PBS/Tween/1% BSA (150 mL/well) for 1 h at RT (see Note 5). 4. Make serial dilutions of the serum samples starting with an 8× dilution of the sample in PBS/Tween/1%BSA. 5. Prepare for the detection of IgG1 8 two-step dilutions and for IgG2a 6 two-step dilutions. Also include wells with only PBS/Tween/1%BSA as blanks to determine the background staining. 6. Add 100 mL of the sample to each well. 7. Incubate for 2 h at room temperature. 8. Wash the plates subsequently 3 times with PBS/Tween. 9. Prepare just before use alkaline–phosphatase conjugated anti mouse-IgG1 or IgG2a (goat-anti-mouse IgG1-AP (SBA 1070-04) and goat-anti-mouse IgG2a-AP (SBA 1080-04) in PBS/Tween/1%BSA. Add 100 mL of AP-conjugated antibody per well and incubate for 1 h at room temperature. 10. Wash wells three times with PBS/Tween and once with diethanolamine buffer. 11. Do not discard diethanolamine buffer in the sink but in a special vessel for halogen poor waste. 12. Add diethanolamine buffer with 1 mg/mL 4-nitrophenylphosphate (100 mL/well). 13. Incubate in the dark at room temperature until a color change is observed (about 30 min). 14. Stop substrate conversion with 50 mL 10% EDTA in bidistilled water per well. 15. Measure the optical density at 405 nm using an automated reader ELX800.

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16. Plot data as optical density per dilution or as maximal dilution at which OD is above background. 3.5. ELISPOT Assay

1. Determine the induction of RA-specific ASC by the treatment of the mice by means of ELISpot assay (5). 2. Prewet the protein blot membranes (Immobilon PVDF Transfer) with methanol in a fume cab (see Notes 6 and 7). 3. Wash the membranes with filtered PBS/Tween (22 mm). 4. Coat the membranes with TNP–BSA (10 mg/mL, 15 mL per blot) in PBS/Tween over night on a shaking table at 4°C. 5. Wash the membranes 2 times with PBS/Tween. 6. Incubate for 1 h at room temperature with PBS/Tween/1%BSA while shaking to block the membrane for a specific protein binding (see Note 8). 7. Wash membranes 2 times with PBS. 8. Place in a special holder and add PBS to the wells (see Note 9). 9. When cell suspensions are ready, discard the PBS from the wells and add 500 mL of cell suspension (500 × 10e3 cells is usually sufficient). 10. Centrifuge the blocks at 230 × g for 7 min without the brake. 11. Incubate the blocks for 4 h at 37°C, 5%CO2. 12. Discard the cells and wash the wells once with PBS. 13. Release the membranes from the holders and wash once more with PBS and twice with PBS/Tween. 14. Incubate the blots with 15 mL AP-conjugated goat anti mouse-IgG1, IgG2a or other isotype (goat-anti-mouse IgG1-AP (SBA 1070-04) and goat-anti-mouse IgG2a-AP (SBA 1080-04), 1/2,000 in PBS/Tween) over night on a shaking table at 4°C. 15. Wash the membranes 4 times with PBS/Tween and once with AP buffer. 16. Incubate the membranes with substrate solution for approximately 15 min while shaking. 17. Do not discard substrate buffer in the sink but in a special vessel for halogen poor waste. 18. Stop the color reaction by rinsing the blots for 15 min with slow running tap water. 19. Dry the membranes between filter paper for about 2 h. 20. Count the spots with a microscope. Spots appear as dark purple spots with a diffuse halo.

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3.6. Flow Cytometry

1. Analyze cell suspensions using flow cytometry for changes in cell subset distribution. This assay gives more clear results for RA–PLNA than for LN cells isolated from oral exposed animals. 2. Plate 150,000 cells per sample in round bottom 96 wells plates in PBS/0.05%BSA/0.1%NaN3. 3. Keep cells at 4°C all the time. 4. Incubate the cells with anti-CD3-, CD8-, and CD4fluorescence labeled mAbs for 30 min at 4°C in order to detect CD4+ (T helper) or CD8+ (cytotoxic) T cells. 5. Wash cells twice in PBS/0.05%BSA/0.1%NaN3 at 230 × g for 5 min at 4°C. Samples can be analyzed immediately or after 3 days maximum. 6. When the samples are not analyzed immediately, fix samples in 100 mL 1% formalin. 7. Detect other cell subsets, like B-cells, macrophages and dendritic cells using the appropriate antibodies.

3.7. Cell Culture

1. Analyze polarization of the T-cell response in the draining lymph node. (a) Culture them with or without a stimulus. (b) Test for IFN-g and IL-4 secretion. 2. Incubate 250,000 per well in a round bottom 96 well plate in a total of 200 mL. 3. Use RPMI with 10% FCS in this assay. 4. Use LPS (2 mg/mL), con A (5 mg/mL), or anti-CD3/antiCD28 antibodies as general stimuli. 5. Incubate cells at 37°C, 5% CO2. 6. Collect the supernatant after 3 days. 7. Store supernatants at −20°C until analysis. 8. Measure IL-4 and IFN-g production using commercially available ELISA kits.

4. Notes 1. Use in both RA–PLNA and oral exposure assay at least three different doses of low molecular weight compound. Do not use solvents that have an intrinsic effect on the immune response, and therefore always include a control group that will receive solvent and RA only.

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2. Use a separate group of animals exposed to vehicle and RA only as negative control group instead of using the contralateral popliteal lymph node (PLN) of the chemical treated mice. 3. Be careful while applying the gavage needle. Harming the esophagus or the stomach can give a false positive test result. 4. The DTH response can also be performed in the hind foot pad. (a) Apply the needle from in-between the toes to the heel. (b) The s.c. injection in the footpad is easier. However, the readout of swelling is less sensitive than in the ear. 5. PBS/Tween with 3% milk powder will also sufficiently block the wells in this assay. 6. Never touch membranes with your fingers because the membranes will also bind proteins from your skin and this can give high background; wear gloves or use pair of forceps. 7. Never leave the membranes without buffer. If membranes are dry you have to prewet them with methanol before you can continue. 8. Coated (and blocked) membranes can be stored in a refrigerator (4°C) for about 2 weeks; dry them between filter paper (for ±2 h) before storage. Before use, prewet the membranes with methanol, wash them once with PBS, and put them in special holder with PBS. 9. There are complete prefab ELISpot plates commercially available. Well surface should be around 2 cm2.

References 1. Albers R, Broeders A, van der Pijl A, Seinen W, Pieters R (1997) The use of reporter antigens in the popliteal lymph node assay to assess immunomodulation by chemicals. Toxicol Appl Pharmacol 143:102–109 2. Pieters R (2007) Detection of autoimmunity by pharmaceuticals. Methods 41:112–117 3. Nierkens S, Aalbers M, Bol M, van Wijk F, Hassing I, Pieters R (2005) Development of an oral exposure mouse model to predict drug-induced hypersensitivity reactions by using reporter antigens. Toxicol Sci 83: 273–281

4. Nierkens S, van Helden P, Bol M, Bleumink R, van Kooten P, Ramdien-Murli S, Boon L, Pieters R (2002) Selective requirement for CD40-CD154 in drug-induced type 1 versus type 2 responses to trinitrophenyl–ovalbumin. J Immunol 168:3747–3754 5. Schielen P, van Rodijnen W, Tekstra J, Albers R, Seinen W (1995) Quantification of natural antibody producing B cells in rats by an improved ELISPOT technique using the polyvinylidene difluoride membrane as the solid support.J Immunol Methods 188: 33–41

Chapter 4 Risk of Autoimmune Disease: Challenges for Immunotoxicity Testing Rodney R. Dietert, Janice M. Dietert, and Jerrie Gavalchin Abstract Autoimmunity represents a potentially diverse and complex category among the range of adverse outcomes for detection with immunotoxicity testing. For this reason, the risk of autoimmune disease is discussed in this overview chapter with additional mention among the later specific protocol chapters. Improvements in clinical diagnostic capabilities and disease recognition have led to a more accurate picture of the extent of autoimmune diseases across different human populations. While the risk of any single autoimmune disease remains modest when compared with that of lung or heart disease, the cumulative prevalence of autoimmune diseases is both significant and increasing. Autoimmune diseases are usually viewed in the context of the damaged tissue or organ (e.g., as a thyroid, gastrointestinal, cardiovascular or neurological disease). But improved recognition that underlying immune dysfunction can connect the risks for these as well as other diseases is critical for optimizing risk assessment. Since autoimmune diseases are chronic in nature with many first appearing in children or in young adults, these diseases exert a serious impact on both health care costs and quality of life. This chapter provides a discussion of the issues that should be considered with immunotoxicity testing for risk of autoimmunity. Key words: Autoimmunity, Autoimmune disease, Target organ, Systemic, T cell populations, Immune dysregulation, Chronic inflammation, Microbial triggers

1. Introduction Autoimmune diseases as a group are recognized as contri buting to a significant portion of chronic diseases in humans. As discussed in a recent review by Fairweather et al. (1), autoimmune diseases affect approximately 5–8% of the popu lation in the United States (2). Included among these are approximately 100 reported or proposed conditions that are either autoimmune or chronic inflammatory in nature (3). Given their prevalence in the human population, the role of toxicants in R.R. Dietert (ed.), Immunotoxicity Testing: Methods and Protocols, Methods in Molecular Biology, vol. 598 DOI 10.1007/978-1-60761-401-2_4, © Humana Press, a part of Springer Science + Business Media, LLC 2010

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risk of disease and the prominent role of immune dysfunction in mediating the diseases, detection of potential autoimmune risk has become an increasing concern in immunotoxicity testing. These diseases can take the form of either systemic conditions (e.g., systemic lupus erythematosus, SLE), or they can target individual organs or tissues (e.g., autoimmune hepatitis). Many of these diseases are most notably identified, characterized and treated based primarily on the organ or tissue that suffers from inflammatory-mediated damage. This is despite the fact that most of these diseases involve some aspect of immune dysregulation. Autoimmune diseases can strike at any age, and a majority of the diseases exhibit an age-of-onset range spanning decades of life. However, most of the diseases first appear in the young adult. Figure 4.1 illustrates a timeline with the approximate age of onset

Fig. 4.1. The timeline illustrates the approximate average age of onset for 18 different autoimmune diseases. Of the diseases listed, most have established autoimmune associations. Hidradenitis suppurativa is a chronic inflammatory condition with a suspected autoimmune basis.

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shown for several established and/or suspected autoimmune diseases and conditions. Note that some of the autoimmune diseases exist in both juvenile and adult forms (e.g., arthritis and celiac dis ease). Additionally, for some of the diseases, males and females differ in their average age of onset. Therefore, these should only be used as approximations. The lack of prior appreciation for the underlying immune dysfunction affecting autoimmunity has been an impediment, not only to a life-long approach to immune management (4, 5), but also to optimizing detection of autoimmune risk factors during immunotoxicity testing. This has happened for several reasons. Importantly, because of the ready organ association of many of these diseases, autoimmune thyroiditis has been viewed as a thyroid–endocrine problem, autoimmune hepatitis as a liver issue, autoimmune myocarditis as a heart disorder, celiac disease as a gastrointestinal disorder, rheumatoid arthritis as a problem of vascular and connective tissues, multiple sclerosis a neurological disorder, and type 1 diabetes mellitus as a pancreas-insulin issue. However, an additional factor is that the identification and understanding of immune cell regulators of autoimmunity have only occurred recently. The identification of specialized T cell populations such as T helper (Th) 17 cells, Th3 cells, and regula tory T cells (Tregs) (e.g., CD4+CD25+Foxp3+ Tregs) has pro vided useful biomarkers for identifying immunotoxic risk for autoimmunity. As will be discussed later in this chapter, a recent trend toward improved recognition of the underlying immune dysfunction that supports autoimmune disease is critical for effective health man agement. In part, this is because the same immune dysfunction (e.g., impaired Treg function) is very likely to alter the risk for other autoimmune and non-autoimmune diseases over the course of a lifetime.

2. The Multifactorial Nature of Autoimmune Disease

Multiple factors go into determining the overall risk for autoim mune disease. Certainly genetic background, age and gender are all significant factors influencing the overall risk. The female–male partitioning of autoimmunity is discussed later in this chapter. But apart from genetic and gender-based considerations, most autoimmune diseases are also under environmental influence. In fact, a recent study across numerous strains of mice found that mercury and silver can predispose for autoantibody production regardless of whether known susceptibility genes are carried within the strain (6). The author suggested that environmental determinants may be at least as important as genetic background in

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the risk of autoimmune disease. For this reason, including the capability of evaluating the risk of autoimmune disease within immunotoxicity testing is a significant issue. Numerous chemicals and drugs have been shown to be capable of elevating the risk of autoimmune disease, and this was recently reviewed by Cooper and Miller (7). Examples of environmental factors reported to increase the risk of autoimmunity in animals and/or humans are: mercury (8–10), lead (11, 12), gold (13), trichloroethylene (14), hexachlorobenzene (15, 16), ethanol (17), silica (18, 19), 2,3,7,8-tetrachlorodibenzo-p-dioxin (20), hydralazine and other demethylating agents (21), cyclosporine A (22) and cigarette smoke (23). Table 4.1 provides examples of environmen tal factors that have been reported to increase the risk of lupus or lupus-like symptoms.

Table 4.1 Examples of environmental factors associated with lupus or lupus-like symptomsa Environmental factor

References

2,3,7,8-tetrachlorodibenzo-p-dioxin

(20)

Aniline

(64)

Asbestos

(65)

Bisphenol-A

(66)

Cigarette smoke

(67)

Diethylstilbesterol

(68, 69)

Estrogen

(70)

Infectious agents

(71, 72)

Lead

(12)

Mercury

(10, 73)

Organochlorine pesticides

(74)

Polychlorinated biphenyls

(75)

Prolactin

(76)

Silica

(18, 77)

Sunlight/UVB radiation

(78)

Trichloroethylene

(79)

L-Tryptophan

(80)

From studies in rodents and/or humans. In some studies, induction of lupus-like symptoms was the endpoint measured. In other studies, acceleration of symptom onset or exacerbation of symptoms was detected in autoimmune-prone strains of mice a

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It should be noted that some autoimmune diseases have both juvenile and adult forms, which may differ slightly in presentation (e.g., celiac disease and rheumatoid arthritis). However, even adult forms generally arise no later than middle age. As a result, these diseases are likely to require extensive and expensive health care management over decades of life. This can place considerable strain on patients and the health care community and negatively impact patients’ quality of life. Since the periods of childhood, adolescence and young adulthood are particular focal points for disease onset, they have implications for immunotoxicity assessment. It means that the windows of greatest interest for exposure to potential toxicants occur before adulthood. As a result, exposure-assessment of the non-adult can be critical in assessing the risk for autoimmunity. This is also discussed in the chapter on developmental immuno toxicity (DIT).

3. Environmental Toxicant Exposure and Skewed Host Responses to Microbes as a Problem

Microbial exposure and host response to microbes have long been recognized as important considerations in the risk of autoimmu nity. Historically, much of the focus has been on molecular mim icry in which pathogen-associated molecules that have overlapping epitopes with self molecules elicit host immune responses of a specificity that damage tissues. Suggestions for the involvement of molecular mimicry can be seen with streptococcal infections, autoimmune responses in rheumatic fever (24) and trypanosome infection with chronic Chagas disease cardiomyopathy (25). While there are many avenues to get to autoimmune disease, recent research points toward one that involves skewed host responses to microbes that may trigger autoimmunity. This is dis cussed by Rose (26) and presented as an adjuvant effect in which immune co-factors can skew a response to an infectious agent resulting in tissue pathology that facilitates autoimmunity. But it is important to recognize that from an environmental/toxicant standpoint, the adjuvant component can just as easily be embedded in the dysfunctional host immune response present after earlier environmentally induced immune insult. So, excessive proinflam matory production by immune cells could be an outcome of prior heavy metal-, alcohol-or dioxin-induced immunotoxicity that caused skewed responses to: 1) bacteria contributing to thyroiditis or 2) viruses contributing to myocarditis (27–29). In fact, Vas et al. (30) recently described a model in which low-level mercury expo sure combined with exposure to natural killer T (NKT) cells and toll-like receptor (TLR) ligands from microbes combine to facili tate a break in tolerance and elevated the risk for autoimmunity.

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Thus specific microbial activation of NKT cells could enhance the risk of mercury-induced autoimmunity. This has been discussed by Cooke (31) who noted the likely importance of dysfunctional host responses to microbes as trig gers for both autoimmunity and childhood leukemia. Infections are likely to serve as triggers for many autoimmune diseases, and the underlying inappropriate or dysfunctional host response to the microbe may trigger a critical step leading to autoimmunity. For example, Mattner et al. (32) found that an accumulation of a common bacterium, Novosphingobium aromaticivorans, in the liver and subsequent responses by NKT cells could lead to auto immune liver disease. Environmental modulation of Th17 cells vs. Tregs seems to be a prominent pathway to facilitate autoimmunity. Veldhoen et al. (33) discussed the opportunity for aryl-hydrocarbon (Ah) receptor agonists like 2,3,7,8 tetrachlorodibenzo-p-dioxin to alter Th17 activity and promote autoimmunity.

4. Sex-Specific Issues in Autoimmunity and Testing

Differences among women and men in risk of disease are only now beginning to be fully appreciated (34). While many immune dysfunction-associated diseases can show some evidence of sex bias, either in prevalence of the disease or timing of onset, autoimmune diseases are in a novel category of their own when it comes to the importance of gender. The vast majority of autoimmune diseases are not equally distributed among the sexes. In fact, greater than 75% of autoimmune disease occurs in women (1, 35). While a handful of specific autoimmune diseases are either of equal risk among sexes or slightly more common in men (36, 37), overall autoimmunity is a major women’s health issue (38, 39). The importance of age and sex in autoimmunity is also supported in animal models of these diseases (40–42). There are three main hypotheses for the bases of the female bias in autoimmunity: (1) the effects of sex steroids on environment-gene-immune cell interaction (42, 43), (2) microchimerism among lymphocytes (44, 45), (3) X chromosome monosomy (46), and/or sex chro mosome complement (47). It should be noted that while X chromosome monosomy has been seen with some female-pre dominant autoimmune diseases, it was not found in a recent study with SLE (48). Whatever toxicant-hormone-chromosome-gene interactions distinguish the sexes as per risk for a specific autoimmune disease, these interactions result in detectable differences in immune capacity and host responses to challenge that are connected to the

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increased specific disease risk (49–51). For immunotoxicity test ing, it also means that the specific testing for autoimmunity needs to use relevant immune endpoints and evaluation in both sexes.

5. The Challenge of Identifying Immunotoxic Risk Factors for Autoimmune Disease

6. Role of Autoimmune-Prone Strains in Immunotoxicity Testing?

It should be noted that detection of risk factors for specific auto immune diseases has been a difficult challenge for immunotoxic ity testing until recently. Risk assessment of pharmaceuticals for autoimmunity was recently discussed (52) as were principles for evaluating chemicals in risk of autoimmunity (37). In many cases, identification of an immunotoxicant as con tributing to risk of autoimmunity resulted as either a secondary observation of other immunotoxic endpoints (enhanced immune responses) or was from the association of a specific autoimmune disease in humans with administration of a specific drug [e.g., minocycline and autoimmune hepatitis (53, 54)]. There are several reasons why detection of xenobiotic risk fac tors for autoimmune disease has proven to be a challenge. Most surround the fact that several different xenobiotic–host interac tion mechanisms can contribute to an elevated risk of autoimmu nity. First, some xenobiotics or their metabolites may be molecular mimics of tissue antigens eventually leading to a loss of tolerance against the self-antigen. Alternatively, the xenobiotic may be complex with self components and form new antigens where no prior tolerance has been established. Finally, exposure to a xeno biotic may alter the immune cell populations increasing the risk of loss of tolerance. The example of disruption of Tregs vs. Th17 balance was previously discussed. However, the actual problem atic immune responses may not occur until the immune system receives a relevant challenge (e.g., an infection). A further prob lem is that the disease-associated reactions may only arise among certain genetic backgrounds. The genetic background can influ ence both the metabolism and distribution of the xenobiotic as well as the susceptibility of the host immune system for a prob lematic response to a specific environmental insult. Obviously, it can be difficult to ensure that a routine immunotoxicity screening strategy can account for all of these possibilities.

There are several unresolved issues in approaching immunotoxicity testing for the risk of autoimmunity. However, among the more significant is whether the necessary information needed for

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hazard characterization can be obtained using conventional animal strains (also used for other toxicity testing), or whether specialized strains of rodents or other test species must be used to fully assess the risk of xenobiotic-induced autoimmunity. While numerous scientific reviews have discussed this issue and presented options, there has yet to be a definitive answer relative to safety testing (37). The issue is connected to the role of genetic background and xenobiotic-associated epigenetic effects leading to autoimmunity. Since induction of autoimmunity can involve several different mechanisms, it is likely that a single test strain of a species will not effectively model all the chemical- or drug- induced routes to autoimmunity that may be relevant for a specific test compound. In contrast, strains with spontaneously occurring autoimmunity may be pertinent for only some autoimmune outcomes and mechanisms. There, it is usually autoimmune exacerbation that can be evaluated most effectively instead of disease induction. Some F1 hybrids between autoimmune prone and control strains have been used to ensure that a genetically permissible back ground is present in the test animals but to avoid the high level of spontaneous disease occurrences present in the parental strain. While this may help the modeling of risk for a single autoimmune disease (e.g., SLE) or a category of diseases, the question still remains whether a single test strain (specialized or not) can pro vide the needed information. Examples of autoimmune-prone experimental animal models frequently used for the study of xenobiotic-promoted autoimmu nity (37, 55–57) include: the Brown Norway, Lewis and biobreeding diabetes prone (BB-DP) strains of rats, the New Zealand Black – New Zealand White F1 hybrid (NZB X NZW F1), the MRL-lpr strain and non-obese diabetic (NOD) strain of mice and the Obese strain of chickens.

7. Tiered Approach As discussed in the World Health Organization Environmental Health Criteria Report titled “Principles and Methods for Assessing Autoimmunity”(37) and in the prior section of this chapter, there are numerous animal models available for examin ing the ability of chemicals and therapeutics to either induce or to exacerbate autoimmunity. However, the reality is that these highly specialized animal models are unlikely to be used in a first tier of screening for general hazard identification and characterization. Beyond research tools, their use in immunotoxicity testing would more likely come after some initial immune-related out come was obtained using more routine test animals or alternatives

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(e.g., in vitro tests). Of course, the concern or limitation is whether a more general test animal or cell line will have a suffi ciently permissive genetic background to model the risk for auto immune reactions that may exist across all genotypes. Among screening tools more feasible to be employed in an initial tier, there are several biomarkers or indicators of immune alterations that should cause a heightened concern for risk of autoimmunity. These are also discussed in more detail in the chapters on sensitization, the local lymph node assay (LLNA) and inflammation. Test compound-induced enhancement of endpoints measured in the LLNAs, the delayed-type hypersen sitivity (DTH) response, and the T-dependent antibody response (TDAR) should be of potential concern. Additionally, inflammation is an important component of most autoimmune conditions, and test compound-induced promotion of inflam mation is also a possible indicator for a heightened concern of autoimmunity. Finally, significant changes in T lymphocyte populations involved in regulation, tolerance maintenance and inflammation are likely to play a role in risk of autoimmune responses. Additional endpoints that could be useful within a general screening assessment would be quantitation of anti-DNA and anti-histone antibodies and measures of immune complex formation (37). These could be added to more general screening protocols. But as is discussed in the other chapters, these may be more readily observed during the course of responses following challenge (e.g., infection or immunization) of the immune system. In fact, because microbial triggers are thought to be associated with a significant incidence of human autoimmune disease, there may be benefit in using a natural infection/antigen model in more general immunotoxicity testing. Infection challenge models are more likely to facilitate the inclusion of add-on indicators of autoimmune reactions (e.g., measures of inflammatory mediators, autoantibodies and immune complexes).

8. Conclusions Risk of autoimmune disease is multifactorial and includes not only genetic background, sex and age as factors, but also exposure to immunotoxic chemicals and therapeutic agents (58). Both sexes are affected although most autoimmune diseases are predominant in females (59, 60). However, toxicant-induced promotion of autoimmune disease may itself be sex-biased. Therefore, it is important to include both females and males in any screening.

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Since autoimmune diseases can be either systemic or organspecific, can be mediated by different immune-related responses and can take many forms, reliable detection of autoimmunepromoting environmental exposures is a formidable challenge. While numerous specialized animal models are available to study the environmental-autoimmune interactions, autoimmune-prone strains are unlikely to be used in more generalized safety testing screening. Therefore, biomarkers that may be useful toward the identification of autoimmune responses should be included in the more general screens. These include enhanced antibody, cellmediated and inflammatory responses as well as misregulated or misdirected inflammatory responses. Added measures of autoan tibodies (61), immune complexes and specialized T cell popula tions may be useful. But these may be best assessed using a challenged immune system including exposure to infectious agents (62, 63). The development of new tools to apply to screen ing for risk of autoimmunity is needed and should be a goal of future immunotoxicity testing. References 1. Fairweather D, Frisancho-Kiss S, Rose NR (2008) Sex differences in autoimmune disease from a pathological perspective. Am J Pathol 173:600–609 2. The Autoimmune Disease Coordinating Committee of the National Institutes of Health (2005) Report to Congress, March. NIH Publication No. 05–5140 3. Shoenfeld Y, Selmi C, Zimlichman E, Gershwin ME (2008) The autoimmunologist: geoepide miology, a new center of gravity, and prime time for autoimmunity. J Autoimmun 31:325–330 4. Dietert RD, Piepenbrink MS (2008) The man aged immune system: protecting the womb to delay the tomb. Hum Exp Toxicol 27:129–134 5. Dietert RR, Zelikoff JT (2009) Pediatric immune dysfunction and health risks following early-life immune insult. Curr Pediatr Rev 5(1):36–51 6. Abedi-Valugerdi M (2009) Mercury and silver induce B cell activation and anti-nucleolar autoantibody production in outbred mouse stocks: are environmental factors more impor tant than the susceptibility genes in connection with autoimmunity? Clin Exp Immunol 155:117–124 7. Cooper GS, Miller FW (2008) Environmental influences on autoimmunity and autoimmune disease. In: Luebke R, House R, Kimber I (eds) Immunotoxicology and immunopharmacol ogy, 3rd edn. CRC Press, Boca Raton, FL, pp 437–453

8. Silbergeld EK, Silva IA, Nyland JF (2005) Mercury and autoimmunity: implications for occupational and environmental health. Toxicol Appl Pharmacol 207:282–292 9. Havarinasab S, Bjorn E, Ekstrand J, Hultman P (2007) Dose and Hg species determine the T-helper cell activation in murine autoimmu nity. Toxicology 229:23–32 10. Pilones K, Lai ZW, Gavalchan J (2007) Prenatal HgCl(2) exposure alters fetal cell phenotypes. J Immunotoxicol 4:295–301 11. Bunn TL, Marsh JA, Dietert RR (2000) Gender differences in developmental immu notoxicity to lead in the chicken: analysis fol lowing a single early low-level exposure in ovo. J Toxicol Environ Health A 61:677–693 12. Hudson CA, Cao L, Kasten-Jolly J, Kirkwood JN, Lawrence DA (2003) Susceptibility of lupus-prone NZM mouse strains to lead exac erbation of systemic lupus erythematosus symptoms. J Toxicol Environ Health A 66: 895–918 13. Havarinasab S, Johansson U, Pollard KM, Hultman P (2007) Gold causes genetically determined autoimmune and immunostimu latory responses in mice. Clin Exp Immunol 150:179–188 14. Blossom SJ, Doss JC (2007) Trichloroethylene alters central and peripheral immune function in autoimmune-prone MRL(+/+) mice fol lowing continuous developmental and early life exposure. J Immunotoxicol 4:129–141

Risk of Autoimmune Disease: Challenges for Immunotoxicity Testing 1 5. Cripps DH, Peters HA, Gocman A, Dogramici I (1984) Porphyria turcica due to hexachlorobenzene: a 20 to 30 year followup study on 204 patients. Br J Dermatol 111:412–422 16. Michielsen CC, van Loveren H, Vos JG (1999) The role of the immune system in hexachlo robenzene-induced toxicity. Environ Health Perspect 107:783–792 17. Vidali M, Stewart SF, Rolla R, Daly AK, Chen Y, Mottaran E, Jones DE, Leathart JB, Day CP, Albano E (2003) Genetic and epigenetic factors in autoimmune reactions toward cyto chrome P4502E1 in alcoholic liver disease. Hepatology 37:410–419 18. Brown JM, Pfau JC, Pershouse MA, Holian A (2005) Silica, apoptosis, and autoimmunity. J Immunotoxicol 1:177–187 19. Otsuki T, Maeda M, Murakami S, Hayashi H, Miura Y, Kusaka M, Nakano T, Fukuoka K, Kishimoto T, Hyodoh F, Ueki A, Nishimura Y (2007) Immunological effects of silica and asbestos. Cell Mol Immunol 4:261–268 20. Mustafa A, Holladay SD, Goff M, Witonsky SG, Kerr R, Reilly CM, Sponenberg DP, Roddick GT, Jr M (2008) An enhanced post natal autoimmune profile in 24 week-old C57BL/6 mice developmentally exposed to TCDD. Toxicol Appl Pharmacol 232:51–59 21. Zhou Y, Lu Q (2008) DNA methylation in T cells from idiopathic lupus and drug-induced lupus patients. Autoimmun Rev 7:376–383 22. Barendrecht MM, Tervaert JW, van Breda Vriesman PJ, Damoiseaux JG (2002) Susceptibility to cyclosporin A-induced auto immunity: strain differences in relation to auto regulatory T cells. J Autoimmun 18:39–48 23. Brix TH, Hansen PS, Kyvik KO, Hegedus L (2000) Cigarette smoking and risk of clinically overt thyroid disease: a population-based twin case-control study. Arch Intern Med 160:661–666 24. Guilherme L, Fae KC, Oshiro SE, Tanaka AC, Pomerantzeff PM, Kalil J (2007) T cell response in rheumatic fever: crossreactivity between streptococcal M protein peptides and heart tis sue proteins. Curr Protein Pept Sci 8:39–44 25. Cunha-Neto E, Bilate AM, Hyland KV, Fonseen SG, Kalil J, Museum ED, Galleries A (2006) Induction of cardiac autoimmunity in Chagas heart disease: a case for molecular mimicry. Autoimmunity 39:41–54 26. Rose NR (2008) The adjuvant effect in infec tion and autoimmunity. Clin Rev Allergy Immunol 34:279–282 27. Croker BA, Lawson BR, Berger M, Rutschmann S, Berger M, Eidenschenk C,

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Blasius AL, Moresco EM, Sovath S, Cengia L, Shultz LD, Theofilopoulos AN, Pettersson S, Beutler BA (2008) Inflammation and autoim munity caused by a SHP1 mutation depend on IL-1, MyD88, and a microbial trigger. Proc Natl Acad Sci USA 105:15028–15033 28. Rose NR (2008) Autoimmunity in coxsackie virus infection. Curr Top Microbiol Immunol 323:293–314 29. Larizza D, Calcaterra V, Martinetti M, Negini R, De Silvestri A, Cisternino M, Iannone AM, Solcia E (2006) Helicobacter pylori infection and autoimmune thyroid disease in young patients: the disadvantage of carrying the human leukocyte antigen-DRB1*0301 allele. Clin J Endocrinol Metab 91:176–179 30. Vas J, Mattner J, Richardson S, Ndonye R, Gaughan JP, Howell A, Monestier M (2008) Regulatory roles for NKT cell ligands in envi ronmentally induced autoimmunity. J Immunol 181:6779–6788 31. Cooke A (2009) Infection and autoimmunity. Blood Cells Mol Dis 42:105–107 32. Mattner J, Savage PB, Peung P, Oertelt SS, Wang V, Trivedi O, Scanlon ST, Pendem K, Teyton L, Hart J, Ridgway WM, Wicker LS, Gershwin ME, Bendelac A (2008) Liver auto immunity triggered by microbial activation of natural killer T cells. Cell Host Microbe 3:304–315 33. Veldhoen M, Hirota K, Westendorf AM, Buer J, Dumoutier L, Renauld JC, Stockinger B (2008) The aryl hydrocarbon receptor links TH17-cell-mediated autoimmunity to envi ronmental toxins. Nature 453:106–109 34. Ober C, Loisel DA, Gilad Y (2008) Sexspecific genetic architecture of human disease. Nat Rev Genet 9:911–922 35. Fairweather D, Rose NR (2004) Women and autoimmune diseases. Emerg Infect Dis 10:2005–2011 36. Calin A, Brophy S, Blake D (1999) Impact of sex on inheritance of ankylosing spondylitis: a cohort study. Lancet 354:1687–1690 37. World Health Organization. Environmental Health Criteria 236 (2006) Principles and methods for assessing autoimmunity associ ated with exposure to chemicals. WHO Publications, Geneva, Switzerland 38. Gleicher N, Barad DH (2007) Gender as risk factor for autoimmune diseases. J Autoimmun 28:1–6 39. Zandman-Goddard G, Peeva E, Shoenfeld Y (2007) Gender and autoimmunity. Autoimmun Rev 6:366–372 40. Gause WC, Jackson JV, Dietert RR, Marsh JA (1985) Autoanti-thyroglobulin production in

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51. Sinha S, Kaler LT, Procter TM, Teuscher C, Vandenbark AA, Offner H (2008) IL-13mediated gender difference in susceptibility to autoimmune encephalomyelitis. J Immunol 180:2679–2685 52. Hastings KL (2006) Risk assessment in drug development: autoimmunity. J Toxicol Environ Health A 69:893–898 53. Goldstein NS, Bayati N, Silverman AL, Gordon SG (2000) Minocycline as a cause of drug-induced autoimmune hepatitis: report of four cases and comparison with autoimmune hepatitis. Am J Clin Pathol 114:591–598 54. Chamberlain MC, Schwartzenberg SJ, Akin EU, Kurth MH (2006) Minocycline-induced autoimmune hepatitis with subsequent cirrho sis. J Pediatr Gastroenterol Nutr 42:232–235 55. Schuurs AH, Dietrich H, Gruber J, Wick G (1992) Effects of sex steroid analogs on spontaneous autoimmune thyroiditis in obese strain chickens. Int Arch Allergy Immunol 97: 337–344 56. Lam-Tse WK, Lernmark A, Drexhage HA (2002) Animal models of endocrine/organspecific autoimmune diseases: do they really help us to understand human autoimmunity? Springer Semin Immunopathol 24:297–321 57. Morrel L (2004) Mouse models of human autoimmune diseases: essential tools that require the proper controls. PloS Biol 2:e241 58. Cooper GS, Gilbert KM, Greidlinger EL, James JA, Pfau JC, Reinlib L, Richardson BC, Rose NR (2008) Recent advances and oppor tunities in research on lupus: environmental influences and mechanism of disease. Environ Health Perspect 116:695–702 59. Gerosa M, De Angeslis V, Roboldi P, Meroni PL (2008) Rheumatoid arthritis: a female challenge. Womens Health (Lond) 4:195–201 60. Lleo A, Battezzati PM, Selmi C, Selmi C, Gershwin ME, Podda M (2008) Is autoimmunity a matter of sex? Autoimmun Rev 7:626–630 61. Rose NR (2008) Predictors of autoimmune dis ease: autoantibodies and beyond. Autoimmunity 41:419–428 62. Plot L, Amiltal H (2009) Infectious associa tions of Celiac disease. Autoimmun Rev 8(4):316–319 63. Ercolini AM, Miller SD (2009) The role of infections in autoimmune disease. Clin Exp Immunol 155:1–15 64. Tabuenca JM (1981) Toxic-allergic syndrome caused by ingestion of rapeseed oil denatured with aniline. Lancet 2:567–568 65. Noonan CW, Pfau JC, Larson TC, Spence MR (2006) Nested case-control study of autoimmune disease in an asbestos-exposed population. Environ Health Perspect 114:1243–1247

Risk of Autoimmune Disease: Challenges for Immunotoxicity Testing 66. Yurino H, Ishikawa S, Sato T, Akadegawa K, Ito T, Ueha S, Inadera H, Matsushima K (2004) Endocrine disruptors (environmental estrogens) enhance autoantibody production by B1 cells. Toxicol Sci 81:139–147 67. Costenbader KH, Kim DJ, Peerzada J, Lockman S, Nobles-Knight D, Petri M, Karlson EW (2004) Cigarette smoking and the risk of systemic lupus erythematosus: a meta-analysis. Arthritis Rheum 50:849–857 68. Burke L, Segall-Blank M, Lorenzo C, Dynesius-Trentham R, Trentham D, Mortola JF (2001) Altered immune response in adult women exposed to diethylstilbestrol in utero. Am J Gynecol 185:78–81 69. Fenaux JB, Gogal RM Jr, Lindsay D, Hardy C, Ward DL, Saunders G, Ahmed SA (2005) Altered splenocyte function in aged C57BL/6 mice prenatally exposed to diethylstilbestrol. J Immunotoxicol 2:221–229 70. Grimaldi CM (2006) Sex and systemic lupus erythematosus: the role of the sex hormones estrogen and prolactin on the regulation of autoreactive B cells. Curr Opin Rhematol 18:456–461 71. Poole BD, Templeton AK, Guthridge JM, Brown EJ, Harley JB, James JA (2009) Aberrant Epstein-Barr viral infection in sys temic lupus erythematosus. Autoimmun Rev 8:337–342 72. Lunardi C, Tinazzi E, Bason C, Dolcino M, Corrocher R, Puccetti A (2008) Human par vovirus B19 infection and autoimmunity. Autoimmun Rev 8:116–120 73. Vas J, Monestier M (2008) Immunology of mercury. Ann N Y Acad Sci 1143:240–267

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Chapter 5 Markers of Inflammation Dori R. Germolec, Rachel P. Frawley, and Ellen Evans Abstract Inflammation is a complex and necessary component of an organism’s response to biological, chemical or physical stimuli. In the acute phase, cells of the immune system migrate to the site of injury in a care fully orchestrated sequence of events that is mediated by cytokines and acute phase proteins. Depending upon the degree of injury, this acute phase may be sufficient to resolve the damage and initiate healing. Persistent inflammation as a result of prolonged exposure to stimulus or an inappropriate reaction to self molecules can lead to the chronic phase, in which tissue damage and fibrosis can occur. Chronic inflam mation is reported to contribute to numerous diseases including allergy, arthritis, asthma, atherosclerosis, autoimmune diseases, diabetes, and cancer, and to conditions of aging. Hematology and clinical chemis try data from standard toxicology studies can provide an initial indication of the presence and sometimes location of inflammation in the absence of specific data on the immune tissues. These data may suggest more specific immune function assays are necessary to determine the existence or mechanism(s) of immunomodulation. Although changes in hematology dynamics, acute phase proteins, complement factors and cytokines are common to virtually all inflammatory conditions and can be measured by a variety of techniques, individual biomarkers have yet to be strongly associated with specific pathologic events. The specific profile in a given inflammatory condition is dependent upon species, mechanisms, severity, chronicity, and capacity of the immune system to respond and adapt. Key words: Acute phase proteins, Basophil, Chemokine, Clinical pathology, Complement, Cytokine, Eosinophil, Hematology, Inflammation, Lymphocyte, Macrophage, Monocyte, Neutrophil, Platelet

1. Introduction Inflammation is a complex and necessary component of an organism’s response to biological, chemical or physical stimuli. In the acute phase, leukocytes, primarily granulocytes, migrate along a chemo tactic gradient to the site of injury in a carefully orchestrated effort that is mediated by cytokines and acute phase proteins (APPs) to remove the stimulus (e.g., infectious agent, foreign material) or cells damaged by injury and to initiate healing. Depending upon R.R. Dietert (ed.), Immunotoxicity Testing: Methods and Protocols, Methods in Molecular Biology, vol. 598 DOI 10.1007/978-1-60761-401-2_5, © Humana Press, a part of Springer Science + Business Media, LLC 2010

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the degree of injury, this acute cellular phase may be sufficient to resolve any damage. Persistent inflammation as a result of prolonged exposure to stimulus or an inappropriate reaction to self molecules can lead to the chronic phase in which the active immune cell pop ulations shift to include a mononuclear phenotype, and tissue dam age and fibrosis can occur. Chronic inflammation is reported to contribute to numerous diseases including allergy, arthritis, asthma, atherosclerosis, autoimmune diseases, diabetes, and cancer, and to conditions of aging. The inflammatory process involves multiple physiological systems with the immune system playing a central role (1–3). Detailed information on the specific cells, cell surface molecules and soluble mediators of the inflammatory response is beyond the scope of this overview, and the reader is referred to chapters which cover specific aspects of the immune response or topic-specific reviews cited below for additional details.

2. General Considerations from Standard Toxicology Studies

Hematology data (including erythrocyte parameters, platelet count, total number of leukocytes, and leukocyte differentials and morphology), coagulation (clotting times, fibrinogen) and clini cal chemistry data (total protein, albumin and globulin, liver enzymes, renal parameters, electrolytes, bilirubin) are included in standard toxicology studies. These clinical pathology data can provide an initial indication of the presence and sometimes loca tion of inflammation in the absence of specific data on immune tissues. When possible, pretest samples should be collected for nonrodent studies so that the experimental data can be inter preted in comparison to a baseline; for all species, data should be compared with age-matched concurrent and historical controls. Hematology and serum chemistry may provide information on both innate and acquired immunity, and in addition to basic infor mation on immune cells, these endpoints provide baseline informa tion on other organ systems that may affect or be affected by the immune system. For example, changes in erythrocyte parameters or leukocyte counts may indicate altered bone marrow function and the potential for decreased production of immune cells or precur sors, and decreases in globulins may signal decreased antibody syn thesis, particularly if the albumin/globulin ratio is increased. Increased fibrinogen may suggest an inflammatory process, even in the absence of an inflammatory leukogram. It is important that these data be considered along with other available information such as clinical observations and histological changes when avail able, and to attempt to distinguish those changes that represent direct effects of a chemical agent on the immune system (such as a shift in leukocyte populations as a result of destruction of bone

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marrow progenitors or lymphocytes) from those that may be a secondary consequence of immune system perturbation (such as a shift in leukocyte populations as a result of infection). The clinical pathology data may suggest more specific immune function assays that are necessary to determine the existence or mechanism(s) of immunomodulation. However, these data alone are not always reliable predictors of immunotoxicity; for example, circulating leukocyte numbers may be within normal values even when there are extreme changes in immune function, such as those observed in chronic, well established infection and in some children with primary immunodeficiencies. Conversely, effects on leukocyte trafficking unrelated to the immune system may affect circulating numbers of individual white blood cell types. 2.1. Cells of the Inflammatory Response

In the acute phase of inflammation, platelets and granulocytic cells such as basophils/mast cells, neutrophils and eosinophils are activated and in turn produce and release a number of soluble mediators that stimulate and regulate the inflammatory response.

2.1.1. Neutrophils

Neutrophils, which are sometimes referred to as polymorphonu clear neutrophils (PMNs), are the primary cellular mediators of the acute inflammatory response. Their granules contain a variety of enzymes, peptides, and proteins and also undergo a respiratory burst. The intent of their armamentarium is to destroy and digest organisms and foreign material following phagocytosis, but gran ule contents may also be released and damage tissues at the inflam matory site. Measurement of some neutrophil products, notably myeloperoxidase, may be used to assess severity of inflammation (4). Vasodilation and increased vascular permeability following basophil/mast cell degranulation, complement activation, or release of prostaglandins and leukotrienes allows neutrophils to migrate from the blood to the site of injury, and this mobilization usually results in an increase of circulating neutrophils (Fig. 5.1). However, there are numerous causes of increased numbers of circulating neutrophils (neutrophilia), and some of these may not directly relate to immune status, which underscores the need to integrate all of the data from a toxicology study rather than assessing individual components separately. Two examples of neutrophil trafficking effects that are not directly immune systemrelated include excitement and stress: excitement with epineph rine release results in demargination and an increased mobilization of neutrophils from bone marrow storage pools; stress and its resultant corticosteroid release results in increased release from bone marrow and decreased migration to tissues. In both cases, an increase in circulating mature neutrophils is seen. In contrast, neutrophilia as a consequence of inflammation is typically characterized by a shift toward immature cell types

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Fig. 5.1 Mediators in the process of inflammation. This figure summarizes the roles of the various mediators important in the process of inflammation from the acute to chronic phase. PGE prostaglandin E, VIP vasoactive intestinal polypeptide, LTB leukotriene B, LTD leukotriene D, PAF platelet activating factor, IFN interferon, IL interleukin.

(called a “left shift”) with increased numbers of bands or earlier neutrophil stages (myelocytes, promyelocytes, ring forms in rodents) as the bone marrow depletes its reserve of mature neu trophils to meet the demand. It should be noted that immature forms are less likely to be seen in chronic, established infections. Specific morphologic changes such as Döhle bodies, basophilia, toxic granulation or vacuolation (known collectively as “toxic change”) may be seen in any situation of accelerated myelopoiesis in the bone marrow. The term “toxic change” is somewhat of a misnomer in that “toxicity” (either from a drug, chemical, or bac terial toxin) is not necessary to bring about these morphologic changes. 2.1.2. Basophils

Basophils and mast cells contain cytoplasmic granules that serve as reservoirs for soluble mediators that function in many aspects of the inflammatory response. Early phase reactants released from mast cells such as histamine and serotonin, and prostaglandin and leukotriene products of arachidonic acid metabolism mediate the vasodilation and increased vascular permeability characteristic of the acute vascular response. The secretion of platelet activating factor (PAF) by mast cells also increases vascular permeability and stimulates the release of inflammatory mediators from platelets and the activation of neutrophils. Enzymes such as B-glucoronidase, amylosidase and chymase released from mast cells play significant roles in tissue damage and repair. While basophil counts are routinely included in leukocyte differentials, their low numbers (<1% of the total leukocytes in

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health) make them the most difficult of the leukocytes to enumerate accurately by manual differential. Unfortunately, the determina tion of basophil counts by automated differential instruments is challenging as well. The size and staining characteristics of baso phils make them difficult for instruments to categorize them, and basophils may overlap with other cell types. It has been dem onstrated that automated instruments tend to underestimate the counts during true basophilia (5). Furthermore, increases in baso phils are rarely significant enough to appreciably affect the total leukocyte count. Manually scanning peripheral blood smears may be useful to identify cases in which the instrument underestimates basophil numbers. It has been suggested that circulating numbers of basophils may be reduced in conditions of chronic inflamma tion due to active recruitment of the cells to the site of injury (6). However, since reference intervals for basophils generally start at 0 cells/mL, basopenia is usually not a practical entity for diagnos tic purposes. 2.1.3. Eosinophils

Eosinophils predominate in inflammatory sites associated with hypersensitivity responses and clearance of parasitic infections. Eosinophils are recruited to the site of inflammation by a number of factors including interleukin (IL)-5, IL-2, IL-16, histamine, neutrophil peptides and some complement proteins. Activation of eosinophils results in the release of eosinophil peroxidase, major basic protein (MBP), eosinophil-derived neurotoxin and eosinophil cationic proteins (ECP). In respiratory hypersensitivity responses, these mediators induce rapid vasoconstriction followed by increased vascular permeability and pulmonary edema. MBP and ECP are highly basic proteins that degrade nearby cells such as the tracheal epithelium. Automated hematology analyzers may vary in ability to accurately identify eosinophils across species due to variability in granulation. However, changes in eosinophil numbers are easy to identify in manual leukocyte differentials, as their cytoplasmic granules have a high affinity for acidic dyes that result in a characteristic pink/red staining. In certain allergic con ditions such as asthma and rhinitis, the numbers of circulating eosinophils may be correlated to disease severity (7). However, eosinophilic inflammatory responses may not always result in increased numbers in peripheral blood (8). MBP and ECP are becoming important clinical markers for eosinophil activation and allergic disease, and can be evaluated in serum, sputum and pulmonary lavage fluid.

2.1.4. Platelets

Platelets are anucleate circulating cell fragments consisting of membrane-bound cytoplasm derived from megakaryocytes, which in turn are derived from the common myeloid progenitor cells in the bone marrow. They are rapidly deployed to sites of vascular compromise, injury and infection and are locally activated.

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Although they are highly differentiated for hemostasis, platelets also have inflammatory and antimicrobial functions, linking clotting and immune events (9). Platelets release reactive oxygen species, which may contrib ute to tissue damage, and mediators such as heparin and sero tonin, which promote the acute vascular response. However, the primary function of platelets is to facilitate clot formation and prevent leakage from damaged blood vessels. Platelets work in concert with APPs of the coagulation system such as fibrinogen and vitronectin at sites of vascular damage. While peripheral blood platelet counts and morphology may be affected by or part of inflammatory processes, assessment of platelet parameters alone is not helpful in identifying inflammation. A number of therapeutic agents (e.g., procainamide, sulfame thoxaxole, gold salts) can induce immune-mediated thrombo cytopenias and/or aplastic anemias that may result in reduced platelet counts. Immune-mediated thrombocytopenias are gener ally regenerative in nature. That is, mean platelet volume (MPV) is increased and large platelets are noted on blood smears, indicating accelerated release of platelets by megakaryocytes, and megakaryocytes in bone marrow are increased in number. However, if the target of toxicity or an immune-mediated process is an earlier progenitor cell, MPV’s may be normal and mega karyocytes reduced in number. Thrombocytosis (an increased number of platelets in periph eral blood) may be manifested during hemorrhage, acute inflam mation or infections, or develop as a secondary result of some chronic inflammatory conditions such as rheumatoid arthritis, or during liver regeneration following hepatotoxicity or hepatec tomy (10). It has also been suggested that the increased cardio vascular mortality associated with high levels of air pollution may be linked to toxicant-induced thrombocytosis that initiates embo lism formation (11). 2.1.5. Lymphocytes

Lymphocytic infiltration is often a prominent feature in chronic inflammation. Lymphocytes may act as specific effectors of cyto toxicity, or they may secrete antibodies or cytokines that partici pate in tissue damage or inflammatory cell recruitment. Lymphocytes can also participate in nonspecific inflammatory reactions, recruited as a consequence of elevated cytokine and adhesion molecule levels. However, lymphocyte responses, both systemic and local, may have no impact on the numbers of circu lating total lymphocytes or subsets. Conversely, changes in the numbers of circulating lymphocytes are not necessarily reflective of immune stimulation or inflammation and may be the result of physiologic responses mediated by epinephrine (lymphocytosis likely due to diminished homing to peripheral lymphoid tissue), changes in cytokine secretion, or modulation of lymphocyte homing

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and trafficking, such as occurs in stress responses mediated by corticosteroids (lymphopenia due to increased homing to lymphoid tissues or lysis). Peripheral blood lymphocytosis is seen with lymphocytic neoplasia and occasionally occurs with chronic infections (e.g., Rickettsial diseases, fungal infections, etc.) or other chronic antigenic stimulation. For most species, peripheral blood lymphocytosis is not a typical component of the inflammatory leukogram; lymphocytes tend to be more concentrated in lymph nodes or sites of inflammation. In fact, stress-induced lym phopenia is a more typical component of inflammation. However, lymphocytosis is not uncommon in inflammatory responses of rats. In inflammatory conditions or chronic antigenic stimu lation, morphologically reactive lymphocytes may be seen in peripheral blood. 2.1.6. Macrophages

Macrophages tend to accumulate at the site of injury following lymphocytic infiltration and may also be the primary response to chronic diseases such as mycobacterial infections. They serve as antigen presenting cells (APCs) and help drive and perpetuate the immune response, releasing a variety of inflammatory cytok ines and cytokines that stimulate lymphocytes. However, they play a predominant role in the regulation and resolution of the inflammatory response. The lysozomal enzymes contained within the macrophage break down inflammatory mediators and participate in the phago cytic degradation of foreign materials. In addition to resident macrophages in such tissues as lung and liver, which are capable of reproducing, macrophages may be recruited to inflammatory sites from the bone marrow as monocytes, which then continue to differentiate into macrophages (Fig. 5.1). Monocytes and macrophages play a major role in the removal of dead or abnormal cells, and thus, increased numbers of these cells may be indicative of inflammation or tissue necrosis. Macrophages are only rarely seen in peripheral blood, but mono cytes on their way to inflammatory sites may appear activated. Increases of monocytes in peripheral blood can occur with both acute and chronic inflammation. Monocytosis may also be seen as part of a corticosteroid-mediated stress response, but this is a less consistent finding than lymphopenia.

2.1.7. Red Blood Cells

While not considered mediators or primary participants in inflam mation, red blood cells can be affected by and reflective of inflam matory processes. Red blood cells are very susceptible to immune-mediated and other processes that accelerate red blood cell destruction. The immune-mediated process and potentially the red blood cell destruction itself may trigger an inflamma tory response. Therefore, inflammatory leukograms are very

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c ommon in immune-mediated anemias and are sometimes seen in other hemolytic anemias. Hemolytic anemias are typically very regenerative in nature, as indicated by increased reticulocytes, polychromasia and anisocytosis. Inflammation can also cause red blood cell loss if significant hemorrhage occurs, either as part of the inflammatory process or if disseminated intravascular coagulation (DIC) is a sequela of the inflammation. This anemia may or may not appear regenerative, depending upon whether or not the bone marrow has had time (usually 2–5 days) to respond. The most common red blood cell finding with inflammation is a nonregenerative, mild-to-moderate decrease in red blood cells termed “anemia of chronic disease (ACD)” or sometimes “ane mia of inflammatory disease.” Multiple inflammatory cytokines are involved in the pathogenesis of ACD (12). The major purpose of ACD seems to be limiting iron availability, which is beneficial in inflammation by reducing the potential for oxidation and free radical formation and limiting iron-dependent bacterial growth. The cytokines thought to play a role in ACD include IL-1b and Tumor Necrosis Factor (TNF)-a, which cause reduction of eryth ropoietin release by the kidneys. TNF-a is also thought to enhance erythrophagocytosis by macrophages and, along with IL-1b and interferon (IFN)-g, it directly decreases erythroid progenitor pro liferation. In addition, IL-6 increases hepcidin production by the liver. Hepcidin in turn inhibits ferroportin-mediated release of iron stores from macrophages and absorption of iron into circula tion from the intestine, resulting in diminished erythropoiesis due to the lack of iron availability (13, 14). Because ACD can mimic iron deficiency anemias, serum ferritin may be measured. With iron deficiency, serum ferritin (a protein involved in the storage of iron and considered an APP) concentration is typically decreased. In ACD, serum ferritin concentration is normal or increased. 2.1.8. Summary of Inflammatory Cell Evaluation

In summary, the classic peripheral blood hematology profile that suggests inflammation is dominated by neutrophilia or neutrope nia with a left shift, and monocytosis is frequently present (15). Decreases in lymphocytes are a common finding in inflammation in nonrodent species primarily due to secondary stress, but lym phocytosis is frequently seen in rodents. In addition, a nonregen erative, mild-to-moderate decrease in red blood cell parameters may be seen. Serum chemistry information such as increased globulins due to increased immunoglobulin and/or APPs (dis cussed in greater detail in Section 2.3.4), or coagulation informa tion such as fibrinogen may provide additional evidence of the presence of inflammation. The hematology profile represents a snapshot of a very dynamic process, therefore, the classic profile is not always seen. The specific profile in a given situation is dependent

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upon the species, mechanisms, severity, chronicity, and capacity of the immune system to respond and adapt. 2.2. Cell Surface Receptors and Adhesion Molecules

Adhesion molecules control the interactions between leukocytes and endothelial cells during the inflammatory response. These cell-to-cell interactions are the result of a cascade of events stimu lated by the upregulation of cell surface ligands early in inflamma tion. Once inflammatory cells have arrived at the site of injury, further interactions with the endothelium, other parenchymal cells, immune cells and extracellular matrix proteins perpetuate the response leading to resolution or chronic inflammation. Several families of adhesion molecules participate in this cascade of events including the integrins, selectins, vascular addressins and lymphocyte homing receptors (16, 17).

2.2.1. Selectins

Selectins are cell surface molecules that are expressed on leuko cytes (L-selectin), endothelial cells (P-selectin, E-selectin) and platelets (P-selectin). L-selectin initiates the interaction between leukocytes and endothelial cells via binding to mucin-like mole cules called addressins on the vascular endothelium and functions as a homing receptor for lymphocytes. P-selectin is localized to membrane-bound granules in endothelial cells and alpha granules in platelets and is rapidly expressed following mast cell degranula tion. P-selectin facilitates the interactions between PMNs, plate lets and endothelial cells and is important in both clot formation and degradation. E-selectin expression promotes the adhesion of PMNs and is upregulated in the acute phase of the inflammatory response by cytokines and bacterial cell wall products such as lipopolysaccharide (LPS). Expression of selectins on cells changes as the acute phase resolves or progresses into chronic inflammation and can be quan titated using antibody-based labeling techniques for immunohis tochemistry, flow cytometry or microscopy. Soluble forms of the selectins are present in serum and can be measured via ELISA or other immunoassays.

2.2.2. Integrins

The integrins are a widely expressed family of molecules that mediate cell–cell interaction and interactions between cells and the extracellular matrix. Integrins contain an extracellular domain that engages other cell adhesion molecules such as intracellular adhesion molecule-1 (ICAM-1) or adhesive ligands such as fibrin ogen and LPS, and a cytoplasmic domain that interacts with intracellular proteins. The leukocyte integrin lymphocyte func tion-associated antigen-1 (LFA-1) binds to ICAM-1 on endothe lial cells and directs lymphocytes to target tissues during inflammation and promotes cell–cell interactions during Natural Killer cell (NK)- or Cytotoxic T Lymphocyte (CTL)-mediated cytotoxicity. Similarly, Mac-1 binding is important in the adherence

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of monocytes to endothelial cells and also serves as a complement receptor to enhance the phagocytosis of opsonized cells or bacteria. Very late antigens of activation (VLA) are integrins that function during inflammation to bind immune cells to extracel lular matrix proteins such as fibronectin, collagen and laminin and to attract nonimmune cells such as fibroblasts and endothelial cells. Inflammatory cytokines upregulate the expression of cell adhesion molecules that serve as ligands for the integrins ICAM1, ICAM-2 and vascular cell adhesion molecule-1 (VCAM-1) on endothelial cells in a coordinated fashion, so inflammatory cells can be selectively recruited during the various phases of the inflammatory response. As discussed for the selectins above, expression of integrins and their ligands can be quantitated using antibody-based techniques for immunohistochemistry, flow cytometry or microscopy. In addi tion, there are a number of antibody- and ligand-based functional assays that have been used as research tools to assess integrin medi ated cell signaling and the efficiency of cell adhesion (18). 2.3. Soluble Mediators of the Inflammatory Response 2.3.1. Cytokines

Cytokines exhibit a number of general biological effects that are important to understand when considering their utility as markers of inflammation. A single cytokine may affect a number of different cell types or targets and may have both autocrine and paracrine effects. Cytokines also exhibit multiple effects on different cell types, often with synergistic or opposing results. This frequently means that cytokines act as molecular messengers to coordinate the interplay between, and control of, different component cell types in the immune response – an action that is of great importance in the amplification of immune and inflammatory responses. During innate immune responses cytokines are primarily secreted by phagocytic cells and NK cells, but in adaptive immune responses they are mainly produced by APCs and lymphocytes. Although often described separately, crosstalk between the innate and adaptive immune systems is frequent, and cytokines represent a major means of communication between the various arms of the immune system. The balance between production of an effec tive immune response and tissue damage depends on careful regulation of the cytokine network. The short half-life of cytokines suggests that under normal conditions most of these soluble mediators are rapidly eliminated thereby ensuring their limited bioactivity. However, during acute and/or chronic inflammatory conditions, cytokines may be released in sufficient quantity that systemic effects are observed, and toxicants that act on either the mediators themselves or the cells that produce these mediators may alter the mechanisms that regulate cytokine production leading to inflammation and disease.

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This section provides general information on the cytokines that contribute to inflammatory processes. For a more detailed discus sion of the production and function of the various cytokines the reader should consult Chapter 20. Many cytokines and chemokines (see below) contribute to inflammation; some facilitate leukocyte chemotaxis to the site of injury, while others modulate immune cell function (Table 5.1). The cytokines that are best known for stimulating and perpetuat ing inflammatory responses are IL-6, IL-1, IL-2, TNF-a, IFN-g, and transforming growth factor (TGF)-b. 1. IL-6 was originally identified as a B-cell differentiation factor, and increased levels of this cytokine have been associated with polyclonal B cell activation and chronic inflammation. In the initial phases of acute inflammation, IL-6 mediates the acute phase response. IL-6 levels remain high in chronic inflammatory processes leading to enhanced survival and growth of lymphocytes and macrophages that perpetuate inflammation. 2. IL-1 has a number of direct and indirect activities that pro mote inflammation including the stimulation of the produc tion of other cytokines and the release of prostaglandins. These promote the generation of cytotoxic effector cells and synergize with colony stimulating factors to increase the pro duction of inflammatory cells in the bone marrow. 3. IL-2 augments NK cell activity, stimulates the production of inflammatory cytokines such as IL-1 and IFN-g and enhances macrophage cytotoxicity. It also contributes to chronic inflam mation by stimulating the proliferation of antigen specific T- and B-lymphocytes. 4. TNF-a enhances inflammation and is important in the pro cess of removing dead and dying cells through apoptosis. TNF-a has been shown to upregulate the expression of Class I and II major histocompatibility complex (MHC) molecules on certain cell types resulting in cell activation and cytokine release. 5. IFN-g is a potent activator of macrophages. It stimulates the production of IL-1 and TNF-a and enhances the expression of Class II MHC molecules on immune cells and vascular endothelium. The latter is of particular importance in allow ing inflammatory cells to move through the vasculature into tissues or a site of injury. 6. TGF-b is important in the regulation of tissue repair and regeneration following injury. It is produced by a number of immune and nonimmune cell types and is important in the regulation of the inflammatory response by inhibiting the production of proinflammatory cytokines such as IL-2, IFN-g

Table 5.1 Cytokines and chemokines important in inflammation Mediator

Source

Function

Cytokines Interleukin-1 (IL-1)

Macrophages, dendritic cells, Activates T and B lymphocytes T and B cells Increases production of other cytokines and acute phase proteins Induces adhesion molecules

Interleukin-2 (IL-2)

Activated T cells, B cells

Growth and activation of T and B cells Growth and activation of NK cells and macrophages Induces production of proinflammatory cytokines

Interleukin-6 (IL-6)

Macrophages, dendritic cells, Multiple effects on T cells B cells, activated T cells Myeloid cell development Regulation of acute phase proteins

Interferon gamma (IFN-g)

T cells, NK cells, epithelial cells, fibroblasts

Increases MHC expression Enhances CTL, NK and macrophage activity Stimulates production of IL-1 and TNF-a

Transforming growth factor Macrophages, megakaryo beta (TGF-b) cytes, chondrocytes

Inhibits cytokine production and activity Inhibits B cell proliferation Stimulates wound healing

Tumor necrosis factor alpha Macrophages, dendritic cells, Increases MHC expression (TNF-a) lymphocytes, mast cells Activates macrophages Enhances tumor cell killing Chemokines Macrophage Endothelial cells, epithelial chemoattractant protein-1 cells, fibroblasts, (MCP-1) (CC family) monocytes

Attracts monocytes Activates macrophages and T cells Stimulates histamine release

RANTES (CC family)

T cells, endothelial cells, platelets

Attracts macrophages

Gro (a, b, g MSGA) (CXC family)

Macrophages, fibroblasts

Attracts PMNs, angiogenesis

Interleukin-8 (IL-8) (CXC family)

Macrophages, lymphocytes

Mobilizes PMNs from bone marrow

Fractalkine (CX3C family)

Endothelial cells, microglia macrophages

Attracts T cells, monocytes and PMNs in the brain

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and TNF-a. TGF-b is released by platelets following tissue injury. It plays an essential role in wound healing by attracting inflammatory cells, inducing angiogenesis and enhancing the deposition of proteins that make up the extracellular matrix. 2.3.2. Nuclear Factor-kB

2.3.3. Chemokines

The relative importance of any individual soluble mediator in a specific inflammatory condition varies. Because cytokines operate in cascades and networks, target cells may be influenced by mul tiple cytokines. While these molecules regulate production of APPs and other inflammatory mediators, they in turn may be modulated by transcription factors such as Nuclear Factor-kB (NF-kB) (19, 20). NF-kB, which has been called a “master tran scriptional switch in inflammation,” regulates dozens of targets involved in inflammation and other biochemical processes, and changes in NF-kB expression have also been used as a biomarker of inflammation (21). Inflammatory chemokines control the recruitment of effector leukocytes in infection, inflammation, tissue injury and tumors. As part of the inflammatory process, chemokines direct cellular migration, activate macrophages and PMNs, and modulate wound healing through their effects on angiogenesis and the generation of fibrosis (16). Cell migration often occurs along chemokine gradients and changes in chemokine receptor levels on leukocytes and endothelial cells regulate localization of leukocytes to sites of inflammation (22). Chemokines are defined based on their amino acid composi tion, specifically on the presence of a conserved tetra-cysteine motif, and not by their function as chemotactic cytokines. The relative position of the first two consensus cysteines (either sepa rated by a nonconserved amino acid or next to each other) provides the basis for division of chemokines into the two major subclasses, CXC and CC chemokines, respectively (23). (a) In general, CC chemokines serve as chemoattractants for monocytes (RANTES, monocyte chemoattractant proteins (MCP) 1–5), eosinophils (eotaxins 1–3), basophils (MCP 4–5) and lymphocytes (Macrophage inflammatory protein (MIP)-1a and b). (b) Members of the CXC family, which includes IL-8, Platelet factor 4 and Gro a and b, attract PMNs and modulate angio genesis and wound healing. Several homologous molecules are also regarded as chemok ines and fall into two additional subclasses. These are typified by CX3C (Fractalkine or neurotaxin), which has three intervening amino acids between the first cysteines, and XCL1 and XCL2 from the C family of chemokines, which have only a single terminal cysteine.

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Chemokines also induce astrocytic migration and microglial cell proliferation, which promote the transmission of nociceptive pain signals that serve as a warning sign in response to acute tissue damage (24). Chemokines share receptors with other chemo tactic molecules such as PAF, C5a (see below) and Leukotriene B4. Activated effector or memory T cells are the source of mul tiple inflammatory chemokines and, by sustaining effector T cell and macrophage recruitment, inflammatory chemokines control the local production of proinflammatory cytokines (25). Thus, locally produced chemokines influence disease progression and pathogenesis. Increased levels of chemokines have been associated with a number of chronic inflammatory diseases including atherosclero sis and glomerulonephritis. Numerous tools are available for eval uating the role and function of chemokines and their receptors in inflammation and disease. As indicated above, for many localized mediators a frequently used method to define the composition of inflammatory infiltrates is immunohistochemical staining of relevant tissue sections. There are a number of monoclonal antibodies available for the characterization and measurement of chemokines and their receptors. In addition, chemokine function can be measured in cell-based assays that assess the migration of cells across filters or membranes in response to chemoattractants (26). 2.3.4. Acute Phase Proteins

The APPs, notably C-reactive protein (CRP), complement fac tors, and the coagulation factor fibrinogen, are considered classi cal biomarkers of inflammation. Other APPs involved in inflammation include amyloid A, ceruloplasmin, haptoglobin, and alpha 2-macroglobulin. These proteins are regulated in response to inflammatory signals and change rapidly as the condi tion changes, making them good markers of inflammation. APPs are produced primarily by hepatocytes in response to cytokine signals from the site of inflammation. They alter homeostasis and initiate or support defensive or adaptive processes that contribute to healing in the short term, but can lead to chronic inflammation, metabolic disturbances and tissue damage with prolonged stimula tion. Traditionally, these proteins are measured directly in plasma or serum and are used clinically to evaluate the presence, intensity and course of an inflammatory process or disease. For individual APPs, basal levels and the degree of participation in the acute phase response varies across species, so the most relevant marker for one species may not have relevance in another (27, 28). In addition, some available assays are species-specific and reagents may not be available for a given APP in the species of interest. To increase sensitivity, it is best to determine the most appropriate APP for the species in question, or to assess a panel of APPs.

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(a) C-Reactive Protein The most sensitive and commonly recognized diagnostic tool for the acute phase response in humans is CRP. However, CRP is of variable importance in species commonly used for toxi cology studies (27). CRP levels can increase as much as 1,000-fold in response to cell damage (16). Hepatic CRP synthesis is regu lated by proinflammatory cytokines including IL-6, IL-1 and TNF-a so that almost any form of tissue injury, infection, inflam mation or stress can be associated with increased circulating CRP values (29). CRP shares several functions with Ig molecules in that it can opsonize microorganisms such as bacteria and fungi through binding to cell wall polysaccharides as well as facilitate phagocyto sis through its ability to activate and fix complement. Conventional CRP immunoassays have been shown to be more reliable indica tors of acute inflammation than leukocyte counts as blood levels rise and fall rapidly in a reflection of the disease process (30). Although somewhat controversial, CRP has been shown to be a good predictor of cardiovascular disease risk, hypertension and diabetes (31–34). (b) The complement system is a complex network of proteins that participate in the acute inflammatory response through their enzymatic activity, effects on mediator release, chemotaxis and vascular permeability, and the ability to enhance phagocytosis through opsonization of microbes. A highly simplified sum mary of the complement system is presented in Fig. 5.2, and the reader is referred to basic texts such as Janeway’s Immunobiology (35) for a more comprehensive discussion. (i) The classical arm of the complement system is activated by antigen–antibody interactions, which trigger a cascade of reactions, each of which results in the activation of another complement component. There are two antibodyindependent pathways that can trigger complement acti vation. Several lectins, including mannose-binding lectin and ficolin, are structurally similar to the C1 proteins that initiate the classical complement cascade (36). These lec tins can activate complement by cleaving the C2 and C4 proteins in a manner similar to that of C1. This leads to the cleavage of C3, which is a common step in all three path ways of the complement network. (ii) Microbial cell wall proteins such as LPS and zymosan (from yeast) activate the alternate complement pathway by interacting with three triggering factors (initiating fac tor, factor B and factor D) that combine to cleave C3. C3 is the most abundant of the complement proteins and plays a central role in the complement pathway (37). Cleavage of C3 releases a small peptide, C3a, which, along

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Fig. 5.2. Overview of complement activation. This figure summarizes the three major pathways of complement activation and formation of the membrane attack complex. MBL mannose binding lectin, MASP MBL-associated serine protease.

with C5a, increases vascular permeability allowing the influx of inflammatory cells and proteins. The larger cleav age product, C3b, binds to the surface of pathogens as well as to complement receptors on inflammatory cells and enhances cell adhesion and phagocytosis. C3b also binds to the activated products from earlier steps in the classical/ lectin (C4b2a) or alternate (Bb) pathways to form the C3 convertase. The C3 convertase cleaves the C5 protein into C5a, the most potent of the peptide mediators of local inflammation, and C5b, which triggers the terminal events in the complement cascade resulting in the formation of the cytolytic membrane attack complex. The formation of immune complexes in autoimmune diseases activates com plement resulting in chronic inflammation and tissue dam age. Deposition of complement is frequently used as an immunohistochemical marker of inflammation, and the levels of individual complement factors can be quantitated in serum. The activity of serum complement can be deter mined in the laboratory by evaluating its ability to lyse cells in vitro. Assessment of the activity of individual comple ment components is not routinely used as a marker of inflammation but is more commonly used as a diagnostic tool for complement deficiencies.

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(c) Fibrinogen Fibrinogen is an APP that has significant cross-species rele vance and is easily incorporated into standard preclinical toxicity studies. It must be measured in plasma, not serum, as it is con sumed in the formation of the blood clot. Fibrinogen provides an interface between inflammation and coagulation (38). The coag ulation process is similar to that of the complement system in that it operates as a series of transformations of proenzymes to acti vated enzymes. This process culminates in the formation of thrombin, which converts soluble fibrinogen into insoluble fibrin. These enzymatic reactions normally occur on the surface of acti vated endothelial cells and platelets. Cross talk between proteins in the complement and clotting cascades occurs at a number of points, either through interactions with target cells or through processes involving inflammatory mediators other than comple ment effectors (39). Receptors for fibrinogen have been identified on monocytes/macrophages, neutrophils, platelets and NK cells, and the binding of fibrinogen activates signaling pathways that upregulate NF-kB expression and the release of proinflammatory cytokines (40). Peptides formed during the cleavage of fibrinogen increase vasodilation and serve as chemoattractants, allowing fibrinogen to act as a bridging molecule between leukocytes and endothelial cells facilitating the recruitment of these cells to the site of inflammation. Along with CRP, serum levels of fibrinogen have been used as a predictor of cardiovascular disease risk, hypertension and diabetes in humans (34, 41). It should be noted that fibrinogen is utilized at an accelerated rate in situations of DIC, which may occur during severe inflammatory processes. In the face of DIC, plasma fibrinogen decreases. This may mask the increase seen in inflammation and vice versa. The measurement of fibrin degradation products (FDP) or D-dimer, breakdown products of fibrin, along with the assessment of other parameters suggesting inflammation (leukogram, other APPs) or DIC (throm bocytopenia, prolonged clotting times) are useful in interpreting fibrinogen data. (d) “Omics” Recently “omics” technologies have been used to detect APP associated with inflammation in human and animal studies. Consistent with conventional measures of APP, serum and plasma are commonly used matrices for detection of circulating proteins by two-dimensional electrophoresis (2DE) and mass spectrome try (MS). These techniques can provide a more complete picture of the inflammatory process, since they can separate and measure active proteins, precursors, metabolites and degradation products (42, 43). Upregulation of a number of APP, including CRP (42, 44), a-2-macroglobulin (45), complement C3 (45), serum amy loid A (46), and plasminogen (42) have been identified through

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2DE and/or MS in association with inflammatory diseases. Ironically, several of the most abundant APP, including fibrino gen, haptoglobin and a-1-antitrypsin, are often removed prior to proteomic analysis because their strong signals mask that of other proteins (42, 47, 48).

3. Conclusions Although changes in hematology dynamics, APP, complement factors and cytokines are common to virtually all inflammatory conditions, reliable, individual biomarkers associated with specific pathologic events are not yet available. High-content technolo gies, such as gene and protein microarrays, are necessary for gene/protein screening and model development. Using DNA microarrays, Ezendam et al. (49) conducted a multi-organ, multipathway study into the toxic effects of hexachlorobenzene (HCB) in rats. They identified increased transcript levels of markers for granulocytes and macrophages that were consistent with histo pathological findings, as well as upregulation of genes encoding proinflammatory cytokines and chemokines, APPs, and comple ment components, suggesting that HCB induces a strong, sys temic inflammatory response. In addition to the immune response genes, this study also identified altered functions and pathways associated with metabolism, oxidative stress, and reproductive toxicity, and it enabled the relative comparison of effects in spleen, liver, kidney, blood, mesenteric lymph nodes and thymus. Since most APP and complement factors are produced in the liver, rather than directly in the affected tissue, they do not help to identify the specific target tissue(s) or provide an accurate reflec tion of the extent of damage. If damage to target tissue is easily assessed by clinical pathology data (e.g., leakage enzymes in the case of hepatitis), the damaged tissue may be identified. However, blood chemistry is generally unrewarding in identifying sites of inflammation. The protein and gene profiles of inflammation vary with each disease or condition, and several studies indicate that multiple molecules representing different biochemical pathways and pathophysiologic processes offer the best strategy for identifying gene signatures representative of inflammatory changes (50–52). Using protein arrays and a support vector machine algorithm, Tabibiazar et al. (50) identified an inflammatory signature expression pattern, consisting primarily of cytokines and chemok ines, that was successfully used to predict the stage of atheroscle rosis in an independent data set. The investigators evaluated proteins found in serum and validated potential markers using

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uantitative real-time reverse transcriptase polymerase chain reac q tion (RT-PCR) of RNA from aortic vascular wall to discover a profile that is specific to atherosclerosis. The primary members of the biomarker panel were Ccl9, Ccl2, Ccl21, Ccl19, Cxcl1, IL5, Tnfsf11, and Vegfa. More common cytokines, such as IL-6, pro duced in other tissues and promoting general inflammation, were not sufficiently specific in this model (50). Proteomic and genomic technologies allow simultaneous analysis of the complement factors, cytokines and chemokines, apolipoproteins, and other APP, as well as tissue-specific pro teins and proteins associated with other physiological functions to provide a more integrated picture of inflammation, mecha nisms of action and the disease process (48). Transcriptome profiles offer the possibility of unifying traditional diagnostic markers of inflammation with regulatory and immune-related molecules to provide temporal, mechanistic and predictive information.

Acknowledgments This work was supported in part by the Intramural Research Program of the National Institute of Environmental Health Sciences, National Institutes of Health References 1. Gabay C, Kushner I (1999) Acute-phase proteins and other systemic responses to inflammation. N Engl J Med 340:448–455 2. NIA workshop on inflammation, inflammatory mediators, and aging. Sept. 1–2, 2004. Sponsored by National Institute on Aging, National Institutes of Health, Department of Health and Human Services. Workshop sum mary. Available at http://www.nia.nih.gov/ NR/rdonlyres/A8B847EC-8E2B-418DAD39-62D56E8CEAB2/2083/ NIAWorkshoponInflammationMtgRpt61805_ Ed.pdf. Accessed 9 Jan 2009 3. Coussens LM, Werb Z (2002) Inflammation and cancer. Nature 420:860–867 4. Alegre J, Jufresa J, Segura R et al (2002) Pleural-fluid myeloperoxidase in complicated and noncomplicated parapneumonic pleural effusions. Eur Respir J 19:320–325 5. Buttarello M, Plebani M (2008) Automated blood cell counts: state of the art. Am J Clin Pathol 130:104–116

6. Grattan CE, Dawn G, Gibbs S, Francis DM (2003) Blood basophil numbers in chronic ordinary urticaria and healthy controls: diur nal variation, influence of loratadine and pred nisolone and relationship to disease activity. Clin Exp Allergy 33:337–341 7. Zimmermann N, Hershey GK, Foster PS, Rothenberg ME (2003) Chemokines in asthma: cooperative interaction between chemokines and IL-13. J Allergy Clin Immunol 111:227–242 8. Latimer KS, Mahaffey EA, Prasse KW (eds) (2003) Duncan & Prasse’s veterinary labora tory medicine: clinical pathology, 4th edn. Iowa State Press, Ames, IA 9. Weyrich AS, Zimmerman GA (2004) Platelets: signaling cells in the immune continuum. Trends Immunol 25:489–495 10. Kosone T, Takagi H, Horiguchi N et al (2007) Hepatocyte growth factor accelerates throm bopoiesis in transgenic mice. Lab Invest 87:284–291

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Markers of Inflammation 39. Markiewski MM, Nilsson B, Ekdahl KN, Mollnes TE, Lambris JD (2007) Complement and coagulation: strangers or partners in crime? Trends Immunol 28:184–192 40. Sitrin RG, Pan PM, Srikanth S, Todd RF 3rd (1998) Fibrinogen activates NF-kappa B tran scription factors in mononuclear phagocytes. J Immunol 161:1462–1470 41. Paraskevas KI, Baker DM, Vrentzos GE, Mikhailidis DP (2008) The role of fibrinogen and fibrinolysis in peripheral arterial disease. Thromb Res 122:1–12 42. Shen Z, Want EJ, Chen W et al (2006) Sepsis plasma protein profiling with immu nodepletion, three-dimensional liquid chro matography tandem mass spectrometry, and spectrum counting. J Proteome Res 5:3154–3160 43. He QY, Yang H, Wong BC, Chiu JF (2008) Serological proteomics of gastritis: degrada tion of apolipoprotein A-1 and alpha-1-antit rypsin is a common response to inflammation irrespective of Helicobacter pylori infection. Dig Dis Sci 53:3112–3118 44. Cho WC, Yip TT, Chung WS, Leung AW, Cheng CH, Yue KK (2006) Differential expression of proteins in kidney, eye, aorta, and serum of diabetic and non-diabetic rats. J Cell Biochem 99:256–268 45. Hanas JS, Hocker JR, Cheung JY et al (2008) Biomarker identification in human pancreatic cancer sera. Pancreas 36:61–69

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46. Kristiansson MH, Bhat VB, Babu IR, Wishnok JS, Tannenbaum SR (2007) Comparative time-dependent analysis of potential inflam mation biomarkers in lymphoma-bearing SJL mice. J Proteome Res 6:1735–1744 47. Lull ME, Carkaci-Salli N, Freeman WM et al (2008) Plasma biomarkers in pediatric patients undergoing cardiopulmonary bypass. Pediatr Res 63:638–644 48. Wu J, Kobayashi M, Sousa EA et al (2005) Differential proteomic analysis of bronchoal veolar lavage fluid in asthmatics following seg mental antigen challenge. Mol Cell Proteomics 4:1251–1264 49. Ezendam J, Staedtler F, Pennings J et al (2004) Toxicogenomics of subchronic hexachlo robenzene exposure in Brown Norway rats. Environ Health Perspect 112:782–791 50. Tabibiazar R, Wagner RA, Deng A, Tsao PS, Quertermous T (2006) Proteomic profiles of serum inflammatory markers accurately pre dict atherosclerosis in mice. Physiol Genomics 25:194–202 51. Parikh SV, de Lemos JA (2006) Biomarkers in cardiovascular disease: integrating pathophys iology into clinical practice. Am J Med Sci 332:186–197 52. Mehra MR, Feller E, Rosenberg S (2006) The promise of protein-based and gene-based clinical markers in heart transplantation: from bench to bedside. Nat Clin Pract Cardiovasc Med 3:136–143

Chapter 6 Evaluating Macrophages in Immunotoxicity Testing John B. Barnett and Kathleen M. Brundage Abstract Macrophages are the heterogeneous grouping of cells that are derived from monocytes. They have a multitude of functions depending on their final differentiated state. These functions range from phagocy tosis to antigen presentation to bone destruction, to name a few. Their importance in both the innate and acquired immune functions is undeniable. Xenobiotics that degrade their functional status can have grave consequences. In this chapter, we provide an overview of the types of macrophages, their hematopoietic origin and a general discussion of the many different assays that are used to assess their functional status. Key words: Macrophage, Monocyte, Phagocytosis, Reactive oxygen species, Reactive nitrogen species, Osteoclast, Autophagy, Apoptosis, Confocal microscopy, ELISA

1. Introduction The purpose of this chapter is to review the basic biology of the group of cells collectively referred to as “macrophages”. Rather than pro viding detailed methods on how to perform the assays to measure the number and functional status of these cells, we provide a reference(s) to particularly instructive literature or in some instances reference to the techniques manual where detailed methods can be found. This allows us to stretch the boundaries of macrophage functional assess ments into areas of molecular biology that we would not have space for in this chapter if we were to provide detailed protocols.

2. Hematopoietic Origin of Macrophages

Like all immune cells, macrophages originate from the hema topoietic stem cells (HSC) and arrive at their final configuration through a series of maturation steps. Each of the known maturation

R.R. Dietert (ed.), Immunotoxicity Testing: Methods and Protocols, Methods in Molecular Biology, vol. 598 DOI 10.1007/978-1-60761-401-2_6, © Humana Press, a part of Springer Science + Business Media, LLC 2010

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steps can be identified by surface markers. However, it is important to note that there appears to be much that we do not know about this maturation process. In this section, we provide a summary of the known steps as primarily discussed in a review by Weissman and Shizuru (1). As shown in Fig. 6.1, the HSC pro gresses through several intermediate stages of multipotent progenitor cells (MPC) that are progressively triggered by growth and differentiation factors. To make the figure more manageable, only one of these stages is diagrammed. The last stage of this series of progenitor cells provides an important decision point in the hematopoietic process. At this point, depending on the influ ence of known growth/differentiation factors, the cells follow the route to become common lymphoid progenitors (CLP) or common myeloid progenitors (CMP). The surface markers that are unique for each progenitor cell population are listed on Fig. 6.1. CMP cells can become granulocyte–macrophage progenitor (GMP) cells or megakaryote–erythroid progenitor (MEP) cells. Again, each of these progenitors possesses a unique set of surface markers and involves a unique combination of transcription factors. For example, GMP, which are also called granulocyte– macrophage colony forming units (GM-CFU), have PU.1high, GATA-1−, c/EMPahigh phenotype, as shown in Fig. 6.1. Under the influence of macrophage-colony stimulating factor (M-CSF), the GMP cells proceed through a semifinal series of maturation steps that ultimately results in the production of monocytes. Monocytes are circulating blood cells, whereas, macrophages are further differentiated end-cells derived from monocytes that are found in tissues. As previously described, under further influence of specific growth and differentiation factors, the monocytes develop into mature tissue-resident cells. Some of these cells have very special ized activities and names. For example, osteoclasts are respon sible for maintaining normal bone homeostasis, in balance with another cell type, the osteoblast. Osteoclasts differentiate from monocytes under the influence of receptor activator of Nuclear Factor (NF)-kB ligand (RANKL) and macrophage-colony stim ulating factor (M-CSF) (2, 3). As outlined below, several other types of macrophages are formed and are often named for the tissue in which they finally reside (e.g., alveolar macrophages are found in the lung). The maturation pathway that leads to the formation of termi nal (mature) “macrophages” presents the first area suitable for testing in the area of immunotoxicology. Agents that interfere with the maturation process can skew the number of mature cells and in some cases, the anatomical location of the hematopoi etic maturation (4), and this may have detrimental effects on the health of the animal. We will discuss this further in a later section.

Evaluating Macrophages in Immunotoxicity Testing

HSC PU.1low GATA-1− C/EBPαlow

PU.1+ GATA-1low

Mu=Lin-c-Kit+Sca1+Flk2-CD34-Slamf1+ Hu=Lin-CD34+CD38-CD90+CD45RA-

MPC (multiple stages) CLP CMP

PU.1+ GATA-1+ C/EBPα+

Mu=Lin−c-Kit+Sca1−CD34+FCγRlow Hu=Lin−CD34+CD38+IL3RαlowCD45RA−

MEP PU.1high GATA-1− C/EBPαhigh

PU.1− GATA-1high

GMP (GM-CFU) Mu=Lin − c-Kit+Sca1− CD34−FCγR+ Hu=Lin− CD34+CD38+IL3Rα+CD45RA+

M-CFU → Monoblast → Promonocyte PBM

tissue macrophage • spleen • histocyte • alveolar • Kupffer cell • microglial

osteoclast

KEY: CELLS: CLP, common lymphoid progenitor; CMP, common myeloid progenitor; GMP, granulocyte-macrophage progenitor; GM-CFU, granulocyte-macrophage colony forming unit; HSC, hematopoietic stem cell; MEP, megakaryocyte-erythroid progenitor; M-CFU, macrophage coloy forming unit; PBM, peripheral blood monocyte; MPC, multipotent progenitor cell. Transcription factors known to affect cell differentiation are listed in the boxes REFN: Bryder et al., Am J. Path 169:338 (06); Weissman & Shizuru, Blood 112:3543 (07); Iwasaki & Akashi, Immunity 26:726 (07); Mosser & Edwards, Nature Reviews Immunol 8:958 (08)

Fig. 6.1. Hematopoetic maturation scheme for the production of macrophages [see also (1,5,44,45)].

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3. Types of Macrophages As is common among the immune cells, there are several types of macrophages. Classifying these types primarily by tissue or organ of residence provides the following list: peritoneal, splenic, histo cyte, Kupffer cells, alveolar, osteoclast and microglial. Some inves tigators further divide some of these cells, e.g., within in the spleen there appear to be differences depending on whether they reside in the white pulp, red pulp or marginal zone (5). A potentially more important subdivision of macrophage populations has been extensively reported by Mosser and others (reviewed in (5)). Briefly, resident macrophage populations have been divided into three distinct populations based on the charac teristics and criteria discussed below. The first of these popula tions is the “classically-activated” (CA) macrophage. This population is generated in response to either interferon-gamma (IFNg) in combination with tumor necrosis factor-alpha (TNF-a) or to stimuli that induce TNF-a (5, 6). These macrophages pro duce reactive oxygen and nitrogen species, secrete the prototypi cal group of cytokines that are either proinflammatory or affect Th1 cell responses and have antimicrobial activities (5, 6). A second population of macrophages is referred to as “woundhealing” (WH) macrophages (also called “alternatively activated” macrophages). These cells are induced by interleukin (IL)-4 and/ or IL-13 (Th2 cytokines) and do not produce nitric oxide (NO). However, they do have an abundance of mannose receptors (6)as well as uniquely expressing the genes for FIZZ1, a resistin-like secreted protein up-regulated during pulmonary inflammation, and Ym, which is a chitnase-like secretory lectin that is also upregulated during the development of allergy (7). WH mac rophages have been reported in bone marrow (6), lung (8), peritoneal cavity (9) and spleen (10). The third population is generated by the macrophage being activated in the presence of immune complexes (6, 11). These macrophages are referred to as “regulatory” macrophages (previ ously referred to as “Type II-activated” macrophages). Cross link ing of Fcg receptors (FcgR) by the immune complexes results in strong down regulation of IL-12 with a concominant up-regula tion of IL-10 (6). Careful dissection of the biochemical and func tional differences between these three populations has revealed a number of different surface markers, biochemical functions as well as biological activities (6). The review by Mosser and Edwards (5) provides a convenient table of the characteristics of the three dif ferent populations of macrophages. The potential effect of differ ent xenobiotic compounds specifically on any of these three different macrophage populations has not been reported.

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4. Macrophage Functional Assays Figure 6.2 provides a schematic of the many functions that have been described for macrophages or other monocyte-derived cells (e.g., osteoclasts). In many cases, there is a known link between

Confocal microscopy APC×T cell cytokine prod’n

Specialized stains (TRAP)

ELISA

Colony-forming assays Antigen Presentation

Autophagy

Extracellular Protein Expression

Differentiation Immunofluorescence

Killing Surface Molecule Expression

Antimicrobial

Phagocytosis

Flow cytometry Gene Expression

Pathology

Pharmacologic Mediators

Histology

Whole genome sequencing

Apoptosis

mRNA quantn

ELISA ELISA Biochemical assays

TUNEL

Intracellular Signaling

Annexin V

Intracellular relocalization Target gene protein expression Protein phosphorylation Ca2+ influx Enzyme analysis

Fig. 6.2. Macrophages functions that are suitable for testing.

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the function described in a particular outer circle, e.g., the expression of a cell surface receptor that is necessary for initiating antimi crobial activity, which in turn requires a cascade of intracellular signaling. However, for the purposes of this chapter, we will describe (in alphabetical order) assays available to measure each of these characteristics individually with the assumption that each assay is interpreted in the context of the function of the whole cell. Before discussing each assay, a brief discussion of the methods used to isolate macrophages from tissues is provided. Because macrophages reside in tissues, a number of laboratory procedures have been developed to enrich the cell harvest for macrophages (i.e., reduce the number of nonmacrophage cells in the prepara tion). An excellent detailed description of a number of methods is provided by Zhang et al. (12). A key feature and problem with macrophages is their “stickiness”. They can (and do) adhere to glass and plastic surfaces, which is useful in the procedures needed to enrich the population for macrophages, however, in doing so, this initiates a number of intracellular signals. Thus, quiescent, unstimulated primary macrophages are difficult to isolate. Many published reports on the effect of xenobiotics on macrophage function make comparisons between treated versus untreated macrophages isolated in an identical manner to control for this problem. A commonly used source of mouse and rat macrophages is the peritoneal cavity. Two types of macrophages from the perito neal cavity are used, resident and elicited. Resident macrophages are those that are residing in the peritoneal cavity normally and have the advantage of being less biochemically stimulated but are fewer in number than if the cells were elicited. Often, to increase the number of macrophages, a sterile irritant, such as thioglycol late, is injected several days prior to harvesting the cells. The resulting peritoneal cells are referred to as elicited macrophages. While this procedure substantially increases the number of perito neal macrophages, it also has its drawbacks. That is, the irritant can be phagocytosed and, thus, upset the normal baseline physi ology of the cells. Thus, an informed choice must be made in the design of the experiment. A hint that is often overlooked by new comers who wish to elicit the peritoneal macrophages is that the thioglycollate must be aged, that is stored in the dark for several weeks after sterilization prior to use. Isolation procedures for the other macrophage populations are obviously organ dependent. Zhang et al. (12) provide detailed methods for the isolation of murine bone marrow and alveolar macrophages. Others have published some detailed methods of isolating human monocytes and macrophages mainly from either peripheral blood or cord blood (13, 14).

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A hallmark characteristic of macrophages is their ability to kill the engulfed microorganisms. Because of the requirement that the cell actually phagocytose the microorganism, these assays can be combined with assays that assess the phagocytosis of the organ ism. These assays are often the foundation of an immunotoxico logical assessment of the effect of a xenobiotic. Various bacterial species are used to assess the killing efficiency of isolated mac rophages. Care must be exercised to be certain that the assay excludes those bacteria that are bound to the surface of the cell as opposed to bacteria that have been engulfed and are in the phagosome or phagolysosome. To control this potentiality, after an initial incubation period to allow the cells to engulf the bacte ria, the macrophages are washed with a bactericidal antibiotic solution and then reincubated to determine the level of killing over time. As with many macrophage assays, the cells often must be activated in order to stimulate their phagocytic and antimicro bial functions. This activation step is dependent on the method used to obtain the macrophages. That is, if the primary mac rophages are from animals elicited with thioglycollate, then they are partially activated at the time of harvest, and the addition of interferon-gamma (IFNg) completes the activation processes leading to antimicrobial killing. Other methods of macrophage harvest (e.g., using resident macrophages) require an initial activation step with lipopolysaccharide (LPS). These stimuli can be used either sequentially (LPS then IFNg) or simultaneously. There are a number of methods that are suitable for measuring the microbial killing by primary macrophages as well as macrophage-like cell lines. A procedure used by Ustyugova et al., (15) to measure the effects of the herbicide, propanil, on mac rophage function was an adaptation of a technique published by Ouadrhiri et al. (16). THP-1 or peritoneal exudate cells (PEC) were activated with IFNg for 24 h in RPMI 1640 media containing 10% FBS (no antibiotics). This activation does not cause adhesion of THP-1 cells, which continued to grow in loose suspension as described (16). THP-1 cells were washed with PBS, resuspended in PBS containing 1% fetal calf serum (FCA), and infected with 2.5 × 106 Listeria monocytogenes, strain EGD/ml. Activated PEC were trypsinized, washed in PBS, resuspended in PBS containing 1% FCS, and also infected with 2.5 × 106 L. monocytogenes, strain EGD/ml. LPS stimulation and vehicle and DCPA treatments were initiated at the same time as initial infection. Cells were transferred to 5-ml snap-cap tubes and incubated at 37°C in 5% CO2 on a rocker for 1 h to allow for infection. Cells were then washed twice with PBS. The final pellet was resuspended in PBS containing 25 mg/ml gentamicin. Initial infection levels (1 h) were obtained by lysing cells in ice-cold, sterile, distilled water and plating an aliquot of the lysate. Remaining treatment groups were

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returned to incubation conditions in the gentamicin solution. Cell lysates were prepared at 3 and 5 h after infection. Lysates were plated on Trypticase soy agar plates (Difco Laboratories, Detroit, MI) using an Autoplate 4000 (Spiral Biotech, Bethesda, MD) in triplicate. Plates were incubated at 37°C for 24 h, and colonies were counted using a CASBA 4 plate scanner and CIABEN colony imaging and analysis software (Spiral Biotech). Resulting L. monocytogenes colony-forming units (CFU) per 1 ml were calculated and compared across treatments. In these studies, we were able to determine that propanil inhibited the ability of primary macrophages and THP-1 (human monocytic cell line) cells to kill L. monocytogenes. 4.2. Autophagy

Autophagy is one of two methods by which cells degrade pro teins. Degradation by proteosomes (the alternative form) targets polyubiquitinated substrates, however, autophagy targets cell organelles and aggregates of long-lived proteins (17). It is fitting that this technique follows the “antimicrobial killing” section because in macrophages, autophagy is known to play a vital role in restricting viral infections and inhibiting replication of intracel lular bacteria and parasites (18). Of course, this is not its only role; it also functions in morphogenesis, cellular differentiation and tissue remodeling (19). Thus, assays for possible effects of xenobiotics on autophagy would also be an appropriate assay. The number of reports on the role of autophagy in macrophage function have increased slowly over the past few years (84 cita tions in 2008 versus 1 in 2000). However, the small number of citations also means that ways for assessing autophagy are also relatively new and untested by time. Nonetheless, the most recent methodology articles discuss the use of the expression of LC3, which is a mammalian homolog of the yeast autophagy-related protein-8 (Atg8), as a measure of autophagy (19). In macrophages, autophagosomes increase in number as a result of appropriate stimulation (e.g., by a microbe). The microbe is phagocytosed into an autophagosome after which it fuses with a lysosome to form an autophagolysosome (19). The merging of these two structures allows for the contents (e.g., microbe) and the LC3 protein to be degraded. Stimulation of the macrophages also causes an increase in autophagosome numbers in the cell and this increase can also be monitored by assaying LC3 protein levels (19). Swanson et al. (19) discuss the various methods of measuring intracellular LC3 levels in bone marrow macrophages after appropriate stimulation using either fluorescence microscopy to visually detect a LC3GFP signal or by western blotting techniques using an antiLC3 antibody. Given the current hypothesis that autophagy consti tutes an important macrophage antimicrobial function, assays of the effects of various xenobiotics on autophagy would seem to be

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an important measure. Similarly, even without the antimicrobial function, it is conceivable that induction or interference with the normal autophagy functions could affect the operative status of the macrophage. 4.3. Antigen Presentation Assays

Given that macrophages were the first antigen-presenting cell (APC) identified, there is no shortage of methods to measure this function. In spite of the longstanding knowledge of the APC capabilities of macrophages, there are few references that actually measure the effect of a xenobiotic on the APC capabilities of macrophages. In addition, some of the early work may need updating because of our new knowledge of dendritic cells. Macro phages can differentiate into dendritic cells and dendritic cells are superb APC (20). One area with some literature on the effects of xenobiotic insult on a macrophage population’s APC capacity is on alveolar cells (21) and Kupffer cells (22), and in both of these reports the outcome was an up-regulation of the immune (allergic) response. Appropriate methods to measure macrophage APC capacity are provided in Harding (23, 24). In this detailed procedure, either peritoneal macrophages elicited with L. monocytogenes pro teose peptone or Concanavalin A (ConA) or bone marrow mac rophages are used as the APC (23). In a companion chapter, Harding (24) provides a detailed protocol for using these mac rophages in an antigen-presenting assay in which the production of IL-2 by the T cells is the read out for the assay.

4.4. Biochemical Assays

Macrophages produce a number of biochemical substrates that are important to their function. Among these are the reactive oxygen species (ROS), reactive nitrogen species (RNS), prostaglandins (PG), inducible nitric oxide synthetase (iNOS), cyclooxygenase (COX)-1 and COX-2. These assays are so important to assess the function of macrophages, that commercial kits to measure their production are readily available. Using our own work as an exam ple, we have measured concentration-dependent changes in respi ratory burst using a luminol assay, intracellular ROS assessed by the fluorescent dye oxidation using confocal microscopic intensity measurements, iNOS production by western blotting, and NO production using the biochemical test called the Greiss test in elic ited PEC (15). In companion analyses (manuscript in prepara tion), we further measured COX-2 protein production by western blot and COX-2 enzymatic activity by assaying the conversion of arachidonic acid (its substrate) to PGE2. PGE2, as well as the other prostaglandin production, can be measured using any one of a number of commercial ELISA assays that are available. Respiratory burst activity can be effectively measured using Luminol, 5-amino-2-3dihydro-1,4 phthalazinedione (Sigma) to amplify the chemiluminescence (CL) signals generated by activation

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of the NADPH-oxidase (25). The CL signal generated by luminol occurs as the compound accepts an electron from free radical species as they return to ground state. The actual experimental configuration is dependent on the instrument used to measure the fluorescence produced by the respiratory burst. In our work, cells were plated in 35 mm tissue culture dishes, and the respiratory burst activity was elicited by addition of PMA and LPS simultane ously with the luminol plus either vehicle or xenobiotic (15). CL readings were taken every minute for 20 min using a luminometer (Berthold, Co., Wilbad, Germany). CL curves are generated from these readings over time to peak values. Control experiments to determine if the xenobiotic or vehicle alone quenched the CL signal are also included (15). Intracellular ROS can be measured using 5-(and 6)-chloromethyl2¢,7¢-dichlorodihydro-fluorescein diacetate, acetyl ester (CMH2DCFDA, Invitrogen, Carlsbad, CA). CM-H2DCFDA is a nonfluorescent dye which enters the cells by passive diffusion. It becomes fluorescent only when it is oxidized by ROS, particularly by hydrogen peroxide (H2O2), hydroxyl radicals (•OH), or their downstream free radical products. After incubation with the dye, cells are typically washed and the cells are treated with the xeno biotic or vehicle control and simultaneously stimulated with LPS and PMA. In our studies, cultures are examined with a Zeiss Axiovert 100 M microscope equipped with a laser-scanning confocal attachment (model LSM 510; Zeiss) to locate the cells and analyze their images overtime (15). CM-H2DCFDA is excited with the 488-nm line of an argon/krypton mixed-gas laser; emis sion was collected with a 505 nm long-pass filter and images were collected every 2 min for up to 20–30 min (15). Fluorescence emission from 25–30 cells per experiment is analyzed by LPS 510 software (15). NO release from macrophages is typically measured using the Greiss reagent (0.1% naphthylethylenediamine chloride (Sigma) in 60% acetic acid and 1% sulfanilamide (Sigma) in 30% acetic acid. The cells are stimulated with LPS, simultaneously treated with the xenobiotic and incubated for 12, 24 and 48 h for NO induction by monitoring color development at 540 nm with a mQuant plate reader (BioTek Instruments, Inc. Winooski, VT). A standard curve is generated with a serial dilution of sodium nitrite dissolved in culture medium. 4.5. Calcium Influx

The quintessential method of measuring calcium (Ca2+) influx into cells is via patch clamping. This requires very specialized equipment that may not be readily available to many immuno toxicologists but, nonetheless, excellent indications of Ca2+ influx can be obtained using one of several Ca2+-specific indicator dyes, such as Fluo-3 and Fura-2. These dyes have been used with a variety of cell types to monitor Ca2+ influx, frequently using a

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spectrofluorometer with a stirring cuvette to keep the cells in suspension. This method works very well for nonadherent cells (e.g., T cells) (26). Use of these dyes requires that the cells remain in suspension, hence the stir-bar equipped cuvette. Thus, mac rophages with their inherent “stickiness” are not generally amenable to these technique if a spectrofluorometer is the instrument used to measure the fluorescent output. We have adapted a technique that uses a fluorescence-recording inverted microscope with appropriate software to measure the quantity of light. Macrophages are allowed to adhere for 2 h at 37°C in a tissue culture incubator to coverslip bottomed culture chambers specifically designed for fluorescence microscopy (LabTek #1 Borosilicate chambers, Nalge Nunc Intl.). The cells are then loaded with the esterified form of the appropriate dye, which enters the cell and is trapped by intracellular esterases that cleave the dye into an active form. For Fura-2, cells are loaded with Fura-2AM with the addition of pluronic, a detergent which aids in the uptake of the dye for 30 min at 37°C in the dark. A group of control cells is treated with the dye diluent alone to act as a background fluorescent measure during the experiment. The cells are then washed with tissue culture media without phenol red to reduce the autofluoresence of the media. Fields of cells can then be monitored for changes in cytosolic Ca2+ due to increased influx or the release of Ca2+ from the internal stores over several minutes. The effect of a xenobiotic on the Ca2+ influx can be measured in comparison to agents known to cause Ca2+ influx, e.g., thapsi gargin, as well as known channel/pump-specific inhibitors. For calculation of the actual intracellular concentration of cal cium, calibration measurements are taken at the end of the exper iment. These typically consist of permeabilizing the cells with ionomycin and driving intracellular calcium to saturating levels (Rmax) and then lowering it to sub nanomolar levels through the addition of EGTA (Rmin). Detergent treatment can also be used to monitor the uptake of the dye into intracellular compartments which can confound the cytosolic signals. Digitonin is typically used to lyse the plasma membrane to release cytosolic dye while maintaining organelle integrity so that the fluorescent signal from these compartments can be detected. 4.6. Colony Forming Assays

As shown in Fig. 6.1, each of the progenitor cell populations beginning with the HSC has a unique combination of cell surface markers. The relative numbers of each of these cell types varies as the hematopoietic maturation process proceeds. The number of HSC is reported to be 0.05% of bone marrow cells (27). Thus, while it is possible to measure each of these progenitor and HSC populations via flow cytometry, this is not always practical because of the limited availability of reagents to measure each of the pro genitors as well as their low total numbers in the specimen.

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Therefore, to determine the myelo-toxicity of xenobiotics, the ability of specific progenitor populations can be determined using the colony-forming assays. Colony forming units (CFU) are induced with recombinant colony stimulating factors (rCSF) in culture conditions described by Metacalf (28) or similar. CFU-IL-3 (multipotent progenitor cells), CFU-GM (granulocyte/macrophage progenitors) and CFU-M (macrophage [monocyte] progenitors) can be enumer ated after 6 days in culture in semisolid agar cultures. The semi solid agar cultures are prepared using Dulbecco’s minimal essential medium containing 20% prescreened fetal calf serum and 0.3% agarose or 2% methyl cellulose. Colonies of greater than 50 cells are counted using an inverted or dissecting microscopy. Control cultures without rCSF are also prepared and evaluated. This method has been useful in determining the myelo-toxicity of two pesticides (4, 29). 4.7. Confocal Microscopy/ Immunofluorescence

Confocal microscopy has become an indispensable tool for cell biology. There are many different types of confocal microscopes, but the most commonly available is the laser-scanning confocal microscope (30). The basic design of this microscopy involves exciting the specimen in the microscopy with a narrow band laser and capturing the emitted fluorescent image that is returned. The availability of many different dyes (stains) that emit light at different wavelengths provides the ability for multiple staining of a specimen. From this, it is possible to detect the intracellular location of a particular organelle or molecule and compare this to the location of a separate molecule by merging the images to produce a separate color. The combinations of fluorochromes with different antibodies attached are large, and the further use of secondary fluorescent-tagged antibodies make this instrument extremely versatile. Newer models can image living cells making it possible to study the effects of different stimuli on on-going biochemical processes (31). These microscopes have the further ability to “focus” on different planes in the specimen (frequently called the Z-stack), thus, providing a complete picture of the loca tion of molecules throughout the entire cell. Examples of the use of confocal microscopy in immunotoxicology include the work of Henjakovic et al. (32), who reported on the use of precision lung slices for ex vivo immune response testing. Similarly, Neumann et al. (33) reviewed the courses of L. monocytogenes within the macrophage using confocal microscopic techniques. Excellent protocols are available in comprehensive methods manuals, such as Current Protocols in Cell Biology. A general tech nique that has been reported specifically for the immunotoxico logical studies used IC-21 macrophages that were grown on coverslips overnight to approximately 70–90% confluence (34). The following day, cells were either not stimulated or stimulated

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with LPS in the presence of either vehicle, or propanil for 1–4 h. Coverslips were rinsed in phosphate buffered saline (PBS), then the cells were fixed in a 1:1 methanol:acetone solution, followed by blocking in PBS + 5% bovine serum albumin (BSA). Rabbitanti-p65 (Santa Cruz Biotechnology, Santa Cruz, CA) was added to the cells at 100 mg/ml in PBS and incubated at room tempera ture for 1 h. Coverslips were washed 3× in PBS for 5 min each with gentle rotation. Goat-anti rabbit FITC was added at 50 mg/ml in PBS for 1 h at room temperature and kept in the dark. Coverslips were washed 3× in PBS for 5 min each, then mounted to glass slides with Fluormount-G (Southern Biotechnology Associates, Inc, Birmingham, AL). Slides were kept in the dark until visua lized by confocal microscopy (Zeiss LSM 510, Thornwood, NY, equipped with an argon laser). This technique allowed us to follow the translocation of NF-kB p65 into the nucleus of these macrophages (34). In summary, the potential uses of confocal microscopy is vast, progress on the newer types of instruments and software is so rapid, that a comprehensive review of this field is not practical within the limitations of this chapter. 4.8. Enzyme-Linked Immunosorbent Assay (ELISA)

4.9. Flow Cytometry

ELISA methods are commonly used to quantitatively measure the protein levels, and there are many variations on the basic prin cipal of this assay. In its simplest form, the protein to be assayed (analyte) is captured (usually in a 96 well tissue culture plate) by its binding to either its specific antigen (if the analyte is an antibody) or using a specific antibody to the analyte for other proteins. A blocking agent, often an unrelated protein such as bovine serum albumin, to prevent nonspecific binding of the next reagent, is then added. The captured analyte is then detected using a second antibody to which an enzyme, such as horse radish peroxidase (HRP), is conjugated. The last step is to add a sub strate for the HRP enzyme that changes color in direct propor tion to the amount of enzyme present. There are extensive wash steps between each step in the assay to eliminate any nonspecific reactions. Thus, the amount of substrate color developed is directly proportional to the amount of the analyte in the sample. This method works well for a variety of fluids, including culture supernatants and serum. This technique is very specific, quantita tive, reproducible and amiable to high throughput analyses. The ELISA technique has become a standard for measuring a vast array of proteins and there are a huge number of commerciallyavailable ELISA kits available to measure these proteins. However, if it is necessary to develop an ELISA for a specific protein, an excellent protocol is described by Hornbeck et al. (35, 36). Flow cytometry is the measurement of the physical and/or the chemical characteristics of single cells or particles as they pass

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through the measuring apparatus in a fluid stream. It utilizes laser beams that hit the cells or particles in the flow cell. A fluorescent objective lens collects and focuses the fluorescent light emitted onto the individual collection fibers. Fiber optic cables transfer the light to the detector arrays. It has been around for many years and is a valuable tool for not only phenotyping cells but also assaying their metabolic activity, viability, DNA integrity and pro liferative capacity. In order to identify cells of the macrophage lineage in the bone marrow, a series of antibodies are used that are conjugated to different fluorochromes and specific for different surface markers. Figure 6.1 shows the combination of the surface proteins that allow for the identification of HSC, MPC, CMP and GMPs. In addition, M-CFU can be identified by the surface markers c-Kitlo, CD13+, CD45RA+, CD33+, MHC Class II+, CD2+/−, CD7+/− and CD34− and monocytes can be identified by the surface markers CD11b+, CD14+, CD33+, CD2−, CD40−, CD80−, CD86−, CD45RA−, TLR2+, TLR4+, TLR5+ and TLR8+. There are a number of surface markers that can identify monocytes/macrophages in the periphery. In mice, CD11b, CD14 and CD115 are the key markers while in humans CD11b and CD14 are used to identify these cell populations. There are other surface markers that can be used to determine the activation state of the cells such as MHC Class II, CD80 and CD86. Apoptosis or programmed cell death is one process that can be analyzed by flow cytometry using the interaction of annexin V and the membrane phospholipid phosphatidylserine (PS) in the presence of calcium. PSs are not normally exposed on the surface of a cell unless the cell is in the process of apoptosis. It is one of the earliest indicators that the cell is undergoing apoptosis. In this assay annexin V conjugated to a fluorochrome, usually fluorescein isothiocyanate (FITC), is incubated with the cells for several min utes (37–39). Excess annexin V is washed away, and propidium iodide is added to discriminate between cells dying via apoptosis from those dying by necrosis (37–39). There are other flow cytometry based assays that can be used to assess if a cell is undergoing apoptosis including measuring a decrease in mitochondrial membrane potential. This is done using the fluorescent probe 3,3′-dihexyloxacarbocyanine (DiOC2(3)). DiOC2(3) is a cell permeant green fluorescent lipophilic dye that is selective for mitochondria at low levels. Cells are incubated with this dye for a short period of time at 37°C and assayed by flow cytometry. A decrease in green fluorescence is indicative of a decrease in mitochondria membrane potential and thus a cell undergoing apoptosis (38). Another important assay of macrophages that uses flow cytometry is one that looks for the generation of reactive oxygen species (ROS). This is usually performed using the nonpolar

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compound 2,7-dichlorodihydrofluorescein diacetate H2DCFDA. This compound diffuses easily into cells. Once in the cell the acetate group is removed by endogenous intracellular esterases. Thus, H2DCFDA is converted to H2DCF and becomes trapped in the cell. When cells loaded with the indicator dye produce hydrogen peroxide and other peroxidases, these compounds oxidize the H2DCF to DCF (2,7¢-dichlorofluorescein), which is highly fluorescent. The standard protocol is to incubate the cells with 1–30 mM of dye for 5–60 min at 37°C then the unincorpo rated dye is removed (38). The more ROS produced by the cell the higher the fluorescence in the cell. 4.10. Protein Phosphorylation

The process of phosphorylation and dephosphorylation is a common mechanism used by many signaling molecules to switch between the active and inactive states. The process of phosphory lation changes the quaternary configuration of the signaling molecule revealing the active site, and the molecule then has phos phokinase activity with specificity for the next molecule in the cascade. This triggers a signaling pathway. Each signaling molecule can then be, in turn, switched off by a specific phosphatase that removes the phosphate returning the molecule to its inactive state. The technique is frequently used to detect the effects of xenobiot ics on specific signaling molecules by assaying cell extracts for the phosphorylated form versus nonphosphorylated form. Basically the phosphorylated form is detected by developing western blots using antibody specific for the phosphorylated form. For example, primary antibody specific for total c-jun (non phosphorylated form) versus phospho c-jun ser 73 (phosphoryla tion of serine 73) will provide an indication of the effect of a xenobiotic on this signaling molecule (40). If the molecule has multiple phosphorylation sites, then specific antibody for these molecules when phosphorylated on a specific site can also be very informative about the upstream problems that a xenobiotic may have caused because different phosphorylation sites may be phos phorylated by different upstream phosphokinases (40).

4.11. Transcriptional Activation/RNA Quantification

Activation of cells often involves de novo protein production with the accompanying transcription of their genes. Thus, quantifying the action of a xenobiotic on transcription is an excellent place to begin to determine the mechanism of the effects of a xenobiotic. This is done by monitoring the mobilization of transcription fac tors to the nucleus, the binding of transcription factors to pro moter targets or the initiation or rate of transcription (mRNA production). Often more than one assay is performed to obtain a more complete picture of what is happening to the cell, e.g., translocation of a transcription factor to the nucleus, its binding to the promoter and the subsequent mRNA generation. This combination can indicate roughly where an alteration has occurred

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in the signal transduction pathway. A brief description of some standard assays is provided below. Nuclear translocation can be ascertained visually (see Subheading 6.4.7) or by measuring protein levels in specific cellular subcompartments. For example, NF-AT a common tran scription factor for many cytokines is translocated to the nucleus after Ca2+-dependent activation. To monitor this phenomenon, cytoplasmic and nuclear cellular fractions can be prepared after appropriate activation using IFNg and LPS (41). The purity or lack of cross contamination of each of the fractions must be con firmed by measuring the organelle- or compartment-specific markers (e.g., Oct1 for nuclear contents and GAPDH for cyto plasm). Usually this is done by western blot. Similarly, the protein of interest, in this case NF-AT, can be measured using western blotting techniques. This technique is especially informative when there are “qualitative” differences between the two states, (i.e., the transcription factor is absent in the nucleus prior to activation and present in the nucleus after activation). Similar methods can be used to measure the other transcription factors when different forms of the transcription factor are present pre- and postactiva tion (e.g., we have used this approach to measure activation of NF-kB in stimulated IC-21 cells) (34). Use of a gene reporter assay is an excellent way to monitor the transcriptional activation. As an example, to monitor NF-kB acti vation, macrophages can be transiently transfected with a NF-kBdependent plasmid (PBIIX) using a transfection agent such as FuGene6 (Boehringer Manneheim, Indianapolis, IN) (34). This reporter plasmid contains two kB sites from the Ig-k enhancer upstream of a luciferase reporter gene (42). Cells were stimulated with LPS in the presence or absence of the xenobiotic for the appropriate times. Whole cell lysates were made using luciferase assay lysis buffer (Promega, Madison, WS), and luciferase activity was measured using luciferase assay reagent (Promega, Madison, WS) in a luminometer. Protein concentration of the lysates must be determined to normalize luciferase activity. The relative luciferase activity is a direct indicator of NF-kB activation (42). Two methods to monitor binding of a transcription factor to its promoter target are available. The first method is the electro phoretic mobility shift assay (EMSA) which monitors the shift in migration of the transcription factor in a gel with and without binding to the DNA. The use of this assay for macrophages is described in Frost et al (34) and Klinke et al. (41). To perform this assay, NF-kB consensus oligonucleotides (e.g., sc-2505, Santa Cruz Biotechnology, Santa Cruz, CA) are labeled with g-32P-ATP using a Ready-To-Go T4 polynucleotide kinase kit (Amersham Pharmacia Biotech, Piscataway, NJ). Nuclear extracts are incubated with the labeled probe for 30 min at room tem perature in binding buffer to allow formation of complexes.

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Additional complexes (called supershift complexes) are obtained by preincubating nuclear extracts in binding buffer with NF-kB p65 specific antibody (Rockland Immunochemicals, Gilbertsville, PA) or p50 specific antibody (Santa Cruz Biotechnology) before addition of the labeled probe for an additional 30 min at room temperature. Complexes are resolved from free probe in poly acrylamide gels, dried on Whatman 3MM paper and placed on PhosphorImage screens for analysis with a Molecular Dynamics PhosphorImager (Sunnyvale, CA) and/or X-Ray film (Eastman Kodak Company). Typically in this assay (as described above), at least three bands are shown on the gel, one for the free probe (unbound), a shifted band that is the probe bound to the DNA promoter and a supershifted band of the probe bound to the DNA promoter with the addition of the antibody (34). Another method to measure DNA binding of transcription factors has been developed called the Chemicon Nonradioactive Transcription Factor Assay (NTFA) (Millipore, Billerica, MA). This assay avoids the use of radioactive probes and in our labora tory it has been proven to be more reproducible than EMSA for measuring DNA-binding of NF-kB (41). Briefly, nuclear extracts are incubated with biotinylated NF-kB consensus DNA sequences in a transcription assay buffer containing sonicated salmon sperm DNA to allow the formation of protein/DNA complexes. Next, NF-kB protein in the nuclear extracts bound to biotinylated consensus sequences are immobilized to a streptavidin-coated plate and any unbound material washed away. NF-kB protein is detected with a specific rabbit anti-p65 antibody, followed by incubation with anti-rabbit HRP-conjugated antibody plus the HRB substrate, TMB/E (3,3′,5,5′-tetramethybenzidine). Absorbance of the samples is measured with a microplate spec trophotometer. The NTFA experimental results are reported as the ratio of the absorbance measured for a particular condition (i.e., time > 0) relative to the absorbance before treatment (i.e., time = 0). 4.12. Transcriptome Expression Analysis

There have been a number of reviews on the use of gene arrays to determine changes to cells after stimulation. One particularly pertinent study is the gene expression profiling during differen tiation from monocyte to macrophage by Lehtonen et al. (43). During this process, human monocytes were induced to differ entiate into macrophages using GM-CSF and IL-4 (43). Using an Affymetrix oligonucleotide microarray system which queries ~13,000 genes, they found that 340 and 190 genes were upregulated and down-regulated, respectively, during this differentiation process (43). The numbers changed over time after the stimulus to differentiate was added (43). At least some of these genes can be understood from a biological perspective (e.g., up-regulation of the transcription factor C/EBPb) (43).

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The real challenge for these studies is how to assemble all of the data into a coherent story of how the choreography of activa tion occurs (i.e., what gene products control which down-stream genes, what inhibitor genes must be down regulated and when, etc). Although the study by Lehtonen et al. (43) was well designed and controlled, it left many questions unanswered. Part of the reason for this is that the bioinformatics software has lagged behind the technology and chemistry of collecting the raw data and the biology of understanding these relationships. Performing the actual RNA extractions as well as the other steps, needed to do these studies is aided by the very detailed protocols provided by the array chip manufacturer. In some cases, there must be concomitant instrumentation purchased to “read” the chips (e.g., the Affymetrix system). These instruments often include the software for the basic analysis of the data (i.e., which “genes” are up- or down-regulated and by how much). However, further analysis of the data usually is performed using either com mercial software specifically designed for this purpose or open source software, such as Bioconductor, that runs under the R-scripting computer language platform. Bioconductor is avail able free at www.bioconductor.org and includes several modules specifically designed for array analysis, including several specifi cally designed modules to read and analyze the data obtained using the Affymetrix system. Detailed instructions are included on the site on how to install and use this software. R can also be obtained free at http://cran.r-project.org. This technique is further hampered by the lack of agreement on how to best analyze the data. There is an attempt to “ware house” the huge amount of data collected so that it can be used (or mined) by other investigators. Many scientific journals require that array data by stored using the MIAME (minimum informa tion about a microarray experiment) standard. More detailed information on this can be found on the MGED Society web page (http://mged.org). Although not readily possible at the time of this writing, the prediction is that whole genome sequencing will soon be fast and inexpensive enough, that this approach will be used to either sup plement or replace the array chips. References 1. Weissman IL, Shizuru JA (2008) The origins of the identification and isolation of hematopoietic stem cells, and their capability to induce donor-specific transplantation toler ance and treat autoimmune diseases. Blood 112:3543–3553 2. Xing L, Schwarz EM, Boyce BF (2005) Osteoclast precursors, RANKL/RANK, and immunology. Immunol Rev 208:19–29

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and c-jun activation in T cells exposed to the amide class herbicide 3,4-dichloro propionanilide (DCPA). Toxicol Sci 79:98–105 Klinke DJI, Ustyugova IV, Brundage KM, Barnett JB (2008) Modulating temporal control of NF-kappaB activation: implications for therapeutic and assay selection. Biophys J 94:4249–4259 Kopp E, Ghosh S (1994) Inhibition of NF-kappa B by sodium salicylate and aspirin. Science 265:956–959 Lehtonen A, Ahlfors H, Veckman V, Miettinen M, Lahesmaa R, Julkunen I (2007) Gene expression profiling during differentiation of human monocytes to macrophages or dendritic cells. J Leukoc Biol 82:710–720 Iwaskai H, Akashi K (2007) Myeloid lineage commitment from the hematopoietic stem cell. Immunity 26:726–740 Bryder D, Rossi DJ Weissman IL (2006) Hematopoietic stem cells: the pradigmatic tissue-specific stem cell. Am J Pathol 169: 338–346

Part III Immunotoxicity and Host Resistance Models

Chapter 7 Host Resistance Assays Including Bacterial Challenge Models Florence G. Burleson and Gary R. Burleson Abstract Immunotoxicity testing is used to provide safety assessment with the major objective being the avoidance of unacceptable risk of infectious or neoplastic disease. To this end, immunotoxicity testing has employed a variety of host resistance challenge models for measuring both host resistance to disease as well as immune function. This chapter provides an overview of those viral, bacterial, fungal, and parasitic host resistance models that are most commonly used in safety assessment. It also describes in more detail the bacterial challenge models that are employed to address specific host resistance and immune function issues. Key words: Host resistance (HR) assays, Bacterial models, Immune challenge, Marginal zone B lymphocytes, Streptococcus, Listeria, Pseudomonas, Immune biomarkers, T-Independent antibody response, T-dependent antibody response

1. Introduction The purpose of immunotoxicity testing is to obtain data that are meaningful for safety assessment. For immunosuppression the major objective is to determine the significance with respect to increased susceptibility to infectious or neoplastic disease. Host resistance (HR) assays provide the only sure method of examining the influence of test articles on the functional integrity of the immune system and its ability to eliminate pathogenic microorganisms or tumor cells. HR assays are used to evaluate the effect of a test article on clearance of an infectious microorganism in order to assess functional immunocompetence. Immunotoxicity caused by a test compound may result in an impaired clearance of an infectious agent, increased susceptibility to opportunistic

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infections, prevention or ineffective immunization, exacerbation of latent viral infections, or unintended immunostimulation. HR assays provide a means to directly assess the functional reserve of the immune system and the opportunity to measure immune biomarkers (all categories of immunoglobulins as well as inflammatory mediators) that have implications beyond infectious diseases and cancer. HR assays may be classified into either comprehensive HR assays or targeted HR assays. Clearance of an infectious microorganism allows an assessment of immunocompetence and serves as a biomarker of net immunological health. Immunological clearance of the infectious challenge agent is a more sensitive and meaningful measure of immunological function (1–3) than mortality. The number of infectious particles per organ or per gram of organ is quantified. Challenging the immune system with an extremely virulent or with an extremely high titer of infectious agent may overwhelm the immune system, with death occurring before development of the cascade of immunological responses required for clearance. Challenge with a highly virulent agent or with a high titer of infectious agent may reflect a model of sepsis or result in a “cytokine storm.” Titer does not necessarily correlate with mortality; that is, similar titers of virus were reported in the lungs of mice infected with either the mouse-adapted lethal influenza A/ Hong Kong/8/68 virus or the mouse-adapted nonlethal influenza A/Port Chalmers/1/73 virus (3). Viral titers also did not correlate with mortality in studies evaluating the immunotoxicity of TCDD (4). Screening assays to detect immunosuppression are surrogates for functional assays that are surrogates for host resistance assays. Luster and colleagues initiated a series of studies that form the basis of risk assessment in immunotoxicology evaluations. Luster et al. (5–9) evaluated immunological assays that predicted immunotoxicity and reported concordance values using host resistance as the comparator, since host resistance assays are considered to be the ultimate predictor of adverse effects (10). HR assays are the gold standard for immunotoxicological evaluation and there are numerous models available. The major function of the immune system is protection from infectious or neoplastic disease and most immunotoxicologists regard host resistance assays to be the most relevant for: (1) validating the usefulness of other detection methods and (2) extrapolating the potential of a substance, drug, or chemical to alter host susceptibility in the human population (10). In summary, HR assays provide information to determine if a test agent results in an adverse effect (decreased clearance) as well as information concerning the mechanism(s) of the adverse effect (cytokines, innate immune function, or adaptive immunity).

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2. Host Resistance Assays 2.1. Comprehensive Host Resistance Assay 2.1.1. Influenza Virus Host Resistance Assay

The influenza model in mice or rats is used to evaluate the overall health of the immune system, i.e., how the numerous components (Table 7.1) of the functional immune system work together to clear an infection while targeted host resistance assays are available to evaluate specific immunotoxicity questions (Table 7.2). The influenza host resistance assay is discussed in Chapter 8. Clearance of influenza virus requires an intact and functional immune system that incorporates a cascade of immune responses. HR assays serve as biomarkers of net immunological health or immunological well-being. Viral clearance requires all aspects of the immune system to work together and is the ultimate measure of the health of the immune system. Mechanistic immune functions may be included while measuring viral clearance and include: cytokines, macrophage activity, natural killer (NK) cell activity, cytotoxic T lymphocyte (CTL) activity, and influenza-specific IgM and IgG. Measurement of these immunological functions provides an evaluation of innate immunity (macrophage or NK activity), an evaluation of cell-mediated immunity (CMI) (CTL activity), and an evaluation of humoral-mediated immunity

Table 7.1 Comprehensive host resistance model to test the overall health of the immune system Influenza virus host resistance model: Viral clearance – Primary endpoint Mechanistic endpoints: • Cytokines • Interferon activity • Macrophage activity • NK cell activity • CTL activity • Influenza-specific IgM, IgG (IgG1 and IgG2a) – TDAR • Immunophenotyping • Histopathology Mechanistic endpoints may or may not be included

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Table 7.2 Targeted host resistance models for evaluation of immunotoxicity Targeted host resistance models: 1. Evaluation of innate immunity: • Streptococcus pneumoniae pulmonary host resistance model 2. Evaluation of therapeutics affecting neutrophils and/or macrophages: • S. pneumoniae pulmonary host resistance model 3. Evaluation of anti-inflammatory therapeutics: • S. pneumoniae pulmonary host resistance model 4. Evaluation of therapeutics targeting TNFa: • S. pneumoniae pulmonary host resistance model 5. Marginal zone B (MZB) cell evaluation: • Systemic S. pneumoniae host resistance model to evaluate MZB cells 6. Neutrophil defect/Gram negative bacterial model: • Pseudomonas aeruginosa pulmonary host resistance model 7. Intracellular bacterial model for evaluation of liver and splenic macrophages and neutrophils: • Listeria monocytogenes systemic host resistance model 8. Fungal host resistance model: • Candida albicans host resistance model 9. Latent viral reactivation host resistance model: • Murine cytomegalovirus (MCMV) host resistance model 10. Tumor host resistance model: • B16F10 Tumor Model • PYB6 Tumor Model 11. Parasite host resistance model: • Trichinella spiralis • Malaria

(HMI) (influenza-specific IgM or IgG). Measurement of influenzaspecific IgM or IgG also provides a measurement of T-dependent antibody response (TDAR) since influenza is a T-dependent antigen (1, 4, 11–14).

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2.2. Targeted Host Resistance Assays

While the influenza host resistance assay is used to assess the overall health of the immune system, targeted host resistance models are available to answer specific questions concerning the immune system. Targeted host resistance assays may be used if a specific defect has been shown to occur or is of concern. Targeted HR assays will determine whether the decreased immune function is adverse (i.e., does the percent decrease in immunological function translate to a decreased clearance of the infectious agent). Targeted HR models are available to evaluate specific immunotoxicity questions (Table 7.2). These HR models answer specific questions concerning status of the immune system.

2.2.1. Latent Virus Reactivation Model

The murine cytomegalovirus (MCMV) latent viral model is a model to assess reactivation of latent viral disease as a result of immunosuppression. There are many similarities between the viruses responsible for latent/reactivated viral disease. CMV (cytomegalovirus), EBV (Epstein-Barr Virus), and HSV (Herpes Simplex Virus) belong to the Herpesviridae virus family, while BK virus and JC virus belong to the Papovaviridae virus family. All these viruses have double stranded DNA (the human polyoma viruses are circular), are ubiquitous in the human population, and cause mild primary infections followed by a latent viral infection. Additionally, immunosuppression, especially suppressed CMI, results in reactivation of latent viral infection (15). The MCMV reactivation model may be used to evaluate a pharmaceutical agent to determine if suppression of CMI or HMI results in reactivation of latent virus. Reactivation of latent virus may result in a fatal disease such as progressive multifocal leukoencephalopathy (PML).

2.2.2. Fungal HR Model

Candida albicans is a well-characterized fungal host resistance model (16, Burleson personal communication). Candida is administered intravenously and mortality or clearance monitored. Candida-specific IgG and cytokines may also be quantified.

2.2.3. Parasite HR Models

Parasite HR models have also been used for immunotoxicity testing. These include malaria (17) and Trichinella spiralis (18). Parasite models are discussed in Chapter 9.

2.2.4. Tumor HR Models

Tumor HR models have been used for immunotoxicity testing using the syngeneic tumor cell models B16F10 and PYB6 (19). Examples of tumor challenge protocols are presented in Chapter 10.

2.3. Bacterial HR Assays

There are several targeted host resistance models that may be used to answer specific questions concerning immune system status.

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2.3.1. Evaluation of Innate Immunity

The Streptococcus pneumoniae pulmonary HR model has been used in Balb/c and C57BL/6 mice and Fischer (CDF), Lewis, and Sprague Dawley (CD) rats. Animals are infected intranasally and bacterial clearance measured. Bacterial clearance is quantified before the specific, acquired adaptive immune system is operative, and bacterial clearance is evaluated by determining the number of colony forming units (CFU) per gram of lung tissue. Dexamethasone or cyclophosphamide is used as a positive immunomodulatory control that has an immunosuppressive effect on innate immunity and decreases bacterial clearance. Cytokines may also be measured in the streptococcal model. The S. pneumoniae host resistance model in mice has been used in numerous immunotoxicity evaluations and was reported as one of a battery of three host resistance assays to evaluate a small molecule therapeutic targeted for splenic tyrosine kinase (Syk) (20). Likewise, the Streptococcal host resistance model in rat has been used in numerous immunotoxicity evaluations (21). One advantageous feature of the model is that cytokines may be measured in the lung as well as in the serum. Bacterial titers and bacterial clearance are quantified as the number of colony forming units (CFU) per organ or per gram of tissue. Additionally, macrophage and/or neutrophil function assays can be measured as a mechanistic probe if an effect on bacterial clearance is observed. However, the most conclusive single endpoint is bacterial clearance.

2.3.2. Evaluation of Therapeutics Affecting Neutrophils and/or Macrophages

Rodent models for bacterial pneumonia can be used to evaluate immunotoxicity that may predispose to bacterial pneumonia. Macrophages were demonstrated to be important in the clearance of streptococci from the lungs of mice (22) and rats (23). Further studies by Gilmour and Selgrade (23) demonstrated the importance of neutrophils in pulmonary streptococcal disease in rats by pretreatment with an antibody to neutrophils. S. pneumoniae has been used in mice and rats as a pulmonary infection following intranasal infection (22, 23; Burleson and Burleson personal communication) and has been used to evaluate whether pharmaceutical agents have either neutrophil and/or macrophage immunotoxicity.

2.3.3. Evaluation of Inflammatory Therapeutics

The S. pneumoniae host resistance model has been well characterized in mice and rats. Animals are infected intranasally and bacterial clearance measured. Bacterial clearance is evaluated by determining the number of CFU per gram of lung tissue or per lung. Dexamethasone is used as a positive immunomodulatory control as it has a suppressive effect on innate immunity and delays bacterial clearance. Komocsar et al. (24) used the S. pneumoniae pulmonary host resistance model in Lewis rats to assess the effects of anti-inflammatory agents on innate immunity. The

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model was able to predict suppression of the innate immune response to S. pneumoniae after administration of anti-inflammatory test articles. The ability to rank order the severity of innate immune suppression with multiple test articles in the same study enhances the utility of this model for screening potential drug candidates. 2.3.4. Evaluation of Therapeutics Targeting TNF-a

The S. pneumoniae host resistance model is also valuable for evaluating the importance of macrophage cytokines on bacterial host resistance. Human biological therapeutics targeting inhibition of TNF-a have been used to treat inflammatory autoimmune diseases such as rheumatoid arthritis, psoriasis, and Crohn’s disease. Decreased TNF-a as a result of treatment with monoclonal antibodies (mAb) to TNF-a has an effect on several biomarkers of infection (25–28). These studies have reported that treatment of mice with a mAb to TNF-a results in altered levels of TNF-a in the lungs and serum, decreased neutrophils and increased numbers of bacteria (impaired bacterial clearance) with decreased survival in mice infected intranasally with S. pneumoniae. The Streptococcal pulmonary host resistance model is thus an important means to assess the functional immunological capacity of macrophages and neutrophils as well as macrophage cytokines. Therapeutic agents that target TNF-a may be tested using the S. pneumoniae pulmonary host resistance model, and this host resistance assay may be used to choose a lead compound among compounds with equivalent therapeutic efficacy based on immunosuppression. Monoclonal antibody to TNF-a has a dramatic effect on bacterial clearance in this model. Pseudomonas aeruginosa can also be used as a pulmonary bacterial host resistance assay to evaluate the immunotoxicity of therapeutics when immunotoxicity is suspected in neutrophils, macrophages, and/ or TNF-a (29; Burleson and Burleson personal communication). TNF-a also plays an essential role in preventing reactivation of latent tuberculosis (30).

2.3.5. Marginal Zone B Cell HR Evaluation

Bacteria encapsulated with a polysaccharide capsule such as S. pneumoniae or Haemophilus influenzae are blood-borne pathogens that present a different challenge to the immune system. Capsular polysaccharide antigens are thymus-independent type 2 antigens (TI-2) (31) and effective immune responses are dependent on the presence of a functional marginal zone (32–34). Capsular antigens stimulate a T-independent antibody response (TIAR). The marginal zone B (MZB) cell model in mice or rats measures bacterial clearance, hematology, cytokine production, and antibody production in a kinetic fashion over a 14-day period after intravenous infection to create a blood-borne infection. MZB cells in both humans and rodents are considered a critical host defense mechanism directed against encapsulated blood-borne pathogenic

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microorganisms. Immunotoxicity directed against MZB cells not only decreases protection against blood-borne pathogens but also results in a depletion of immunological memory. In summary, T-independent antibody responses (TIAR) are decreased or ablated as a result of MZB cell immunotoxicity (35). Histopathology will detect defects in the splenic marginal zone and special immunophenotyping markers can be included to detect alteration in the number of MZB cells. Should an effect on MZB cells be observed, the pharmaceutical agent may be evaluated in the S. pneumoniae systemic MZB host resistance model for encapsulated bacteria to determine if the effect is adverse. The S. pneumoniae marginal zone B cell model has been characterized in mice and Sprague Dawley rats with a systemic blood-borne infection by intravenous inoculation. Bacteria are quantified by determining the number of CFU in the spleen, liver, lungs, and blood over a 2 week period. Cytokines, hematology, immunophenotyping, and anti-streptococcal antibody (TIAR) are also quantified in this model (Burleson and Burleson personal communication). 2.3.6. Neutrophil/Gram Negative Bacterial HR Model

P. aeruginosa is a Gram negative bacillus that is a human pathogen and primarily causes diseases of the urinary tract, burn patients, septicemia, abscesses, corneal infections, meningitis, bronchopneumonia, and subacute bacterial endocarditis. Treatment often fails and the mortality rate in Pseudomonas septicemia has been reported to be greater than 80%. P. aeruginosa is used as a pulmonary bacterial host resistance model to evaluate the immunotoxicity of therapeutics when an immunotoxic effect is suspected in neutrophils, macrophages, and/or TNF-a (29). TNF-a also is important in bacterial clearance of S. pneumoniae and plays an essential role in preventing reactivation of persistent tuberculosis (30).

2.3.7. Intracellular Bacterial HR Model for Evaluation of Liver and Splenic Macrophages and Neutrophils

The Listeria monocytogenes host resistance model is controlled primarily in the liver and spleen. The L. monocytogenes systemic infection assay is used primarily to evaluate adverse effects on neutrophils and Kupfer cells of the liver and splenic macrophages and neutrophils. NK cells and T lymphocytes also play a role in bacterial clearance. The L. monocytogenes host resistance model has been used to evaluate monoclonal antibodies (mAbs) directed against CD11b to determine whether inhibition of this adhesion molecule would enhance disease susceptibility to listeria and predict whether this anti-inflammatory therapeutic approach would enhance susceptibility to opportunistic infections in humans. CD11b/CD18 (Mac-1) is a leukocyte integrin that plays a critical role in neutrophil adhesion and the initiation of acute inflammatory processes and is therefore a therapeutic anti-inflammatory target. CD11b (alpha M integrin) complexes with CD18 (beta 2 integrin) to form complement receptor type 3 (CR3) heterodimer. Treatment with

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either monoclonal antibody NIMP-R10 or 5C6, both directed against CD11b resulted in decreased clearance of listeria in the liver and spleen with increased mortality (36), Burleson and Burleson personal communication). Neutrophils and monocytes were decreased and mice were unable to control the infectious intracellular bacterial disease. Treatment of mice with a surrogate biological mAb designated NIMP-R10 directed against the CD11b polypeptide of the CD18/CD11b heterodimer exacerbated listeriosis by preventing myelomonocytic cells from focusing at sites of infected hepatocytes in the liver. Under these conditions, an otherwise sublethal listeria inoculum grew unrestricted within hepatocytes and caused death in 3 days (36). The results obtained with NIMP-R10 are similar to those reported with a different anti-CD11b mAb (5C6) (37, 38).

3. Summary The influenza comprehensive HR assay is able to evaluate the overall health of the immune system as well as to evaluate mechanistic immunological function of the innate, CMI, and HMI of the immune system. HR assays measure the functional integrity of the immune system and are able to evaluate functional immunological reserve. The immunotoxicity safety assessment is difficult to assess when the biological significance of “X” percent change in a particular immune function is not known. Functional immune assessment must evaluate all facets of the total immune system and include innate immunity, CMI, and HMI. Targeted host resistance assays are available to evaluate specific immunotoxicity concerns including: therapeutics affecting neutrophils and/or macrophages, anti-inflammatory therapeutics, as well as therapeutics targeting TNF-a in the treatment of autoimmune disorders such as rheumatoid arthritis, psoriasis, and Crohn’s disease. Targeted host resistance assays also include evaluations of therapeutics affecting splenic marginal zone B (MZB) cells. Immunotoxicity of MZB cells may result in an increase in the number of infections with encapsulated bacteria, blood–borne infections, and bacterial pneumonias. A number of bacterial infections is possible if the antibody response to T-independent antigens is depleted or suppressed. The L. monocytogenes targeted host resistance assay can be used to evaluate systemic immunotoxicity involving Kupffer cells and neutrophils of the liver and splenic neutrophils and macrophages. Therapeutics that suppress CMI or HMI should be further evaluated with a latent virus reactivation model. Suppression of CMI can result in recrudescence of latent viral disease with resultant serious herpes virus disease, cytomegalovirus disease or

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reactivation of JC virus causing PML. Suppression of antibody production may result in a lower threshold with an increased susceptibility to the reactivated virus disease if concomitant suppression of CMI has resulted in reactivation of latent virus. Suppression of the humoral arm of the immune system may not only affect susceptibility to opportunistic infections, but may also result in ineffective immunizations, which can also be tested in HR models. There are several strategies to evaluate the potential immunotoxicity of therapeutic compounds. HR assays allow the evaluation of the overall health of the immune system, allow specific questions to be evaluated by the use of targeted HR assays, allow the use of sufficient numbers of animals to assure the statistical power to detect immunotoxicity, and allow the inclusion of positive controls necessary to confirm negative findings. It is crucial to evaluate all arms of the functional immune response in order to derive data that are useful in performing a meaningful immunotoxicity safety assessment.

Acknowledgment The authors thank Janice Dietert for her editorial suggestions. References 1. Burleson GR (1995) Influenza virus host resistance model for assessment of immunotoxicity, immunostimulation, and antiviral compounds, Chapter 14. In: Burleson GR, Dean JH, Munson AE (eds) Methods in immunotoxicology, vol 2. Wiley, New York, pp 181–202 2. Selgrade MJK, Daniels MJ (1995) Host resistance models: murine cytomegalovirus, Chapter 15. In: Burleson GR, Dean JH, Munson AE (eds) Methods in immunotoxicology, vol 2. Wiley, New York, pp 203–219 3. Lebrec H, Burleson GR (1994) Influenza virus host resistance models in mice and rats: utilization for immune function assessment and immunotoxicology. Toxicology 91:179–188 4. Burleson GR (1996) Pulmonary immunocompetence and pulmonary immunotoxicology, Chapter 7. In: Smialowicz R, Holsapple MP (eds) Experimental immunotoxicology. CRC, Boca Raton, FL, pp 113–135 5. Luster MI, Munson AE, Thomas PT, Holsapple MP, Fenters JD, White KL Jr, Lauer LD, Germolec DR, Rosenthal GJ, Dean JH (1988) Development of a testing battery

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to assess chemical-induced immunotoxicity: National Toxicology Program’s guidelines for immunotoxicity evaluation in mice. Fundam Appl Toxicol 10(1):2–19 Luster MI, Pait DG, Portier C, Rosenthal GJ, Germolec DR, Comment CE, Munson AE, White K, Pollock P (1992) Qualitative and quantitative experimental models to aid in risk assessment for immunotoxicology. Toxicol Lett 64–65:71–78 Luster MI, Portier C, Pait DG, White KL, Gennings C, Munson AE, Rosenthal GJ (1992) Risk assessment in immunotoxicology. I. Sensitivity and predictability of immune tests. Fundam Appl Toxicol 18(2):200–210 Luster MI, Portier C, Pait DG, Rosenthal GJ, Germolec DR, Corsini E, Blalock BL, Pollock P, Kouchi Y, Craig W, White KL, Munson AE, Comment CE (1993) Risk assessment in immunotoxicology. II. Relationships between immune and host resistance tests. Fundam Appl Toxicol 21(1):71–82 Luster MI, Portier C, Pait DG, Rosenthal GJ, Germolec DR (1995) Immunotoxicology and risk assessment, Chapter 5. In: Burleson GR,

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Dean JH, Munson AE (eds) Methods in immunotoxicology, vol 1. Wiley, New York, pp 51–68 Germolec DR (2004) Sensitivity and predictivity in immunotoxicity testing: immune endpoints and disease resistance. Toxicol Lett 149:109–114 Burleson GR (2000) Models of respiratory immunotoxicology and host resistance. Immunopharmacology 48:315–318 Burleson GR, Burleson FG (2007) Testing human biologicals in animal host resistance models. J Immunotoxicol 5:1–9 Burleson GR, Burleson FG (2007) Influenza virus host resistance model. Methods 41: 31–37 Burleson GR, Burleson FG (2008) In: Herzyk DJ, Bussiere JL (eds), Immunotoxicology strategies for pharmaceutical safety assessment. Wiley, Hoboken, NJ, pp 163–177, Chapter 5.1 Burleson GR (2008) MCMV host resistance model to detect latent viral reactivation immunotoxicity. Int J Toxicol 27(6):417 Herzyk DJ, Gore ER, Polsky R, Nadwodny KL, Maier CC, Liu S, Hart TK, Harmsen AG, Bugelski PJ (2001) Immunomodulatory effects of anti-CD4 antibody in host resistance against infections and tumors in human CD4 transgenic mice. Infect Immun 69(2): 1032–1043 Luebke RW (1995) Assessment of host resistance to infection with rodent malaria. In: Burleson GR, Dean JH, Munson AE (eds). Wiley, New York, pp 221–242, vol 2, Chapter 16 Van Loveren H, Luebke RW, Vos JG (1995) Assessment of immunotoxicity with the parasitic infection model Trichinella spiralis. In: Burleson GR, Dean JH, Munson AE (eds), Wiley, New York, pp 243–271, vol 2, Chapter 17 McCay JA (1995) Syngeneic tumor cell models: B16F10 and PYB6. In: Burleson GR, Dean JH, Munson, AE (eds), Methods in immunotoxicology, vol 2. Wiley, New York, pp 143–157, Chapter 11 Zhu Y, Herlaar E, Masuda ES, Burleson GR, Nelson AJ, Grossbard EB, Clemens GR (2007) Immunotoxicity assessment for the novel spleen tyrosine kinase inhibitor R406. Toxicol Appl Pharmacol 221:268–277 Steele TD, Geng W, Burleson F, Burleson G (2005) Enfuvirtide does not impair host resistance to infection in rats. Toxicologist 84:178 Gilmour MI, Park P, Selgrade MK (1993) Ozone-enhanced pulmonary infection with Streptococcus zooepidemicus in mice. Am Rev Respir Dis 147:753–760

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23. Gilmour MI, Selgrade MK (1993) A comparison of the pulmonary defenses against streptococcal infection in rats and mice following O3 exposure: differences in disease susceptibility and neutrophil recruitment. Toxicol Appl Pharmacol 123:211–218 24. Komocsar W, Burleson G, Wierda D (2007) The optimization of an acute rat model to evaluate effects on innate immunity induced by anti-inflammatory agents. Toxicologist 96:357 25. Van der Poll T, Keogh CV, Buurman WA, Lowry SF (1997) Passive immunization against tumor necrosis factor-a impairs host defense during pneumococcal pneumonia in mice. Am J Crit Care Med 155:603–608 26. Takashima K, Tateda K, Matsumoto T, Iizawa Y, Nakao M, Yamaguchi K (1997) Role of tumor necrosis factor alpha in pathogenesis of pneumococcal pneumonia in mice. Infect Immun 65:257–260 27. Benton KA, VanCott JL, Briles DE (1998) Role of tumor necrosis factor alpha in the host response of mice to bacteremia caused by pneumolysin-deficient Streptococcus pneumoniae. Infect Immun 66(2):839–842 28. O’Brien DP, Briles DE, Szalai AJ, Tu A-H, Sanz I, Nahm MH (1999) Tumor necrosis factor alpha I is important for survival from Streptococcus pneumoniae infections. Infect Immun 67(2):595–601 29. Gosselin D, DeSanctis J, Boule M, Skamene E, Matouk C, Radzioch D (1995) Role of tumor necrosis factor alpha in innate resistance to mouse pulmonary infection with Pseudomonas aeruginosa. Infect Immun 63(9):3272–3273 30. Mohan VP, Scanga CA, Keming Y, Scott HM, Tanaka KR, Tsang E, Tsai MC, Flynn JI, Chann J (2001) Effects of tumor necrosis factor alpha on host immune response in chronic persistent tuberculosis: possible role of limiting pathology. Infect Immun 69(3):1847–1855 31. Mond JJ, Lees A, Snapper CM (1995) T cellindependent antigens type 2. Annu Rev Immunol 13:655–692 32. Amlot PL, Grennan D, Humphrey JH (1985) Splenic dependence of the antibody response to thymus-independent (TI-2) antigens. Eur J Immunol 15:508–512 33. Harms G, Hardonk MJ, Timens W (1996) In vitro complement-dependent binding and in vivo kinetics of pneumococcal polysaccharide TI-2 antigens in the rat spleen marginal zone and follicle. Infect Immun 64:4220–4225 34. Guinamard R, Okigaki M, Schlessinger J, Ravetch JV (2000) Absence of marginal zone B cells in Pyk-2-deficient mice defines their role in the humoral response. Nat Immunol 1:31–36

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35. Burleson FG (2008) Systemic Streptococcus pneumoniae host resistance model to evaluate marginal zone B (MZB) cell immunotoxicity. Int J Toxicol 27(6):416 36. Conlan JW, North RJ (1992) Monoclonal antibody NIMP-R10 directed against the CD11b chain of the type 3 complement receptor can substitute for monoclonal antibody 5C6 to exacerbate listeriosis by preventing the focusing of myelomonocytic cells at infectious foci in the liver. J Leukocyte Biol 52(1):130–132

37. Rosen H, Gordon S, North RJ (1989) Exacerbation of murine listeriosis by a monoclonal antibody specific for the type 3 complement receptor of myelomonocytic cells. Absence of monocytes at infective foci allows Listeria to multiply in nonphagocytic cells. J Exp Med 170(1):27–37 38. Conlan JW, North RJ (1991) Neutrophilmediated dissolution of infected host cells as a defense strategy against a facultative intracellular bacterium. J Exp Med 174(3): 741–744

Chapter 8 Viral Host Resistance Studies Wendy Jo Freebern Abstract A foremost objective of preclinical immunotoxicity testing is to address whether or not a drug or environmental toxicant causes adverse effects on net immune health, expressly the host’s ability to mount an appropriate immune response to clear infectious organisms. Given the complex interactions, diverse molecular signaling events, and redundancies of immunity that has itself been subdivided into interdependent arms, namely innate, adaptive, and humoral, the results of single immune parameter testing may not reflect the final outcome of a drug or toxicant’s effect on net immune health. The most comprehensive experimental approach to ascertain this information is utilization of host resistance models. Herein, application of viral host resistance models in rodents and non-human primates is described. Although brief descriptions of numerous viral models are discussed including reovirus, EpsteinBarr virus, cytomegalovirus, and lymphocryptovirus, the most well-characterized viral host resistance model, rodent influenza, is emphasized. Key words: Immunotoxicology, Host resistance, Viral, Influenza, Latent viral models, Viral clearance, Non-human primate, Rodent, Immunosuppression, Gastrointestinal viral model

1. Introduction This chapter focuses on the use of viral host resistance models to assess the effects of a drug or environmental toxicant on host net immune status. As discussed in other chapters of this book, there are numerous assessments (e.g., CTL, NK, respiratory burst, phagocytosis, and T-cell dependent antibody response [TDAR]) that can be performed to evaluate the immunotoxic potential of a drug or environmental toxicant. However, does inhibition of one of these assessments, for example, TDAR to keyhole limpet hemocyanin, in a rat model provide enough evidence for labeling a drug or potential environmental toxicant as an immunotoxicant? How much inhibition of TDAR correlates with a host being immunocompromised? The immune system and its complex R.R. Dietert (ed.), Immunotoxicity Testing: Methods and Protocols, Methods in Molecular Biology, vol. 598 DOI 10.1007/978-1-60761-401-2_8, © Humana Press, a part of Springer Science + Business Media, LLC 2010

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networking of cell-to-cell communications and molecular signaling events does not allow for easy answers to these questions. The aforementioned assays provide useful, often mechanistic, information; however, they do not address the ability of a host to mount a productive immune response to an invading microbe. Perhaps, the assumption that net immune status is compromised when a drug is associated with decreased numbers of lymphocyte counts or inhibition of CTL, NK, innate immune function, and/ or TDAR would be the most conservative conclusion for risk assessment, but often one or more immune functions can be affected without having an effect on a host’s ability to surmount infection. A host resistance study can provide more evidence of a drug’s effect on host net immune status, as clearance of an infectious reagent requires the integration of innate, adaptive, and humoral immunity (1–9). Factors that would support the use of a viral host resistance study include: targeting of a drug or class of toxicants that indicates inhibition of immune function and/or previous testing demonstrating inhibited TDAR, profound decrease in lymphocytes, or inhibited CTL or NK function. Once the decision is made to utilize a viral host resistance model, it is imperative that the investigator select the model system to best fit the overall evaluation. Rodent host resistance models are most commonly used for the investigation of drug or environmental toxicant effects on immunocompetence. However, non-human primate retroviral and herpesviridae models have demonstrated xenobiotic immunosuppressive-effects on viral proliferation and reactivation (10). Mouse models offer the most versatility for additional assessments (e.g., isolation of viral-specific CD8+ cell response (3)), but a rat model may be more appropriate if rat was used for previous toxicity testing or the drug achieves higher systemic exposures in rat. Other considerations for animal selection include sex and age. If, for example, a drug is indicated for the geriatric population, utilization of aged animals should be considered. Duration of the host resistance studies is dependent on the virus challenge and drug or environmental toxicant and/or timing of previously observed potential immunotoxicity. Although the foremost question set forth to be answered when using a viral host resistance model is clearance or viral proliferation, addition of assessments such as CTL, NK, phenotyping, and viral-specific antibody response can not only show consistency of previous findings (if particular endpoints were assessed in prior studies), but may also reveal a mechanistic basis for any potential aberrancies in host resistance. The following sections will discuss a variety of viral host resistance models with greatest emphasis on influenza host resistance as it is the most common viral model utilized and many of the general concepts of study design are similar in other viral host resistance models.

Viral Host Resistance Studies

2. Influenza: A Viral Host Resistance Model

Rodent influenza host resistance models are the most wellcharacterized host resistance models utilized in immunotoxicity testing. A typical study design for a rat influenza model is shown in Fig. 8.1. The dosing concentrations and duration should be based on what and when previous aberrancies in immune parameters were observed upon drug administration or environmental toxicant exposure. If unknown, the length of dosing phase should be justified by the molecular target of the drug, pharmacodynamics/pharmacokinetics, and/or expected length of human exposure. Addition of groups of “positive-control” animals and noninfected naive animals to the study is strongly recommended. Examples of immunosuppressants that have been administered to positive-control animals include dexamethasone, cyclophosphamide, and cyclosporin A. A decrease in viral clearance in the positive-control group ensures that the assay is capable of detecting immunosuppression. Noninfected naive animals are utilized as negative controls for viral titer assessment, specific-antibody response assays, and/or sentinel animal evaluation. Influenza infection is usually via intranasal administration and in rats a common viral challenge is 2 × 105 plaque forming units (PFU). Considerations for viral challenge include: strain of influenza and rodent, size of animals, and age of animals. If a new passage of virus is indicated for use or the animal assessment includes juvenile or aged animals (extremes of life span), or for example, a transgenic animal not previously used in an influenza host resistance study, an in vivo viral titration with the appropriate animal model is strongly recommended before performing a study introducing drug or toxicant into the model. For mortality studies, at least 75% survival of infected-control

2.1. General Study Design

First Dose

Necropsies

Necropsy

Necropsy

Dosing phase

Fig. 8.1. Schematic of influenza host resistance study design.

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animals is advisable. If too many animals die, then interpretation of drug effect on total immunocompetance may be compromised due to acute mortality which most likely would occur prior to the complex integration of innate, adaptive, and humoral immunity. Clearance, not survival, has been shown to be a more robust endpoint (4). Early necropsies are necessary to calculate viral titers as well as to determine the drug effect on NK, neutrophil, and/or macrophage innate-immune function. Several necropsies are normally performed postinfection to appropriately determine viral clearance. Recommended necropsy days postinfection are shown in Fig. 8.1. Although viral clearance supports the ultimate determination of net immune health status, additional assessments should be considered and are discussed in a later section. If previous immunotoxicities have been demonstrated for a drug or environmental agent, it may be important to confirm that the immunotoxicity is observed prior to infection. 2.2. Viral Clearance Assessment

There are several methods utilized to determine viral titer described in the literature, but it is important that the method selected quantitatively measures the infectious virus. Figure 8.2 summarizes a virus plaque assay, a common method for quantitative assessment of infectious virus and viral clearance (4, 8).

2.3. Additional Assessments

Elicitation of interferons is generally the first hallmark of immune defense against viral infection and, in the intranasal influenza viral host resistance model, it can be easily measured in bronchoalveolar lavage or lung homogenate by ELISA (7, 8). Interferon peaks within the first 36h postinfection (7). Expression of other cytokines including IL-1a, IL-1B, IL-6, TNFa, and GM-CSF indicate a robust innate immune response and generally peak 2–3 days postinfection, although IL-6 remains elevated for approximately 1 week postinfection.

2.3.1. Elicitation of Cytokines

Homogenize lung Add serial dilutions of lung homogenate to monolayers of MDCK cells Cover cells with an agarose overlay Incubate for 36-48 hours Fix with buffered formalin and stain with crystal violet Count viral plaques and calculate viral titer

Fig. 8.2. Outline of viral clearance assessment methodology. MDCK Madin Darby Canine Kidney.

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In addition, G-CSF and M-CSF peak 4–6 days postinfection (4, 11). Multiplex technologies, including Luminex’s xMAP®, have increased the efficiency of evaluating expression of multiple cytokines in a small volume of lavage fluid or homogenate. 2.3.2. Natural Killer Cell Activity

Investigating the functional activities of innate immune cells that produce the aforementioned chemical mediators can also provide an insight into mechanism of potential drug effects on viral clearance. Natural killer cell (NK) cytotoxic activity peaks approximately 2 days postinfection. At a necropsy 2–3 days postinfection, NK activity can be measured in lung homogenate using a traditional Chromium release assay (4, 8) or a fluorometric flow cytometric-based method (12). Briefly, leukocytes from the lung homogenate are incubated with labeled (either Cr51 or fluorometric dye) target cells (an NK-sensitive cell-line such as YAC-1). After incubation, the release of Cr51 or increase in exclusion dye positive (propidium iodide, 7-AAD) target cells is quantitated.

2.3.3. Alveolar Macrophages

Alveolar macrophages are prevalent in the lung (approximately 90% of cells obtained by bronchoalveolar lavage) and are instrumental in the clearance of airborne microbes as well as environmental toxicants. Not only are the macrophages important in eliminating virus by phagocytosis of infected cells, but are also important in antigen presentation for specific arms of immunity. The least complex procedures for evaluating alveolar macrophage function in an influenza host resistance study include cytokine expression analyses and histological assessment (evidence of phagocytosis function) of lung tissue. Ex vivo phagocytosis and respiratory burst functional assays, either flow cytometric or fluorometric plate-based methods, can be performed on bronchoalveolar lavage to further assess macrophage function.

2.3.4. Cytotoxic T-Lymphocyte Activation

Testing of cytotoxic T-lymphocyte (CTL) activation provides information concerning potential drug effects on adaptive immunity following viral challenge. CTL response to influenza is dependent on viral antigen presentation in the context of major histocompatibility class I (MHC I) to CD8+ T-cells and peaks between 4 and 9 days postinfection (4, 8, 13, 14). Activation of CTL specific for viral antigen can be tested ex vivo by measuring the cytotoxic function of CTL on target cells 4–8 days postinfluenza infection; the labeled target cells must present viral antigen in the context of MHC I to demonstrate specificity of the response (4, 8). CTL response is described in greater detail in a later chapter. The population size of influenza-specific CD8+ cells, which correlates with the necessary expansion for an appropriate adaptive immune response, can also be evaluated in a mouse influenza model utilizing the tetramers of MHC I containing viral peptides in flow cytometric immunostaining protocols (3, 15, 16).

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2.3.5. Influenza Specific T-Cell Dependent Antibody Response

Last for discussion, but certainly not least, is the investigation of influenza-specific TDAR. TDAR assessments as discussed in Chapter 12, provide information on antigen presentation, T-cell activation and B-cell function. Influenza-specific immunoglobulins can be simply detected via an ELISA-based method. In general, influenza-specific IgM peaks 4–6 days postinfection and IgG 14–21 days postinfection. Further evaluation of influenza-specific IgG subtypes can provide information on drug effects on TH1 and TH2 responses (17, 18).

3. Additional Viral Models 3.1. Reovirus Gastrointestinal Rodent Model

A reoviral host resistance model tests the effects of drugs or environmental toxicants on gastrointestinal (GI) immune competence (19, 20). In a healthy animal, enteric reovirus infection is selflimiting with viral clearance from the GI tract within 7–14 days (21). Viral clearance or lack thereof is indicative of GI immune status in a reoviral host resistance model. After oral gavage with reovirus, viral titers can be determined in feces by virus plaque assays, thus multiple necropsy days are not necessary. Appropriate reovirus-specific IgA and IgG responses, as well as, cell-mediated activity and associated cytokine production have been shown to be important in reoviral clearance (22–24). Therefore, assessment of CTL activity, reovirus-specific IgA (peaks after 8 days postinfection) and IgG (peaks after 21 days postinfection) concentrations, and cytokine levels may provide useful information on a possible mechanism(s) for observed decreases in reoviral host resistance.

3.2. Latent Viral Rodent Models

Latent viral rodent models provide a useful tool to investigate the effects of drugs or environmental toxicants on the reactivation of latent viruses. Immunosuppression can potentially lead to reactivation of latent viruses with numerous possible sequelae including lymphoproliferative disorders and solid tumors. Epstein-Barr virus (EBV) and cytomegalovirus (CMV), Herpesviridae family members, are two common viruses involved in immunotoxicity testing. Molecular biological advances have resulted in the increasing use of EBV host resistance models utilizing xenochimeric mice to predict drug or toxicant effects on the potential for EBV associated-lymphoproliferative disease. In these models, human B-cells harboring the EBV genome are introduced into the mouse and the incidence of lymphoproliferative disease, expressly B-cell lymphomas, can be evaluated by standard clinical and histologic pathology (25, 26). For investigation of drug effect on CMV reactivation, rodent CMV host resistance models have been well-characterized and, in general, mimic human CMV infection (6, 27–29). Mice or rats infected with mouse CMV (MCMV) or rat CMV (RCMV),

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respectively, present a primary infection with viral replication occurring in multiple organs. In a healthy animal, viral replication is halted by the complex integration of molecular signaling events and immune functions after which latency is established. Test article effect on immune competence can then be evaluated in these animals, as immunosuppression results in reactivation of the virus. Evaluation of T-cell and NK populations, their respective functions (see above sections), and expression of associated cytokines should be considered in the latent viral model as both cell types have been implicated in immunosurveillance (30–33). 3.3. Serendipitous MMTV Host Resistance Model

The mouse mammary tumor virus (MMTV) could be discussed in the previous section above since mice commonly harbor this virus yet never develop mammary tumors until either immunosuppressed through test articles, stress, or age. However, in reviewing the literature on viral host resistance models for immunotoxicity testing, MMTV is not often mentioned. When testing a drug or potential environmental toxicant in standard or investigative toxicology studies, if an increased incidence of mammary tumors occurs then presence of MMTV by either PCR or immunohistochemistry should be determined. Increased expression of MMTV may indicate potential immunotoxicity, thus its serendipitous nature as a host resistance model.

3.4. Non-human Primate Viral Host Resistance Models

The use of non-human primate host resistance models for immunotoxicity testing is not common, but has increased due to the rise in biologic therapeutic reagents that are not efficacious in rodent models and for which rodent orthologues are not available. Lymphocryptovirus (LCV) infection in monkeys models EBV infection and viral persistence in humans (34). Primary infection with LCV, like EBV in humans, manifests as an acute viremia followed by an asymptomatic persistence in healthy individuals. If the animal becomes immunosuppressed, viral replication and lymphomagenesis may ensue. Thus, LCV carriers are effective latent viral host resistance models. For immunotoxicity investigations, test article effects on immunosuppression in this model are demonstrated by presentation of LCV-induced lymphoproliferative disease. LCV-lymphoproliferative disease is diagnosed by standard histological evaluation followed by a screening for virus in lesions via in situ hybridization techniques and/or immunostaining with intermediate and late viral proteins. As with MMTV virus in mouse, LCV is common in monkeys and presentation of lymphoproliferative lesions in standard toxicity testing results in a serendipitous host resistance model indicating the immunotoxicity potential of a test article. Other opportunistic viral infections that are common in monkeys and, as with LCV, can indicate immunotoxicity include adenovirus, simian virus 40 (SV40), rhesus rhadinovirus (RRV), and CMV (35).

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Acknowledgments The author extends gratitude to Florence G. Burleson and Gary R. Burleson for their thoughtful review of this chapter and their scientific guidance in immunotoxicology. References 1. Burleson GR, Burleson FG (2008) Testing human biologicals in animal host resistance models. J Immunol 5:23–31 2. Shornick LP, Wells AG, Zhang Y, Patel AC, Huang G, Takami K, Sosa M, Shukla NA, Agapov E, Holtzman MJ (2008) Airway epithelial versus immune cell Stat 1 function for innate defense against respiratory viral infection. J Immunol 180(5):3319–3328 3. Head JL, Lawrence BP (2008) The aryl hyrdocarbon receptor is a modulator of antiviral immunity. Biochem Pharmacol 77(4): 642–653 4. Burleson GR, Burleson FG (2007) Influenza host resistance model. Methods 41(1):31–37 5. Vorerstrasse BA, Cundiff JA, Lawrence BP (2006) A dose-response study of the effects of TCDD on the immune response to influenza A virus. J Toxicol Environ Health A 69(6): 445–463 6. Scalzo A, Corbett AJ, Rawlinson WD, Scott GM, Degli-Esposti MA (2007) The interplay between host and viral factors in shaping the outcome of cytomegalvirus infection. Immunol Cell Biol 85(1):46–54 7. Burleson GR (1996) Pulmonary immunocompetences and pulmonary immunotoxicology. In: Smialowicz R, Holsapple MP (eds) Experimental immunotoxicology. CRC, Boca Raton, FL, pp 113–135 8. Burleson GR (1995) Influenza virus host resistance model for assessment of immunotoxicity, immunostimulation, and antiviral compounds. In: Burleson GR, Dean JH, Munson AE (eds) Methods in immunotoxicology. Wiley, New York, pp 181–202 9. Burns LA, Bradley SG, White KL, McCay JA, Fuchs BA, Stern M, Brown RD, Musgrove DL, Holsapple MP, Luster MI et al (1994) Immunotoxicity of nitrobenzene in female B6C3F1 mice. Drug Chem Toxicol 17(3):271–315 10. Haustein SV, Kolterman AJ, Sundblad JJ, Fechner JH, Knechtle SJ (2008) Nonhuman primate infections after organ transplantation. ILAR J 49(2):209–219

11. Hennet T, Ziltener HJ, Frei K, Peterhans E (1992) A kinetic study of immune mediators in the lungs of mice infected with influenza A virus. J Immunol 149:932–939 12. Kim GG, Donnenberg, Donnenberg AD, Gooding W, Whiteside TL (2007) A novel multiparametric flow cytometry-based cytotoxicity assay simultaneously immunophenotypes effector cells; Comparisons to a 4 h 51 Cr-release assay. J Immunol Methods 325:51–66 13. Yap KL, Ada GL, McKenzie IFC (1978) Transfer of specific cytotoxic T lymphocytes protects mice inoculated with influenza virus. Nature 273:238–240 14. Wells MA, Albrecht P, Ennis FA (1981) Recovery from a viral respiratory infection. I. influenza pneumonia in normal and T-deficient mice. J Immunol 126:1036–1041 15. Mitchell KA, Lawrence BP (2003) Exposure to 2, 3, 7, 8-tetrachlorodibenzo-p-dioxin (TCCD) renders influenza virus-specific CD8+ T cells hyporesponsive to antigen. Toxicol Sci 74:74–84 16. Belz GT, Xie W, Doherty PC (2001) Diversity of epitope and cytokine profiles for primary and secondary influenza A virus specific CD8+ T cell responses. J Immunol 166(7): 4627–4633 17. Finkelman FD, Holmes J, Katona IM, Urban JF, Beckmann MP, Park LS, Schooley KA, Coffman RL, Mosmann TR, Paul WE (1990) Lymphokine control of in vivo immunoglobulin isotype selection. Annu Rev Immunol 8:303–333 18. Mutwiri G, Benjamin P, Soita H, Townsend H, Yost R, Roberts B, Andrianov AK, Babiuk LA (2007) Poly[di(sodium carboxylatoethylphenooxy) phosphazene] (PCEP) is a potent enhancer of mixed Th1/Th2 immune responses in mice immunized with influenza virus antigens. Vaccine 25(7):1204–1213 19. Cuff CF, Fulton JR, Barnett JB, Boyce CS (1998) Enteric reovirus infection as a probe to study immunotoxicity of the gastrointestinal tract. Toxicol Sci 42:99–108

Viral Host Resistance Studies 20. Maoxiang L, Cuff CF, Pestka J (2005) Modulation of murine host response to enteric reovirus infection by the trichothecene deoxynivalenol. Toxicol Sci 87(1):134–145 21. Barkon ML, Haller BL, Virgin HWIV (1996) Circulating immunoglobulin G can play a critical role in clearance of intestinal reovirus infection. J Virol 70:1109–1116 22. London SD, Rubin DH, Cebra JJ (1987) Gut mucosal immunization with reovirus serotype 1/L stimulates viral specific cytotoxic T cell precursors as well as IgA memory cells in Peyer’s patches. J Exp Med 165:830–847 23. Major AS, Cuff CF (1996) Effects of the route of infection on immunoglobulin G subclasses and specificity of the reovirus-specific humoral immune response. J Virol 70:5968–5974 24. Silvey KJ, Hutchings AB, Vajdy M, Petzke MM, Neutra MR (2001) Role of immunoglobulin A in protection against reovirus entry into murine Peyer’s patches. J Virol 75:10870–10879 25. Fuzzati-Armentero MT, Duchosal MA (1998) hu-PBL-SCID mice: and in vivo model of Epstein-Barr virus-dependent lymphoproliferative disease. Histol Histopathol 13(1): 155–168 26. Yajima M, Imadome K, Nakagawa A, Watanabe S, Terashima K, Nakamura H, Ito M, Shimizu N, Honda M, Yamamoto N, Fujiwara S (2008) A new humanized mouse model of Epstein-Barr virus infection that reproduces persistent infection, lymphoproliferative disorder, and cell-mediated and humoral immune responses. J Infect Dis 198(5):673–682 27. Selgrade MJK, Daniels MJ (1995) Host resistance models: murine cytomegalovirus. In: Burleson GR, Dean JH, Munson AE (eds) Methods in immunotoxicology. Wiley, New York, pp 203–219 28. Garssen J, van der Vliet H, De Klerk A, Goettsch W, Dormans JA, Bruggeman CA,

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Osterhaus AD, van Loveren H (1995) A rat cytomegalovirus infection model as a tool for immunotoxicity testing. Eur J Pharmacol 292(3–4):223–231 Ross PS, de Swart RL, van der Vliet H, Willemsen L, De Klerk A, van Amerongen G, Groen J, Brouwer A, Schipholt I, Morse DC, van Loveren H (1997) Impaired cellular immune response in rats exposed perinatally to Baltic Sea herring oil or 2, 3, 7, 8-TCDD. Arch Toxicol 71(9):563–574 Polic B, Hengel H, Krmpotic A, Trgovcich J, Pavic I, Lucin P, Jonjic S, Koszinowski UH (1998) Hierarchical and redundant lymphocyte subset control precludes cytomegalovirus replication during latent infection. J Exp Med 188:1047–1054 Suvas S, Azkur AK, Rouse BT (2006) Qa-1b and CD94-NKG2a interaction regulate cytolytic activity of herpes simplex virus-specific memory CD8+ T cells in latently infected trigeminal ganglia. J Immunol 176(3):1703–1711 Pappworth IY, Wang EC, Rowe M (2007) The switch from latent to productive infection in Epstein-Barr virus-infected B cells is associated with sensitization to NK cell killing. J Virol 81(2):474–482 Stowig T, Brilot F, Arrey F, Bougras G, Thomas D, Muller WA, Munz C (2008) Tonsilar NK cells restrict B cell transformation by the Epstein-Barr virus via INF-gamma. PLoS Pathog 4(2):e27 Rivailler P, Carville A, Kaur A, Rao P, Quink C, Kutok JL, Westmoreland S, Klumpp S, Simon M, Aster JC, Wang F (2004) Experimental rhesus lymphocryptovirus infection in immunosuppressed macaques: an animal model for Epstein-Barr virus phathogenesis in the immunosuppressed host. Blood 104(5):1482–1489 Sasseville VG, Diters RW (2008) Impact of infections and normal flora in nonhuman primates on drug development. ILAR J 49(2): 179–190

Chapter 9 Parasite Challenge as Host Resistance Models for Immunotoxicity Testing Robert W. Luebke Abstract Identification of potentially immunosuppressive compounds typically involves assessing a combination of observational endpoints as surrogates for functional endpoints and functional endpoints as surrogates for resistance to infectious or neoplastic disease. Host resistance assays are considered to be the “gold standard” against which suppression of immune function at the molecular or cellular level can be judged, because resistance to infection, regardless of the actual pathogen, involves multiple pathways of effector function to neutralize or eliminate pathogens. Resistance to infection with the parasitic nematode Trichinella spiralis has been used to assess immune function following exposure to a variety of immunotoxicants at the whole animal level. The various immunological mechanisms that are responsible for resistance to different phases of the life cycle are well documented, as are the effects of immunosuppression on the outcome of infection. This chapter describes methods to assess elimination of adult parasites from the small intestine, body burdens of larvae, as well as antibody responses and lymphocyte responses to parasite antigens Key words: Trichinella spiralis, Host resistance, Immunotoxicity, Immunosuppression, Susceptibility to infection, Parasite infections, Methods

1. Introduction 1.1. Background

Identification of potentially immunosuppressive compounds typically involves assessing a combination of observational endpoints as surrogates for functional endpoints and functional endpoints as surrogates for resistance to infectious or neoplastic disease. Host resistance assays are not generally considered suitable for screening purposes because of the costs associated with dedicated sets of animals, the additional safety measures required to work with pathogens, and reduced sensitivity when compared

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to functional assays (1). However, host resistance assays are considered to be the “gold standard” against which suppression of immune function at the molecular or cellular level can be judged because resistance to infection, regardless of the actual pathogen, involves multiple pathways of effector function to neutralize or eliminate pathogens. Challenge with an infectious agent or tumor cell line, chosen to exploit a suspected defect, can provide easily appreciated biological context for changes in observational or functional endpoints. This approach has generally been successful in linking changes in host resistance to suppressed immune function in animal models (2) and in humans (3). Although less straightforward, a lack of agreement between suppressed immune system endpoints and host resistance assay outcomes may suggest that alternative pathways of resistance exist that are not affected in exposed animals, thus implying that the identified functional defect is less likely to translate into an increased disease risk. For example, Keil et al. (4) reported adequate resistance to Listeria monocytogenes infection in mice exposed to doses of dexamethasone that suppressed cell mediated immune function that normally is central to clearing infection. In this case, dexamethasone treatment also increased the production of neutrophilic granulocytes, phagocytic leukocytes that also phagocytize the organism. The authors hypothesized that the increased numbers of neutrophils were sufficient to provide protection at all but the highest dose of dexamethasone. A variety of infectious agents, including viruses, bacteria, protozoans, and metazoans have been used in challenge assays to generate qualitative and quantitative data that reflect the overall competence of the host’s immune system. Challenge agents may be chosen because they are associated with significant human diseases, or because the mechanisms of resistance are well documented. Our current understanding of host resistance is based on decades of studies in animal models expressing inherited or induced immune system defects that have been linked to resistance or an increased susceptibility. Detailed discussions of these resistance mechanisms can be found in introductory immunology textbooks. Because specific immune system defects are linked to an increased susceptibility, challenge agents used in host resistance assays should be matched to the suspected immune system defect to avoid falsely concluding that exposure did not affect resistance to infection. This review is focused on the nematode parasite Trichinella spiralis (Tsp) infection model, although parasitologists and immunologists have characterized the host response to many nematode parasites of humans and animals. Thus, alternative nematode host resistance models could be developed using techniques described in the parasitology literature.

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Humans are susceptible to a number of nematode infections, including T. spiralis, Ascaris (large roundworms), pin worms, whip worms, and a variety of hookworms. Unless infected with large numbers of parasites, infection is not particularly life-threatening, although significant morbidity may occur with chronic or repeated infection. However, Strongyloides stercoralis, which causes only mild infection in immunocompetent individuals, can cause a life threatening hyperinfection syndrome in immunocompromised individuals; a deficient immune response to the infection allows larval parasites to re-infect the host by penetrating the intestinal mucosa rather than being eliminated from the host in feces. T. spiralis is primarily a mammalian parasite that is transmitted by the consumption of raw or undercooked meat from animals that harbor encapsulated T. spiralis muscle larvae. Transmission between animals occurs through predation or consumption of carrion containing viable Trichinella larvae. Modern farming practices in developed nations prohibit feeding uncooked meat scraps to commercially produced hogs, and reduce their access to potentially infected rodents. As a result, the incidence of Trichinella infection in developed countries declined dramatically starting in the last half of the twentieth century. Most outbreaks in the developed countries are now associated with consumption of improperly cooked game, including bear, walrus, and wild hogs (5, 6). In regions that employ less stringent farming practices, pork still poses an infection threat as does the consumption of horse meat if infected rodent carcasses contaminate feed. Given the relatively low incidence of infection risk, T. spiralis is not a significant public health risk in the West. However, the immune response to infection is well understood and resistance to the various life cycle stages requires the participation of innate, cellular, and humoral immunity (see below). In addition, the life cycle of the organism is “dead end” because no infectious stages of the parasite are released into the environment, thus greatly reducing the possibility of animalto-animal transmission in a laboratory setting. Resistance to T. spiralis infection has been used to evaluate the immunotoxicity of environmental contaminants, drugs, and radiation in mice and rats and, in the case of tributyltin oxide (TBTO), resistance data generated by Vos et al. (7) were used by the U.S. Environmental Protection Agency’s Integrated Risk Information System to set the reference dose for human TBTO exposure (http://www.epa.gov/iris/subst/0349.htm). This model has also been used to evaluate immunosuppression in animals exposed to diethylstilbestrol (8), 2,3,7,8-tetrachlorodibenzo-p-dioxin (9-11) ultraviolet light (12) and a food coloring additive (13). 1.2. Biology of the Parasite

Ingested larvae hatch in the stomach of the host, migrate to the small intestine and burrow into the mucosa where mating takes place. Female parasites release live larvae that are distributed via

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the blood and lymphatics to most tissues. Only those larvae that reach striated muscle are able to encyst and persist for years inside a protective capsule (“nurse cell”); the life cycle is completed when encysted larvae are consumed in raw or undercooked muscle. An intense inflammatory response is initiated in response to larvae that reach the brain and heart, and it is this phase of the infection that causes the greatest host morbidity and mortality. In rats and mice, fecundity of female parasites decline over the course of infection, and adult parasites are expelled from the intestine after 12–14 days. Migration of the first stage larvae to striated muscle and formation of cysts is complete after approximately 30 days. 1.3. Resistance to Infection

The mechanisms of resistance to T. spiralis infection have been the subject of study for many years. Several detailed reviews of the host response to infection have been published (14-16). As noted above, morbidity is ultimately influenced by the number of migrating newborn larvae. Thus, effective immunity is characterized by responses that limit female parasite fecundity, damage or destroy the newborn larvae and mediate the elimination of adult parasites from the gut. Inflammation of the bowel is first evident about 6 days after a primary infection in rats and mice and is virtually absent in congenitally athymic animals (17, 18). In contrast, expulsion of a primary infection is not altered in animals which lack the ability to produce antibody (19). Studies in rodents with targeted gene disruptions or following exogenous cytokine administration have shown that infection stimulates a strong T-helper (Th2) cell response; additional research established that the expulsion of worms is initiated by interleukin-4 (IL-4) and IL-13 activation of the transcription factor signal transducer and activator of transcription 6 (STAT6) via IL-4 receptor a ligation (20). Signaling via STAT6 increases IL-4 and IL-13 production and thus the induction of intestinal mastocytosis, which has a central role in adult parasite elimination (20). Mast cell degranulation increases the permeability of intestinal epithelial cells, which is one of the ultimate effector mechanisms responsible for parasite expulsion (21). Signaling via STAT6 induces intestinal smooth muscle cell hypercontractility, which acts to propel adult parasites out of the intestine (22). Hypercontractility is under T cell control and does not occur in athymic mice or in animals with major histocompatibility class II or CD4+ cell deficiencies (23, 24). STAT6 activation and subsequent direct effects on the gut appears to be a common pathway for eliminating intestinal nematodes, because elimination of the nematode Nippostrongylus brasiliensis requires STAT6 signaling that is independent of STAT6 effects on lymphocytes and mast cells (20).

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As is true for gut tissue infected with adult T. spiralis, striated muscle tissue infected with muscle larvae shows an inflammatory response that is T-cell dependent. In athymic animals, inflammation around encysted larvae is very scarce when compared to what is observed in immunocompetent animals, and the number of encysted larvae is considerably higher. The increased burden of larvae in host muscle depends to a great extent on the failure of the infected, immunocompromised host to expel the adult worms from the gut (18). It has been clearly established that antibodies, particularly IgG, are an important component of the rapid expulsion (RE) response that quickly clears the infectious larvae during a second infection in rats (25). In addition to de novo development of RE in infected adults, RE is also transferred from dams to the suckling offspring (26). A role for IgA in this process has not been definitely identified, although Tsp-susceptible C3H mice fail to mount IgA responses to infection, whereas NIH mice, which are much more resistant, mount IgA responses to surface components of the parasite that are closely correlated with the expulsion of the worms (19). It has also been suggested that IgA may have a role in preventing reinfection by inhibiting penetration of the intestinal mucosa by infectious larvae (27). Both IgM and IgG antibodies may be involved in resistance as both classes of antibodies are formed during infection (18). IgG antibody also participates in an important form of systemic resistance to the migrating larvae, via antibody dependent cellular cytotoxicity (ADCC) effected by neutrophils, eosinophils, and monocytes. Evidence for this mechanism of killing was derived from in vitro studies, which established the fact that eosinophilic granulocytes adhere to and kill the antibody-coated newborn larvae (28). A similar response has been described using human cells (29). Furthermore, newborn larvae, injected into the peritoneal cavity or incubated with the blood of previously-infected rats are killed by adherent cells (30). However, isolated intestinal lamina propria cells from humans and rats that included an enriched eosinophil population, although very slow (days) to kill the newborn larvae, avidly bound newborn larvae, and prevented maturation of larvae when transferred back into a naïve host (31). Finally, IgE is thought to play a pivotal role in the immune response to T. spiralis (32). Binding of T. spiralis-specific IgE to heavy chain receptors on mast cells induce granule release after cross-linking with the specific antigens. The granules contain eosinophil chemotactic factors that recruit more eosinophils and enhance eosinophilic cytotoxic activity. Furthermore, treatment with antibody to IgE diminished eosinophil involvement in inflammatory responses around the encapsulated larvae in striated muscle, as well as increased numbers of muscle larvae (33).

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1.4. Trichinella spiralis Infection as a Host Resistance Model

The inflammatory response generated in response to circulating newborn larvae causes the greatest damage to host tissues. Limiting the body burden of encysted larvae provides the greatest protection to the host and is the ultimate indicator of an effective response to infection. Effective immunity to this phase of infection is evaluated by estimating either the total body burden of larvae or the number of larvae per gram of muscle tissue. Multiple immune and non-immune effector mechanisms limit the number of larvae that survive the migration to host tissues. As such, evaluating the rate of parasite expulsion, female parasite fecundity, and parasite-specific antibody titers provide insight into the underlying cause of increased larvae burdens in animals exposed to a test article, as well as a “severity index” of suppression observed in traditional tests of cellular, humoral, and innate immune function. When used to evaluate host immunocompetence, experimental groups are typically comprised of an untreated control and two or three doses of the test article. A positive control group of animals treated with a known immunosuppressive drug (e.g., dexamethasone or cyclophosphamide) may also be included. Parasite expulsion is evaluated by isolating the adult parasites from the small intestine, typically 14 days after infection, by which time control animals will have eliminated most or all adults. Female parasite fecundity is assessed by isolating females from the small intestine prior to expulsion, typically on day 9 or 10 after the animals are infected. Muscle burdens of encysted larvae are determined by digesting skeletal muscle or the tongue (a preferred site of encystment) 30 days or more after infection, after the completion of larvae migration. Blood for the determination of specific antibody titers can be obtained at the time of sacrifice for any of the above procedures. In addition, ex vivo evaluation of antigen-specific lymphocyte proliferation may also be done with single cell suspensions of lymphocytes that have been isolated from the spleen or mesenteric lymph nodes when animals are killed for adult or larvae parasite counts. Effects of chemical exposure on acquisition of immunity to infection can be assessed by a second round of infection, with or without continued chemical exposure, and evaluation of expulsion, larvae counts and female parasite fecundity.

1.5. Outline of Major Procedures

A stock of infected animals is kept as a source of infectious larvae. Infection is initiated by recovering viable larvae from the muscle of stock animals, adjusting the larvae to set concentration, and administering the larvae to experimental animals by gavage. Adult parasites are recovered from the small intestine to assess parasite expulsion and larvae are isolated from muscle tissue to determine the body burden. Alternatively, larvae burdens may be assessed by counting numbers of larvae in stained sections of muscle tissue.

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Gravid female parasites may be isolated from the small intestine over the course of infection to determine parasite fecundity. Parasite-specific responses may be evaluated at the cellular level by culturing lymphocytes from infected animals with an extract of infectious larvae; the same extract is useful for evaluating the humoral response to infection by measuring antibody titers, using standard ELISA techniques.

2. Materials Warning: T. spiralis is a human pathogen. The US National Institutes of Health group the organism as a Class 2 pathogen; that is, it is of ordinary potential hazard, capable of causing disease after accidental exposure (ingestion) but can be controlled under normal laboratory conditions. Safe handling of the organism depends on a level of skill equivalent to that of university departments of microbiology. Protective gloves must be worn when handling larvae and countertops should be wiped down with disinfectant after handling larvae or dissection of infected animals. Infected carcasses must be incinerated, and a suitable quantity of disinfectant should be added to all the left-over liquids containing viable larvae before disposal. Note that a temperature of −20°C will kill the isolated muscle larvae in about a week or less; as such, larvae stored in a normal lab freezer for later preparation of antigen are unlikely to pose an infection threat. Infected animals do not represent an infection hazard to cage mates or to other animals under normal conditions and thus may be kept in the same room as other experimental animals without the danger of spreading infection. 2.1. Isolation of Larvae

1. Scissors and forceps.

2.1.1. Equipment

2. Commercial duty blender with 250 mL stainless steel container. 3. Magnetic stirring plate and stirring bar. 4. Beakers (500 mL/mice, 2 L/rats). 5. 37°C Environmental room or incubator with electrical outlet. 6. #80 and #200 mesh stainless steel sieves (McMaster Carr, New Brunswick, NJ). 7. Squeeze bottle for saline.

2.1.2. Reagents

1. Digestion fluid: 1% (w/v) pepsin A (Sigma Chemical Co., St. Louis, MO), 1% (v/v)13 N HCl in warm (approximately 37°C) tap water (see Note 1). 2. 0.85% NaCl solution (saline, at 37°C).

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2.2. Infection of Experimental or Stock Animals

1. Magnetic stirrer.

2.2.1. Equipment

4. 1 mL Glass syringe.

2. Stirring bar. 3. 500-mL Beaker. 5. 18 g Curved oral gavage needle. 6. 37°C Water bath. 7. Plaque viewer (Bellco, Vineland, NJ) or dissection microscope.

2.2.2. Reagents and Supplies

1. Viable infectious Tsp larvae from subheading 9.2.2. 2. Nutrient broth/2% gelatin (both from Difco, Detroit, MI) (see Note 2). 3. 50 mL Screw-capped glass centrifuge tubes. 4. 15 mL Screw-capped culture tubes. 5. Glass microscope slides. 6. Disposable 1-mL pipette.

2.3. Adult Parasite Counts

1. Wire cage inserts for mice or rats.

2.3.1. Equipment

3. Acrylic plate 4 × 12 × 3/16.

2. Scissors and forceps. 4. Small iris scissors with one point blunted or #11 surgical blade. 5. Incubator at 37°C. 6. 500-mL beaker for disinfectant. 7. Large bore funnel. 8. 50 mL centrifuge tubes (disposable screw-capped polypropylene). 9. Centrifuge with carriers for 50 mL tubes. 10. Plaque viewer (Bellco, Vineland, NJ) or dissection microscope. 11. 0.5 mL tubes for freezing serum if collected. 12. Freezer for storing serum.

2.3.2. Reagents and Supplies

1. 0.85% NaCl solution (saline) containing 250 mg gentamicin/mL. 2. 0.85% NaCl solution (saline) without gentamicin. 3. Disinfectant. 4. 5 N NaOH. 5. Pasteur pipettes. 6. Petri dishes (100 mm, disposable plastic). 7. #10 surgical blades if serum is to be collected. 8. Serum separator tubes and centrifuge if serum is collected for antibody titers.

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9. 0.5 mL tubes for freezing serum. 10. Freezer for storing serum. 2.4. Larvae Counts

1. Electronic balance with 0.1 mg sensitivity.

2.4.1. Equipment

2. Disposable weighing boats, approximately 2.5 × 2.5 cm. 3. Stomacher, Model 80 (Tekmar, Fisher Scientific, Raleigh, NC). 4. 37°C rocking water bath or rocking platform in an enclosure at 37°C. 5. Centrifuge with carriers for 15 mL tubes. 6. Acrylic plates, 4 × 12 × 3/16 inches. 7. Plaque viewer (Bellco, Vineland, NJ) or dissection microscope. 8. Mechanical or electronic hand tally for counting larvae. 9. Serum separator tubes and centrifuge if serum is collected for antibody titers. 10. 0.5 mL tubes for freezing serum. 11. Freezer for storing serum.

2.4.2. Reagents and Supplies

1. 0.85% NaCl solution (saline) containing 250 mg gentamicin (Gibco)/mL. 2. Pepsin/HCl digestion fluid (see Subheading 9.2.1.2). 3. Bags for Stomacher (Tekmar, Fisher Scientific, Raleigh, NC). 4. Plastic Petri dishes. 5. #11 surgical blade, #10 surgical blade (if serum is to be collected). 6. 6 well tissue culture plates. 7. Pasteur pipettes or disposable tips for micropipeter. 8. Serum separator tubes and centrifuge if serum is collected for antibody titers. 9. 0.5 mL tubes for freezing serum. 10. Freezer for storing serum.

2.5. Parasite Fecundity

1. Wire cage inserts for mice or rats.

2.5.1. Equipment

2. Acrylic plate 4 × 12 × 3/16 inches. 3. 37°C incubator. 4. Small iris scissors with one blunt tip or #11 surgical blade. 5. Inverted microscope, 40× magnification. 6. Serum separator tubes and centrifuge if serum is collected for antibody titers. 7. 0.5 mL tubes for freezing serum. 8. Freezer for storing serum.

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2.5.2. Reagents and Supplies

1. Saline containing 250 mg/mL gentamicin. 2. Plastic Petri dishes. 3. 50 mL plastic centrifuge tubes with cap. 4. 6 well tissue culture plates. 5. 96 well tissue culture plates. 6. RPMI 1640 medium. 7. Fetal bovine serum. 8. Gentamicin. 9. #10 surgical blade (if serum is to be collected). 10. Serum separator tubes (if serum is to be collected). 11. 0.5 mL capacity tubes (if serum is to be collected).

2.6. Preparation of Tsp Antigen (T. spiralis extract, TsE)

A crude saline extract of isolated muscle larvae is used in ELISA assays to determine class-specific antibody titers or to stimulate parasite antigen-specific lymphocyte proliferation.

2.6.1. Equipment

1. pH meter. 2. Sonifier with micro tip (e.g., Branson Sonic Power Co., Model W-350, Danbury CT). 3. Glass tissue homogenizer. 4. Magnetic stirrer and stirring bar. 5. Refrigerated centrifuge capable of 10,000 g. 6. Microscope capable of 400× magnification. 7. Ice containers.

2.6.2. Reagents and Supplies

1. Shaved ice. 2. Freshly isolated or frozen Tsp larvae. 3. Saline. 4. 10× Phosphate buffered saline: 76 g NaCl, 14.8 g Na2HPO4, 4.3 g KH2PO4 in 900 mL distilled or deionized water; adjust to pH 7.4, and bring final volume to 1 L. 5. 1× PBS with protease inhibitors (PBSpi): 370 mg iodoacetamide (final [10 mM]), 3.03 mg Na-p-tosyl-L-arginine methyl ester (final [40 mM]) and 2.95 mg Na-p-tosyl-Llysine chloromethy ketone (final [40 mM]) in 200 mL of 1:10 dilution (in distilled or deionized water) of 10× PBS. 6. 15 mL centrifuge tubes. 7. Microscope slides and 22-mm2 cover slips. 8. 50 mL Polypropylene conical tubes. 9. Pasteur Pipettes. 10. 10,000 molecular weight cut-off dialysis tubing (Kirkegaard and Perry, Gaithersburg, MD).

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11. 45 mm pore size filter for syringe. 12. Sterile 1 mL O-ring freezing vials (Nunc). 13. Lowry protein assay kit. 2.7. Determination of IgM and IgG Antibody Titers to Trichinella Antigens

1. Pipettes (1, 5,10, 25 mL).

2.7.1. Equipment

5. pH meter.

2. Pipetman or equivalent (P20, P200, P1000). 3. 8-Channel pipette (various commercial sources). 4. Glass bottles for reagents (100 mL, 500 mL). 6. Plate washer (various commercial sources) or squeeze bottle. 7. Spectrophotometer.

2.7.2. Reagents and Supplies

1. 96 well ELISA plates (Costar, Corning, NY). 2. Paper towels or equivalent. 3. Reusable plastic troughs for loading multi-channel pipette. 4. Pipette tips (200 mL). 5. Plastic tubes (12×75, 75×100, 15 mL conical, 25 mL conical). 6. Wash Solution: H2O + 0.05% Tween 20. 7. Dilution/Blocking Buffer: PBS (see Subheading 9.2.6) containing 0.5% (w/v) bovine serum albumin and 0.05% (w/v) Tween 20 pH 7. 8. Coating Buffer: 0.1 M Na2CO3 (sodium carbonate) pH 9.6. 9. Stop solution: 5% EDTA (disodium salt) in water. 10. T. spiralis extract (TsE) prepared from muscle larvae. 11. Pooled normal rat serum (NRS, from non-infected animals). 12. Anti-Rat IgG–biotin labeled (Kirkegaard and Perry, 16-16-02). 13. Anti-Rat IgM–biotin labeled (Kirkegaard and Perry, 16-16-03). 14. Streptavidin (SAV), phosphatase labeled (Kirkegaard and Perry, 15-30-00). 15. Phosphatase substrate system for ELISA (p-nitrophenylphosphate tablets and diethanolamine buffer) (Kirkegaard and Perry, 14-30-00).

3. Methods 3.1. Maintenance of Stock Larvae Donors

Because freezing kills the infectious larvae, they cannot be frozen for storage. Instead, maintain a stock of larvae donors, usually in rats. 1. Pretreat donors with cyclophosphamide (20 mg/kg/d for rats, 80–100 mg/kg/d for mice, injected i.p., in sterile saline for injection, USP) for the 4 days preceding infection to

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suppress resistance to infection to increase the number of encysted larvae. Note that cyclophosphamide is extremely immunosuppressive, suppresses bone marrow function, and is a likely human carcinogen. Wear gloves and a mask when handling this drug. 2. Infect donor rats with approximately 2,500 larvae and 1,000 larvae for mice. Donors can be kept for up to one year, although it is preferable to use donors infected for not more than 6–8 months. 3.2. Isolation of Infectious Larvae

1. Euthanize donors by cervical dislocation or CO2 asphyxiation followed by cervical dislocation. 2. Skin, decapitate and eviscerate the carcass. Remove the feet and tail as well. 3. Remove the tongue and include in the digestion step. 4. Cut mouse carcasses into ten equal pieces. 5. Remove muscle tissue from the legs and backs of rat donors with scissors. Combine with the diaphragm and rib cage for processing. 6. Place the mouse carcass pieces or rat tissues in the blender cup with approximately 100 mL of digestion fluid. Cap securely and cycle the blender on and off for about 5 s/cycle (to avoid overheating). Repeat this step until the contents are reduced to small pieces (approximately ten cycles). 7. For mice, pour the fluid into a 500-mL beaker and rinse the blender cup with another 100 mL of digestion fluid. For rats, pour the initial 100 mL volume into a 2-L beaker, then rinse the blender cup with 100 mL of fluid and add another 800 mL (approximately) to the beaker. 8. Place the beaker on a magnetic stirrer located in an environmental room or incubator at 37°C. Stir at moderate speed until soft tissue is digested (approximately 2 h). 9. At the end of the incubation period, isolate the larvae by pouring the digestion fluid through a #80 stainless steel mesh screen to trap the undigested material, which is stacked on a #200 mesh to trap the larvae. 10. Invert the #200 mesh screen over a beaker and backwash larvae off the screen with a stream of 37°C saline from a squeeze bottle. 11. Transfer the larvae in saline into a 50 mL conical glass centrifuge tube. Allow to settle for 15 min at 37°C. 12. Using a clean Pasteur pipette, transfer the larvae into another 50-mL tube containing approximately 25 mL of saline and allow to settle for 15 min at 37°C. These transfers act as a

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washing step. Slowly add the larvae to the top of the saline column and allow them to settle completely to the bottom of the tube. 13. After the second wash, immediately use the larvae designated for infection. 3.3. Infection of Experimental or Stock Animals

1. Put one 30 mL and two 5 mL tubes of nutrient broth/gelatin in a 37°C water bath at the outset of the digestion step to insure that it will be liquefied when needed. 2. Add a portion of the isolated larvae to a 50 mL centrifuge tube containing 30 mL of nutrient broth/gelatin. Add 150 mL of settled larvae for infecting mice or 300 mL settled larvae for infecting rats. 3. Maintain the nutrient broth/gelatin at 37°C to prevent gelling. Keep the tube in the water bath as much as possible. 4. Suspend the larvae by gently inverting the tube approximately ten times (to prevent bubbles from forming, do not shake the tube). 5. Fill a 1 mL glass syringe fitted with a 1.5″ × 18 g oral gavage needle with larvae suspension and empty three times to make certain that all air has been excluded from the syringe. Glass syringes are preferable to disposable plastic syringes because the former provides greater precision in dispensing small volumes. 6. After the third filling, draw the larvae suspension into the syringe and expel all but 50 mL. Transfer the suspension to a microscope slide or Petri dish and count the larvae using the 17.5× magnification setting on a plaque viewer or dissecting microscope. Repeat this procedure until three successive counts of larvae are within 10% of each other. 7. Add or remove (after unit gravity sedimentation) larvae as needed to obtain the desired concentration of larvae. For experimental purposes, infect rats with 1,000 ± 100 larvae in 0.5 mL. Infect mice with 200 ± 20 larvae in 0.2 mL. For rats, adjust the concentration of larvae to 2,000/mL and for mice, adjust to 1,000/mL. 8. Repeat the counting procedure after half of the animals have been infected and adjust the concentration of larvae if required. Count the larvae suspension again at the end of the infection procedure to ensure that the animals were infected with a similar number of larvae. 9. When infecting animals, take care to guarantee that all larvae are deposited in the esophagus. 10. Infect one animal from group #1, one from group #2…. group #n until one animal from each group has been infected.

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After every four animals, remix the larvae by inverting the tube 5–6 times. 11. Reverse the order of infection, beginning with the last group infected and working backwards to group #1. By following this infection order, any effects of slight animals distress or changes in larvae viability or concentration will be evenly distributed among all the animal groups. 12. After a single cycle of infection (i.e., group 1…n, n…1) or a maximum of eight animals, rinse the needle and syringe with distilled water by filling and emptying it 4–5 times. 13. Groups of 6–8 inbred mice or rats are usually sufficient per treatment group for host resistance studies, although larger numbers of outbred animals may be required due to greater variability of results between animals of the same group. Stock animals only: Infect stock rats with 2,500 larvae in 0.5 mL and stock mice with 1,000 larvae in 0.2 mL. 14. Larvae not used to infect animals should be frozen for preparation of larvae extract for use in ELISA and lymphocyte proliferation assays (Subheading 9.3.8 and 9.3.9). 15. Allow the unused larvae to settle to the bottom of a 15 mL conical test tube. Transfer to a Nunc tube using a Pasteur pipette. 16. Wash larvae that had been suspended in nutrient broth/ gelatin by sedimentation through three changes of saline. 17. Store larvae frozen at −20°C. 3.4. Adult Parasite Counts

1. Starve the animals overnight in a cage containing a wire insert without bedding or other chewable material to help clear debris from the intestine. 2. The following morning, sacrifice the animals. Remove the small intestine and place it on an acrylic plate. (Note: If animals are to be bled for antibody titers, they should be anesthetized and bled as described in subheading 9.3.5 before removing the intestine, to prevent the blood from clotting before sampling). 3. Divide the rat intestine into anterior and posterior pieces of roughly equal length to facilitate handling. Process each section separately. 4. Open the intestine lengthwise using a #11 surgical blade or small iris scissors that have had the point of one blade blunted to reduce the likelihood of puncturing the intestine while slitting. 5. Rinse the opened intestine gently under running tap water and cut into piece 3–5 cm in length.

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6. Place the pieces in a Petri dish containing 30 mL of 0.85% NaCl + 250 mg/mL gentamicin and incubate for 4 h at 37°C in 5% CO2. 7. After incubation, grasp the pieces of intestine with forceps, agitate briefly in the incubation medium and then discard into a beaker of disinfectant. 8. Decant the saline/gentamicin into a 50 mL screw-capped plastic centrifuge tube using a large bore funnel. With the funnel still in place, rinse the plate with 10–15 mL of salinegentamicin into the 50 mL tube. (If necessary, cap and refrigerate the tubes overnight at this point). 9. Cap the tubes, centrifuge at 250×g for 5 min and remove all but 3 mL of the saline/gentamicin by aspiration. 10. To count the parasites, add approximately 0.5 mL of 5 N NaOH to the tube, mix well and transfer to an acrylic plate by making multiple streaks down the length of the plate using a Pasteur pipette. 11. Rinse the tube with approximately 1 mL of saline to recover any remaining parasites and add to the plate for counting. 12. Count the adults using a plaque viewer or dissection microscope at a magnification of 17.5×. In our experience, female B6C3F1 mice begin to expel adult parasites by day (d) 6 of infection; female C57BL/6J and F344 rats begin on d 7 or d 8. Control animals typically eliminate all but a few parasites by d 14 of infection. Note that rodents with a Th2 cytokine bias (e.g., BALB/c mice and Brown Norway rats) expel the parasites more rapidly. Because host strain will influence resistance to infection, plot studies should be done before infecting the experimental animals to determine expulsion kinetics. 13. Present results as the number of adult worms recovered. 14. Alternatively, infect an additional group of unexposed animals along with the experimental groups and sacrifice them after 5–6 days of infection to determine the number of parasites prior to the onset of expulsion. Calculate percentage of expulsion by dividing the number of parasites recovered on d 14 by the number recovered on d 5 or 6. 3.5. Collection of Blood for Antibody Titers

1. Anesthetize animals with CO2, isoflurane, or Nembutal. 2. Grasp mice by the nape of the neck and cut the vessels that travel along the side of the neck using a #10 blade. Take care not to transect the trachea. 3. Immediately place an opened serum separator tube under the neck to collect blood. Sacrifice mice by cervical dislocation after blood collection is completed.

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4. Open the peritoneal cavity of the anesthetized rats by inserting scissors through the top (ventral) portion of the diaphragm and cut the aorta. 5. Collect the blood from the thoracic cavity via a Pasteur pipette and transfer the sample to a serum separator tube. 6. Keep the samples on ice until all have been collected. 7. Centrifuge serum separator tubes according to the manufacturer’s instructions. 8. Collect serum with a clean Pasteur pipette and transfer to duplicate 0.5 mL tubes. 9. Store sample tubes in 50 mL polypropylene centrifuge tubes at −20°C that are capped and sealed with Parafilm. 3.6. Larvae Counts

1. Prepare digestion fluid as described in subheading 9.2.1. Each sample requires 12 mL. Calculate the volume to make by multiplying the number of samples by 12 mL. Round this value off to the next highest 50 mL. 2. Collect blood as described in subheading 9.3.5 if antibody titers will be determined. 3. After sacrificing the animal, remove and weigh the tongue to the nearest mg. Place the tongue in a weighing boat and macerate using two #11 surgical blades. 4. Transfer the macerated tissue into a Stomacher bag containing 6 mL of digestion fluid. Process for 45 s and transfer to a 20 mL glass scintillation vial using a 10-mL pipette. 5. Rinse the bag with an additional 6 mL of digestion fluid and add rinse fluid to the vial. 6. Tightly cap and Parafilm the vial and place in a suitable holder on a rocking platform or rocking water bath at 37°C for approximately 4 h or until there are no visible pieces of muscle tissue remaining. (Note: There are indigestible portions of the tongue. Thus, while all muscle tissue will be digested, some particulate matter will remain after digestion.) 7. Transfer the vial contents into a 15 mL centrifuge tube. Rinse the vial with 3 mL of saline and pellet the larvae by centrifugation (250–300×g for 5 min). 8. Aspirate the liquid from the tube being careful not to disturb the pelleted larvae. 9. Resuspend the pellet in 2–3 mL of saline (no gentamicin). 10. Add approximately 0.5 mL of 5 N NaOH to solubilize the precipitate. 11. Mix well with a Pasteur pipette and transfer the larvae suspension to an acrylic plate by making multiple streaks down the length of the plate.

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12. Rinse the tube with approximate 1 mL of saline and transfer to the counting plate. 13. Count larvae at 17.5 to 40× magnification using either an electronic or manual hand tally. Express results as larvae/g of tissue. (We typically recover approximately 3,000–5,000 larvae/g of tongue from rats infected with 1,000 larvae and 1,000–2,000 larvae/g from mice infected with 200 larvae). 3.7. Counts of Larvae in Tissue Sections

This is the procedure used by the Dutch National Institute for Public Health and Environment (RIVM), and was provided by Prof. Dr. Henk van Loveren. The method requires standard histopathology methods and morphometric analysis. 1. Remove the tongues and fix in 10% buffered formalin. Embed in paraffin. 2. Cut section 5 mm thick and stain by the periodic acid-Schiff method. 3. Count the number of muscle larvae in two sections using a morphometric analysis system (e.g., an eye piece or more advanced automated systems). Express results as the number of larvae per square millimeter. Using this method, approximately 10–50 larvae per squared millimeter of rat tongue are detected after infection with 1,000 larvae. Historic data are available from RIVM for mice.

3.8. Determination of Parasite Fecundity

1. Isolate the adult parasites as described in subheading 9.3.4. 2. Place sections of intestine in a Petri dish containing 30 mL of 0.85% NaCl plus 250 mg/mL gentamicin and incubate for 4 h at 37°C in 5% CO2. 3. After incubation, remove the pieces of intestine. 4. Place the Petri dish on the stage of a dissection microscope and collect the female worms that have migrated out of the intestine. Females are approximately 3.5 mm long vs. 1.5 mm long for males. 5. Rinse the isolated females (8–10 from each Petri dish) by placing them into one well of a 6 well tissue culture place containing 5 mL of saline/250 mg/mL gentamicin. 6. Transfer individual females to separate wells of a 96 well tissue culture plate containing 100 mL of RPMI 1640 media supplemented with 10% fetal calf serum plus 250 mg/mL gentamicin. Parasites should be taken up in the least possible volume of saline to avoid dilution of the culture medium. As an aid to counting, scratch a “cross-hair” on the bottom of each well prior to worm transfer. 7. Add RPMI 1640/10% FBS to all wells (including unused wells) to bring the final volume to approximately 200 mL.

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8. Incubate plates for 18 h at 37°C in 5% CO2. 9. Place the 96 well plate on the stage of an inverted microscope and count newborn larvae at 40× magnification. It is only necessary to count 3–4 wells each from the anterior and posterior bowel segments of rats although more complete results are obtained if 6–8 wells are counted for both portions of bowel. Data are expressed as the average number of newborn larvae per female parasite 3.9. Preparation of Trichinella Antigens (T. spiralis Extract, TsE)

1. Thaw the frozen larvae. 2. Suspend 2–3 mL of packed larvae in approximately 10 mL of PBS-pi in a 15 mL plastic centrifuge tube. 3. Disrupt larvae using one minute periods of sonication at a 50% duty cycle with a power setting of five. Allow approximately 30 s between cycles for cooling. Keep the tube of larvae in a small beaker of shaved ice during sonication to prevent overheating. 4. After seven cycles, remove a small volume of the suspension, place on a microscope slide, add a cover slip and observe at 40× magnification to determine how thoroughly the larvae have been disrupted. 5. Continue sonication until most of the larvae have been ruptured. 6. Transfer the larvae preparation to glass-to-glass tissue homogenizer set in shaved ice. Homogenize for ten up and down cycles. 7. Allow debris to settle for about one minute, pour off the supernatant into a 50 mL plastic centrifuge tube (on ice), and add an additional 5 mL PBS-pi to the pellet. Repeat the homogenization procedure for a total of 5 cycles of homogenization. 8. Add final homogenization liquid plus larvae debris to the 50 mL tube. Cap the tube and mix by end-over-end rotation for 30 min at 4°C. 9. Centrifuge the mixture for 60 min at 50,000×g, 4°C to remove debris from the suspension. 10. Rehydrate 10,000 mw cutoff dialysis tubing in PBS-pi according to package instructions. 11. Place 10 mL of supernatant into the dialysis tubing. Fill the tubing no more than 60%. Securely clamp each end of the tubing. 12. Dialyze 4°C against 200 volumes PBS-pi for approximately 48 h. Change PBS-pi twice daily, in the morning and the afternoon.

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13. After dialysis, collect TsE with a Pasteur pipette and pass through a 45 mM filter to sterilize. 14. Store 0.5 mL aliquots in sterile Nunc vials at −20°C. 15. After 24 h, thaw one vial and determine the protein concentration using a commercial Lowry protein assay kit. 3.10. Determination of IgM and IgG Antibody Titers to Trichinella antigens (see Note 3)

1. Coat microtiter plate wells overnight at 4°C by adding TsE in a volume of 100 mL. 2. The following morning, dump coating solution from the plates and wash three times with washing buffer using a plate washer or a squeeze bottle. After the final wash, tamp plates on a pile of paper towels to remove the excess wash solution. 3. Add 300 mL/well of warmed (37°C) blocking buffer to all wells. Cover plates with lids and incubate at 37°C for 1 h. 4. Dump blocking buffer from plates and tamp plates on paper towels to remove any remaining liquid. 5. Add 100 mL of dilution buffer to all except the first column of wells. 6. Add 200 mL of diluted experimental sera to wells A–G of column 1, and 200 mL of diluted NRS to well 1H. Use an 8-channel pipette to prepare serial 1:2 dilutions through column 10. Reserve columns 11 and 12 for blanks (all assay components except serum sample of noninfected rat serum). Take care not to contaminate the blank wells while doing the dilutions. 7. Cover the plates with lids and incubate at 37°C for 1 h. 8. Discard plate contents, wash three times as described above in step 2, and add 100 mL of the appropriate dilution of detection antibody to each well of the plates. 9. Incubate the plates at 37°C for 1 h, followed by three washes. 10. Add 100 mL of diluted SAV to each well and incubate at 37°C for 1 h followed by three washes. 11. Prepare the diethanolamine buffer with reagent quality water according to the manufacturer’s instructions. Allow 10 mL per plate. 12. Prepare substrate by adding d p-nitrophenyl-phospate tablets to the diethanolamine buffer just prior to use. 13. Remove excess liquid from the first plate by tamping on paper towels. 14. Add 100 mL of substrate to all wells.

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15. Incubate for 10 min then read the absorbance at 405 nm using an automated plate reader. Convert absorbance values to titer values using commercial software (e.g., Softmax or equivalent). 3.11. Determination of Total IgE Concentration

Detection of Tsp-specific IgE antibodies is difficult. As a result, we measure total IgE antibodies using paired monoclonal antibodies specific for IgE heavy chains in a capture assay. Antibodies and antibody pairs are available commercially (e.g., BD-Pharmingen, Zymed, and others) and typically include detailed instructions for reagent preparation and running the ELISA. Because the concentration of total IgE is typically very low in noninfected animals and increases dramatically in infected animals, differences in total IgE between the treated and control animals provide indirect evidence that chemical exposure attenuated the response.

3.12. Parasite-Specific Lymphoproliferative Responses

1. Lymphocytes that have been isolated from spleens or mesenteric lymph nodes of infected animals will undergo blastogenesis when cultured with the larvae extract (TsE, subheading 9.3.9) used to coat plates for the ELISA assay. 2. Standard techniques used to evaluate lymphocyte transformation in response to mitogens are suitable to evaluate the response. As such, a description of the method will not be presented here, although details of the method have been published (11). 3. Determine the optimal concentrations of TsE prior to use in an actual experiment using cell donors infected with the same number of larvae used to challenge animals for adult and larvae counts.

4. Notes 1. Prepare 200 mL of digestion fluid for each mouse carcass, or 1,000 mL for each rat carcass. Typically, 30 mice or rats can be infected with the larvae obtained from each donor. To prepare the digestion fluid, place a stirring bar in an appropriately sized beaker containing warm tap water and place on a magnetic stirrer. While stirring, gradually add the pepsin and acid. 2. Preparation of Nutrient broth/Gelatin. Add 2.8 g nutrient broth powder and 7 g of powdered gelatin to 350 mL distilled water. Add a stirring bar and heat gently with stirring until in solution. Dispense most of the solution as 30 mL into 50 mL screw-capped glass centrifuge tubes; a small portion should

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be stored as 5–7 mL volumes in 15 mL screw-capped culture tubes in case a small volume is needed. Cap all tubes loosely and autoclave for 15 min at 121°C, 15 psi. After cooling the tubes, tighten the caps and store at 4°C. 3. This assay has been optimized to detect T. spiralis specific IgG and IgM in rat serum. (This assay does not work for the detection of T. spiralis-specific IgA or IgE.) The assay is performed using standard ELISA techniques, and other combinations of detection antibodies and substrates may be used, although those listed in subheading 9.2.7 were found to be optimal in our lab. Concentrations of TsE used to coat plates, initial serum dilutions, and appropriate dilutions of heavy chain-specific antibodies should be optimized before analyzing samples from experiments. In our experience, 0.5 mg TsE/well works well for coating wells. A range of working dilutions for experimental serum samples can be determined by using the serum obtained from control animals infected for 7, 14, and 28 days, beginning at an initial dilution of 1:4 and continuing over 8–10 log2 dilutions. As a quality control measure, include one row of serum from noninfected animals, that has been diluted over the same log2 range as experimental samples, on each plate. The OD value in the “middle” of the noninfected serum curve (i.e., the midpoint between the lower and upper deflections of a 4-parameter curve) is calculated for each plate; the values should be similar for all plates. As an optional QA step, serum from infected control or nontreated animals can be pooled and stored in small aliquots and analyzed to establish titers of IgM, IgG and total IgE concentration, and analyzed as known positive in future assays.

Acknowledgments Thanks to Mr. Carey Copeland and Ms. Debbie Andrews for excellent technical assistance in the development of the protocols, and to Drs. Christal Bowman and Marsha Ward for helpful comments and suggestions on the manuscript. References 1. Luster MI, Portier C, Pait DG, Rosenthal GJ, Germolec DR, Corsini E, Blaylock BL, Pollock P, Kouchi Y, Craig W, White KL, Munson AE, Comment CE (1993) Risk assessment in immunotoxicology II. Relation ships between immune and host resistance tests. Fundam Appl Toxicol 21:71–82

2. Germolec DR (2004) Sensitivity and predictivity in immunotoxicity testing: immune endpoints and disease resistance. Toxicol Lett 149:109–114 3. Luebke RW, Parks C, Luster MI (2004) Suppression of immune function and susceptibility to infections in humans: association

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Luebke of immune function with clinical disease. J Immunotoxicol 1:15–24 Keil D, Luebke RW, Pruett SB (2001) Quantifying the relationship between multiple immunological parameters and host resistance: probing the limits of reductionism. J Immunol 167:4543–4552 Pozio E (1998) Trichinellosis in the European Union: Epidemiology, ecology and economic impact. Parasitol Today 14:35–38 Roy SL, Lopez AS, Schantz PM (2003) Trichinellosis Surveillance – United States, 1997–2001. MMWR Surveill Summ 52(6):1–8 Vos JG, De Klerk A, Krajnc EI, van Loveren H, Rozing J (1990) Immunotoxicity of bis(tri-n-butyltin)oxide in the rat: effects on thymus-dependent immunity and on nonspecific resistance following long-term exposure in young versus aged rats. Toxicol Appl Pharmacol 105:144–155 Luebke RW, Luster MI, Dean JH, Hayes HT (1984) Altered host resistance to Trichinella spiralis infection following subchronic exposure to diethylstilbestrol. Int J Immunopharmacol 6:609–617 Luebke RW, Copeland CB, Deliberto JJ, Akubue PI, Andrews DL, Riddle MM, Williams WC, Birnbaum LS (1994) Assessment of host resistance to Trichinella spiralis in mice following preinfection exposure to 2, 3, 7, 8-TCDD. Toxicol Appl Pharmacol 125:7–16 Luebke RW, Copeland CB, Andrews DL (1995) Host resistance to Trichinella spiralis infection in rats exposed to 2, 3, 7, 8-tetrachlorodibenzo-p-dioxin (TCDD). Fund Appl Toxicol 24:285–289 Luebke RW, Copeland CB, Andrews DL (1999) Effects of aging on resistance to Trichinella spiralis infection in rodents exposed to 2, 3, 7, 8-tetrachlorodibenzo-pdioxin. Toxicology 136:15–26 Goettsch W, Garssen J, Deijns A, de Gruijl FR, van Loveren H (1994) UV-B exposure impairs resistance to infection by Trichinella spiralis. Environ Health Perspect 102:298–301 Houben GF, Penninks AH, Seinen W, Vos JG, van Loveren H (1993) Immunotoxic effects of the color additive caramel color III: immune function studies in rats. Fundam Appl Toxicol 20:30–37 Bell RG (1988) The generation and expression of immunity to Trichinella spiralis in laboratory rodents. Adv Parasitol 41:149–217 Khan WI (2008) Physiological changes in the gastrointestinal tract and host protective immunity: learning from the mouse-Trichinella spiralis model. Parasitology 135:671–682

16. Knight PA, Brown JK, Pemberton AD (2008) Innate immune response mechanisms in the intestinal epithelium: potential roles for mast cells and goblet cells in the expulsion of adult Trichinella spiralis. Parasitology 135:655–670 17. Ruitenberg EJ, Elgersma A (1976) Absence of intestinal mast cell response in congenitally athymic mice during Trichinella spiralis infection. Nature 264:250–260 18. Vos JG, Ruitenberg EJ, Van Basten N, Buys J, Elgersma A, Kruizinga W (1983) The athymic nude rat IV. Immunocytochemical study to detect T-cells, and immunological and histopathological reactions against Trichinella spiralis. Parasite Immunol 5:195–215 19. Almond NM, Parkhouse RM (1986) Immunoglobulin class specific responses to biochemically defined antigens of Trichinella spiralis. Parasite Immunol 8:391–406 20. Urban JF Jr, Schopf L, Morris SC, Orekhova T, Madden KB, Betts CJ, Gamble HR, Byrd C, Donaldson D, Else K, Finkelman FD (2000) Stat6 signaling promotes protective immunity against Trichinella spiralis through a mast cell- and T cell-dependent mechanism. J Immunol 164:2046–2052 21. McDermott JR, Bartram RE, Knight PA, Miller HRP, Garrod DC, Grencis RK (2003) Mast cells disrupt epithelial barrier function during enteric nematode infection. Proc Nat Acad Sci USA 100:7761–7766 22. Khan WI, Vallance BA, Blennerhassett PA, Deng Y, Verdu EF, Matthaei KI, Collins SM (2001) Critical role for signal transducer and activator of transcription factor 6 in mediating intestinal muscle hypercontractility and worm expulsion in Trichinella spiralis-infected mice. Infect Immun 69:838–844 23. Vallance BA, Croitoru K, Collins SM (1998) T lymphocyte-dependent and independent intestinal smooth muscle dysfunction in the T. spiralis-infected mouse. Am J Physiol 275:G1157–G1165 24. Vallance BA, Galeazzi F, Collins SM, Snider DP (1999) CD4 T cells and major histocompatibility complex class II expression influence worm expulsion and increased intestinal muscle contraction during Trichinella spiralis infection. Infect Immun 67:6090–6097 25. Bell RG, Appleton JA, Negrao-Correa DA, Adams LS (1992) Rapid expulsion of Trichinella spiralis in adult rats mediated by monoclonal antibodies of distinct IgG isotypes. Immunology 75:520–527 26. Appleton JA, McGregor DD (1984) Rapid expulsion of Trichinella spiralis in suckling rats. Science 226:70–72

Parasite Challenge as Host Resistance Models for Immunotoxicity Testing 27. Inaba T, Sato HA, Kamiya H (2003) Impeded establishment of the infective stage of Trichinella in the intestinal mucosa of mice by passive transfer of an IgA monoclonal antibody. J Vet Med Sci 65:1227–1231 28. Ruitenberg EJ, Buys J, Teppema JS, Elgersma AZ (1983) Rat mononuclear cells and neutrophils are more effective than eosinophils in antibodymediated stage-specific killing of Trichinella spiralis in vitro. Z Parasitenk 69:807–815 29. Venturiello SM, Giambartolomei GH, Costantino SN (1993) Immune killing of newborn Trichinella larvae by human leucocytes. Parasite Immunol 15:559–564 30. Wang CH, Bell RG (1988) Antibodymediated in-vivo cytotoxicity to Trichinella spiralis newborn larvae in immune rats. Parasite Immunol 10:293–308

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31. Lee TD, Befus D (1989) Effects of rat and human intestinal lamina propria cells on viability and muscle establishment of Trichinella spiralis newborn larvae. J Parasitol 75: 124–128 32. Gurish MF, Bryce PJ, Tao H, Kisselgof AB, Thornton EM, Miller HR, Friend DS, Oettgen HC (2004) IgE enhances parasite clearance and regulates mast cell responses in mice infected with Trichinella spiralis. J Immunol 172:1139–1145 33. Dessein AJ, Parker WL, James SL, David JR (1981) IgE antibody and resistance to infection I. Selective suppression of the IgE antibody response in rats diminishes the resistance and the eosinophil response to Trichinella spiralis infection. J Exp Med 153:423–436

Chapter 10 Tumor Challenges in Immunotoxicity Testing Sheung Ng, Kotaro Yoshida, and Judith T. Zelikoff Abstract Syngeneic murine tumor models have been widely used by researchers to assess changes in tumor susceptibility associated with exposure to toxicants. Two common tumor models used to define host resistance against transplanted tumors in vivo are EL4 mouse lymphoma cells (established from a lymphoma induced in a C57BL/6 mouse by 9,10-dimethyl-1,2-benzanthracene) and B16F10 mouse melanoma cells (derived through variant selection from a B16 melanoma arising spontaneously in C57BL/6 mice). While C57BL/6 mice are commonly used as the syngeneic host for these tumor models, other mouse strains such as B6C3F1 (C57BL/6 × C3H) can also be used. Tumor challenge of the host can be done by subcutaneous (sc) or intravenous (iv) injection, depending upon whether the effects are to be examined on local tumor development or experimental/artificial metastasis. Materials and methodologies for injection of both tumor cell models are described in detail in the subsequent sections. Key words: Tumor challenge, Tumor cell models, B16F10 melanoma cell model, EL4 lymphoma cell model, Murine model

1. Introduction Syngeneic murine tumor models have been widely used by researchers (particularly, immunologists and immunotoxicologists) to assess changes in tumor susceptibility associated with exposure to toxicants (1–3). The central concept for this model is based on a notion known as the “immune surveillance” hypothesis that was first discussed over a century ago and reintroduced by Burnet in the late 1950s (4). After losing momentum for a number of years, this theory, which postulates that the immune system plays a central role in the resistance against the development of detectable tumors, has been given new life over the last decade. Tumor challenge assays using syngeneic animal models can help to illuminate the different components of the immune surveillance R.R. Dietert (ed.), Immunotoxicity Testing: Methods and Protocols, Methods in Molecular Biology, vol. 598 DOI 10.1007/978-1-60761-401-2_10, © Humana Press, a part of Springer Science + Business Media, LLC 2010

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hypothesis in vivo. By studying the roles of specific immune cell types that mediate tumor growth and metastasis in laboratory animals, the immune response of the host against tumors can be more clearly defined. Expression of major histocompatibility complex (MHC) class 1 molecules on the surface of tumor cells, as well as antigen-presenting cells (APC) such as macrophages and dendritic cells (DC) that present processed self-peptides, are important components of anti-tumor mechanisms that help to recruit effector T-lymphocytes to the tumor microenvironment (5–8). Once targeted for apoptosis or necrosis, tumor cells are killed by the coordinated anti-tumor activities of cytotoxic T-lymphocytes (CTL) and T-helper cells (i.e., Th1 and Th2) that are part of the adaptive immune system, as well as by innate immune cells, including natural-killer (NK) cells and cytolytic macrophages. Both the innate and adaptive arms of the immune system are vital for a successful immune response against a growing tumor (9, 10). Recent investigations, however, have demonstrated the ability of tumor cells to actively escape immune surveillance and thus prolong its survival in the host (11). One prominent mechanism employed by tumor cells is to reduce or lose the expression of MHC class I molecules on its surface, thus rendering it undetectable by circulating lymphocytes (12–14). Another tumor avoidance strategy involves the active migration of immunosuppressive regulatory T-cells (Treg) into the tumor microenvironment. The influx of Treg cells can inhibit the antitumor immune response by blocking the activity and proliferation of effector T-lymphocytes (15, 16). Imbalance between anti-tumor effector functions and immunosuppression could result in changes in tumor incidence, growth rate, and/or risk of metastasis (17). The immunological effects of murine challenge with validated syngeneic tumor cell models can be assessed by a number of wellestablished methods. Flow cytometry, for example, can be employed to measure the changes in specific immune cell profiles (compared with control levels) within the tumor microenvironment, blood, or peripheral lymphoid organs such as the thymus or spleen. Enzyme-linked immunosorbent assays (ELISA) can measure the plasma or intratissue levels of chemokines and cytokines that are associated with suppressing or promoting tumor growth, such as transforming growth factor beta (TGF-b) and interleukin (IL)-10 (18, 19). A histological assessment of lymphoid tissues, such as thymic epithelium or splenic white pulp, can also illuminate the organ-specific effects that a growing tumor can have on the immune system. Investigators can also provide a relationship between tumor dose and a specific immune endpoint (e.g., degree of thymic atrophy) by utilizing several different tumor cell concentrations.

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Two common syngeneic tumor models used to define host resistance against transplanted tumors in vivo are EL4 mouse lymphoma and B16F10 mouse melanoma cells. Both tumor cell models have successfully demonstrated the critical role that the immune system plays in the induction, growth, and metastasis of induced-tumors. A prominent advantage of this model system is that it can assess the effects of a variety of toxic chemicals (e.g., metals, pesticides, polycyclic hydrocarbons) on the functional integrity of the intact immune system necessary for the protection of the host against nascent tumors. Murine host resistance models of tumor cell rejection are highly reproducible, and results can be correlated with outcomes seen in vitro making them ideal for assessing immunotoxic risk. However, such models are also sensitive and influenced by a number of variables, including: the rodent strain and gender, specific tumor cell model and injection dose, and the temporality of chemical exposure. These types of assays also require a large number of animals to be used per treatment group for adequate statistical power. Therefore, a thorough knowledge of the host organism as well as a clear understanding of the pathogenic or carcinogenic process of tumor-induction must be obtained prior to testing.

2. Materials 2.1. Maintenance of Tumor Cell Cultures

1. EL4 mouse lymphoma cells (ATCC, Manassas, VA) 2. Falcon polystyrene serological pipets (BD Labware, Franklin Lakes, NJ) 3. Falcon 50 ml centrifuge tubes (BD Labware, Franklin Lakes, NJ) 4. Hemacytometer Hampton, NH)

(Fisher

Scientific

International,

Inc.,

5. Trypan blue stain (0.4%) (Invitrogen Corporation, Carlsbad, CA) 6. High-speed refrigerated (4°C) Wilmington, DE) set at 350×g

centrifuge

(DuPont,

7. Dulbecco’s Modified Eagle’s Medium (DMEM) (Invitrogen Corporation, Carlsbad, CA) 8. Horse serum (Invitrogen Corporation, Carlsbad, CA) 9. Penicillin-streptomycin (Sigma-Aldrich, St. Louis, MO) 10. l-glutamine (200 mM) (Invitrogen Corporation, Carlsbad, CA)

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11. Falcon 75 cm2 tissue culture flasks (BD Labware, Franklin Lakes, NJ) 12. CO2 water-jacketed incubator (Nuaire, Plymouth, MN) set at 37°C (5% CO2) 13. B16F10 mouse melanoma cells (ATCC, Manassas, VA) 14. Fetal bovine serum (FBS) (Invitrogen Corporation, Carlsbad, CA) 15. Trypsin (0.25%) with ethylenediamine tetraacetate (EDTA) 4Na (Invitrogen Corporation, Carlsbad, CA) 2.2. Preparation for In Vivo Challenge

1. Dulbecco’s Phosphate Buffered Saline (PBS) (Invitrogen Corporation, Carlsbad, CA)

2.3. Tumor Cell Injection

1. Mouse tail illuminator (Braintree Scientific, Inc., Braintree, MA) 2. Tailveiner® (Braintree Scientific, Inc., Braintree, MA) 3. One milliliter syringe (BD Labware, Franklin Lakes, NJ) fitted with a 23-G or 27-G needle (BD Labware, Franklin Lakes, NJ) 4. Forceps (George Tiemann & Company, Hauppauge, NJ)

2.4. Measurements and Endpoints

1. Caliper (Fisher Scientific International, Inc., Hampton, NH) 2. Pentobarbital sodium (Sleepaway) (Fort Dodge Laboratories, Fort Dodge, IA) 3. Bouin’s fixative solution (saturated picric acid/formaldehyde/acetic acid) (Sigma-Aldrich, St. Louis, MO)

3. Methods The tumor models that are described in the following section are syngeneic to C57BL/6 mice. Other strains of mice such as B6C3F1 (C57BL/6× C3H) can also be used (see Note 1). Mice should be ordered and allowed to acclimate for at least 1 week prior to preparation of cell cultures (see Note 2). Tumor challenge is performed most commonly by either subcutaneous (sc) or intravenous (iv) injection, depending upon whether the effects are to be examined on local tumor development or experimental/artificial metastasis. Spontaneous metastasis refers to the formation of a primary tumor at the site of transplantation followed by a distant metastasis. The formation of tumor colonies at a target organ after tumor cells are injected directly into the circulation (either by iv or intraperitoneally [ip]) is

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described as experimental/artificial metastasis (20). Although many different tumor models can be used in this challenge system (e.g., PYB6 fibrosarcoma cells), EL4 lymphoma (established from a lymphoma induced in a C57BL/6 mouse by 9,10-dimethyl-1,2benzanthracene) and B16F10 melanoma cells (derived through variant selection from a B16 melanoma arising spontaneously in C57BL/6 mice) are routinely used to assess changes in tumor susceptibility in response to toxicant exposure. Both the tumor cell types can be used in murine hosts, using either injection protocol. To obtain reliable and reproducible results, it is critical that all the reagents are sterile and all the procedures are performed under aseptic conditions in a biological safety hood. As EL4 lymphoma cells grow in suspension and B16F10 melanoma cells grow as adherent cell cultures, each requires somewhat different culturing procedures. Both the cell types should be grown and maintained at their logarithmic growing range in order to avoid a decrease in tumor cell viability. Thus, it is important to monitor tumor cell concentration at each passage. Prior to host challenge, both the tumor cell types should be passaged at least twice prior to animal injection. It is also recommended that cells have the same passage history for each experiment in order to obtain comparable results between studies. The injection dose will vary depending upon the tumor cell type, injection route, and desired tumor incidence. Thus, a preliminary study performed prior to the actual experiment is recommended to define the exact concentration of tumor cells needed (see Note 3). 3.1. Maintenance of Tumor Cell Cultures 3.1.1. EL4 Lymphoma Cells

1. Thaw EL4 mouse lymphoma cells from the frozen ampule (stored in liquid nitrogen at –80°C) in a warm (37°C) water bath 2. Pipet the contents of the ampule into a 50 ml centrifuge tube 3. Determine the exact cell concentration and cell viability by hemacytometer counting and trypan blue exclusion, respectively (see Note 4) 4. Centrifuge the cells (at 4°C) for 5 min at 350×g and discard the supernatant 5. In the same centrifuge tube, resuspend the tumor cells in 10 ml growth medium (DMEM, supplemented with 10% horse serum, 1% penicillin-streptomycin, and 1% l-glutamine) 6. Gently vortex the tube 7. Transfer the cell suspension into a 75 cm2 screw-cap tissue culture flask 8. Place the tissue culture flask in a humidified, 37°C incubator containing 5% CO2

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9. When the cell concentration in the tissue culture flask reaches 1 × 106 cells/ml (see Note 5), pipet the entire contents of the flask into a 50 ml centrifuge tube 10. Repeat steps 3–6 11. Resuspend the cells (in a new 75 cm2 culture flask) to a final concentration of 2 × 105 cells/ml in a total of 10 ml growth medium 12. Place the flask at 37°C in a CO2 incubator (5% CO2) 3.1.2. B16F10 Melanoma Cells

1. Follow steps 1 through 8 (Subheading 10.3.1.1), using DMEM supplemented with 10% FBS, 1% penicillin-streptomycin, and 1% l-glutamine 2. When the attached cells reach approximately 90% confluency, decant growth medium and add 3 ml of 0.25% trypsin-EDTA solution to the flask 3. After incubation (at 37°C) for approximately 3 min, tap the flask repeatedly until the cells detach from the internal surface 4. Pipet 7 ml of supplemented growth media (see step 1 above) into the tissue culture flask to stop the digestive action of the trypsin 5. Rinse the internal surface of the flask several times until all the cells are completely detached 6. Pipet the contents into a 50 ml centrifuge tube 7. Centrifuge the cells at 350×g for 5 min (at 4°C) and discard the supernatant 8. In the same centrifuge tube, resuspend the cells in 10 ml of growth medium 9. Gently vortex the tube 10. Pipet 2 ml of tumor cell suspension and 8 ml of growth medium into a new 75 cm2 tissue culture flask (i.e., a subcultivation ratio of 1:5 is suggested for 90% confluency to be reached in ~2–3 days) 11. Incubate the flask in a humidified 37°C incubator containing 5% CO2

3.2. Preparation for In Vivo Challenge

1. Two to three days after the last cell passage, pipet the entire contents of the flask into a 50 ml centrifuge tube about 30 min prior to animal inoculation (see Note 6) 2. Centrifuge the cells for 5 min (at 4°C) at 350×g and discard the supernatant 3. In the same centrifuge tube, resuspend the tumor cells with 10 ml of PBS and then gently vortex the tube

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4. Repeat step 2 5. Resuspend the cells with 1 ml of PBS and gently vortex the tube 6. Determine the cell concentration and viability by hemacytometer counting and trypan blue exclusion, respectively (see Note 4) 7. Add PBS to obtain the desired tumor cell concentration (volume of PBS added depends upon the original cell concentration) (see Note 3) 8. Vortex the centrifuge tube containing the cell suspension and keep on ice until needed 3.3. Tumor Cell Injection

1. Place the mouse into the restraining tube of a mouse tail illuminator or tailveiner®

3.3.1. Subcutaneous (sc) Injection

2. Gently pull the tail through the slot of the sliding door, then slide and lock the tapered plug to accommodate the size of the mouse 3. Vortex the centrifuge tube containing the previously prepared tumor cell suspension (see Subheading 10.3.2) 4. Load 0.1 ml of the tumor cell suspension into a 1 ml syringe affixed with a 23-G needle (see Note 7) 5. Open the sliding door of the restrainer or tailveiner® and pull the right back leg straight out 6. While pulling the thigh skin upwards with a forceps, inject 0.1 ml of tumor cells (see step 4 above) subcutaneously (sc) into the right rear thigh

3.3.2. Intravenous (iv) Injection

1. Follow steps 1 and 2 in Subheading 10.3.3.1 using a mouse tail illuminator 2. Place the tail in the illuminated slot until the tail vein dilates 3. Vortex the centrifuge tube containing the previously prepared tumor cell suspension (see Subheading 10.3.2) 4. Load 0.1 ml of the tumor cell suspension into a 1 ml syringe affixed with a 27-G needle (see Note 7) 5. Inject 0.1 ml of tumor cells (c.f. step 4 above) intravenously (iv) into the visible tail vein

3.4. Measurements and Endpoints 3.4.1. Subcutaneous (sc) Challenge

1. Palpate each mouse daily at the injection site and record the first day when a tumor/mass becomes palpable. Use these data to determine “time to tumor formation” 2. Measure tumor size daily using a ruled caliper for 60 days postinjection or until the tumor reaches 20 mm in size. Use these data to determine mean “tumor growth rate (mm/day)” (see Note 8)

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3. Determine “tumor incidence” at the end of the 60 day observation period (see Note 9) 4. Sacrifice tumor cell-injected mice either at the end of the observation period or when the tumor reaches about 20 mm in size (tumors >20 mm can interfere with animal movement and quality of life) by intraperitoneal (ip) injection of 175 mg/ kg pentobarbital sodium 5. Monitor mice having no palpable tumor after 30 days every other day. If no tumor is palpable 60 days post-injection, consider the outcome as negative and note it as “no palpable tumor” 3.4.2. Intravenous (iv) Challenge Pilot Study

1. Inject a group of 20 naïve mice with a concentration of tumor cells previously determined in a preliminary dose-response experiment (see Note 3) 2. Sacrifice 2 mice every other day 3. Remove the appropriate “metastatic” target organ (i.e., lungs for B16F10 cells and liver for EL4 cells) and count visible nodules on the organ surface 4. Determine the day post-injection when tumor nodules reach a macroscopic/countable size is determined as the appropriate “day of sacrifice” in the actual tumor challenge study (see Note 10)

3.4.3. Actual Tumor Challenge Study

1. Euthanize the mice (ip injection of 175 mg/kg pentobarbital sodium) on the selected “day of sacrifice” (see step 4 above) 2. Remove the appropriate “metastatic” target organ (i.e., lungs for B16F10 cells and liver for EL4 cells) and place it in a tube containing 2 ml of Bouin’s fixative solution for 3 days 3. Wash the organ thoroughly with 70% ethanol 4. Count the total number of tumor cell colonies (nodules) on the entire surface of the organ using a dissecting microscope (see Note 11)

4. Notes Notes are given in a list as follows. A tumor challenge study performed by Ng et al. (21) is used to provide relevant examples (italicized) throughout this section. 1. The tumor models described in this protocol (i.e., B16F10 melanoma and EL4 lymphoma cell lines) were derived originally from C57BL/6 mice. Thus, both the tumor models are syngeneic to C57BL/6 mice and to most other parental crosses

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involving the C57BL/6 strain as a single parent. Host resistance assays using these particular tumor models are most commonly carried out using C57BL/6 mice as well as B6C3F1 mice, a laboratory mouse strain produced from a parental cross between C57BL/6 and C3H mice. 2. Mice should be acclimated in the “home” laboratory for at least 1 week prior to tumor challenge. It is recommended that the animal source, husbandry conditions, and handling procedures be standardized in order to obtain comparable results between experiments. It is also suggested that at least 25 mice be assigned to each experimental (control and treatment) group and another 10–15 mice be used for the vehicle control group. For example, in a study investigating the effects of prenatal cigarette smoke exposure on tumor susceptibility in the offspring, 9-11-wk-old pathogen-free B6C3F1 female mice (purchased from The Jackson Laboratory [Bar Harbor, MA]) were mated, and 28 male offspring each from smoke- and air-exposed female mice were injected subcutaneously (sc) at 5-wk-of-age with EL4 lymphoma cells. Ten offspring from each exposure group were injected with PBS to determine any spontaneous and/or vehicle-induced tumors.

For challenge studies involving intravenous (iv) injection and examination of tumor nodules on organ surfaces, additional mice are needed to serve as “sentinels” for defining actual “day of sacrifice” (see Subheading 10.3.4.2 and Note 10).

Prior to the actual experiment, a pilot study should be performed that employs at least 4 different concentrations of transplanted tumor cells. This study will help to establish the optimal concentration for the particular tumor cell type and the route of injection being used. Some tumor cell concentrations used in other challenge studies are shown in Table 10.1.

One of four concentrations of EL4 lymphoma cells (i.e., 5,000, 50,000, 200,000, and 500,000 tumor cells/mouse) were injected subcutaneously (sc) into the right rear thigh of juvenile B6C3F1 mice to determine the dose of tumor cells which yielded a 20-40% tumor incidence (TI) in naïve mice (i.e., 50,000 cells); this (particular) TI was used so that a toxicant-induced change in either direction (i.e., higher or lower than control) could be observed.

3. Tumor cell concentration and viability can be determined by transferring 20 ml of the previously vortexed tumor cell suspension into a 1.5 ml Eppendorf tube already containing 80 ml of Trypan blue (0.04%), and by immediately placing 10 ml of the mixed solution onto a hemacytometer. Count the numbers of viable (bright) and injured/dead (dark blue) cells on the hemacytometer using a light microscope (40×) and calculate final

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Table 10.1 Suggested cell concentrations used in tumor challenge studies Tumor model

Route of injection

Suggested tumor cell concentrations/mouse (references)

EL4

Subcutaneous

5 × 104 (21)

Intravenous

5 × 105 (22), 1 × 106 (23)

Subcutaneous

3 × 103 (24)

Intravenous

2 × 105 (25)

B16F10

tumor cell concentration and viability. Cells with <80% viability should not be used for the challenge study. Tumor cell concentration = (number of viable cells) (dilution ratio)(104) = (40)(5)(104) = 2 ´106 viable cells / ml Viability = (number of viable cells) / (total number of cells) ´ 100 = 38 / 40´100 = 95% 4. Doubling time for EL4 lymphoma cells (when maintained between 1 × 105 and 1 × 106 cells/ml) is approximately 24 h. “Starting concentration” refers to the initial concentration in the cell culture flask, not the frozen ampule.

The day of passage after initiation of the culture (n) in our study was calculated by:

Highest optimal growing concentration = Starting concentration × 2n 1 × 106 cells/ml = 2 × 105 cells/ml × 2n n = 2.32

Based on these calculations, tumor cells in this laboratory were passaged every 2 or 3 days after initiation of the culture.

5. Calculate the total number of tumor cells needed for animal injections for the entire experiment. Calculations are based upon the: (a) desired tumor challenge dose (i.e., number of tumor cells/mouse), (b) number of experimental groups, and (c) number of mice per experimental and vehicle control group.

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Table 10.2 Tumor size (diameter) of 5-week-old naive male offspring following subcutaneous (sc) injection with cultured EL4 lymphoma cells Tumor measurement Mouse number

Parameter

Day 1a

Day 2

Day 3

n–1b

n c

Mean

1

Diameter (mm)

4.52

5.84

7.03

17.66

20.18

–

Growth (mm)

–

1.32

1.19

–

2.52

2.26

Diameter (mm)

5.41

7.02

9.13

18.54

21.22

–

Growth (mm)

–

1.61

2.11

–

2.68

2.39

Diameter (mm)

5.16

7.07

8.95

17.85

19.99

–

Growth (mm)

–

1.91

1.88

–

2.14

2.04

Diameter (mm)

4.65

6.88

8.49

18.14

20.65

–

Growth (mm)

–

2.23

1.61

–

2.51

2.27

2

3

4

Mean

2.24

Size measurements began on Day 1 when the tumor first became palpable and continued until 60 days postinjection or when the tumor reached ~20 mm in diameter b n–1 = one day prior to final measurement c n = last day of tumor size measurement a

Table 10.3 Tumor incidence of 5-, 10-, and 20-week-old naive male offspring after subcutaneous (sc) injection with cultured EL4 lymphoma cells Age (week)

5

10

20

Number of mice injected (sc) with EL4 cells

28

28

20

Number of mice with palpable tumors

8

8

9

Tumor incidence (%)

29

29

45

Number of tumor cells needed = 5 × 104 cells/mouse × 2~group × 28 mice/group = 2.8 × 106 cells A 75 cm2 tissue culture flask contains about 1 × 106 cells/ml (in 10 ml growth medium) at logarithmic growth. Thus, 1 flask provided about 1 × 107 cells, which was sufficient for our experiment.

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6. Proceed quickly to the next injection steps to avoid settling and uneven distribution of tumor cells in the syringe. 7. Determine the daily tumor growth (i.e., tumor diameter [mm] on dayn – tumor diameter [mm] on dayn–1) for a single mouse. All individual values from a single mouse are averaged to determine the mean daily tumor growth rate (mm/day). Tumor growth rate for an entire treatment group can be calculated by averaging the mean daily tumor growth rates for each mouse in a given treatment group. Examples of actual EL4-induced tumor diameters and calculation of individual tumor growth rate are shown in Table 10.2. 8. Tumor incidence can be determined by dividing the number of mice that develop a palpable tumor by the total number of mice injected with tumor cells. The actual number of mice with palpable tumors and the calculation of tumor incidence are demonstrated in Table 10.3. 9. The purpose of the proposed pilot study is to avoid the mice being sacrificed before the tumor nodules in the target organ are countable. During the actual tumor challenge study, four extra mice could be injected per experimental group and sacrificed early to assure adequate nodule size. 10. After staining with Bouin’s solution, the organ tissue exhibits a yellowish-gold color. The nodules can be black or white, depending on the tumor cell type used for challenge.

Acknowledgements This work was supported (in part) by the New York University (NYU) National Institute of Environmental Health Sciences (NIEHS) Center Grant (ES00260) and the Institute For Science and Health (07-1500-01RFA05). References 1. Luster MI, Munson AE, Thomas PT, Holsapple MP, Fenters JD, White KL Jr, Lauer LD, Germolec DR, Rosenthal GJ, Dean JH (1988) Development of a testing battery to assess chemical-induced immunotoxicity: National toxicology program’s guidelines for immunotoxicity evaluation in mice. Toxicol Sci 10:2–19 2. Luster MI, Portier C, Gayla Pait D, White KL, Gennings C, Munson AE, Rosenthal GJ (1991) Sensitivity and predictability of immune tests. Toxicol Sci 18:200–210

3. Luster MI, Portier C, Gayla Pait D, Rosenthal GJ, Germolec DR, Corsini E, Blaylock BL, Pollock P, Kouchi Y, Craig W, White KL, Munson AE, Comment CE (1993) Relationships between immune and host resistance tests. Toxicol Sci 21:71–82 4. Burnet FM (1957) Cancer – a biological approach. Br Med J 1:779–786, 841–847 5. Townsend AR, Rothbard J, Gotch FM, Bahadur D, Wraith D, McMichael AJ (1986) The epitopes of influenza nucleoprotein recognized by cytotoxic T lymphocytes can

Tumor Challenges in Immunotoxicity Testing be defined with short synthetic peptides. Cell 28:959–968 6. Ljunggren HG, Karre K (1990) In search of the “missing self”: MHC molecules and NK cell recognition. Immunol Today 11:237–244 7. Pike MC, Snyderman R (1976) Depression of macrophage function by a factor produced by neoplasms: A mechanism for abrogation of immune surveillance. J Immunol 117: 1243–1249 8. Vacari AP, Caux C, Trinchieri G (2002) Tumor escape from immune surveillance through dendritic cell inactivation. Sem Cancer Biol 12:33–42 9. Boon T, Cerottini JC, van den Eynde D, van der Bruggen P, van Pel A (1994) Tumor antigens recognized by T lymphocytes. Annu Rev Immunol 12:337–365 10. Trowsdale J (2001) Genetic and functional relationships between MHC and NK receptor genes. Immunity 15:363–374 11. Igney FH, Krammer PH (2002) Immune escape of tumors: Apoptosis resistance and tumor counterattack. J Leukocyte Biol 71:907–920 12. Garrido F, Cabrera T, Concha A, Glew S, Ruiz-Cabello F, Stern PL (1993) Natural history of HLA expression during tumor development. Immunol Today 14:491–493 13. Garrido F, Ruiz-Cabello F, Cabrera T, PerezVillar J, Lopez-Botet M, Duggan-Keen M, Stern PL (1997) Implications for immunosurveillance of altered class I phenotypes in humans tumors. Immunol Today 18:89–95 14. Hicklin DJ, Marincola FM, Ferrone S (1999) HLA class I antigen downregulation in human cancers: T cell immunotherapy revives an old story. Mol Med Today 5:178–186 15. Kiessling R, Wasserman K, Horiguchi S, Kono K, Sjoberg J, Pisa P, Petersson M (1999) Tumor-induced immune dysfunction. Cancer Immol Immunother 48:353–362 16. Siegmund K, Feueuer M, Siewert C, Ghani S, Haubold U, Dankof A, Krenn V, Schon MP, Scheffold A, Lowe JB, Hamann A,

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Syrbe UM, Huehn J (2005) Migration matters: Regulatory T-cell compartmentalization determines suppressive activity in vivo. Blood 106:3097–3104 Suri-Payer E, Amar AZ, McHugh R, Natarajan K, Marquiles DH, Shevach EM (1999) Postthymectomy autoimmune gastritis: Fine specificity and pathogenicity of anti-H/K ATPase-reactive T cells. Eur J Immunol 29:669–677 Markowitz SD, Roberts AB (1996) Tumor suppressor activity of the TGF-beta pathway in human cancers. Cytokine Growth Factor Rev 7:93–102 Moore KW, O’Garra A, de Waal MR, Vieira P, Mosmann TR (1993) Interleukin-10. Ann Rev Immunol 11:165–190 Elkin M, Vlodavsky I (2001) Tail vein assay of cancer metastasis. Curr Protoc Cell Biol S12, Unit 19.2 Ng SP, Silverstone AE, Lai ZW, Zelikoff JT (2006) Effects of prenatal exposure to cigarette smoke on offspring tumor susceptibility and associated immune mechanisms. Toxicol Sci 89:135–144 Li C, Bai X, Wang S, Tomiyama-Miyaji C, Nagura T, Kawamura T, Abo T (2004) Immunopotentiation of NKT cells by lowprotein diet and the suppressive effect on tumor metastasis. Cell Immunol 231: 96–102 Salem ML (2005) Systemic treatment with n-6 polyunsaturated fatty acids attenuates EL4 thymoma growth and metastasis through enhancing specific and non-specific antitumor cytolytic activities and production of TH1 cytokines. Int Immunopharmacol 5:947–960 Ng SP, Zelikoff JT (2008) The effects of prenatal exposure of mice to cigarette smoke on offspring immune parameters. J Toxicol Environ Health A 71:445–453 Grundy MA, Zhang T, Sentman CL (2007) NK cells rapidly remove B16F10 tumor cells in a perforin and interferon-gamma independent manner in vivo. Cancer Immunol Immunother 56:1153–1161

Part IV Testing Protocols in Rodents and Other Laboratory Animals

Chapter 11 The T-Dependent Antibody Response to Keyhole Limpet Hemocyanin in Rodents Lisa M. Plitnick and Danuta J. Herzyk Abstract Central to the evaluation of potential immunotoxicants is the concept that measurement of multiple parameters is required for the determination of toxicity toward the immune system. A carefully considered integration of endpoints involved in the immune response should be used to determine an immunotoxic effect. A functional evaluation, specifically the rodent T-cell-dependent antibody response (TDAR) model developed for regulated immunotoxicity evaluations, has been established to detect potential immunotoxicity, especially immunosuppression, caused by chemicals and novel pharmaceuticals in development. This chapter provides an overview and detailed procedures involved in the TDAR assay that measures the immune response (i.e., antibody production) to an introduced antigen (i.e., keyhole limpet hemocyanin (KLH)) in rats or mice treated with a chemical (e.g., a known immunotoxicant and/or a new drug candidate). The TDAR model of competent immune function requires the participation of multiple effector cells such as antigen presenting cells, T lymphocytes, and B lymphocytes to produce the final product, the antigen-specific antibody response. Thus, alterations in the level of antibody production to the specific antigen may reflect effects on any or all of the cell populations involved in TDAR. Key words: Immune function, Humoral immunity, TDAR, KLH, Rat- and mouse anti-KLH antibodies, IgG, IgM, ELISA

1. Introduction The immune system is a tightly regulated and very complex network of various lymphoid and other cell types interacting by cell-to-cell contact and communicating via soluble mediators such as cytokines. This multicellular organ system consists of granulocytes, macrophages, lymphocytes, and dendritic cells with various functions and unique phenotypic characteristics. The immune responses represent a series of complex events that occur following the introduction of foreign antigenic material into the body.

R.R. Dietert (ed.), Immunotoxicity Testing: Methods and Protocols, Methods in Molecular Biology, vol. 598 DOI 10.1007/978-1-60761-401-2_11, © Humana Press, a part of Springer Science + Business Media, LLC 2010

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Since the immune system is very dynamic and involves cell proliferation, differentiation, activation, and maturation, it is vulnerable to agents that disrupt any of these cellular processes. At the same time, immune cells are in a constant state of selfrenewal that contributes to the functional immunological reserve. An intact and functional immune system that includes local and systemic, innate and acquired, as well as humoral and cell-mediated immunity is required for protection from and to eliminate infectious and neoplastic disease. The main goal of the immunotoxicological evaluation of chemicals and pharmaceuticals that can affect the immune system is to detect true immunotoxic effects that may result in an increased disease susceptibility. While strong immunotoxicity signals can be detected by hematology and/or lymphoid tissue histopathology evaluation on standard toxicity studies, potential moderate immunotoxicity as a consequence of immune dysregulation by xenobiotics will manifest itself only at the functional level during an immune response to a challenge with an antigen (e.g., foreign protein, pathogen, toxin etc.). Thus, evaluation of the functional immune system requires studies involving “activated” immune cells, organs, or entire hosts in response to encountered “enemy.” T-cell-dependent antibody response (TDAR) to an antigen is a comprehensive immune function assay evaluating various aspects of immune responses, including antigen processing and presentation, B and T lymphocyte interactions, antibody production, and cytokine-dependent isotype class switch (i.e., IgM to IgG specific antibody response). The use of a TDAR assay is endorsed by regulatory agencies as a “default” immune function test for evaluating the immunotoxicity potential of new drug candidates, because any compound-induced alteration in antigen processing, presentation, cell proliferation, differentiation and/or secretion of interleukins, antibody production, and cytokine-dependent isotype class switch is thought likely to modify the response (1). In recent years, TDAR assay development and application has focused on the use of keyhole limpet hemocyanin (KLH) as a T-cell dependent antigen (2). KLH is a soluble and highly immunogenic protein and provides a better standardized and characterized reagent compared with sheep red blood cells (SRBC) used for many years in TDAR tests (3). Antibody responses to KLH, including anti-KLH IgM and anti-KLH IgG, are measured by enzyme-linked immunosorbent assay (ELISA). The TDAR test as a stand-alone functional evaluation is mainly used for hazard identification (i.e., unintended immunosuppression). 1.1. Rat KLH TDAR Assay- General Study Design Background

A typical immunotoxicology study that employs KLH TDAR in rats consists of up to five treatment groups. The first group is the KLH control. It is used to demonstrate the level of anti-KLH response after the administration of KLH by a single injection to rats on the study. This group does not receive the test article.

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The level of anti-KLH antibody production in this group is used to calculate the magnitude of potential test article-related reduction of anti-KLH antibody production (immunosuppressive effects) in test article-treated groups on the study. The second control group includes rats treated with the vehicle in which the test article is diluted, using the same dosing frequency and route of administration as for the test article. It is used to control for any potential effects not related to the treatment with the test article. The vehicle control group receives a single injection of phosphate buffered saline (PBS) in place of KLH administration. In general, there are three groups of rats treated with the test article at three dose levels. Doses of the test article are typically selected based on the existing information from other toxicology studies with the given test article. In most cases, the test article dose levels to be used in a KLH TDAR study should include the following considerations: (1) the previously observed or suspected immunotoxic effects that occur at the highest dose that is not overtly toxic, (2) a dose–response relationship should be evaluated, and (3) a no observed effect level (NOEL) should be identified. In some cases, this may be accomplished in fewer than the three dose levels typically used on routine toxicity studies. The route and frequency of dosing should be consistent with the standard toxicity studies conducted with the test article. An additional group of rats treated with a known strong immunosuppressant (e.g., Cyclosporine A, Dexamethasone, or Cyclo-phosphamide), which is considered as a positive control, may also be included in the study design if desired. If not included in each study, a positive control should be tested intermittently to ensure the consistency of the model. KLH is administered to test article groups, the positive control group and the KLH control group, and PBS (in place of KLH) to the vehicle control group on the study day previously determined to be appropriate for evaluating immunotoxic effects of the test article. The age of the rats used in the TDAR may vary depending on the duration of the study but is typically approximately 8 weeks at the beginning of the study. A routine TDAR study is 28 days in duration with the KLH administered on day 14 to enable blood sample collection for anti-KLH IgG antibody evaluation on the last day of the study. However, the duration may vary, because the test article must be administered to the rats for a period of time appropriate for evaluating potential immunosuppressive effects based on signals from the initial toxicity studies. For example, if findings such as an increased incidence of infections were observed following 4 weeks of dosing, the animals should be dosed for at least 4 weeks prior to the administration of KLH. Serum is then collected from all the animals for the analysis of anti-KLH IgM and IgG antibodies, using ELISA or another appropriate immunoassay at predetermined intervals following KLH

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administration. Kim et al. (4) demonstrated through statistical analysis that several different TDAR methods are acceptable to detect immunosuppression resulting from known immunosuppressants. Therefore, there may be slight variations in how this assay is performed in different laboratories. The procedures for the rat KLH TDAR method described here are based on studies conducted by the Immunotoxicology Laboratory at Merck and Co., Inc. The series of experiments included the evaluation of various doses of KLH given through several routes of administration in the presence and absence of an adjuvant (ALHYDROGEL™) in two strains of rats, Sprague– Dawley (SD) and Wistar–Hannover (WH) (Plitnick et al., unpublished data). SD rats were administered KLH intravenously (IV) or intraperitoneally (IP) at doses of 100, 500, 1,000, or 2,000 mg/ animal. An additional group was administered KLH IP at doses of 100, 500, and 1,000 mg/animal in the presence of the adjuvant (it was not possible to study 2,000 mg/animal because of the limitations of the adjuvant dose). Days of sampling for IgM (6 days following KLH administration on Study Day 1) and IgG (18 days following KLH administration on Study Day 1) were determined in previous studies. The results of this study suggested that the response to KLH when administered IP was less variable than when administered IV. In addition, variability tended to decrease as the dose increased in both the routes of administration, although the overall response did not appear to be dose-dependent up to 2,000 mg/animal. The inclusion of an adjuvant resulted in a slight additional decrease in variability but did not appreciably increase the overall response to KLH. An additional study was conducted in which the response to KLH in the presence or absence of the adjuvant was tested following treatment with a strong immunosuppressant (Cyclosporine A). This study suggested that the sensitivity of detection of suppression of the KLH response was somewhat blunted in the presence of an adjuvant. Based on these results as well as the ICH S8 guidance (5), which suggests that adjuvants are not considered appropriate for use in rodent TDAR assays, the adjuvant was not included in the final study design. A similar study was conducted in WH rats. KLH was administered at 500, 1,000, and 2,000 mg/animal via IV, SC, and IP routes. A time course was also included to determine whether or not the days of sampling for anti-KLH IgM and IgG previously selected for SD rats would be acceptable. WH rats were bled on Study Days 6, 7, and 8 for anti-KLH IgM analysis and Study Days 17, 18 and 19 for anti-KLH IgG analysis. All the routes of administration and dose levels of KLH exhibited increases in the anti-KLH IgM and IgG responses on all study days evaluated. However, upon examination of the variability between groups and genders and the overall response, KLH dosed IP at 1,000 µg/animal

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appeared optimal on Study Day 6 and Study Day 19 for anti-KLH IgM and IgG respectively. As determined by the studies discussed earlier, the optimal dose of KLH for both SD and WH rats is 1,000 mg/animal via IP administration. Serum is collected from all the animals 5 and 17 (SD) or 18 (WH) days following the administration of KLH for the evaluation of anti-KLH IgM and IgG antibodies, respectively, using ELISA. 1.2. Mouse KLH TDAR Assay – General Study Design Background

When potential immunotoxicity may be observed or suspected in standard toxicity studies, follow-up TDAR assays should be conducted in the same species and strain of animals in which the original finding was observed. Therefore, conducting a TDAR in mice may be necessary. There may be slight variations in how this assay is performed, and the method described here is based on studies conducted by the Immunotoxicology Laboratory at Merck and Co., Inc. Dose–response and time course studies utilizing various routes of administration were conducted (Plitnick, et. al., unpublished data). Mice were administered 100, 500, 1,000, or 2,000 mg of KLH/animal intravenously (IV) or intraperitoneally (IP) once on Study Day 1. Mice were bled for anti-KLH IgM analysis on Study Days 7, 8, and 9 and for anti-KLH IgG analysis on Study Days 14, 15, and 16. This study demonstrated that the IV route at a dose of 2,000 mg of KLH/animal was the optimal route and dose for KLH. The time course portion indicated that the optimal blood sampling days for the detection of anti-KLH IgM and IgG antibodies were day 6 (Study Day 7) and day 14 (Study Day 15) following KLH administration, respectively. In a series of follow-up TDAR studies mice were administered a known immunosuppressant (Cyclosporine A) at a dose of 30 mg/ kg/day given by subcutaneous injection or doses of 5, 15, 30, or 100 mg/kg/day by oral gavage. In contrast to rats, in which approximately 100% suppression of the response to KLH is observed at an oral dose of 20 mg/kg/day, suppression of the KLH response in mice was observed only in the group orally administered 100 mg/ kg/day. Therefore, careful consideration should be given to the dose selected for positive controls prior to use on study. Materials and Methods for both rat and mouse TDAR ELISA are the same with a few exceptions described in the following section.

2. Materials 2.1. Rat KLH TDAR

1. Rat options (a) Sprague–Dawley rats, Crl:CD®(SD), approximately 8 weeks of age at the beginning of the study, (Charles River Laboratories, Raleigh, NC).

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(b) Wistar–Hannover rats, CRL: WI(HAN), approximately 8 weeks of age at the beginning of the study, (Charles River Laboratories, Raleigh, NC). 2. Keyhole Limpet Hemocyanin (KLH) (Pierce Biotechnology, Rockford, IL). 3. Phosphate Buffered Saline (PBS), diluent for KLH. 4. Serum Separator tubes. 5. Micro plate reader. 6. 96 well Nunc Maxisorb Immunoplates. 7. Filter, 0.22 mm NYL membrane. 8. Polyoxethylenesorbitan monolaurate, Tween 20. 9. Gelatin. 10. Secondary antibodies: (a) HRP-mouse anti-Rat IgM antibody (Zymed, South San Francisco, CA). (b) HRP-mouse anti-Rat IgG antibody (Leinco Technologies, Inc., St. Louis, MO). 11. o-phenylendiamine dihydrochloride (OPD), Store at 4°C. 12. Citric acid monohydrate. 13. Hydrogen peroxide 30% (w/w) Solution, H2O2, Store at 4°C and cover in tinfoil. 14. Sulfuric acid 4.0N. 15. Distilled H2O. 16. 10N NaOH. 17. Negative Serum (see Note 1). 18. Positive Serum (see Note 2). 19. Elisa Reagents: (a) Blocking Buffer: PBS with 1% Gelatin/0.05% Tween 20 (see Note 3). (b) Wash Solution: PBS with 0.05% Tween 20. Store at RT up to 2 months. (c) Secondary Antibodies (see Note 4): i. HRP-conjugated mouse anti-rat IgM antibody: 1:5,000 dilution in blocking buffer. Prepare immediately prior to use and discard. ii. HRP-conjugated mouse anti-rat IgG antibody: 1:10,000 dilution in blocking buffer. Prepare immediately prior to use and discard. (d) 0.1 M Citric Acid buffer, pH 4.8. Store at RT up to 2 weeks.

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(e) Color Developer: o-phenylendiamine dihydrochloride (OPD; 1 mg/mL in 0.1 M citric acid). Prepare immediately prior to use and discard. (f) Stop Solution: 2N H2SO4. Store at RT for 1 month. 2.2. Mouse KLH TDAR

1. Mice: Crl:CD1(ICR), approximately 6 weeks of age at the beginning of the study (Charles River Laboratories, Raleigh, NC). 2. Blocker Casein Blocking Buffer. 3. Secondary Antibodies: (a) HRP-goat anti-Mouse IgM antibody ImmunoResearch, West Grove, PA).

(Jackson

(b) HRP-goat anti-Mouse IgG antibody ImmunoResearch, West Grove, PA).

(Jackson

4. Negative Serum: CD-1 mouse serum (Bioreclamation, Liverpool, NY) (see Note 1). 5. Positive Controls: Mouse anti-KLH IgM, mouse anti-KLH IgG (BD Pharmingen, San Diego, CA) (see Note 2). 6. Elisa Reagents (a) Blocking Buffer: Blocker Casein Buffer (1% w/v casein in PBS), Store at 4°C. (b) Assay Buffer: Casein buffer/0.05% Tween 20 (see Note 5). (c) Secondary Antibodies (see Note 4): i. HRP-conjugated goat anti-mouse IgM antibody: 1:6,000 dilution in assay buffer. Prepare immediately prior to use and discard. ii. HRP-conjugated goat anti-mouse IgG antibody: 1:2,000 dilution in assay buffer. Prepare immediately prior to use and discard.

3. Methods 3.1. Rat KLH TDAR 3.1.1. In Vivo Dosing and KLH Administration

1. Administer test article (and positive control, if included) to rats via an appropriate route and at an appropriate dose and frequency. 2. Administer KLH IP following the test article at a dose of 1,000 mg/animal in 0.4 mL PBS to KLH control and all the treatment and positive control groups and administer vehicle to the vehicle control group at a time point beyond which the previously noted immunotoxic effect is expected to occur.

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3. Collect a minimum of 300 mL of blood from all the animals into serum separator tubes for the analysis of anti-KLH IgM (5 days following KLH administration) and IgG (17 or 18 days following KLH administration for SD or WH rats, respectively) antibodies via ELISA or other appropriate immunoassay (see Note 6). 4. After collection, allow the blood to clot at room temperature for at least 30 min. 5. After clotting, centrifuge blood for 10 min at 2,500 × g, RT. 6. Collect the serum. 7. Store in aliquots and keep frozen at −70°C until tested. 3.1.1. ELISA (see Note 7)

Although other immunoassays may be used to measure antibodies to KLH, the ELISA method will be described here. 1. Quality Control Preparation (a) Prepare the negative QC by diluting the Negative Control Pool 1:200 or 1:50 for IgM or IgG, respectively, in blocking buffer. (b) Prepare Low and High concentrations of Positive QC samples in blocking buffer for IgM and IgG. 2. Screening Assay Study samples are initially screened at the minimal dilution (IgM-1:200 or IgG-1:50) in blocking buffer. Samples with a mean OD reading ³NCO are considered positive and are further titrated. (a) Approximately 15–24 h prior to test, coat micro titer plates with 2.5 mg/ml of KLH, 100 mL/well. (b) Thaw frozen KLH (10 mg/mL) immediately prior to use and dilute in cold PBS. (c) Cover all the plates with sealing tape and incubate 16–24 h at 4°C. Do not stack the plates. (d) Wash 3× with wash solution. Tap the plates on paper towels after wash. (e) Dispense 200 mL/well blocking buffer. (f) Cover all the plates with sealing tape and incubate 2 h at RT. (g) For IgM Samples: i. Prepare a 1:200 dilution of test serum samples in duplicate and a 1:200 dilution of Negative control samples (8 wells/plate). ii. Include a high positive and low positive in duplicate on each plate.

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(h) For IgG Samples: i. Prepare a 1:50 dilution of test serum samples in duplicate and 1:50 dilution of Negative control samples (8 wells/plate). ii. Include a high positive and low positive in duplicate on each plate. (i) When blocking step is complete, decant blocking buffer from plates. Finish by tapping the plates on paper towels. (j) Transfer 100 mL of the sample dilutions to the plate wells. (k) Cover with sealing tape and incubate at RT for 2 h. (l) Wash 3× with washing buffer. Tap the plates on paper towels after wash. (m) Dispense 100 mL/well HRP-conjugated mouse anti-rat IgM diluted 1:5,000 or HRP-conjugated mouse anti-rat IgG diluted 1:10,000 in blocking buffer in appropriate wells. (n) Cover with sealing tape and incubate at RT for 1 h. (o) Wash 3× with washing solution. Tap the plates on paper towels after wash. (p) Prepare color developer reagent, then dispense 100 mL/ well. (q) Incubate at RT for 10 min, and then dispense 100 mL/ well stop solution. (r) Measure absorbance at 490 nm on Micro plate reader. 3. Titration Assay (a) Follow steps (a)–(f) of the screening assay procedure. (b) For IgM samples: i. Further titrate study samples that are positive at the 1:200 dilutions in twofold increments in duplicate and determine end point titer. ii. Dilute Negative Controls 1:200 (8 wells/plate). iii. Include a high positive and a low positive in duplicate on each plate. (c) For IgG samples i. Further titrate study samples that are positive at the 1:50 dilution in twofold increments in duplicate and determine end point titer ii. Dilute Negative Controls 1:50 (8 wells/plate).

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iii. Include a high positive and a low positive in duplicate on each plate. (d) Follow steps (i)–(r) of the screening assay procedure. 4. Calculations and Interpretation (a) Interpretation i. Define the negative cut-off (NCO) as the mean plus 2 standard deviations of the OD values of the negative control wells on a given plate. ii. Consider any OD value greater than the NCO to be positive and titrate further. iii. Determine the titers as endpoint titers (i.e., the value of the last positive in the dilution series) and report as ln (endpoint titer) values. iv. Any sample with an endpoint titer below 50 (for antiKLH IgG antibodies) or below 200 (for anti-KLH IgM antibodies) is considered negative and given an arbitrary endpoint titer of 25 (IgG) or 100 (IgM), for the calculation of Geometric Mean Titer, in each treatment group. (b) Criteria for the Quality Controls i. Negative Control: The mean OD of the negative control pool must be
Overall, the procedure for in vivo dosing and KLH administration in the mouse TDAR Assay is similar to the rat TDAR Assay with the following exceptions: 1. Administer KLH IV at a dose of 2,000 mg/animal in 0.25 mL PBS. 2. Collect a minimum of 200 mL of blood from all animals into serum separator tubes for the analysis of anti-KLH IgM (6 days following KLH administration) and IgG (14 days following KLH administration) antibodies (see Notes 6 and 8).

ELISA (see Note 7)

Although other immunoassays may be used to measure antibodies to KLH, the ELISA method will be described here. Overall, the mouse TDAR ELISA is similar to the rat TDAR ELISA with the following exceptions:

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1. Quality Control Preparation (a) Prepare the negative QC by diluting the Negative Control Pool 1:100 in assay buffer containing 1% CD-1 mouse serum for IgM and 1:50 in assay buffer containing 2% serum for IgG. (b) Prepare Low and High concentrations of Positive QC samples in assay buffer for IgM and IgG. 2. Screening Assay As in the rat assay described earlier, initially screen study samples at the minimal dilution (IgM – 1:100 or IgG – 1:50) in assay buffer. Consider samples with a mean OD reading ≥NCO positive and titrate further. (a) Coat micro titer plates with 2.5 mg/ml (IgM) or 5 mg/ml (IgG) of KLH (b) During the blocking step, incubate the plates on a plate shaker 1–1.5 h. (c) For IgM Samples – Prepare a 1:100 dilution of test serum samples in duplicate and a 1:100 dilution of Negative control samples (8 wells/plate). (d) For IgG samples – Prepare a 1:50 dilution of test serum samples in duplicate and 1:50 dilution of Negative control samples (8 wells/plate). (e) Incubate samples and controls on a plate shaker at RT for 2 h. (f) For secondary antibodies, dilute HRP-conjugated goat anti-mouse IgM or IgG 1:6,000 or 1:2,000, respectively, in assay buffer. 3. Titration Assay (a) For IgM samples: i. Further titrate study samples that are positive at the 1:100 dilution in twofold increments in duplicate and determine end point titer. ii. Dilute negative controls 1:100 (8 wells/plate). iii. Include a high positive and a low positive in duplicate on each plate. (b) For IgG samples: i. Further titrate study samples that are positive at the 1:50 dilution in twofold increments in duplicate and determine end point titer. ii. Dilute negative controls 1:50 (8 wells/plate). iii. Include a high positive and a low positive in duplicate on each plate.

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4. Calculations and Interpretation (a) Interpretation i. Consider any sample with an endpoint titer below 100 (for anti-IgM antibodies) or 50 (for anti-IgG antibodies) negative, and assign an arbitrary endpoint titer of 50 (IgM) or 25 (IgG), for the calculation of the Geometric Mean Titer, in each treatment group. (b) Criteria for the Quality Controls i. Negative Control: The mean OD of the negative control pool must be
4. Notes 1. Serum from 8–12 week old naïve animals is pooled and analyzed for the presence of antibodies against KLH (IgM and IgG) following the methods described in this Procedure. Negative (Titer < NCO) sera are pooled, aliquoted, and stored at −70°C. Alternatively, naïve serum (pooled or individual) may be purchased from a qualified vendor and tested as described earlier prior to aliquoting and storage at −70°C. 2. Serum is collected from previously-immunized 8–12 week old animals. Individual serum samples are checked for the presence of antibodies against KLH (IgM and IgG) following the methods described in this Procedure. Samples with high titers are pooled separately for IgM or IgG, aliquoted, and stored at −70°C. Purified rat or mouse anti-KLH antibodies are also available commercially (BDPharmingen, San Diego, CA) and are tested as described earlier prior to storage at −70°C. 3. Filter gelatin prior to use. Store at room temperature (RT) for a maximum of 2 days. Store up to 1 month at 4°C. Buffer containing gelatin should be used at temperatures no lower than 20°C to avoid potential solidification, which could lead to inconsistent assay results. 4. As there may be lot-to-lot variability, new lots of antibodies should be tested at several dilutions to confirm the optimal performance prior to use.

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5. Store at RT for a maximum of 2 days. Store up to 1 month at 4°C. 6. Blood samples may be collected via the tail vein, orbital sinus, or vena cava (at study termination) into serum separator tubes for the isolation of serum. 7. Commercially available rat and mouse anti-KLH IgM and IgG kits are also available (Life Diagnostics, West Chester, PA). 8. In order to utilize the same animals for multiple time points, mice may be bled via the tail artery (“tail nick”) up to twice (approximately 0.2 mL/bleed) followed by a vena cava bleed (maximum amount possible) at the termination of the study.

Acknowledgments The authors thank Mark Thompson and Lyudmila Denisova for their technical assistance and Cindy Pauley for establishing the described ELISA methods for the detection of anti-KLH antibodies at Merck and Co., Inc. References 1. Dean JH, House RV, Luster MI (2007) Immunotoxicology: effects of and response to drugs and chemicals. In: Wallace Hayes A (ed) Principles and methods of toxicology, 5th edn. Taylor and Francis, Philadelphia, pp 1761–1793 2. Gore ER, Gower J, Kurali E, Sui J-L, Bynum J, Ennulat D, Herzyk DJ (2004) Primary antibody response to keyhole limpet hemocyanin in rat as a model for immunotoxicity evaluation. Toxicology 197:23–35 3. Temple L, Kawabata TT, Munson AE, White KL (1993) Comparison of ELISA and plaqueforming cell assays for measuring the humoral immune response to SRBC in rats and mice

treated with benzo(a)pyrene or cyclophosphamide. Fund Appl Toxicol 21:412–419 4. Kim CJ, Berlin JA, Bugelski PJ, Haley P, Herzyk DJ (2007) Comparison of immune functional tests using T-dependent antigens in immunotoxicology studies: a meta-analysis. Per Exp Clin Immunotoxicol 1:60–73 5. International Conference on harmonisation of technical requirements for registration of pharmaceuticals for human use (2006) ICH harmonised tripartite guideline: immunotoxicity studies for human pharmaceuticals S8. (http://www.ich.org/cache/compo/276254-1.html)

Chapter 12 The Sheep Erythrocyte T-Dependent Antibody Response (TDAR) Kimber L. White, Deborah L. Musgrove, and Ronnetta D. Brown Abstract The sheep erythrocyte T-dependent antibody Response (TDAR) evaluates the ability of animals sensitized in vivo to produce primary IgM antibodies to sheep erythrocytes (sRBC). The assay enumerates the number of antigen specific IgM antibody producing cells in the spleen. When exposure to the test material takes place in vivo, as does sensitization, the actual quantification of the number of antibody producing cells occurs ex vivo. Following the animal being euthanized, a single cell suspension of spleen cells is prepared. These spleen cells containing the IgM secreting plasma cells are incubated in a semisolid matrix of agar, sheep erythrocytes, and guinea pig serum as a single cell layer between a Petri dish and glass cover slip. After a 3 h incubation period, lysis of sRBCs around each of the IgM secreting antigen specific plasma cells results in the formation of a clear plaque, which can easily be counted. The TDAR has been found to be the most sensitive functional assay for evaluating effects on the immune system, particularly the humoral immune component. The TDAR to sheep erythrocytes still remains the gold standard for evaluating the potential adverse effects of xenobiotics on the immune system. Key words: TDAR, Plaque assay, Antibody forming cell response, T-dependent antigen SRBC, Sheep erythrocytes, Plaque forming cell response

1. Introduction For several decades, the IgM plaque forming cell response to the T-dependent antigen sheep erythrocytes (sRBCs), or T-dependent antibody Response (TDAR), has been the gold standard as a functional assay for evaluating the effects of xenobiotics on the immune system and specifically on the humoral immune component of the immune response. The assay originally referred to as the Jerne Plaque Assay (1) has undergone several improvements to make the assay more sensitive (2). Furthermore, the Plaque assay to sRBCs has been validated in numerous national and R.R. Dietert (ed.), Immunotoxicity Testing: Methods and Protocols, Methods in Molecular Biology, vol. 598 DOI 10.1007/978-1-60761-401-2_12, © Humana Press, a part of Springer Science + Business Media, LLC 2010

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international “ring studies” where the same compounds were evaluated in multiple laboratories (3–6). In recent years, there has been a movement away from the Plaque assay with the substitutions of ELISA based assays to sRBC or other T-dependent antigens such as chicken gamma globulin, tetanus toxoid, and keyhole limpet hemocyanin (KLH), with the last antigen being the most frequently used in the ELISA immunotoxicological evaluations (7–9). While the use of evaluating the humoral immune response using ELISA has many advantages when compared to the Plaque assay, recent publications have demonstrated that the sRBC and KLH ELISA are less sensitive than the conventional Plaque assay (6, 10). As a result, some of the regulatory agencies in the United States have chosen not to accept the KLH ELISA data for the evaluation of the primary IgM TDAR. In keeping with the “3 Rs” in reducing the use of animals in immunotoxicological testing, the KLH ELISA does have the advantage that in addition to evaluating primary IgM, the primary IgG and secondary IgG responses can also be evaluated in the same animals and, thus, reduce the number of animals needed. Whether the decrease in sensitivity of the primary IgM response to KLH is outweighed by the collection of additional data from the same animals is dependent on the questions being asked. This issue is a question that regulatory agencies and the stakeholder will have to resolve. In this situation, the stakeholder can be an industrial client or an academic researcher working on a limited budget. The objective of this chapter is to provide a detailed step-bystep procedure for conducting the TDAR using the Plaque assay to sRBCs. Included in Subheading 12.4 are many of the “insights” we have learned in conducting this assay for over 32 years.

2. Materials 2.1. Solutions

1. Earle’s balanced salt solution (EBSS) with HEPES (4-(2-hydroxyethyl)-1-Piperazineethanesulfonic acid) (Invitrogen, Gibco Product No. 15630). 2. EBSS without HEPES (Sigma, Cat. #A3551). 3. DEAE-Dextran (Sigma, Cat. #D 9885). Stock solution is 30 mg/ml in 0.9% NaCl, adjusted to pH 6.9 when dissolved. This is stored at 2–8°C. Made sterile for storage. 4. Alsever’s solution (Sigma, Cat. # A3551).

2.2. Reagents

1. Guinea Pig complement (GPC) (Accurate Chemical, Cat. #CL5000) (see Note 1). 2. “Bacto-Agar” (Fisher, Cat. #DF0140-15-4).

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3. Zap-o-globin II (Beckman-Coulter Part # 7546138). 4. Streck cytometry sheath fluid (Fisher, Cat #11-716-336). 2.3. Plastic and Glassware

1. 250 ml Pyrex flask. 2. 12 × 75 mm plastic test tubes with caps (Fisher, Cat. #14959-2A). 3. Frosted microscope slides (Fisher, Cat. #12-552). 4. Pasteur pipettes (Fisher, Cat. #13-678-20A). 5. 100 × 15 mm petri dishes (Fisher, Cat. #08-757-100D). 6. 45 × 50 mm cover slips (1 1/2 oz.) (Fisher, Cat. #12-544F). 7. 60 × 15 mm petri dishes (Fisher, Cat. #08-757-100B). 8. 12 × 75 mm borosilicate glass tubes (Fisher, Cat. #14-925D). 9. 10 ml pipettes (VWR #53283-708). 10. Accuvette vials (Beckman-Coulter Part # 8320592). 11. 15 ml conical tubes (Fisher, Cat #05-539-1).

2.4. Equipment

1. Pipette bulbs (Fisher, Cat. #13-678-9B). 2. Adjustable micropipettes and tips. 3. 46–48°C water bath. 4. Bellco plaque viewer. 5. Coulter counter.

3. Methods The TDAR measures the number of antigen specific antibody secreting B cells (plasma cells) producing IgM antibody directed against sheep erythrocytes (sRBCs). This is accomplished by removing the spleens from sensitized animals (e.g., those which had previously been injected with sRBCs) and preparing single cell suspensions from each spleen. In order to enumerate the number of antigen specific plasma cells, a semisolid matrix is prepared using agar and sRBCs. In addition to being the target for the secreted IgM antibody, the sRBCs serve another purpose by providing a red background due to the hemoglobin present in the red blood cell. This lawn of red blood cells is a single cell layer sandwiched between the Petri dish on the bottom and the cover slip on the top. IgM antibody being produced by the plasma cells diffuses through the semisolid agar matrix coming in contact with the surrounding red blood cells. By adding guinea pig serum as a source of complement, when the IgM antibody binds to epitopes on the surface of the sheep erythrocyte, the conformational

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change in the antibodies’ constant region resulting from the antigen–antibody complex, activates the first factor of the Classical Complement Pathway, C1q, and the subsequent complement components, which ultimately results in the lysis of the sheep erythrocyte. When the erythrocyte lyses, the hemoglobin diffuses away through the agar matrix, resulting in a clear area or “plaque” on the red lawn of sRBCs. If one looks at the plaque under a microscope, what is seen is a single cell in the middle of the clear area. This single cell is the plasma cell which secreted the antisRBC IgM antibody resulting in the lysis of the neighboring sRBCs and thus the plaque. By knowing the spleen cell number, the dilution of spleen cells added and the number of plaques counted, one can calculate the number of plaque forming cells per million spleen cells (Specific Activity) and the number of plaque forming cells per spleen (Total Spleen Activity). By comparing these values obtained from control animals to those treated with drugs or exposed to chemicals, one can determine if the drug or chemical suppresses or enhances the humoral immune response (see Fig. 12.1). 3.1. Labeling of Tubes, Petri Dishes, Reagent Bottles, and Data Collection Forms

1. Before the assay day, label 12 × 75 mm snap cap tubes with the rodent I.D. number. 2. Label Petri dishes, 100 × 15 mm, with animal numbers and the dilution of the spleen cell suspension which will be made (1:30 – plate tops and 1:120 – plate bottoms for mice and 1:50 – plate tops and 1:150 – plate bottoms for rats) for evaluation on day 4.

Fig. 12.1. Female B6C3F1 mice were administered 1,2:5,6 Dibenzanthracene, diluted in a corn oil vehicle (VH) by s.c. injection once daily for 28 days. On day 25, animals were sensitized with 7.5 × 107 SRBCs administered i.v. (0.2 ml of 3.75 × 108 cells/ml). On day 29, animals were euthanized and spleens removed, and the TDAR conducted as described above. In the left panel is the data presented as Specific Activity (IgM AFC/106 Spleen Cells). In the right panel the data are presented as Total Spleen Activity (IgM/Spleen). Each group consisted of eight animals. The data are presented as the mean ± standard error of the mean. Data were evaluated by ANOVA followed by Dunnett’s Test. ** = p £ 0.01 vs. vehicle treated animals.

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3. Complete the other preparation as much as possible before the assay day including reagent tubes, bottles and the appropriate paper work. 3.2. Preparation of Sheep Erythrocytes for Sensitization

1. For both mice and rats, 4 days before the sacrifice, immunize the animals with the appropriate number of sRBCs . Day 4 is the peak response day for the TDAR in rodents (8) (see Notes 2 and 3). 2. Use sRBCs that are no older than 2 weeks from the time they were drawn. 3. Obtain blood, drawn using aseptic techniques, supplied in Alsever’s solution and shipped cold on ice packs. 4. Store blood at 2–8°C upon receipt at the laboratory. 5. Remove blood from the supplied preparation using sterile techniques. 6. Use approximately 15–20 ml of blood to immunize 100–120 mice; 50–75 ml of blood to immunize 40–50 rats. 7. Place blood in a conical tube and centrifuge for approximately 10 min at 2,200 ± 100 rpm (1,150 × g), 2–8°C. 8. Remove and discard the Alsever’s solution. 9. Wash blood three times in EBSS with HEPES in centrifuge for 10 min at 2,200 ± 100 rpm (1,150 × g), 2–8°C. 10. After the first wash, remove the buffy coat with a Pasteur pipette and discard. 11. After the last wash, resuspend the pellet in 10 ml EBSS with HEPES. 12. Count the cells using a Coulter Counter. 13. Resuspend to the appropriate cell number in EBSS with HEPES. 14. When sRBCs cannot be prepared on the day of sensitization, dilute to desired cell count with Alsever’s and store refrigerated. Store for up to 2 weeks from the date the blood was drawn from the sheep. 15. On the day of sensitization, mark the volume on the side of the tube. 16. Centrifuge at 2,200 ± 100 rpm (1,150 × g) for approximately 10 min at 2–8°C. 17. Resuspend in EBSS with HEPES up to the marked volume. 18. Recount to confirm cell concentration.

3.3. Preparation of Animal for Injection of Sheep Erythrocytes

1. To facilitate the tail vein intravenous injection of the sRBCs, warm both rats and mice by placing them in plastic cages (containing no bedding).

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2. Place the cages on heating pads, such as those obtained from drug stores for use by patients for back pain or leg cramps (see Note 4). 3. Warm rodents on the high setting of the heating pads for approximately 20 min, during that time, maximum dilation of the tail veins is observed. If the animals are heated longer (i.e., 45 min), the dilation of the tail veins will actually decrease, thus, making the injections harder to conduct. 4. After the tail veins have been appropriately dilated, select an animal from a warming cage. 5. Place the animal in a different cage with the tail being held outside the cage through the metal bars of the cage top. 6. Wipe the tail with 70% ethanol to enhance seeing the tail veins and to prevent possible infection from the injection. 7. For mice, inject the animals with 0.2 ml per animal; inject rats with 0.5 ml of sRBCs per animal (see Note 5). 8. After the rats are appropriately warmed and the tail veins are easily identified, remove the rats and place into the “White’s Rat Injection Box (Box)” (see Note 6). 9. Use a new, 25 gauge needle for mice, 23 gauge needle for rats for each injection. 10. Attach the needles to 1 ml syringes to obtain the optimum accuracy of the injection volume (see Note 7). 3.4. Preparation of Spleen Cells

1. On day 4 after sensitization, weigh the mice or rats and sacrifice by using CO2 anesthesia or using a procedure acceptable to your IACUC. 2. Remove, trim, and weigh spleens, placing them in approximately 3 ml of EBSS with HEPES in a 5 ml plastic capped test tube for mice and in approximately 6 ml of EBSS with HEPES in a 15 ml plastic capped test tube for rats. 3. Prepare spleen suspensions by first pouring the contents of the spleen tube into a Petri dish, and then pressing the spleen between the frosted ends of two microscope slides into the dish until a single cell suspension is obtained (see Note 8). 4. Wash slides with the tube’s buffer using a Pasteur pipette. 5. Transfer the suspension into a 5 ml plastic capped test tube for mice and the 15 ml tube for rats (see Note 9). 6. Centrifuge the cell suspension at 1,200 ± 100 rpm (350 × g) for approximately 10 min. 7. Resuspend in 3 ml of cold EBSS with HEPES for mice and 6 ml of cold EBSS with HEPES for rats.

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8. Hold samples on ice for assay and determination of cell number. 9. Determine the spleen cell numbers for each animal by counting on a Coulter Counter while the Petri dishes are being incubated. 3.5. Plaquing Procedures

1. In conducting the actual plaquing of the samples, add the following components together in a 12 × 75 mm borosilicate glass tube: agar-dextran solution, sheep erythrocytes, complement, and spleen cells. 2. Prepare a working agar solution containing 0.5% Bacto-Agar and 0.05% DEAE-dextran in EBSS without HEPES the morning of the assay. 3. Prepare by adding the agar (0.5 g per 100 ml) to the EBSS without HEPES and dissolving while heating to a boil (Do Not Reboil). The heating can be done with a Bunsen burner or a microwave (the latter being the safer of the two). 4. Add the DEAE-dextran to the agar (1.6 ml stock solution per 100 ml) (see Note 10). 5. Dispense the warm agar in 0.50 ml aliquots into 12 × 75 mm disposable glass tubes that are held in a 47 ± 1°C in the water bath. 6. Prepare sheep erythrocytes and add to the tubes in the water bath containing the liquid agar-dextran solution. The sRBCs are prepared as described in Subheading 12.3.2.7 above, except after the last centrifugation of cells the supernatant is removed and a volume of EBSS with HEPES equal to the volume of packed cells is added (see Note 11). 7. From the sRBC solution, add 25 µl to each agar-dextran containing tube. The sRBCs should not be added to more than 16 tubes at a time. 8. Prepare the spleen cell suspensions at two dilutions for the IgM assay. For the mouse a 1:30 dilution is prepared as is a 1:120 dilution in cold EBSS with HEPES. For rats, both a 1:50 dilution and a 1:150 dilution are prepared using the same cold media. 9. Hold all diluted samples on ice. 10. From the appropriate dilution tube, add 0.1 ml of the test cells to the test tubes in the water bath. 11. Remove the test tube from the water bath. 12. Quickly add 25 µl of appropriately diluted GPC. 13. Gently vortex the tube. 14. Pour into a 100 × 15 mm Petri plate (see Note 12).

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15. Quickly cover the agar spot with a 45 × 50 mm cover slip. 16. Set up one sample dilution on the top and the other dilution at the bottom of the plate (see Note 13). 17. When the agar is solid (approximately 5–10 min), gently put the plates back together. 18. Incubate the plates at 37 ± 1°C for approximately 3 h in a non-CO2 incubator. 19. Count the plaques using the Bellco Plaque Viewer. 20. Record the results on the appropriate forms. The plates may be stored in the refrigerator overnight for any necessary recounts. 21. Count either dilution plate, making the choice by observing which plate has approximately 100–300 plaques (see Note 14). 22. Document that the results are the result of an average of the two plates after adjusting for the proper dilution factor. 23. At a minimum, record the following parameters for the assay: body weight of the rodents on day of the assay, spleen weight, cell count of original suspension, dilution used to obtain IgM plaques, plaque count, water bath and incubator temperatures and incubation time. 24. Report the results of the assay as both Specific Activity and Total Spleen Activity (see Note 15).

4. Notes 1. Each lot of GPC needs to be evaluated to establish the appropriate dilution to use in the TDAR. Once the lypholized complement is reconstituted and proper dilution is established, it should be aliquoted into appropriate volumes for assay use and stored at −20°C. The reconstituted complement is good for 2 months when stored in this manner. 2. The source of sRBCs used in the TDAR is critically important. Most laboratories purchase the sRBCs from a commercial vendor. It has been our experience that the age of the animal and the number of times the animal has recently been bled can affect the antigenicity of the sRBCs. We routinely screen several sheep to identify one which produces a TDAR response consistent with our historical controls. When blood is collected from the sheep, the blood should be drawn using aseptic techniques to insure the sterility of the blood which will be injected into the animals. Routine sterility testing of the blood received from the vendor is a good laboratory practice.

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3. The number of sRBCs administered to mice and rats, to obtain a response, will differ. In addition, different strains of mice and rats will also differ in the optimum number of sRBCs to be used for sensitization. Routinely, we have found that 7.5 × 107 sRBCs administered i.v. (0.2 ml of 3.75 × 108 cells/ ml) is optimum for B6C3F1 mice. For ICR mice we use 1 × 108 sRBC/mouse administered i.v. (0.2 ml of 5 × 108 cells/ml). The concentration we use in Sprague Dawley rats is 2 × 108 administered in a volume of 0.5 ml. As indicated above, a sRBC cell number response evaluation should be conducted to establish the appropriate concentration for the strain and species you are using in your study. 4. When setting up for warming animals, be sure to place a barrier, i.e., cardboard, surgical blue pads or lab counter top paper under the heating pads, between the bottom of the heating pads and the table or cart, to reduce heat loss which is absorbed by the table or cart. This is especially critical if the cages are placed on metal tables or carts which can absorb heat more efficiently than the plastic cages which can result in insufficient heating of the animals. 5. It is much easier to inject the lateral tail veins instead of the dorsal vein. We have one injector that begins on one side, starting from the distal end of the lateral tail vein and moves forward toward the base of the tail, if unsuccessful in initial injection attempts. If the animal is not successfully injected, a different injector will sensitize the animal using the lateral vein on the other side of the tail. As a last resort, we will inject in the dorsal vein, however, this requires deeper penetration, a different angle of injection, and a more highly skilled individual. 6. This box is a simple inexpensive plastic Plano® fishing tackle box which costs about $4.00. The box has been modified by cutting out a semicircle on the top edge of one side of the bottom of the box. The cut out section is sanded and smoothed so that it will not cause abrasion of the rat’s tail which is placed through the opening. When the rat is removed from the warming cage and placed in the Box, the top is closed and the animal is in total darkness. It has been our experience that the rats “enjoy” the darkness of the box and remain relatively immobile as they evaluate their new surroundings. During this time, one individual holds the rat’s tail next to the box, as well as the Box itself. The injector, wipes the tail with 70% ethanol to enhance seeing the tail veins and to prevent possible infection from the injection, and the rat is injected with sRBC. The individual holding the tail releases their hold to allow the injected cells to travel up the vein. While applying direct pressure to the site of the injection,

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the animal is then removed from the Box and placed in its appropriate cage. The box is wiped clean of urine and feces, if present, and the next animal is placed in the box for injection. Using this procedure we can inject four rats every minute. 7. Injections are best done before noon so the assay can be conducted in the morning. Also, this is the time the animals are less rambunctious since it is the beginning of their sleep cycle. 8. While this procedure will work for both mice and rat spleens, cutting up the rat spleens before mashing is helpful. Additionally the use of a Stomacher, to prepare rat spleens is highly recommended if the assay is done for a large number of animals. 9. Spleen capsule and connective tissue is left behind by tilting the Petri dish to the side and pipetting off only the splenocytes. 10. The DEAE-Dextran plays a critical role in the assay as it inactivates complement proteins which can inactivate the GPC which is necessary for the lysis of the sheep erythrocytes (2). 11. For example, if the volume of packed cells is 2 ml, then 2 ml of EBSS with HEPES is added to the tube, essentially resulting in a blood solution with a hematocrit of 50%. We have found this to be the optimum procedure to obtain a consistent red background of sRBCs in the assay. 12. It has been our experience that the most efficient way to quantitate the proper dilution of GPC for use in the TDAR, is to purchase a large quantity of the same lot number of the lypholized complement. The lypholized material is good for approximately 1 year, when left un-reconstituted. Reconstitute one of the bottles from the lot and conduct a plaque assay using different dilutions of the guinea pig serum. Once the appropriate dilution is determined, when a lypholized bottle is reconstituted, it is allocated in the volume needed for a TDAR assay. The reconstituted serum is usable for 2 months after reconstitution when stored at −20°C. 13. We have found that if one “preheats” the cover slip by bringing it to your mouth and “huffing” on it, that is exhaling onto the cover slip, that the cover slip produces consistent spreading of the agar spot and a more uniform single cell layer of sRBCs, which serves as the red background for the assay. The clever idea of warming the cover slips by “huffing” on them was that of Thomas Kawabata, Ph.D., during his post-doc period in our laboratory. On a similar point, if the counter top on which the Petri dishes are placed is cold, such as when the assay is conducted in winter near a window, the

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agar may cool too quickly and it does not produce a uniform spreading under the cover slip. 14. Avoid <100 or >300 plaques if possible for accurate counting. If there are more than 300 plaques on the first dilution plate, count the other dilution plate and multiply that number times the appropriate dilution and average the count from the two plates. If there are less than 100 plaques on the more concentrated plate, follow the same procedure as described above. 15. Specific Activity is the number of Antibody Forming Cells (AFC) also referred to as Plaque Forming Cells (PFC) per million spleen cells (AFC/106 Spleen Cells) or (PFC/106 Spleen Cells). In addition, a second parameter, Total Spleen Activity, e.g., AFC/Spleen or PFC/Spleen, is also reported whenever a TDAR is conducted. This is critically important since some compounds, such as low doses of dexamethasone, will not have an effect when evaluated as Specific Activity, but since this corticoid steroid causes a massive decrease in spleen cell number, the immunosuppressive effect is obvious when expressed as Total Spleen Activity. Reporting only either Specific Activity or Total Spleen Activity is an inappropriate way to present the results of the assay. Shown in Fig. 12.1 are typical Plaquing results obtained from the female B6C3F1 mice. As can be seen, following exposure to the aromatic hydrocarbon, 1,2:5,6 Dibenzanthracene, a dose-dependent decrease in both Specific Activity and Total Spleen Activity is observed in the female B6C3F1 mice.

Acknowledgments Supported in part by NIEHS Contract ES 05454. References 1. Jerne NK, Nordin AA, Henry C (1963) The agar plaque technique for recognizing antibody-producing cells. In: Amos B, Koprowski H (eds) Cell-bound antibodies. Wistar Institute Press, Philadelphia, PA, pp 109–125 2. Hubner KF, Gengozian N (1968) Critical variables of the Jerne plaque technique as applied to rodent antibody-forming systems responding to heterologous red cell antigens. J Immunol 102:155–167 3. Luster MI, Munson AE, Thomas PT, Holsapple MP, Fenters JD, White KL Jr, Lauer LD, Germolec DR, Rosenthal GJ, Dean

JH (1988) Development of a testing battery to assess chemical-induced immunotoxicity: National Toxicology Program’s guidelines for immunotoxicity evaluation in mice. Fundam Appl Toxicol 10:2–19 4. White KL Jr, Gennings C, Murray MJ, Dean JH (1994) Summary of an international methods validation study, carried out in nine laboratories, on the immunological assessment of cyclosporine A in the Fischer 344 rat. Toxicol In Vitro 8:957–961 5. Dayan AD, Kuper F, Madsen C, Smialowicz RJ, Smith E, Van Loveren H, Vos JG, White KL Jr

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(1998) Report of validation study of assessment of direct immunotoxicity in the rat. Toxicology 125:183–201 6. Loveless SE, Ladics GS, Smith C, Holsapple MP, Woolhiser MR, White KL Jr, Musgrove DL, Smialowicz RJ, Williams W (2007) Interlaboratory study of the primary antibody response to sheep redblood cells in outbred rodents following exposure to cyclophosphamide or dexamethasome. J Immunotoxicol 4(3):233–238 7. Temple L, Kawabata TT, Munson AE, White KL Jr (1993) Comparison of ELISA and plaque-forming cell assay for measuring the humoral immune response to SRBC in animals treated with benzo(a)pyrene or

cyclophosphamide. Fundam Appl Toxicol 21:412–419 8. Smith HW, Winstead CJ, Stank KK, Halstead BW, Wierda D (2003) A predictive F344 rat immunotoxicology model: cellular parameters combined with humoral response to NP-CgG and KLH. Toxicology 194:129–145 9. Gore ER, Gower J, Kurali E, Sui JL, Bynum J, Ennulat D, Herzyk D (2004) Primary antibody response to keyhole limpet hemocyanin in rat as a model for immunotoxicity evaluation. Toxicology 197:23–35 10. White KL Jr, Sheth CM, Peachee VL (2007) Comparison of primary immune responses to SRBC and KLH in rodents. J Immunotoxicol 4(2):153–158

Chapter 13 The Delayed Type Hypersensitivity Assay Using Protein and Xenogeneic Cell Antigens Rodney R. Dietert, Terry L. Bunn, and Ji-Eun Lee Abstract The delayed-type hypersensitivity (DTH) assay has a lengthy history in immunotoxicity testing since it was one of the original functional assays included in the National Toxicology Program (NTP) immunotoxicology test panel. Based on NTP data analysis, the DTH assay is among the most predictive immunotoxicity tests when included with at least two other immune parameters. The DTH assay has the advantage of being: (1) a useful measure of cell-mediated immunity, (2) an in vivo assay where there is less opportunity for ex vivo confounders and (3) a clinically significant human correlate to the tuberculin test. Disadvantages of the DTH assay are that it is potentially labor-intensive to perform, it is somewhat resistant to automation and, when compared with the cyctotoxic T lymphocyte (CTL) assay, it is a relatively crude measurement. However, some groups have been attempting to address the limitations of the DTH assays (see Note 1). The assay is related to the contact hypersensitivity response (CHR), which is covered in another chapter. The DTH response has been used as an indicator of cell-mediated immune status and is dependent upon both T helper 1(Th1)-driven responses as well as cell recruitment and chemotaxis to a local site. As a result, the DTH functional response may be influenced by disruption of either Th1-driven, antigen-dependent T cell development or mobilization of sensitized T cells to a local site. The present chapter describes four common protocols with consideration restricted to protein and xenogeneic cell immunogens. Key words: Delayed type hypersensitivity response, DTH assay, Cell-mediated immunity (CMI), In vivo assay, Risk assessment, Protein antigens, Xenogeneic cells, Sensitization, Elicitation

1. Introduction An important part of immunotoxicity testing involves the capability to assess the potential impact of a xenobiotic on cellmediated immune (CMI) responses. This provides an immune assessment complement to humoral immune activity measurements such as the plaque-forming cell assay. Antigen-specific CMI R.R. Dietert (ed.), Immunotoxicity Testing: Methods and Protocols, Methods in Molecular Biology, vol. 598 DOI 10.1007/978-1-60761-401-2_13, © Humana Press, a part of Springer Science + Business Media, LLC 2010

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responses are key components of host defense against virallyinfected cells, tumor cells and certain fungal infections. In addition to the cytotoxic T lymphocyte (CTL) response, the delayed-type hypersensitivity (DTH) response is one of the assays that has been used for CMI evaluation. The DTH assay, as used in chemical safety evaluation and preclinical drug testing, has the advantage of being a direct in vivo assay with no opportunity for ex vivo confounders to influence the interpretation. This diminishes the need to extrapolate from in vitro- or ex vivo-obtained data back to a whole animal or human to discern the functional ramification. The DTH assay as conducted in test species is a direct animal assay correlate to the human assay used for diagnosis of recent or prior Mycobacterium tuberculosis infection. The human DTH diagnostic test is called the tuberculin test or, alternatively, the Mantoux test (1). DTH assessment in humans extends well beyond the detection of tuberculosis since the DTH assay has been advocated as an effective measure of secondary immune responses in humans (2). For this reason, DTH assessment has been used in humans for a wide range of applications requiring the evaluation and/or monitoring of immune status. These include monitoring of patients with multiple sclerosis during testosterone therapy (3), immune monitoring of the effectiveness of dendritic cell-based anticancer vaccines (4), evaluating vaccination against recurrence of breast cancer (5), and optimizing vitamin A supplementation in children (6). Additionally, the DTH response has been used in humans as a measure of the balance and optimum dosing needed for immunosuppressive therapy among stable transplant patients (7). 1.1. History of the DTH Assay in Immunotoxicity Testing

The DTH assay has been historically used for immunotoxicity testing (8) and was among a seminal functional assay panel evaluated for predictive effectiveness in the risk assessment performed on the National Toxicology Program (NTP) data set (9). One indication of its utility, when used in combination with the other immune parameters, was shown in the NTP data analysis. Eight combinations of three immune functional assays produced 100% concordance in predicting immunotoxicity. Of these eight, the DTH response was a component of seven of the combinations. By comparison, the plaque forming cell (PFC) assay was a component of 4 of the 8 three-way combinations (9). These results suggest that the DTH assay, or more generally a CMI assay, can be very effective for predicting immunotoxicity particularly when it is used as a component of multi-functional immunotoxicity evaluation.

1.2. Nature of the DTH Response

The DTH response is largely a T helper 1 (Th1) functionallydriven CMI response that is also classically known as the type IV hypersensitivity reaction. The overall response requires (1)

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antigen-specific priming or sensitization of T lymphocytes and (2) elicitation of an inflammatory response to the distal challenge site such as a rat foot pad resulting in induration of the challenge site. The DTH response involves the recruitment of several different leukocyte populations including T lymphocytes, as well as dendritic cells, macrophages and neutrophils to the local site of antigenic challenge. Upon challenge, the earliest waves of infiltrated cells are usually neutrophils, but the resultant cellular infiltrate is composed primarily of T lymphocytes and macrophages with some variation among species. Chemokines are important in the elicitation response and include interleukin-8, monocyte chemotractant protein-1, macrophage inflammatory protein 1 alpha, tumor necrosis factoralpha, and granulocyte–macrophage colony-stimulating factor (10). Of particular importance in the response are effector memory T cells. Doebis et al. (11) showed that both P- and L-selectins are involved in the multi-cellular infiltrates characteristic of the DTH response. However, the selectin-associated binding of Th1 cells was most critical to the inflammation process and most significantly determined the extent of foot pad swelling in a rodent DTH response. Suppression of the DTH response may or may not imply that Th1 function is suppressed. The DTH response, as usually evaluated, requires effective cell trafficking and chemotaxis to distal reaction sites. Some xenobiotics appear capable of suppressing the magnitude of the DTH response by interfering with the chemotactic migration of the antigen-primed T lymphocytes rather than by suppressing the early steps in Th1 priming (12). It is important to recognize that immunotoxic modulation of the DTH response can be targeted at effector cells or may involve cells that can regulate this response. Among those are the T regulatory cells (Tregs). Modulation of these cell populations may also exert an influence on the DTH response. For example, control of the DTH response is influenced by CD8+ Tregs (13), and the DTH response can be suppressed by CD4+ CD25 low adaptive Tregs to induce transplant tolerance (14). 1.3. Examples of DTH Assay Applications

The DTH assay has been employed for a wide range of immunotoxicity testing including both adult and early-life environmental exposures. It has been used in a variety of species including the mouse (15), rat (16, 17), dog (18), chicken (19), swine (20), monkey (21, 22), and human (23). Of note, either suppressed DTH responses or enhanced DTH responses can be important in determining the potential adverse outcomes following xenobiotic exposures. Examples where depressed DTH responses were observed include exposures to 2,3,7,8-tetrachlorodibenozo-pdioxin (24), acrylamide (25), atrazine (26), cadmium (27), dexamethasone (28), dihydrocucurbitacin B (29), and lead (30, 31).

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In contrast, elevated DTH responses were seen in at least one gender after exposure to aminoglutethemide (27), cyclophosphamide (32, 33), and trichloroethylene (34). The following sections provide four examples of DTH protocols used in immunotoxicity assessment. The examples are drawn from the mouse, rat, and chicken, and include the use of sheep red blood cells (SRBCs), keyhole limpet hemocyanin (KLH), and bovine serum albumin (BSA) as antigens.

2. Materials 2.1. SRBCs Used in Mice

1. Commercially bred mice. 2. Sheep erythrocytes (SRBCs) in Alsever’s solution (Cambrex BioScience, Walkersville, MD). 3. Phosphate buffered saline (PBS) (Cellgro, Mediatech, Herndon, VA). 4. Spring loaded calipers (The Dyer Company, Lancaster, PA) to measure foot pad swelling.

2.2. KLH Used in Rats Without Adjuvant

1. Commercially bred juvenile, young, or older adult rats. 2. Keyhole limpet hemocyanin (KLH) (Calbiochem, La Jolla, CA). 3. PBS (Sigma Chemical Co., St. Louis, MO). 4. Spring loaded calipers (The Dyer Company, Lancaster, PA). 5. Water bath.

2.3. BSA Used in Rats and Mice with Adjuvant

This protocol from the Immunotoxicology Division of the EPA Health Effects Research has been used with several strains of rats and mice. 1. Bovine serum albumin (BSA) (Fraction V, Sigma Chemical Co., St. Louis). 2. Freund’s Complete Adjuvant (FCA) Distributions, Inc., Kansas City, MO).

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3. 0.9% injectable sodium chloride (Abbott Labs, North Chicago, IL). 4. Isoflurane (Mallinckrodt Specialty Chemicals Co., St. Louis, MO) as an inhalent anesthetic. 5. Electronic calipers (Model Lab of the National Health and Environmental Effects Research Laboratory of the U.S. Environmental Protections Agency, Research Triangle Park, NC). 6. Inhalant chamber with a cone mask and a vaporizer are used for anesthesia. 7. Water bath.

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1. Commercially bred chickens. 2. BSA (Sigma Chemical Co., St. Louis (Fraction V)). 3. PBS (Sigma Chemical Co., St. Louis, MO). 4. Spring loaded calipers (The Dyer Company, Lancaster, PA). 5. Water bath.

3. Methods The methods vary based primarily on the number of immuni zations prior to challenge. The SRBC and BSA with adjuvant applications use a single sensitization injection. The KLH and BSA procedures without adjuvant employ two immunizations separated by a week. 3.1. Method 1 SRBCs Used in Mice

The procedure follows that of Peden-Adams et al. (34) similar to Kim (35). 1. Prepare a suspension of SRBCs in PBS at a concentration of 2 × 107 cells/mL. 2. Inject mice subcutaneously (s.c.) with 0.1 mL of the SRBC suspension. 3. Four days later, prepare a suspension of SRBCs at 4 × 109 cells/ mL in PBS. 4. Anesthetize mice with metafane. 5. Inject 0.025 mL of the SRBC suspension into the right rear footpad and inject 0.025 mL of PBS into the left rear footpad as the control. 6. 24 h later, measure the swelling in each footpad using springloaded calipers. 7. Use the difference between control vs. antigen injected swelling as the indicator of the antigen-specific response.

3.2. Method 2 KLH in Rats Without Adjuvant

This protocol follows that of Exon et al. (36). Note that adjuvant is not employed, and two sensitizing immunizations are given since this can also be used in protocols to obtain antigen-specific antibody quantitation. A similar protocol was used by Chen et al. (12) and Bunn et al. (37) (see Notes 2 and 3). 1. Prepare a solution of KLH in sterile water at a concentration of 5 mg/mL. 2. Cover the container with tin foil and stir continuously for 1 h prior to use. 3. Inject 0.2 mL of 5 mg/mL KLH solution into the caudal tail fold on days 1 and 8. 4. Inject the same volume of sterile water into control rats.

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5. On day 14, prepare KLH at a concentration of 20 mg/mL in PBS and heat at 80ºC for 1 h. 6. After the solution has cooled, load syringes and inject 0.1 mL of the heat aggregated KHL into the right footpad. Inject 0.1 mL of PBS into the left footpad to serve as a reaction control. 7. After a 24-h interval, measure both footpads with spring loaded calipers. The difference in the swelling between the KLH injected and PBS injected footpads provides an indication of the DTH response. 3.3. Method 3 BSA in Rats and Mice with Adjuvant 3.3.1. BSA with Adjuvant in Rats

This procedure is based on the methods in Henningsen et al. (38) and Gehrs et al. (24) for the rat. A similar protocol was used by Gerhs and Smilaowicz (39), and a modification of the procedure was used by Rooney et al. (26). In mice, a similar protocol was used by DeWitt et al. (40). 1. Dissolve 10 mg of BSA in 5 mL of saline and emulsify 1:1 in Freund’s Complete Adjuvant using two luer-lock syringes equipped with a female–female interlocked adapter. 2. Store the resulting emulsification (1 mg/mL BSA) at 4ºC for up to 24 h. 3. Anesthetize the animal and then inject 0.1 mL (0.1 mg BSA/ rat) s.c. into the caudal tail fold of a rat for sensitization. 4. A week later, dissolve 200 mg of crystalline BSA in 10 mL saline creating a 2% solution and heat in a water bath at 75ºC for 1 h with intermittent stirring. 5. Wash the gelatinous BSA in saline and recollect by centrifugation at 450×g for 10 min. 6. Store the heat aggregated BSA overnight at 4ºC. 7. Challenge the previously sensitized rat by s.c. injection in the center of the right hind footpad with 0.1 mL of heat aggregated BSA in saline. Orient the foot for injection such that the foot is facing the researcher and the bevel of the needle is up. 8. Inject physiological saline (0.1 mL) s.c. using the same orientation into the middle of the left hind footpad. 9. 24 h later, measure the thickness of each footpad with electronic calipers (see Note 4). 10. Determine the difference in thickness between the control and the antigen-challenged footpads.

3.3.2. BSA with Adjuvant in Mice

1. Dissolve 20 mg BSA in 5 mL of saline and emulsify 1:1 in Freund’s Complete Adjuvant using two luer-lock syringes equipped with a female–female interlocked adapter.

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2. Store the resulting emulsification (2 mg/mL BSA) at 4ºC for up to 24 h. 3. Anesthetize the animal and inject 0.05 mL (0.1 mg BSA/ mouse) s.c. into the caudal tail fold of a mouse for sensitization. 4. A week later, dissolve 800 mg of BSA in 10 mL saline creating an 8% solution. 5. Heat in a water bath at 75ºC for 1 h with intermittent stirring. 6. Wash the gelatinous BSA in saline and recollect by centrifugation at 450×g for 10 min. 7. Store the heat aggregated BSA overnight at 4ºC. 8. Challenge the previously sensitized mouse by s.c. injection in the center of the right hind footpad with 0.025 mL of heat aggregated BSA in saline. Ensure that for injection, the foot is facing the researcher and the bevel of the needle is up. 9. Inject physiological saline (0.025 mL) s.c. using the same orientation into the middle of the left hind footpad of the mouse. 10. 24 h later, measure the thickness of each footpad with electronic calipers (see Note 4). 11. Determine the difference in thickness between control and antigen-challenged footpads. 3.4. Method 4 BSA in Juvenile Chickens

This protocol follows Lee et al. (41). Note that the wing web can be used for DTH assessment in both juvenile and adult chickens. In older chickens (e.g., 8 weeks or older), the wattle is also used as a convenient site for DTH challenge. 1. Inject s.c. near the breast muscle 0.1 mL of a 20 mg/mL solution of BSA in sterile water to 3 week old chickens. 2. Give a second identical injection 1 week later. 3. Perform a preinjection web measurement on day 14. 4. Inject the left wing web with 0.1 mL of heat aggregated (80ºC 1 h) BSA in PBS (20 mg/mL). 5. Inject the right wing web with 0.1 mL of PBS as the control. 6. 24 h after injection (Day 15), measure the wing web thickness using the same spring loaded calipers. 7. Evaluate antigen-specific DTH by subtracting the ratio of the postinjection PBS web thickness divided by the preinjection PBS web thicknesses from the ratio of the postinjection BSA web thickness divided by the preinjection BSA web thicknesses.

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4. Notes 1. It should be noted that the opportunities for potential automation of the DTH assessment are being examined by several groups (as discussed by Price et al. (10)), and this may further enhance the utility of the assay in immunotoxicity testing panels. 2. While several of the previously-described DTH protocols were conducted using juvenile animals or young adult animals, the same procedures have been applied to older adult animals as well. 3. Several of the protocols that did not describe rodent anesthetization at the time of final DTH measurement were performed at the termination of experiments immediately following euthanasia. 4. The basis for using electronic calipers in recent procedures was the usefulness of reducing potential pressure exerted by the spring-loaded calipers during the measurement of swelling.

Acknowledgments The authors thank Dr. Robert Luebke of the U.S. Environmental Protection Agency Health Effects Research Laboratory for providing details on his DTH protocols. References 1. Hill PC, Jackson-Sillah DJ, Fox A, Brookes RH, de Jong BC, Lugos MD, Adetifa IM, Donkor SA, Aiken AM, Howie SR, Corrah T, McAdam KP, Adegbola RA (2008) Incidence of tuberculosis and the predictive value of ELISPOT and Mantoux tests in Gambian case contacts. PLoS One 3:e1379 2. Vukmanovic-Stejic M, Reed JR, Lacy KE, Rustin MH, Akbar AN (2006) Mantoux test as a model for a secondary immune response in humans. Immunol Lett 107:93–101 3. Gold SM, Chalifoux S, Giesser BS, Voskuhl RR (2008) Immune modulation and increased neurotrophic factor production in multiple sclerosis patients treated with testosterone. J Neuroinflammation 5:32 4. Aarntzen EH, Figdor CG, Adema GJ, Punt CJ, de Vries IJ (2008) Dendritic cell vaccination and immune monitoring. Cancer Immunol Immunother 57:1559–1568

5. Holme JP, Gates JD, Benavides LC, Hueman MT, Carmichael MG, Patil R, Craig D, Mittendorf EA, Stajaninovic A, Ponniah S, Peoples GE (2008) Optimal dose and schedule of an HER-2/neu (E75) peptide vaccine to prevent breast cancer recurrence: from US Military Cancer Institute Clinical Trials Group Study I-01 and I-02. Cancer 113:1666–1675 6. Diness BR, Fisker AB, Roth A, Yazdanbakhsh M, Sartono E, Whittle H, Nante JE, Lisse IM, Ravn H, Rodrigues A, Aaby P, Benn CS (2008) Effect of high-dose vitamin A supplementation on the immune response to Bacille Calmette–Guerin vaccine. Am J Clin Nutr 86:1152–1159 7. Van Besouw NM, van der Mast BJ, van der Wetering J, Rischen-Vos J, Weimar W (2008) Tapering immunosuppressive therapy significantly improves in vivo cutaneous delayed type hypersensitivity responses. Transplant Immunol 19:229–234

The Delayed Type Hypersensitivity Assay Using Protein and Xenogeneic Cell Antigens 8. Faith RE, Luster MI, Kimmel CA (1979) Effect of chronic developmental lead exposure on cell-mediated immune functions. Clin Exp Immunol 35:413–420 9. Luster MI, Portier C, Pait DG, White KL Jr, Gennings C, Munson AE, Rosenthal GJ (1992) Risk assessment in immunotoxicology. I. Sensitivity and predictability of immune tests. Fundam Appl Toxicol 18:200–210 10. Price K (2008) Chapter 3.1.3 Cellular immune response in delayed type hypersensitivity tests. In: Herzyk DJ, Bussier JL (eds) Immunotoxi cology strategies for pharmaceutical assessment. Wiley, Hoboken, NJ, pp 87–101 11. Doebis C, Siegmund K, Loddenkemper C, Lowe JB, Issekutz AC, Hamann A, Huehn J, Syrbe U (2008) Cellular players and role of selectin ligands in leukocyte recruitment in a T-cell-initiated delayed-type hypersensitivity reaction. Am J Pathol 173:1067–1076 12. Chen S, Golemboski KA, Sanders FS, Dietert RR (1999) Persistent effect of in utero meso2,3-dimercaptosuccinic acid (DMSA) on immune function and lead-induced immunotoxicity. Toxicology 132:67–79 13. Cone RE, Chattopadhyay S, O’Rourke J (2008) Control of delayed-type hypersensitivity by ocular-induced CD8+ regulatory T cells. Chem Immunol Allergy 94:138–149 14. Xu Q, Lee J, Jankowska-Gan E, Schultz J, Roenneburg DA, Haynes LD, Kusaka S, Sollinger HW, Knechtle SJ, VanBuskirk AM, Tottealba JR, Burlingham WJ (2007) Human CD4+ CD25 low adaptive T regulatory cells suppress delayed-type hypersensitivity during transplant tolerance. J Immunol 178: 3983–3995 15. Ohga K, Takezawa R, Arakida Y, Shimizu Y, Isikawa J (2008) Characterization of YM-58483/ BTP2, a novel store-operated Ca(2+) entry blocker, on T cell-mediated immune responses in vivo. Int Immunopharmacol 8(13–14): 1787–1792 16. Dewitt JC, Copeland CB, Luebke RW (2007) Immune function is not impaired in SpragueDawley rats exposed to dimethyltin dichloride (DMTC) during development or adulthood. Toxicology 232:303–310 17. Thangasamy T, Subathra M, Sittadjody S, Jeyakumar P, Joyee AG, Mendoza E, Chinnakkanu P (2008) Role of l-carnitine in the modulation of immune response in aged rats. Clin Chim Acta 389:19–24 18. Tamura K, Yamada M, Isotani M, Arai H, Yagihara H, Ono K, Washizu T, Bonkobara M (2008) Induction of dendritic cell-mediated immune responses against canine malignant melanoma cells. Vet J 175:126–129

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19. Minozzi G, Parmentier HK, Bedhom B, Mivielle F, Gourichon D, Pinard-Vander Laan MH (2008) Delayed-type hypersensitivity response to KLH in F2 and backcrosses of two immune selected chicken lines: effect of immunisation and selection. Dev Biol (Basel) 132:267–270 20. de Groot J, de Jong IC, Prelle IT, Koolhaas JM (2002) Immunity in barren and enriched housed pigs differing in baseline cortisol concentration. Physiol Behav 71:217–223 21. Bleavins MR, de la Inglesia FA (1995) Cynomolgus monkeys (Macaca fascicularis) in preclinical immune function safety testing: development of a delayed-type hypersensitivity procedure. Toxicology 95:103–112 22. Martin PL, Oneda S, Treacy G (2007) Effects of an anti-TNF-alpha monoclonal antibody, administered throughout pregnancy and lactation, on the development of the macaque immune system. Am J Reprod Immunol 58:138–149 23. Mathew S, Bauer KL, Fischoeder A, Bhardwaj N, Oliver SJ (2008) The anergic state in sarcoidosis is associated with diminished dendritic cell function. J Immunol 181:746–755 24. Gehrs BC, Riddle MM, William WC, Smialowicz RJ (1997) Alterations in the developing immune system of the F344 rat after perinatal exposure to 2,3,7,8-tetrachlorodibenzo-p-dioxin: II. Effects on the pup and the adult. Toxicology 122:229–240 25. Zaidi SI, Raisuddin S, Singh KP, Jafri A, Husain R, Husain MM, Mall SA, Seth PH, Ray PK (1994) Acrylamide induced immunosuppression in rats and its modulation by 6-MFA, an interferon inducer. Immunopharmacol Immunotoxicol 16:247–260 26. Rooney AA, Matulka RA, Luebke RW (2003) Developmental atrazine exposure suppresses immune function in male, but not female Sprague-Dawley rats. Toxicol Sci 76:366–375 27. Lall SB, Dan G (1999) Role of corticosteroids in cadmium induced immunotoxicity. Drug Chem Toxicol 22:401–409 28. Dietert RR, Lee JE, Olsen J, Fitch K, Marsh JA (2003) Developmental immunotoxicity of dexamethasone: comparison of fetal versus adult exposures. Toxicology 194:163–176 29. Escandell JM, Recio MC, Manez S, Giner RM, Cerda-Nicolas M, Gil-Benso R, Rios JL (2007) Dihydrocucurbitacin B inhibits delayed type hypersensitivity reactions by suppressing lymphocyte proliferation. J Pharmacol Exp Ther 322:1261–1268 30. Miller TE, Golemboski KA, Ha RS, Bunn T, Sanders FS, Dietert RR (1998) Developmental

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exposure to lead causes persistent immunotoxicity in Fischer 344 rats. Toxicol Sci 42:129–135 31. McCabe MJ Jr, Singh KP, Reiners JJ Jr (1999) Lead intoxication impairs the generation of a delayed type hypersensitivity response. Toxicology 139:255–264 32. Ben Efraim S (2001) Immunomodulating anticancer alkylating drugs: targets and mechanisms of activity. Curr Drug Targets 2:197–212 33. Brode S, Cooke A (2008) Immunepotentiating effects of the chemotherapeutic drug cyclophosphamide. Crit Rev Immunol 28:109–126 34. Peden-Adams MM, Eudaly JG, Heesemann LM, Smythe J, Miller J, Gilkeson GS, Keil DE (2006) Developmental immunotoxicity of trichloroethylene (TCE): studies in B6C3F1 mice. J Environ Sci Health A Tox Hazard Subst Environ Eng 41:249–271 35. Kim JH (2000) Effect of biphenyl dimethyl dicarboxylate on the cellular and nonspecific immunotoxicity by ethanol in mice. Biol Pharm Bull 23:1206–1211 36. Exon JH, Bussier JL, Mather GG (1990) Immunotoxicity testing in the rat: an improved

multiple assay model. Int J Immunopharmacol 12:699–701 37. Bunn TL, Parsons P, Kao E, Dietert RR (2001) Gender-based profiles of developmental immunotoxicity to lead in the rat: assessment in juveniles and adults. J Toxicol Environ Health A 64:223–240 38. Henningsen GM, Koller LD, Exon JH, Talcott PA, Osborne CA (1984) A sensitive delayedtype hypersensitivity model in the rat for assessing in vivo cell-mediated immunity. J Immunol Methods 70:153–165 39. Gehrs BC, Smialowicz RJ (1999) Persistent suppression of delayed-type hypersensitivity in adult F344 rats after perinatal exposure to 2,3,7,8-tetrachlorodibenzo-p-dioxin. Toxicology 134:79–88 40. Dewitt JC, Copeland CB, Strynar MJ, Luebke RW (2008) Perfluorooctanoic acid-induced immunomodulation in adult C57BL/6J or C57BL/6N female mice. Environ Health Perspect 116:644–650 41. Lee JE, Chen S, Golemboski KA, Parsons PJ, Dietert RR (2001) Developmental windows of differential lead-induced immunotoxicity in chickens. Toxicology 156:161–170

Chapter 14 The Cytotoxic T Lymphocyte Assay for Evaluating Cell-Mediated Immune Function Gary R. Burleson, Florence G. Burleson, and Rodney R. Dietert Abstract Evaluation of cell-mediated immunity (CMI) is a significant component in any assessment designed to predict the full range of potential immunotoxic risk underlying health risks. Among measures of CMI, the cytotoxic T Lymphocyte (CTL) response is recognized as perhaps the most relevant functional measure that reflects cell-mediated acquired immune defense against viral infections and cancer. The CTL response against T-dependent antigens requires the cooperation of at least three different major categories of immune cells. These include professional antigen presenting cells (e.g., dendritic cells), CD4+ T helper lymphocytes, and CD8+ T effector lymphocytes. It is also among the few functional responses dependent on and, hence, capable of evaluating effective antigen presentation via both class I and class II molecules of the major histocompatibility complex (MHC). For this reason the CTL assay is an excellent candidate for evaluation of potential immunotoxicity. This chapter provides an example of a mouse CTL assay against influenza virus that has been utilized for this purpose. Key words: Cell-mediated immune function, CMI, Cytotoxic T lymphocyte, CTL, Influenza virus, Natural infection, Mouse

1. Introduction Cell-mediated immunity (CMI) represents a major component of the host response against both intracellular pathogens and tumor cells. Among the most commonly used measures of CMI are the delayed type hypersensitivity (DTH) assay (described in another chapter in this book) and the cytotoxic T lymphocyte (CTL) assay. The CTL response is acquired or adaptive and requires the interaction of at least three major categories of immune cells. These include professional antigen presenting cells such as dendritic cells and/or macrophages, CD4+ T lymphocytes that provide necessary help for response to T-dependent antigens, and R.R. Dietert (ed.), Immunotoxicity Testing: Methods and Protocols, Methods in Molecular Biology, vol. 598 DOI 10.1007/978-1-60761-401-2_14, © Humana Press, a part of Springer Science + Business Media, LLC 2010

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CD8+ T lymphocytes that develop into antigen-specific cytotoxic effector cells. Equally important is the requirement for antigen presentation using both class I and class II molecules of the major histocompatibility complex (MHC) to generate effector CTLs. For this reason, the CTL response is distinguished from the DTH response or the T-dependent antibody response (TDAR) both of which require only class II presentation of antigen. Further differences between the DTH and CTL have been investigated when protein or nonreplicating antigens are studied. Nonreplicating antigens can induce either optimum DTH or antibody-mediated responses depending on the antigen dose. For example, the induction of antibody to an antigen “A” may result in animals specifically unresponsive for the induction of DTH to “A” (1). This cross-regulation does not allow an optimum DTH and humoral antibody response in the same animal, and makes the immunotoxicological analysis of DTH and TDAR in the same animal more difficult with a nonreplicating or protein antigen. This phenomenon has been studied extensively and is referred to as immune deviation (2). Conversely, the induction of DTH to an antigen could result in unresponsiveness for the induction of antibody (3). Mice optimally immunized to produce antibody exhibit no DTH but have antigen-specific CD4+ CD8− T cells that suppress the induction of DTH (4, 5). This population of cells coordinately inhibits the DTH and facilitates the antibody response (i.e., they switch the response induced by antigen from a cellmediated to a humoral mode) (6–8). Mice induced to express a strong DTH response that were unresponsive to the induction of antibody have antigen-specific CD4− CD8+ T cells that inhibit the antibody response (5, 9). Differences between DTH and CTL were also demonstrated in studies with influenza CTL. It should be noted that although DTH is a cell-mediated immune response, DTH and CTL are not synonymous (10). Class I-restricted DTH-mediating T cells generated in mice infected with virus convey protection (11). However, class II-restricted, DTH-mediating T cells generated by immunization with virus augments disease (12). Therefore, T cells exhibiting DTH in viral disease measures both T cells that are beneficial and those associated with immunopathology. Therefore, a decreased DTH may either be adverse (immunotoxicology) or beneficial (immunopharmacology). CTLs use two distinct pathways to kill virally infected cells, tumors cells or allografts. One process involves the exocytosis of granules into the cellular synaptic space (13). The granules contain various granzymes and perforin that compromise the integrity of target cell membranes resulting in necrosis. Numerous components of granules are important in affording protection. For example, for protection against influenza virus infection in the lung, granzyme A, B, and K are all necessary (14).

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A second process involves the induction of apoptosis in target cells via the cell surface expression of Fas-ligand on CTLs (15). Both preformed Fas-ligand and subsequently expressed Fasligand appear to be involved in the CTL action (16). Some viral infection studies suggest that the level of infection may influence the balance of the two CTL killing mechanisms (17). As used to evaluate immunotoxicity, the CTL response is influenced by the functional status of a wide variety of immune and other cell types. Dendritic cell maturation, status, and distribution are all important in the CTL response (18) as is the need for T helper 1 (Th1) conditioning at some point in the process of CD4+ Th cell support (19, 20). Th1 associated cytokines such as interleukin (IL-) 2 interferon-gamma (IFN-gamma), and IL-12 support CTL activity. Natural killer (NK) cells can help to stabilize the needed Th1 polarization to support CTL generation (21). In contrast, a variety of studies suggest that T helper 2 (Th2)-driven responses including the production of cytokines such as IL-4, IL-10, and IL-13 tend to suppress CTL generation and/or activity (22–25). Other immune cell types and cytokines are also important for CTL function. The level of T regulatory cell (Treg) function has a significant impact on the CTL response against both viruses (26) and tumors (27). Th1-dependent cytokines such as IL-12 are critical for CTL generation (28). But other cytokines are important as well. IL-15 can significantly elevate CTL activity (29). IL-18, while not absolutely required for CTL induction, accelerates CTL responses (30). IL-21 is produced by natural killer T lymphocytes (NKT), CD4+ T cells, and Th17 cells (31). This cytokine can expand CD8+ T cells and the level of CTL responses (32, 33) at least in part by reducing the levels of Foxp3 expressing cells (33) and enhancing memory T cell responses (34). Additionally, IL-23 can facilitate CTL production (35) and IL-27 can play a role in elevating the expression of T-bet (a Th1specific T box transcriptional factor), an IL-12 receptor and granzyme B via STAT1 (36). Innate immune and inflammatory status can also impact CTL activity, since redox status has been shown to modulate CTL function (37). Additionally, IL-4 induced production of arginase-1 among myeloid cells can suppress CTL activity (38). Metabolism status can impact CTL generation as many gene expression changes in the early generation of CTL are highly glucose-dependent (39). The CTL assay has been used extensively in immunotoxicology research and immunotoxicity testing evaluation. Various forms of the CTL assay have been used in mice for studies of immunotoxicity induced by cigarette smoke (40), 2,3,7,8-tertrachlorodibenzo-p-dioxin (41–43), N,N-diethylaniline (44), oligonucleotide drugs (45), polychlorinated biphenyls (46), synthetic

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androgens (47), herbicides (48), pesticides (49), mycotoxins (50), ultraviolet radiation (51), tyrosine kinase inhibitors (52), and phosgene inhalation (53). In a risk assessment analysis of the National Toxicology Program (NTP) database concerning the predictability of immune assays in the identification of immunotoxicants, there were 8 three-way combinations of immune parameters that achieved 100% concordance for accurate immunotoxicant identification (54). Of these eight combinations of assays, acquired cell-mediated immune functional measures were included in seven of these eight combinations. In contrast, a humoral measure, the TDAR assay, was represented in four of the successful combinations. This suggests that CMI evaluation is an important component of immunotoxicity testing and that immunotoxicity testing based on a multiparameter approach should include a functional measure of CMI to optimize predictability. This chapter describes the CTL assay as performed on influenza virus-infected mice using spleen cells as the source for CTL evaluation. However, it should be noted that the same assay has been performed using rats as well (53, 55). Additionally, CTL activity within mucosal sites such as the bronchus-associated lymphoid tissue (BALT) or whole-lung homogenate can be evaluated by using collagenase-treated lungs from infected rodents as the source of cells for the CTL assay (53, 55). This feature enables systemic and local CTL activity levels to be compared from the same animals. Finally, this approach can be adapted for use in developmental immunotoxicity testing with assessment in the juvenile or young adult (56). Numerous influenza viruses have been used for immunotoxicity testing and include Influenza A/PR8/34 (H1N1), Influenza A/ Taiwan/1/64 (H2N2), Influenza A/Aichi (H3N2), Influenza A/ HKx31 (H3N2), Influenza A/Hong Kong/8/68 (H3N2), and Influenza A/Port Chalmers/1/73 (H3N2) (reviewed by 55, 57, 58).

2. Materials 1. Mice options: Purchase 7–8 week old male and female mice (Jackson Laboratories, Bar Harbor, MA). (a) C57BL/6 mice (H-2b). (b) BALB/c (H-2d). (c) CBA/H (H-2k). (d) C3HeB/FeJ (H-2k). (e) CBA mice (H-2k).

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2. Virus options: All from American Type Culture Collection, Manassas, VA (a) Influenza A/PR8/34 (H1N1) virus (ATCC # VR-95). (b) Influenza A/Taiwan/1/64 (H2N2) virus (ATCC # VR-295). (c) Influenza A/Hong Kong/8/68 (H3N2) virus (ATCC # VR-544). (d) Influenza A/Port Chalmers/1/73 (H3N2) virus (ATCC # VR-810). 3. Cell culture plastic ware (Greiner Bio-One, Inc, Longwood, FL). 4. 51Chromium: sodium-51Cr-chromate (PerkinElmer Life and Analytical Sciences, Boston, MA). 5. Dulbecco’s Minimum Essential Medium (Invitrogen Life Sciences, Carlsbad, CA).

(D-MEM)

6. Fetal bovine serum (FBS) (Invitrogen Life Sciences, Carlsbad, CA). 7. Target cells: MHC-Class I matched to mouse strain H-2 type (American Type Culture Collection, Manassas, VA). (a) EL-4 (H-2b) target cells (ATCC # TIB-39). (b) P815 (H-2d) target cells (ATCC # TIB-64). (c) L929 (H-2k) target cells (ATCC # CCL-1).

3. Methods 3.1. Animals, Viruses, and Target Cells

1. Acquire mice, 7–8 week old male and female of strain of choice. 2. Acclimate mice 1 week prior to infection. 3. Infect mice intranasally with 50 mL virus (approximately 5 × 105 pfu) under light isoflurane sedation. 4. Purchase appropriate MHC-Class I matched target cells (see Note 1) for the mouse strain.

3.2. Prepare Effector Cells

Prepare single cell suspensions from mice or rats via the use of spleen, lungs, lymph nodes, bone marrow, or Ficoll–Hypaque peripheral blood lymphocytes in prepared D-MEM media. Effector to target ratios (E:T) (see Note 2).

3.3. Target Cell Infection

1. Grow target cells in D-MEM with 10% FBS. 2. Ensure target cells are passaged before use in experiment.

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3. Use freshly passaged target cells (within 2–3 days after passage of cells). 4. Collect cells and resuspend in approximately 3 ml of D-MEM. 5. Perform cell count. 6. Perform a quick thaw of influenza virus. 7. Prepare a mixture of the cells, virus, and media to achieve a final cell concentration of 2 × 106 c/ml and a final virus concentration of 2 × 107 pfu/ml. This will achieve a multiplicity of infection (MOI) of 10 (see Note 3). 8. Add the cell:virus mixture (2–4 ml/well) to a 12 well, flatbottom tissue culture plate. 9. Incubate for 12–24 h at ~37°C/5%CO2 prior to labeling the target cells. 3.4. Labeling of Target Cells

1. Transfer infected target cells into a 15 ml polypropylene centrifuge tube. 2. Centrifuge at 900 × g for 5 min, at room temperature. 3. Remove the supernatant. 4. Add 1 ml of prepared D-MEM media. 5. Aspirate the pellet with a 1 ml pipette 6. Add another 1 ml of the prepared D-MEM media and vortex. 7. Determine viability and perform a cell count. 8. Transfer the desired number of target cells to a 15 ml polypropylene centrifuge tube (see Note 4). 9. Centrifuge the cells at 900 × g for 5 min (at room temperature). 10. Carefully remove the supernatant. 11. Tap the 15 ml tube to loosen the pellet. 12. Resuspend in cells.

51

Chromium at a rate of 100 µCi per 1 × 106

13. Incubate the cells for 90 min at ~37°C/5% CO2 with periodic mixing. 14. Remove the 15 ml tube from the incubator. 15. Add 10 ml of the prepared D-MEM media to the tube. 16. Centrifuge at 900 × g for 5 min (at room temperature). 17. Decant supernatant. 18. Resuspend cells in 10 ml of the prepared D-MEM media. 19. Incubate for 30 additional minutes at 37°C/5% CO2.

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20. Centrifuge cells at 900 × g for 5 min following the 30 min incubation. 21. Resuspend cells in 1 ml of the prepared D-MEM media. 22. Determine viable cell count using a coulter counter, and/or hemacytometer. 23. Resuspend cells to a concentration of 1 × 105 cells/ml. 3.5. CTL Assay

1. Add 100 µl of effector cells to triplicate wells in round bottom microtiter plates. 2. Add 100 µl of labeled target cells (1 × 105 c/ml) to each well for a total of 200 µl per well. 3. Prepare 6–12 wells containing 100 µl of target cells in 100 µl of prepared D-MEM media. These wells will serve as the spontaneous release (S) controls. 4. Prepare 6–12 wells containing 100 µl of target cells plus 100 µl of 0.25% Triton X-100. These wells will serve as the total 51Cr release (T) controls. 5. Centrifuge plates at 250 × g (at room temperature) for 5 min. 6. Incubate plate(s) at ~37°C/5% CO2 for 4 h. 7. Remove plate(s) from incubator. 8. Centrifuge plates at 250 × g (at room temperature) for 5 min. 9. Harvest supernatants by transferring 100 µl–1.0 ml to polystyrene tube strips or polypropylene cluster tubes (see Note 5). 10. Place the tubes (or Skatron collection filters) into 5 ml, 12 × 75 mm style round bottom tubes. 11. Load these tubes into gamma counter cassettes for counting.

3.6. Calculate the Percent CTL Lysis (51Cr release)

Calculate the percentage of CTL Lysis (51Cr release) using the formula [(E − S)/(T − S)] × 100. E is the 51Cr released from target cells in the presence of effector cells, S is the spontaneous release of 51Cr from target cells alone, and T is the maximum release of 51 Cr from target cells in the presence of Triton X-100.

4. Notes 1. Matching target cells to mouse strain. (a) For C57BL/6 mice: H-2b choose virus-specific CTL kill MHC-matched virus-infected EL-4 (H-2b) target cells but not MHC-mismatched mastocytoma P815 (H-2d) target cells.

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(b) For BALB/c mice: H-2d choose virus-specific CTL kill MHC-matched virus-infected P815 (H-2d) target cells but not MHC-mismatched EL-4 (H-2b) target cells. (c) For CBA/H, C3HeB/FeJ and CBA mice: H-2k choose virus-specific CTL kill MHC-matched virus-infected L929 (H-2k) target cells but not MHC-mismatched mastocytoma P815 (H-2d) or EL-4 (H-2b) target cells. 2. Effector to target cell ratios (E:T) may vary (e.g., in order to achieve a 50:1 ratio the effector cells would be adjusted to 5 × 106 cells/ml). 3. A minimum of 2 ml per well will be prepared, but the total volume prepared will depend on the size of the experiment. 4. It is recommended that, for each 96-well assay plate required, 3 × 106 total cells be labeled. 5. Alternatively, the supernatants may be collected using a Skatron Supernatant collection system.

Acknowledgments The authors thank Janice Dietert for her editorial suggestions. It is noted that R.R. Dietert serves as a consultant in developmental immunotoxicology for Burleson Research Technologies, Inc. References 1. Bretscher PA (1981) Significance and mechanisms of the cellular regulation of the immune response. Fed Proc 40:1473–1478 2. Asherson CR, Stone SH (1962) Selective and specific inhibition of 24 hour skin reactions in the guinea pig. I. Immune deviation: description of the phenomenon and the effect of splenectomy. Immunology 9:205–217 3. Parish CR (1972) The relationship between humoral and cell-mediated immunity. Transplant Rev 13:35–66 4. Ramshaw IA, Bretscher PA, Parish CR (1976) Regulation of the immune response. I. Suppression of delayed-type hypersensitivity by T cells from mice expressing humoral immunity. Eur J Immunol 6:674–679 5. Ramshaw IA, Bretscher PA, McKenzie IFC (1977) Discrimination of suppressor T cells of humoral and cell-mediated immunity by anti-Ly and anti-Ia sera. Cell Immunol 31:364–369 6. Bretscher PA (1983) In vitro analysis of the cellular interactions between unprimed lymphocytes responsible for determining the class of response an antigen induces: specific T cells

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switch a cell-mediated response to a humoral response. J Immunol 131:1103–1107 Bretscher PA (1983) Regulation of the immune response induced by antigen. I. Specific T cells switch the in vivo immune response from a cell-mediated to a humoral mode. Cell Immunol 81:345–356 Tuttosi S, Bretscher PA (1992) Antigen-specific CD8+ T cells switch the immune response induced by antigen from an IgG to a cell-mediated mode. J Immunol 148:397–403 Ramshaw IA, Bretscher PA, Parish CR (1977) Regulation of the immune response. II. Repressor T cells in cyclophosphamideinduced tolerant mice. Eur J Immunol 7:180–185 Yap KL, Ada GL (1978) Transfer of specific cytotoxic T lymphocytes protects mice inoculated with influenza virus. Nature 273:238–239 Leung KN, Ada GL (1981) Effect of helper cells on the primary in vitro production of delayed-type hypersensitivity to influenza virus. J Exp Med 153:1029–1043

The Cytotoxic T Lymphocyte Assay for Evaluating Cell-Mediated Immune Function 12. Leung KN, Ashman RB, Ertl HCJ, Ada GL (1980) Selective suppression of the cytotoxic T cell response to influenza infection in mice. Eur J Immunol 10:803–810 13. Menasche G, Menager MM, Lefebvre JM, Deutsche E, Athman R, Lambert N, Mahlaoui N, Court M, Garin J, Fischer A, de Saint Basile G (2008) A newly identified isoform of Slp2a associates with Rab27a in cytotoxic T cells and participates to cytotoxic granule secretion. Blood 112:5052–5062 14. Jenkins MR, Trapani JA, Doherty PC, Turner SJ (2008) Granzyme K expressing cytotoxic T lymphocytes protects against influenza virus in granzyme AB-/- mice. Viral Immunol 21:341–346 15. Zelinskyy G, Balkow S, Schimmer S, Schepers K, Simon MM, Dittmer U (2004) Independent roles of perforin, granzymes, and Fas in the control of Friend retrovirus infection. Virology 330:365–374 16. He JS, Ostergaard HL (2007) CTLs contain and use intracellular stores of FasL distinct from cytolytic granules. J Immunol 179:2339–2348 17. Zelinskyy G, Balkow S, Schimmer S, Werner T, Simon MM, Dittmer U (2007) The level of friend retrovirus replication determines the cytolytic pathway of CD8+ T-cell-mediated pathogen control. J Virol 81:11881–11890 18. Pascual DW, Wang X, Kochetkova I, Callis G, Richardi C (2008) The absence of lymphoid CD8+ dendritic cell maturation in L-selectin-/- respiratory compartment attenuates antiviral immunity. J Immunol 181:1345–1356 19. Sato M, Chamoto K, Tsuji T, Iwakura Y, Togashi Y, Koda T, Nishimura T (2001) Th1 cytokine-conditioned bone marrow-derived dendritic cells can bypass the requirement for Th functions during the generation of CD8+ CTL. J Immunol 167:3687–3691 20. Chattopadhyay S, Chakraborty NG (2005) Continuous presence of Th1 conditions is necessary for longer lasting tumor-specific CTL activity in stimulation cultures with PBL. Hum Immunol 66:884–891 21. Mailliard RB, Son YI, Redlinger R, Coates PT, Giermasz A, Morel PA, Storkus WJ, Kalinski P (2003) Dendritic cells mediate NK cell help for Th1 and CTL responses: twosignal requirement for the induction of NK cell helper function. J Immunol 171:2366–2373 22. Chavin KD, Qin L, Yon R, Lin J, Yagita H, Bromberg JS (1994) Anti-CD2 mAbs suppress cytotoxic lymphocyte activity by the generation of Th2 suppressor cells and receptor blockade. J Immunol 152:3729–3739

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23. Terabe M, Park JM, Berzofky JA (2004) Role of IL-13 in regulation of anti-tumor immunity and tumor growth. Cancer Immunol Immunother 53:79–85 24. Bachuk N, Utermohlen O, Gugel R, Warnecke G, Karow U, Paulsen D, Brombacher F, Kronke M, Deppert W (2007) Interleukin-4 impairs granzyme-mediated cytotoxicity of Simian virus 40 large tumor antigen-specific CTL in BALB/c mice. Cancer Immunol Immunother 56:1625–1636 25. Nishimura M, Sato H, Okazaki H, Satake M, Tadokoro K (2008) Interleukin-10 containing normal human serum inhibits granzyme B release but not perforin release from alloreactive and EBV-specific T cell clones. Cell Immunol 251:31–36 26. Fernendez MA, Puttur FK, Wang YM, Howden W, Alexander SI, Jones CA (2008) T regulatory cells contribute to the attenuated primary CD8+ and CD4+ T cell responses to herpes simplex virus type 2 in neonatal mice. J Immunol 180:1556–1564 27. Xu L, Xu W, Jiang Z, Zhang F, Chu Y, Xiong S (2009) Depletion of CD4(+)CD25(high) regulatory T cells from tumor infiltrating lymphocytes predominantly induces Th1 type immune response in vivo which inhibits tumor growth in adoptive immunotherapy. Cancer Biol Ther 8(1):66–72 28. Chen J, Wang J, Li J, Wu Q, Chu Lim F, Yang P, Hsu HC, Curiel DT, Mountz JD (2008) Enhancement of cytotoxic T-lymphocyte response in aged mice by a novel treatment with recombinant AdIL-12 and wild-type adenovirus in rapid succession. Mol Ther 16:1500–1506 29. Tang F, Zhao LT, Jiang Y, de Ba N, Chi LD, He W (2008) Activity of recombinant human interleukin-15 against tumor recurrence and metastasis in mice. Cell Mol Immunol 5:189–196 30. Itoh H, Fujimora Y, Tsutsui H, Matsui K, Sugihara A, Terada N, Hada T, Kakishita E, Okamura H, Hara N, Nakanishi K (2004) Involvement of interleukin-18 in acute graftversus-host disease in mice. Transplantation 78:1245–1250 31. Spolski R, Warren WJ (2008) The Yin and Yang of interleukin-21 in allergy, autoimmunity and cancer. Curr Opin Immunol 20:295–301 32. Ansen S, Butler MO, Berezovskaya A, Murray AP, Stevenson K, Nadler LM, Hirano N (2008) Dissociation of its opposing immunologic effects is critical for the optimization of antitumor CD8+ T-cell responses induced by interleukin 21. Clin Cancer Res 14:6125–6136

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33. Li Y, Yee C (2008) IL-21 mediated Foxp3 suppression leads to enhanced generation of antigen-specific CD8+ cytotoxic T lymphocytes. Blood 111:229–235 34. Smyth MJ, Hayakawa Y, Creney E, Zerafa N, Sivakumar P, Yagita H, Takeda K (2006) IL-21 enhances tumor-specific CTL induction by anti-DR5 antibody therapy. J Immunol 176:6347–6355 35. Ha SJ, Kim DJ, Baek KH, Yun YD, Sung YC (2004) IL-23 induces stronger sustained CTL and Th1 immune responses than IL-12 in hepatitis C virus envelope protein 2 DNA immunization. J Immunol 172:525–531 36. Morishima N, Owaki T, Asakawa M, Kamiya S, Mizuguchi J, Yoshimoto T (2005) Augmentation of effector CD8+ T cell generation with enhanced granzyme B expression by IL-27. J Immunol 175:1686–1693 37. Sklavos MM, Tse HM, Piganelli JD (2008) Redox modulation inhibits CD8 T cell effector function. Free Radic Biol Med 45:1477–1486 38. Bronte V, Serafini P, De Santo C, Marigo I, Tosello V, Mazzoni A, Segal DM, Staib C, Lowel M, Sutter G, Colombo MP, Zanovello P (2003) IL-4-induced arginase 1 suppresses alloreactive T cells in tumor-bearing mice. J Immunol 170:270–278 39. Cham CM, Driessens G, O’Keefe JP, Gajewski TF (2008) Glucose deprivation inhibits multiple key gene expression events and effector functions in CD8+ T cells. Eur J Immunol 38:2438–2450 40. Ng SP, Silverstone AE, Lai ZW, Zelikoff JT (2006) Effects of prenatal exposure to cigarette smoke on offspring tumor susceptibility and associated immune mechanisms. Toxicol Sci 89:135–144 41. Hanson CD, Smialowicz RJ (1994) Evaluation of the effect of low-level 2, 3, 7, 8-tetrachlorodibenzo-p-dioxin exposure on cell mediated immunity. Toxicology 88:213–224 42. Kerkvliet NI, Baecher-Steppan L, Shepard DM, Oughton JA, Dekrey GK (1996) Inhibition of TC-1 cytokine production, effector cytotoxic T lymphocyte development and alloantibody production by 2, 3, 7, 8-tetrachlorodibenzo-p-dioxin. J Immunol 157:2310–2319 43. Lawrence BP, Roberts AD, Neumiller JJ, Cundiff JA, Woodland DL (2006) Aryl hydrocarbon receptor activation impairs the priming but not the recall of influenza virus-specific CD8+ T cells in the lung. J Immunol 177:5819–5828

44. Li Q, Hirata Y, Piao S, Minani M (2000) Immunotoxicity of N, N-diethylaniline in mice: effect on natural killer activity, cytotoxic T lymphocyte activity, lymphocyte proliferation response and cellular components of the spleen. Toxicology 150:179–189 45. Henry SP, Levin AA, White K, Mennear JH (2006) Assessment of the Effects of ISIS 2302, an Anti-Sense Inhibitor of Human ICAM-1, on Cellular and Humoral Immunity in Mice. J Immunotoxicol 3:199–211 46. De Krey GK, Bauecher-Steppan L, Deyo JA, Smith B, Kerkvliet NI (1993) Polychlorinated biphenyl-induced immune suppression: castration, but not adrenalectomy or RU 38486 treatment, partially restores the suppressed cytotoxic T lymphocyte response to alloantigen. J Pharmacol Exp Ther 267:308–315 47. Karrow NA, McCay JA, Brown R, Musgrove D, Munson AE, White KL Jr (2000) Oxymetholone modulates cell-mediated immunity in male B6C3F1 mice. Drug Chem Toxicol 23:621–644 48. Sheil JM, Frankenberry MA, Schell TD, Brundage KM, Barnett JB (2006) Propanil exposure induces delayed but sustained abrogation of cell-mediated immunity through direct interference with cytotoxic T-lymphocyte effectors. Environ Health Perspect 114: 1059–1064 49. Blaylock BL, Soderberg LS, Gandy J, Menna JH, Denton R, Barnett JB (1990) Cytotoxic T-lymphocyte and NK responses in mice treated prenatally with chlordane. Toxicol Lett 51:41–49 50. Yamada A, Kataoka T, Nagai K (2000) The fungal metabolite gliotoxin: immunosuppressive activity on CTL-mediated cytotoxicity. Immunol Lett 71:27–32 51. Loser K, Apelt J, Voskort M, Mohaupt M, Balkow S, Schwarz T, Grabbe S, Beissert S (2007) IL-10 controls ultraviolet-induced carcinogenesis in mice. J Immunol 179: 365–371 52. Mumprecht S, Matter M, Pavelic V, Ochsenbein AF (2006) Imatinib mesylate selectively impairs expansion of memory cytotoxic T cells without affecting the control of primary viral infections. Blood 108: 3406–3413 53. Ehrlich JP, Gunnison AF, Burleson GR (1989) Influenza virusspecific cytotoxic T-lymphocyte activity in Fischer 344 rat lungs as a method to assess pulmonary immunocompetence: Effect of phosgene inhalation. Inhal Toxicol 1:129–138 54. Luster MI, Portier C, Pait DG, White KL Jr, Gennings C, Munson AE, Rosenthal GJ

The Cytotoxic T Lymphocyte Assay for Evaluating Cell-Mediated Immune Function (1992) Risk assessment in immunotoxicology. I. Sensitivity and predictability of immune tests. Fundam Appl Toxicol 18:200–210 55. Burleson GR, Burleson FG (2007) Influenza virus host resistance model. Methods 41:31–37 56. Dietert RR (2008) Influenza host resistance assay to assess developmental immunotoxicity (DIT). Int J Immunotoxicol 27(6):416

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Chapter 15 NK Cell Assays in Immunotoxicity Testing Qing Li Abstract It is well known that natural killer (NK) cells are involved in defense against viruses and some tumors. NK cells kill target cells by the directed release of cytolytic granules that contain perforin, granzymes, and granulysin. It is increasingly important to evaluate NK cell function in immunotoxicity testing. NK cell function can be evaluated by determining cytolytic activity against target tumor cells by the 51Cr-release assay and also by determining the number of NK cells in peripheral blood in humans and in the spleen in animals using flow cytometry. Recently, the intracellular levels of perforin, granzymes, and granulysin determined by flow cytometry have also been used in the evaluation of NK cell function. This chapter will describe the methods for NK cell assays in immunotoxicity testing. Key words: Chromium release, Flow cytometry, Granulysin, Granzymes, NK activity, Perforin

1. Introduction Natural killer (NK) cells commonly express cell surface markers such as CD16 and CD56 in humans and NK-1.1 and/or CD49b/ Pan-NK in mice. It is well known that NK cells are involved in defense against viruses and some tumors. NK cells induce tumor or virus-infected target cell death by two main mechanisms (1–8). The first mechanism is the direct release of cytolytic granules that contain the pore-forming protein perforin, granzymes (1–6), and granulysin (5, 7) by exocytosis to kill target cells. The second mechanism is mediated by the Fas ligand (FasL)/Fas pathway, in which FasL (CD95L), a surface membrane ligand of the killer cell, cross-links with the target cell’s death receptor Fas (CD95) to induce apoptosis of the target cells (1, 2, 8). NK cell function can be evaluated by determining cytolytic activity against target tumor cells by the 51Cr-release assay in mice (3, 8–13) and in humans (3, 5, 6, 13–18) and, also by determining the number of R.R. Dietert (ed.), Immunotoxicity Testing: Methods and Protocols, Methods in Molecular Biology, vol. 598 DOI 10.1007/978-1-60761-401-2_15, © Humana Press, a part of Springer Science + Business Media, LLC 2010

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NK cells by flow cytometry (11, 16–20). Recently, it was found that the intracellular levels of perforin, granzymes, and granulysin show good correlations with NK activity in humans (16–18) and that determining the intracellular levels of perforin, granzymes, and granulysin by flow cytometry are useful parameters for evaluating the effect of chemicals on human NK cell function (5, 6, 14). It is increasingly important to measure NK cell function when evaluating chemical-induced immunotoxicity (1–3, 5, 6, 8, 11–15, 19). This chapter will describe the methods for NK cell assays in immunotoxicity testing.

2. Materials 2.1. Cell Lines as Target Cells

1. YAC-1 target cells, a Moloney murine leukemia virus (Mo-MuLV) induced T lymphoma cell line, originally from A/Sn mouse (ATCC, Manassas, VA, USA) and maintained in RPMI 1640 medium containing 10% FBS at 37°C in 5% CO2 are used in murine NK activity assays (3, 8–13). 2. K-562 target cells, a human erythroleukemia line (ATCC, Manassas, VA, USA) are maintained in RPMI 1640 medium containing 10% FBS at 37°C in 5% CO2, and are used in human NK activity assays (3, 6, 13–18).

2.2. Cell Culture and NK Activity Assay

1. Alpha minimum essential medium (a-MEM) without ribonucleosides and deoxyribonucleosides and RPMI 1640 medium (Sigma, St. Louis, MO, USA). 2. Inositol, 2-mercaptoethanol (2-ME), folic acid, l-glutamine, and gentamicin (Sigma, St. Louis, MO, USA). 3. Fetal bovine serum (FBS, JRH Biosciences, Lenexa, KS, USA) is heat-inactivated at 56°C for 30 min prior to use. 4. Tris–NH4Cl buffer solution for lysing erythrocytes: mix 0.83% NH4Cl solution and 2.06% Tris–HCl buffer solution at pH 7.65 at a ratio of 9:1, and then filter through a 0.22 µm sterilized filter. 5. Dimethyl 2,2-dichlorovinyl phosphate (DDVP, dichlorvos) (Wako Pure Chemical Industries, Osaka, Japan). 6. Sodium 51Cr-chromate (Perkin Elmer, Boston, MA, USA). 51 Cr should be stored behind lead shielding at 4°C and is radiation hazard. The half life of 51Cr is 27.7 days.

2.3. Flow Cytometry

1. Phycoerythrin (PE) labeled-mouse anti-mouse NK-1.1 (mouse IgG2a, PK136) and PE-CD49b/Pan-NK (rat IgM, DX5) were used for staining mouse NK cells, and PE-mouse IgG2a and PE-rat IgM were used as negative isotypic controls (BD PharMingen San Diego, CA, USA) (see Note 1).

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2. Purified rat anti-mouse CD16/32 (IgG2b) was used to block Fc-mediated non-antigen-specific binding of antibodies and PE to mouse Fc gamma II/III receptors that are expressed on murine NK cells (BD PharMingen, San Diego, CA, USA). 3. PE-mouse anti-human CD56 and PE-CD16 (IgG1) were used for staining human NK cells and PE-IgG1 was used as the negative isotypic control (BD PharMingen, San Diego, CA, USA). 4. Fluorescein isothiocynate (FITC)-labeled mouse anti-human perforin (mouse IgG2b) and FITC-mouse IgG2b negative isotype control (BD PharMingen, San Diego, CA, USA). 5. FITC-mouse anti-human granzyme A (GrA, IgG1) and FITC-mouse IgG1 for the negative isotypic control (BD PharMingen, San Diego, CA, USA). 6. FITC-mouse anti-human granzyme B (GrB, IgG1) (BD PharMingen, San Diego, CA, USA). 7. Mouse anti-human granzyme 3/K (Gr3/K, IgG2b) and mouse IgG2b (Abcom, Cambridge, UK) and stored at −30°C. 8. Rabbit anti-human granulysin (GRN) polyclonal antibody was kindly provided by Prof. Krensky (21) and stored at −80°C. GRN is stable at 4°C for up to 1 year. 9. PE-goat-anti rabbit IgG and PE-goat-anti mouse IgG (Vector Laboratories Inc., Burlingame, CA, USA). 10. Cytofix/Cytoperm and Perm/Wash solutions (BD PharMingen, San Diego, CA, USA) are stored at 4°C in the dark. 11. Propidium iodide (PI) for staining dead cells (Sigma, St. Louis, MO, USA). PI is light sensitive and should be stored at 4°C in the dark. 12. PBS (−): 0.80% NaCl, 0.29% Na2HPO4∙12H2O, 0.02% KCl, 0.02% KH2PO4, and 0.1% sodium azide and is stored at room temperature.

3. Methods 3.1. NK Activity Assay

Human and murine NK activity can be measured in a coculture of the target cells with NK cells (effectors) in a microplate in vitro by determining the proportion of dead target cells induced by NK cells. There are several assays for determining the dead target cells (9, 10, 22–24), however, the best method is still the 51Cr-release assay (3, 8–18).

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3.1.1. Preparation for Murine Splenocytes as Effectors (3, 8, 11–13)

1. Sacrifice the mouse by cervical dislocation. Remove the spleen and transfer to a sterile plastic dish containing 5 ml of RPMI 1640 medium supplement with 20 µg/ml gentamicin, 2.3 mM glutamine and 5 × 10−5 M 2-ME. 2. Cut the spleen into smaller parts and grind between two sterile glass slides. 3. Further disperse clumps by pipetting up and down with a sterile Pasteur pipette. 4. Pass the suspension through a sterile mesh and transfer to a 15 ml centrifuge tube. 5. Centrifuge the cells at 300 × g for 7–10 min and aspirate the supernatant. 6. Resuspend the cell pellets in Tris–NH4Cl buffer and incubate for 3 min on ice to lyse the erythrocytes. 7. Wash the cells twice with RPMI 1640 medium and resuspend in RPMI 1640 medium containing 10% FBS. 8. Count the cells in trypan blue solution. The viability of the cells should be more than 95%. 9. Finally, adjust the cells to 1 × 107 cells/ml in RPMI 1640 medium containing 10% FBS for the subsequent assays (see Note 2).

3.1.2. Preparation for Human Peripheral Blood Lymphocytes as Effectors (16–18)

Human peripheral blood lymphocytes (PBLs) can be separated from peripheral blood with a BD Vacutainer® CPT™ Tube containing sodium heparin (Becton Dickinson, Franklin Lakes, NJ, USA) as follows: 1. Draw peripheral blood by venipuncture and collect 8 ml into the BD Vacutainer® CPT™ Tube (see Note 3). 2. Mix the blood sample immediately prior to centrifugation by gently inverting the tube five times. 3. Centrifuge the BD Vacutainer tube containing the blood sample at room temperature (18–25°C) in a horizontal rotor (swing-out head) for 15 min at 1,500–1,800 × g (see Note 4). 4. After centrifugation, PBLs will form a whitish layer just under the plasma layer. 5. Aspirate approximately half of the plasma without disturbing the cell layer. 6. Collect the cell layer with a Pasteur pipette and transfer to a 15 ml centrifuge tube with a cap (see Note 5). 7. Wash the PBLs with 12 ml of PBS in a 15 ml tube and then recollect the PBLs by centrifugation for 15 min at 300 × g. Aspirate the supernatant without disturbing cell pellet.

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8. Resuspend the cell pellet by gently tapping the tube and wash again with 12 ml of PBS by centrifugation for 10 min at 300 × g. 9. Aspirate the supernatant and resuspend the cell pellet in RPMI 1640 medium containing 10% FBS. 10. Determine the viability of the cells by trypan blue dye exclusion and the viability should be more than 95%. 11. Resuspend the PBLs at 4 × 106 cells/ml in RPMI 1640 medium containing 10% FBS for the subsequent assays. 3.1.3. Preparations for Target cells

1. Label 100 µl of YAC-1 or K-562 target cells with >95% viability at 1 × 107 cells/ml with 100 µCi of sodium 51Cr-chromate solution in a 15 ml tube for 1 h at 37°C in 5% CO2 (see Note 6). 2. Wash the cells four times in RPMI 1640 medium containing 10% FBS (see Note 7). 3. Finally, resuspend the target cells at 1 × 105 cells/ml in RPMI 1640 medium containing 10% FBS for the subsequent NK activity assay.

3.1.4. NK Activity Determined by 51 Cr-Release Assay

1. Plate 100 µl of target cells at 1 × 105/ml into round-bottomed 96-well microplates. 2. Add effector cells (splenocytes at 1, 0.5, or 0.25 × 107cells/ml or PBLs at 4, 2, or 1 × 106 cells/ml) in 100 µl to the wells in triplicate at effector-to-target (E/T) ratios of 100:1, 50:1, and 25:1 for mouse assays, and 40:1, 20:1, and 10:1 for human assays (see Note 8). The total reaction volume should be 200 µl. 3. Controls should include target cells with medium alone for spontaneous release and with 0.2% Triton X-100 for maximum release. Each titration should be performed in triplicate to reduce variation. 4. Following a 4 h incubation at 37°C in 5% CO2, centrifuge the microplates at 300 × g for 5 min. 5. Collect 100 µl of supernatant from each well. 6. Count in a gamma counter (1470 Wizard™, Perkin Elmer). 7. Calculate the cytolysis attributed to NK cells by averaging counts per minute (cpm) for trpilicate wells and applying the following formula: % specific cytolosis = [(cpm of EXP − cpm of SR)/(cpm of MR − cpm of SR)] × 100%, where cpm for the experimental release (EXP) are counts from cocultures of effectors with target cells; the cpm for spontaneous release (SR) are counts from cultures of the target cells only; and the cpm for the maximum release (MR) are measured by adding 0.1 ml of 0.2% Triton X-100, instead of

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the effector cells, to the designated wells containing the target cells (see Note 9). Examples of the results produced are shown in Fig. 15.1 (murine splenic NK activity) (12) and Fig. 15.2 (human NK activity) (13). b

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Fig. 15.2. Human NK activity following a 3 h in vitro coculture treatment with diisopropyl methylphosphonate (DIMP) at 37°C. The concentrations of DIMP were 0, 125, 250, and 500 ppm. Data are presented as the mean ± SE (n = 10). **p < 0.01, *p < 0.05, significantly different from 0 ppm by unpaired t-test. ANOVA: F = 50.45, p = 3.36 × 10−11. The activity values for the 40:1 (effector/target) ratio were used for statistical analysis, but similar results were obtained with effector/ target ratio of 20:1. (Reproduced from ref. (13) with permission from Elsevier Science).

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3.2. Determining the Number of NK Cells 3.2.1. Determining the Murine Splenic NK Cells by Flow Cytometry (12, 19)

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1. Add 50 µl of splenocytes at 1 × 107/ml in PBS (−) containing 2% FBS to a 1.5 ml microcentrifuge tube, then add CD16/32 at 1–2 µg/ml (final concentration) (see Note 10) into the tubes to reduce nonspecific reactions (Fc binding) of FITC and PE, and incubate the mixture for 5 min at 4°C. 2. Add PE-NK-1.1+ or PE-CD49b/Pan-NK at 1–2 µg/ml (final concentration) (see Note 10) to the above tubes and mix, then incubate for 30 min at 4°C in the dark (see Note 11). 3. After staining, wash the cells three times with PBS (−), then pour into FACS tubes in PBS (−) containing 2% FBS in a total volume of 500 µl. 4. Add PI dye at 1 mg/ml (1–2 µl) to stain dead cells before analysis. 5. Perform flow cytometric analysis with a FACScan flow cytometer (Becton Dickinson, San Jose, CA, USA) equipped with a 488 nm argon laser and detectors for forward scatter (FSC), 90° light scatter (side scatter, SSC), and FL1 (band pass filter wavelength 530 nm), FL2 (585 nm), and FL3 (650 nm) fluorescence emission in the green, red/orange, and long red parts of the spectra, respectively. 6. Detect splenocytes stained with PE-NK-1.1+ or PE-CD49b/ Pan-NK using the FL2 filter and identify dead cells stained with PI using FL3. 7. Compensate electronically for fluorescence overlap using splenocytes stained with single colors (PE and PI). Acquire and store 10,000 events for each analysis. 8. Splenocytes can be identified by their characteristic appearance on dot plots of FSC versus SSC and electronically gated to exclude platelets, red cells, or dead cell debris. 9. The gate is the same for both exposed and control mice. Indicate the results as the percentage of positive cells within a gate. Set the fluorescence gate using the negative isotypic control. Calculate the absolute number of each cell population using the percentage of each positive cell and the total number of splenocytes.

3.2.2. Determining the Human NK Cells by Flow Cytometry (16–18, 20)

1. Stain 1 × 106 PBLs in 30 µl PBS (−) in a 1.5 ml microcentrifuge tube with 10 µl PE-CD16 for 30 min at 4°C in the dark (see Note 10). 2. After staining, wash the cells three times with PBS (−) and then pour into FACS tubes in PBS (−) containing 2% FBS in a total volume of 400 µl. 3. Add PI dye at 1 mg/ml (1–2 µl) to stain dead cells before analysis.

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4. Perform flow cytometric analysis with a FACScan flow cytometer as described above. 5. Detect PBLs stained with PE-CD16 using FL2. Detect dead cells stained with PI using FL3. 6. Compensate electronically for fluorescence overlap using PBLs stained with single colors (PE and PI). Acquire and store 10,000 events for each analysis. 7. PBLs can be identified by their characteristic appearance on dot plots of FSC versus SSC and electronically gated to exclude platelets, red cells, or dead cell debris. 8. Indicate the results as the percentage of positive cells within a gate. Set the fluorescence gate using the negative isotypic control. Calculate the absolute numbers of each cell type using the percentage of each positive cell and the total number of PBLs. An example of the results produced is shown in Fig. 15.3 (human CD16+ NK cells). 3.3. Intracellular Levels of Perforin, Granzymes, and Granulysin in NK Cells

Cell line NK-92CI is an IL-2 independent human NK cell line derived from cell line NK-92 by transfection with the human IL-2 cDNA. This cell line expresses high levels of intracellular perforin, GrA, GrB, Gr3/K, and GRN (5, 6, 14, 15) and is cytotoxic to a wide range of malignant cells, killing both K562 (6, 14, 15) and Daudi cells in 51Cr-release assays. In this experiment, instead of human NK cells, the NK-92CI cell line was used to explore the effect of chemicals on the intracellular level of perforin, GrA, GrB, Gr3/K, and GRN.

Fig. 15.3. Human CD16+ NK cells in PBLs determined with PE-CD16 by flow cytometry. The solid histogram shows the control stained with PE-mouse IgG1 (isotypic control) and the blank histograms show the results stained with PE-mouse anti-human CD16. The percentage (36.2%) in the figure shows PE-CD16+ (NK cells) in PBLs. (Li, unpublished data).

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3.3.1. Preparations for NK-92CI Cells

Maintain the NK-92CI cell line (ATCC, Manassas, VA, USA) using a-MEM without ribonucleosides and deoxyribonucleosides with 2 mM l-glutamine adjusted to contain 1.5 g/L sodium bicarbonate and supplemented with 0.2 mM inositol, 0.1 mM 2-ME, 0.02 mM folic acid, and incubation with 10% FBS at 37°C in 5% CO2 (see Note 12).

3.3.2. Treatment with Chemicals

1. Incubate NK-92CI cells at 2–4 × 105/ml with chemicals at various concentrations for different time periods at 37°C in a 5% CO2 incubator. 2. Harvest, wash with PBS, and use for the subsequent flow cytometric analysis.

3.3.3. Cell Staining and Flow Cytometric Analysis

1. Stain NK-92CI cells of 1 × 106 in 30 µl in a 1.5 ml microcentrifuge tube with 15 µl PE-CD56 or PE-IgG1 (isotypic control) for 30 min at 4°C in the dark, then wash twice with PBS (−) (see Note 10). 2. Fix/permeabilize the cells with Cytofix/Cytoperm solution in 250 µl for 20 min at 4°C in the dark, then wash twice with Cytoperm washing solution. 3. Stain the cells in 30 µl with 10 µl FITC-mouse anti-human perforin, FITC-GrA, FITC-GrB, or FITC-labeled mouse IgG1 and IgG2b for the negative isotypic controls, respectively, for 30 min at 4°C in the dark (see Note 10). 4. Stain intracellular GRN with rabbit anti-human GRN polyclonal antibody at 1:800 for 20 min after fixation/permeabilization with Cytofix/Cytoperm solution, then stain with PE-goat anti-rabbit IgG at 0.5 µg/ml for 20 min at 4°C in the dark (see Note 10). 5. Stain intracellular Gr3/K with mouse anti-human Gr3/K monoclonal antibody at 5 µg/ml for 30 min after fixation and permeabilization with Cytofix/Cytoperm solution, and then stain with PE-goat anti-mouse IgG at 1 µg/ml for 30 min at 4°C in the dark (see Note 10). 6. After staining, wash the cells twice with the Cytoperm washing solution and once with PBS (−) containing 2% FBS, then resuspend in 400 µl PBS (−) containing 2% FBS. 7. Perform flow cytometric analysis with a FACScan flow cytometer as described above. 8. Detect NK-92CI cells stained with FITC- and PE-labeled antibodies using FL1 and FL2, respectively. 9. Compensate electronically for fluorescence overlap using NK-92CI cells stained with single colors (FITC and PE). 10. Acquire and store 10,000 events for each analysis.

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11. Indicate the results as the percentage of positive cells within a gate and the intensity of fluorescence of FITC or PE (5, 6). Set the fluorescence gate using the negative isotypic control. An example of the results produced is shown in Fig. 15.4 (5, 6).

4. Notes 1. All FITC/PE-labeled antibodies are light sensitive and should be stored at 4°C in the dark. They are stable at 4°C for up to 1–2 years. 2. Splenocytes may be collected from a mouse exposed to a chemical in vivo. Alternatively, splenocytes from a normal mouse may be exposed to a chemical in vitro in immunoto xicity testing. 3. For best results, the BD Vacutainer® CPT™ Tube with sodium heparin should be kept at room temperature (18–25°C) and

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Fig.15.4. Effect of DDVP on the expression of perforin, GRN, GrA, GrB, and Gr3/K in NK-92CI cells. NK-92CI cells at 4 × 105/ml were incubated with DDVP at the indicated concentrations for 15 h at 37°C in 5% CO2, then harvested, washed with PBS. A: the solid histogram shows the control stained with FITC-mouse IgG2b (isotype control) and the blank histograms show the results stained with FITC-mouse anti-human perforin after treatment with DDVP at 0, 0.055, 0.110, 0.225, and 0.450 mM from right to left, respectively. All experiments were repeated at least 3 times with very similar results. (Reproduced from refs. (5) and (6) with permission from Elsevier Science).

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blood samples should be centrifuged within 2 h of blood collection. 4. A preliminary test is necessary for determining the optimal time and relative centrifugal force of centrifugation. 5. Collection of cells immediately following centrifugation will yield the best results. 6. High viability of the target cells will produce lower spontaneous release. For laboratory controls of the NK activity assay, the same K-562 or YAC-1 cells as the targets should be used in all experiments and always keep the K-562 or YAC-1 cells in the same conditions before the experiments, e.g., use the K-562 or YAC-1 cells 96-h after thawing out the cells. It is important to suspend the cells by gently tapping the tube every 15 min during the labeling. 7. It is important to wash cells four times to obtain lower spontaneous release. 8. The E/T ratios can be changed based on the design of the experiment. 9. SR from target cells should be less than 12% of the respective MR; if SR is over 20%, the experiment should be excluded from further analysis. 10. It is very important to keep the same volume of cells in all experiments of the same series, and the optimal volume (concentration) of antibodies should be determined in preliminary experiments. 11. The NK-1.1 marker is useful in defining NK cells in C57BL, FVB/N, NZB, Swiss NIH, and SJL mice, but not for A, AKR, BALB/c, CBA/J, C3H, C57BR, C58, DBA/1, DBA/2, NOD, and 129. CD49b/Pan-NK is useful in defining NK cells in most strains tested such as A/J, AKR, BALB/c, C3H/HeJ, C57BL/6, C57BL/10, C57BR, C57L, C58, CBA/Ca, CBA/J, DBA/1, DBA/2, SJL, SWR, and 129/J, but not in NOD mice. 12. The NK-92CI cells should be maintained in a-MEM supplemented with 12.5% horse serum and 12.5% FBS in the original method. However, good results have also been obtained in a-MEM supplemented with 10% FBS.

Acknowledgments The parts of experimental data shown in this chapter and performed by author were supported by grants from the Ministry of Education, Culture, Sports, Science and Technology of Japan

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(No. 09877077, No. 10770178, No. 12770206, No. 15590523, and No. 19590602). The author would like to thank Dr. Tomoyuki Kawada (Professor and Chief) at the Department of Hygiene and Public Health, Nippon Medical School, for his advice. References 1. Li Q, Kawada T (2006) The new mechanism of organophosphorus pesticides-induced inhibition of cytolytic activity of killer cells. Cell Mol Immunol 3:171–178 2. Li Q (2007) New mechanism of organophosphorus pesticide-induced immunotoxicity. J Nippon Med Sch 74:92–105 3. Li Q, Nagahara N, Takahashi H, Takeda K, Okumura K, Minami M (2002) Organophosphorus pesticides markedly inhibit the activities of natural killer, cytotoxic T lymphocyte and lymphokine-activated killer: a proposed inhibiting mechanism via granzyme inhibition. Toxicology 172:181–190 4. Hirata Y, Inagaki H, Shimizu T, Li Q, Nagahara N, Minami M et al (2006) Expression of enzymatically active human granzyme 3 in Escherichia coli for analysis of its substrate specificity. Arch Biochem Biophys 446:35–43 5. Li Q, Nakadai A, Ishizaki M, Morimoto K, Ueda A, Krensky AM et al (2005) Dimethyl 2, 2-dichlorovinyl phosphate (DDVP) markedly decreases the expression of perforin, granzyme A and granulysin in human NK-92CI cell line. Toxicology 213:107–116 6. Li Q, Kobayashi M, Kawada T (2008) DDVP markedly decreases the expression of granzyme B and granzyme 3/K in human NK cells. Toxicology 243:294–302 7. Okada S, Li Q, Whitin JC, Clayberger C, Krensky AM (2003) Intracellular mediators of granulysin-induced cell death. J Immunol 171:2556–2562 8. Li Q, Nakadai A, Takeda K, Kawada T (2004) Dimethyl 2, 2-dichlorovinyl phosphate (DDVP) markedly inhibits activities of natural killer cells, cytotoxic T lymphocytes and lymphokine-activated killer cells via the Fasligand/Fas pathway in perforin-knockout (PKO) mice. Toxicology 204:41–50 9. Kiessling R, Klein E, Wigzell H (1975) “Natural” killer cells in the mouse. I. Cytotoxic cells with specificity for mouse Moloney leukemia cells. Specificity and distribution according to genotype. Eur J Immunol 5:112–117

10. Kiessling R, Klein E, Pross H, Wigzell H (1975) “Natural” killer cells in the mouse. II. Cytotoxic cells with specificity for mouse Moloney leukemia cells. Characteristics of the killer cell. Eur J Immunol 5:117–121 11. Li Q, Minami M, Inagaki H (1998) Acute and subchronic immunotoxicity of p-chloronitrobenzene in mice: I. Effect on natural killer, cytotoxic T lymphocyte activities and mitogen-stimulated lymphocyte proliferation. Toxicology 127:223–232 12. Li Q, Hirata Y, Piao S, Minami M (2000) Immunotoxicity of N, N-diethylaniline in mice: Effect on natural killer activity, cytotoxic T lymphocyte activity, lymphocyte proliferation response and cellular components of the spleen. Toxicology 150:181–191 13. Li Q, Hirata Y, Piao S, Minami M (2000) The by-products generated during sarin synthesis in the Tokyo sarin disaster induced inhibition of natural killer and cytotoxic T lymphocyte activity. Toxicology 146:209–220 14. Li Q, Nakadai A, Matsushima H, Miyazaki Y, Krensky AM, Kawada T et al (2006) Phytoncides (wood essential oils) induce human natural killer cell activity. Immunopharmacol Immunotoxicol 28:319–333 15. Li Q, Kobayashi M, Kawada T (2007) Organophosphorus pesticides induce apoptosis in human NK cells. Toxicology 239:89–95 16. Li Q, Morimoto K, Nakadai A, Inagaki H, Katsumata M, Shimizu T et al (2007) Forest bathing enhances human natural killer activity and expression of anti-cancer proteins. Int J Immunopathol Pharmacol 20:3–8 17. Li Q, Morimoto K, Kobayashi M, Inagaki H, Katsumata M, Hirata Y et al (2008) Visiting a forest, but not a city, increases human natural killer activity and expression of anti-cancer proteins. Int J Immunopathol Pharmacol 21:117–128 18. Li Q, Morimoto K, Kobayashi M, Inagaki H, Katsumata M, Hirata Y et al (2008) A forest bathing trip increases human natural killer activity and expression of anti-cancer proteins

NK Cell Assays in Immunotoxicity Testing in female subjects. J Biol Regul Homeost Agents 22:45–55 19. Li Q, Minami M, Hanaoka T, Yamamura Y (1999) Acute immunotoxicity of p-chloronitrobenzene in mice: II. Effect of p-chloronitrobenzene on the immunophenotype of murine splenocytes determined by flow cytometry. Toxicology 137:35–45 20. Li Q, Morimoto K, Nakadai A, Qu T, Matsushima H, Katsumata M et al (2007) Healthy lifestyles are associated with higher levels of perforin, granulysin and granzymes A/B-expressing cells in peripheral blood lymphocytes. Prev Med 44:117–123 21. Hanson DA, Kaspar AA, Poulain FR, Krensky AM (1999) Biosynthesis of granulysin, a novel cytolytic molecule. Mol Immunol 36: 413–422

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22. Ito M, Watanabe M, Kamiya H, Sakurai M (1996) Non-radioactive assay of natural killer cell-mediated cytotoxicity against cytomegalovirus-infected fibroblasts by DNA fragmentation ELISA. J Virol Methods 56:77–84 23. Nagao F, Yabe T, Xu M, Yokoyama K, Saito K, Okumura K (1996) Application of nonradioactive europium (Eu3+) release assay to a measurement of human natural killer activity of healthy and patient populations. Immunol Invest 25:507–518 24. Kim GG, Donnenberg VS, Donnenberg AD, Gooding W, Whiteside TL (2007) A novel multiparametric flow cytometry-based cytotoxicity assay simultaneously immunophenotypes effector cells: comparisons to a 4 h 51Cr-release assay. J Immunol Methods 325: 51–66

Chapter 16 The Local Lymph Node Assay and Skin Sensitization Testing Ian Kimber and Rebecca J. Dearman Abstract The mouse local lymph node assay (LLNA) is a method for the identification and characterization of skin sensitization hazards. In this context the method can be used both to identify contact allergens, and also determine the relative skin sensitizing potency as a basis for derivation of effective risk assessments. The assay is based on measurement of proliferative responses by draining lymph node cells induced following topical exposure of mice to test chemicals. Such responses are known to be causally and quantitatively associated with the acquisition of skin sensitization and therefore provide a relevant marker for characterization of contact allergic potential. The LLNA has been the subject of exhaustive evaluation and validation exercises and has been assigned Organization for Economic Cooperation and Development (OECD) test guideline 429. Herein we describe the conduct and interpretation of the LLNA. Key words: Contact allergy, Local lymph node assay, Hazard identification, Potency

1. Introduction The mouse local lymph node assay (LLNA) was developed initially to provide an alternative to guinea pig tests for the identification of skin sensitization hazards (1). Descriptions of the development, evaluation and eventual validation of the LLNA for this purpose are available elsewhere (1–3), and a survey of that history is unnecessary here. The approach is based on an understanding that the acquisition of skin sensitization requires the induction of a cutaneous immune response to chemical allergen experienced at the skin surface. The immunobiology of skin sensitization is complex and requires an array of cellular/molecular interactions that are orchestrated in time and space. For the purposes of this article, however, it is possible to summarise the key events as follows. If an inherently susceptible subject is exposed R.R. Dietert (ed.), Immunotoxicity Testing: Methods and Protocols, Methods in Molecular Biology, vol. 598 DOI 10.1007/978-1-60761-401-2_16, © Humana Press, a part of Springer Science + Business Media, LLC 2010

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topically to a sufficient quantity of contact allergen then specific priming of the immune system will be provoked that results in sensitization. The inducing allergen is recognized in the skin by one or more of several populations of dendritic cells, including epidermal Langerhans’ cells (LC). These cells have responsibility for internalization and processing of antigen (including chemical allergen) at skin surfaces and for delivery of that antigen to regional lymph nodes draining the site of exposure. Within nodes antigen is presented to responsive T lymphocytes that are triggered to divide and differentiate. The division of antigen-specific T lymphocytes results in selective clonal expansion and the immunological priming that is defined as sensitization. Central to the development of skin sensitization, therefore, is the activation and clonal expression of chemical (hapten)-specific T lymphocytes. We therefore chose to measure the ability of chemicals to cause skin sensitization as a function of their ability in mice to stimulate lymph node cell (LNC) proliferation following topical application (4, 5). This is the approach that became known as the LLNA. In principle the assay is very straightforward. Groups of mice are exposed topically on the dorsum of both ears to various concentrations of the test chemical, or to an equivalent volume of the relevant vehicle alone. Exposure is performed daily for three consecutive days. Five days following the initiation of exposure mice receive a source of radiolabelled thymidine by intravenous injection. Thereafter animals are sacrificed, and draining lymph nodes excised and processed for determination of the vigour of mitosis. Results are recorded in the form of a stimulation index (SI) of LNC proliferation relative to values obtained with the concurrent vehicle control. Chemicals that induce, at one or more test concentrations, an SI of equal to or greater than 3 are classified as having skin sensitization potential. Chemicals that fail to elicit a threefold or greater SI at all concentrations are regarded as lacking significant skin sensitizing activity. Experience with this assay has now been extensive and it has been assigned an Organization for Economic Cooperation and Development (OECD) guideline method (guideline 429). More recently attention has focused on the use of the LLNA not only to provide accurate hazard identification but also to produce an estimation of relative skin sensitization potency. This is predicated on an understanding that LNC proliferation not only has a causal relationship with the acquisition of skin sensitization, but it also correlates closely with the extent to which sensitization will be achieved (that is both a causal and quantitative relationship). For this reason the LLNA is now used also for hazard characterization with regard to relative potency; information that has proven invaluable for the derivation of effective risk assessments and risk management (1, 6).

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2. Materials 1. Chemical to be tested, formulated in an appropriate vehicle and vehicle alone (see Note 1). Gloves should always be worn when handling suspect contact sensitizers. Solutions must be prepared freshly (within 1 h of dosing). 2. Female CBA strain mice (young adults); maintained in semibarriered conditions (see Note 2). 3. Automatic pipet with disposable tips. 4. Tritiated thymidine: 2 Ci/mmol (80 mCi/ml). Store the stock solution at 4°C. Due to the potential for the tritium on the thymidine to exchange onto water molecules, the chemical expiry date provided by the manufacturer must be observed. Gloves must be worn, and the area in which thymidine is used must be swabbed and monitored regularly for radioactive contamination. 5. Phosphate-buffered saline (PBS); pH 7.2–7.4, stored at room temperature. 6. Filters (0.4 mM). 7. Sterile 1 mL graduated syringes. 8. Sterile 25G 5/8 needles. 9. Temperature controlled “hot box” (see Note 3). 10. Mouse restrainer tube with outlet for tail. 11. 200 gauge stainless steel mesh, cut into approximately 3 cm squares, washed in 70% ethanol, autoclaved and edges turned up to form a box. 12. Sterile 5 mL syringes. 13. 5% w/v trichloroacetic acid (TCA) in water; stable at room temperature for 6 months. 14. Scintillation fluid (eg Hi-Safe Optiphase). 15. b-Scintillation counter.

3. Methods After rigorous evaluation at an international level, the LLNA has become recognized as the preferred skin sensitization test method for the evaluation of chemicals. However, in common with all safety assessment methods, inappropriate conduct of the assay can lead to difficulties in data interpretation. Experience with the assay over recent years has demonstrated that strict adherence to

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published procedures and guidelines are critical for the successful outcome of an LLNA. The standard LLNA identifies contact sensitization hazard potential as a function of radiolabeled thymidine incorporation into the lymph nodes draining the site of topical exposure to chemical. There are two protocols that are used routinely for this purpose. In both methods, mice are exposed topically on the dorsum of both ears to various concentrations of test chemical, or to vehicle alone, daily for three consecutive days. Five days after the initiation of exposure, animals receive an intravenous injection of radiolabeled thymidine. Five hours later, the draining auricular lymph nodes are excised, a single cell suspension of lymph node cells (LNC) prepared and thymidine incorporation measured by b-scintillation counting. In the first method, four mice per group are used and lymph nodes are pooled on an experimental group basis. In the second method, five mice per group are used and lymph nodes are pooled for each individual animal. Processing of lymph nodes on an individual animal basis provides the opportunity to assess inter-animal variation in thymidine incorporation and for the application of statistical analyses. The LLNA not only provides reliable data for hazard identification but also allows assessment of relative skin sensitizing potency for the purpose of risk assessment. It is recognized that contact allergens vary substantially (at least four orders of magnitude) with respect to the relative potency with which they are able to induce contact sensitization (6). An understanding of relative potency of contact allergens can provide useful information that can contribute to the risk assessment process. Data derived from the protocol using lymph nodes pooled on a group basis or data derived from the protocol using lymph nodes pooled per individual animal are both suitable for this purpose. 3.1. Topical Exposure to Chemical

1. Allow mice to acclimatize for at least 2 days after arrival in the facility in cages of four animals per group (see Note 4). 2. Identify animals individually (using an indelible marker), weigh and record individual weights (see Note 5). 3. Prepare dosing solutions. Three consecutive concentrations are generally selected from the following: 50, 25, 10, 5, 2.5, 1, 0.5, 0.25 and 0.1% (w/v). Also prepare the appropriate vehicle solution (see Note 6). 4. Dispense 25 mL of chemical, or vehicle alone, onto the dorsum of both ears of each animal (n = 4 per group) (See Note 4) using an automatic pipet with a disposable tip, ensuring an even distribution over the surface of the ear. 5. Perform identical treatment (steps 3 and 4) once daily for the next two consecutive days. 6. Rest the animals for 2 days.

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1. Weigh the animals and record individual weights (see Note 5). 2. Prepare a solution of tritiated thymidine in PBS (80 mCi/mL) and filter it using a 0.4 mM filter until sterile. 3. Place the animals in a temperature controlled “hot box”, one experimental group at a time, for 5 min to allow the veins to dilate. The temperature must not exceed 37°C. 4. One at a time, restrain each mouse using a restraining tube with an outlet for the tail. Inject the tail vein with 0.25 mL of radiolabeled thymidine (80 mCi/mL), dispensed with a 1 mL graduated syringe and 25G 5/8 needle. Care must be taken to ensure syringe is free of air bubbles. (see Note 7). 5. Return the animals to their cages and allow to rest for 5 h. 6. Euthanize the animals, excise draining (auricular) lymph nodes and pool for each experimental group in a small volume (approximately 2 mL) of PBS. (see Notes 8 and 9). 7. Using tweezers, place the nodes onto a square of stainless steel gauze contained within a 60 mm plastic Petri dish with a small volume (approximately 2 mL) of fresh PBS. Prepare a single cell suspension of LNC by gently disrupting the lymph nodes and pushing them through the gauze using the plunger of a 5 mL syringe (mechanical disaggregation). 8. Transfer the LNC from the Petri dish into a 10 mL plastic centrifuge tube, rinsing the gauze and the Petri dish with fresh PBS to ensure that all of the cells are collected before disposal. Wash the LNC twice in fresh PBS by centrifugation at 100×g for 10 min (see Note 10). 9. After the final wash, resuspend the cell pellet in 3 mL of TCA and store overnight at 4°C (see Note 10). 10. Centrifuge the cells at 100×g for 10 min, pipette off the TCA with a disposable pastette and resuspend the cells in 1 mL of fresh TCA. Transfer the suspension to 10 mL of scintillation fluid (eg Hisafe Optiphase) and measure thymidine incorporation by b-scintillation counting. (see Note 11).

3.3. Calculation and Interpretation of Results

1. Record results as total disintegrations per min per node (dpm/node) for each experimental group. Use the vehicle control group as the comparator (see Note 12) in order to derive a stimulation index (SI) according to the following equation SI =

testd pm / node vehicle dpm / node

2. The criterion for positivity in the LLNA is an SI of 3 or more at one or more test concentrations (see Note 13).

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3. Typical results derived using the standard LLNA protocol are shown in Fig. 16.1 in which animals have been exposed to three concentrations of the potent contact allergen paraphenylenediamine (PPD), or to the non-sensitizing skin irritant methyl salicylate, both formulated in the vehicle of choice for the LLNA, AOO. Data are displayed as a function of dpm/ node and as SI relative to concurrent vehicle-treated control values; the latter being the usual method of displaying the results. Exposure to PPD induced a marked dose-dependent increase in proliferation compared with vehicle treated control mice, with all concentrations tested provoking an SI of 3 or greater. This material is, therefore, classified as a skin sensitizer. In contrast, the non-sensitizing skin irritant methyl salicylate, despite testing at doses some 20-fold higher than those utilized for PPD, failed to elicit increases in thymidine incorporation at any concentration examined. This material is, therefore, classified as lacking significant potential to cause skin sensitization.

b

4500 4000 3500 3000 2500 2000 1500 1000 500

proliferation (dpm/node)

proliferation (dpm/node)

a

Stimulation Index (SI)

c

0.25 0.5 1 concentration (%w/v)

30

d

30

25

Stimulation Index (SI)

0

4500 4000 3500 3000 2500 2000 1500 1000 500

25

20 15 10 5 0

0.25 0.5 1 concentration (%w/v)

0

5 10 20 concentration (%w/v)

0

5 10 20 concentration (%w/v)

20 15 10 5

Fig. 16.1. Local lymph node assay responses to paraphenylenediamine (PPD) and methyl salicylate CBA strain mice (n = 4) received 25 ml of various concentrations of the contact allergen PPD (a, c) or the non-sensitizing skin irritant methyl salicylate (b, d), or vehicle (AOO) alone on the dorsum of both ears daily for three consecutive days. Five days after the initiation of exposure, draining auricular lymph nodes were excised, pooled on a treatment group basis and processed for b-scintillation counting. Results are expressed as mean dpm/node (a, b) or as a stimulation index (SI) relative to vehicle-treated control groups (c, d). An SI of 3 (current threshold value for a positive in the LLNA) is illustrated as a broken line.

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4. Typical results derived using the LLNA protocol for the assessment of relative skin sensitizing potency are shown in Table 16.1. Animals have been exposed to five concentrations of the potent contact allergen PPD, or to the more moderate skin sensitizer hexyl cinnamic aldehyde (HCA), both formulated in the vehicle of choice for the LLNA, AOO. Data are displayed as dpm/node and as SI relative to concurrent vehicle-treated control values. Both chemicals provoked marked dose-dependent increases in proliferation compared with vehicle treated control mice. Both materials are therefore classified as skin sensitizers. The EC3 value (Estimated Concentration of chemical required to induce an SI of 3 relative to concurrent vehicle treated controls) is derived by linear interpolation according to the following equation EC3 = c + [(3 − d ) / (b − d )](a − c )

where the data points lying immediately above and below the SI value of 3 have the co-ordinates (a, b) and (c, d), respectively The vehicle-treated control (SI = 1) cannot be used for the (c, d) data point.

Table 16.1 Local lymph node assay responses to paraphenylenediamine (PPD) and hexyl cinnamic aldehyde (HCA): determination of EC3 values CBA strain mice (n = 4) received 25 ml of various concentrations of the contact allergens PPD or HCA, or vehicle (AOO) alone on the dorsum of both ears daily for three consecutive days. Five days after the initiation of exposure, draining auricular lymph nodes were excised, pooled on a treatment group basis and processed for b-scintillation counting. Results are expressed as mean dpm/node and as a stimulation index (SI) relative to vehicle-treated control groups Conc. (%w/v)

PPD dpm/node (SI)

Conc. (%w/v)

HCA dpm/node (SI)

0

155 (1)

0

324 (1)

0.05

349 (2.2)

2.5

517 (1.6)

0.1

669 (4.2)

5

690 (2.3)

0.25

2,180 (13.7)

10

1,111 (3.7)

0.5

3,203 (20.8)

25

2,724 (9.1)

1

4,036 (25.3)

50

4,576 (14.2)

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In each case, concentrations of chemical have been selected that elicit SI values either side of an SI of 3, thus the data are appropriate for the derivation of EC3 values. PPD EC3 = 0.05 + [(3 – 2.2)/(4.2 – 2.2)] ( 0.1 – 0.05) = 0.07% HCA EC3 = 5 + [(3 – 2.3)/(3.7 – 2.3)] (10 – 5) = 7.5% According to the recommendations of Kimber et al. (6), PPD is categorized as an extreme contact sensitizer, whereas HCA is classified as a moderate contact sensitizer (see Note 14).

4. Notes 1. Many organic vehicles may be used. Water, however, is inappropriate as a result of its high surface tension that makes it impossible to apply evenly and to remain in contact with the surface of the skin for a sufficient period of time for absorption. Experience indicates that, in order of preference, the vehicles of choice are: 4:1 [v:v] acetone:olive oil (AOO), methylethyl ketone, dimethylformamide and dimethylsulfoxide. Vehicle selection is dictated by the relative solubility of the test material. For most purposes, AOO is suitable (7). 2. Young adult (6–12 weeks old) female CBA strain mice are used for regulatory LLNA studies. For non-regulatory studies, mice up to 20 weeks old may be used. 3. An alternative approach to improve tail vein dilation is to hold the tails under warm running tap water. 4. In the alternative protocol in which lymph nodes from individual animals are processed, n = 5. 5. Body weight changes of the animals are monitored over the course of the study to confirm their health status. A decrease in body weight of 10% or greater may be an indication of systemic toxicity and should call into question the validity of the data for that group (8). 6. The dose selection is based on providing the highest possible test concentration to maximize the potential for hazard identification. Test concentration selection will be limited by compatibility with the vehicle chosen and the suitability of the resultant preparation for unoccluded dermal application and should preclude local (dermal) trauma or systemic toxicity. Dosing levels may be set on the basis of oral toxicity data, but

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when dealing with a new chemical, it is advisable to perform preliminary sighting studies using limited numbers of animals (1 per dose group is sufficient). In the sighting study, the health status and weight of the animals is recorded, on the fifth day the animals are terminated and the draining (auricular) lymph nodes examined visually for signs of activation (increase in size). 7. There are two appropriate veins on the tail of the mouse. The needle should be introduced at a shallow angle (less than 45o) about halfway down the tail and should be tracked up the vein for most of the length of the needle. Once inside the vein, less resistance should be felt. 8. Quiescent (vehicle-treated) lymph nodes can be difficult for the novice to find. Exposure to a potent contact sensitizer such as 2,4-dinitrochlorobenzene results in a marked increase (approximately tenfold) in lymph node size, thus, it is helpful to identify these activated lymph nodes initially. The lymph nodes are found with surrounding adipose tissue, the lymph nodes can be teased away from the adipose tissue using a pair of forceps. In some animals the auricular lymph node may appear to be a chain of several lymph nodes rather than a single lymph node; the whole lymph node chain should be isolated in this case. 9. In the alternative protocol in which lymph nodes from individual animals are processed, each pair of lymph nodes are pooled on an experimental animal basis and processed. 10. Clumping of LNC is avoided by ensuring the pellet is first completely resuspended in a small volume of liquid before making up to final volume. Round bottomed test tubes are preferable as these allow a more even resuspension of the cell pellet. When washing the LNC by centrifugation, it is advisable to pipette off the liquid from the cell pellet rather than tipping off the waste PBS as this may disturb the cell pellet. Even for vehicle-treated control lymph nodes, the cell pellet should be clearly visible at the bottom of the test tube after every centrifugation step. Loss of the cell pellet may indicate a cracked test tube or that the centrifugation step has been compromised. 11. It is possible that the TCA solution may become contaminated and may react with the scintillation fluid giving unacceptable backgrounds. It is good practice to incorporate concurrent controls of scintillation fluid alone and scintillation fluid plus 1 mL of 5% TCA and process these for b-scintillation counting along with the experimental samples. Disintegrations per minute (dpm) recorded for these controls should be less than 100 dpm.

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12. The interpretation of LLNA data requires that the proliferation achieved in test groups compared with the background activity in concurrent vehicle-treated controls so that the SI can be derived. Clearly, the use of the vehicle treated control value as the denominator in the calculation for deriving the SI means that if this value is abnormally high or abnormally low, it will impact markedly on the calculated SI. An abnormal level of thymidine incorporation may be due to technical difficulties with the assay (such as failed intravenous injections) or may reflect the health status of the animals; for example relatively high levels of basal proliferation accompanied by increased lymph node size in the vehicle-treated control group can be indicative of a viral infection or a lymphoproliferative disorder. It is apparent that for experienced competent laboratories, control levels of proliferation in the LLNA are relatively consistent, although the absolute values vary according to the testing laboratory (9). Some biological variation and interlaboratory variation is to be expected in basal levels of thymidine incorporation. However, in the hands of experienced technicians, very little inter-experimental variation in derived SI values is observed generally for a given chemical (10). It is necessary therefore to establish a normal range of control values for a given vehicle and to monitor any excursions from this range in any laboratory conducting the LLNA. 13. In many experiments, a typical dose response curve will be observed for chemical allergen-induced proliferation, with dose-dependent increases in proliferation that may or may not reach a plateau. In some cases, a bell shaped curve is recorded, with thymidine incorporation reaching maximal levels at intermediate doses and at higher doses, less vigorous proliferation observed. This effect may be due to systemic toxicity at high doses, which may be monitored by general health checks such as the daily recording of weight gain on an individual animal basis. The existence of a bell-shaped curve should not affect the interpretation of the LLNA with respect to hazard identification or relative potency assessment. In some cases, the proliferative response to chemical may show a dose-dependent increase in thymidine incorporation that does not attain an SI of 3 at the maximum dose tested. In this case, the most judicious interpretation is that the chemical has potential to cause contact allergy and, if possible, a repeat assay using higher concentrations of material or an alternative vehicle should be performed. 14. It is advisable for a laboratory to demonstrate regularly that competence has been achieved with the assay. This can be effected by the use of a positive control material such as HCA, which is recommended currently by the OECD as a positive

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control for tests for contact sensitization. This chemical has been evaluated thoroughly in the LLNA, both in the context of inter-laboratory trials and with respect to the stability of responses with time and the comparative data are available in the literature (10–12). These experiments have demonstrated that HCA is identified routinely as a contact sensitizer in the LLNA, with comparable EC3 values obtained between laboratories and for repeat assays within the same laboratory. Suggested doses for HCA (formulated in AOO) for determination of EC3 values are: 50%, 25%, 10%, 5% and 2.5%. The EC3 value for this material is approximately 10%. References 1. Kimber I, Dearman RJ, Basketter DA, Ryan CA, Gerberick GF (2002) The local lymph node assay: past, present and future. Contact Dermatitis 47:315–328 2. Dearman RJ, Basketter DA, Kimber I (1999) Local lymph node assay: use in hazard and risk assessment. J Appl Toxicol 19:299–306 3. Basketter DA, Evans P, Fielder RJ, Gerberick GF, Dearman RJ, Kimber I (2002) Local lymph node assay – validation, conduct and use in practice. Food Chem Toxicol 40:393–398 4. Basketter DA, Gerberick GF, Kimber I, Willis CM (1999) Toxicology of contact dermatitis. Wiley, Chichester 5. Kimber I, Dearman RJ, Basketter DA, Ryan CA, Gerberick GF, McNamee PM, Lalko J, Api AM (2008) Dose metrics in the acquisition of skin sensitization: thresholds and importance of dose per unit area. Regul Toxicol Pharmacol 52:39–45 6. Kimber I, Basketter DA, Butler M, Gamer A, Garrigue J-L, Gerberick GF, Newsome C, Steiling W, Vohr H-W (2003) Classification of contact allergens according to potency: proposals. Food Chem Toxicol 41:1799–1809 7. Dean JH, Twerdok LE, Tice RR, Sailstad DM, Hattan DG, Stokes WS (2001) ICCVAM evaluation of the murine local lymph node

8.

9.

10.

11.

12.

assay. Conclusions and recommendations of an independent scientific peer review panel. Regul Toxicol Pharmacol 34:258–273 Basketter DA, Gerberick GF, Kimber I (2001) Skin sensitisation, vehicle effects and the local lymph node assay. Food Chem Toxicol 39:621–627 Basketter DA, Glimour NJ, Briggs D, Ludwig U, Gerberick GF, Ryan CA, Dearman RJ, Kimber I (2003) Utility of historical control data in the interpretation of the local lymph node assay. Contact Dermatitis 49:37–41 Dearman RJ, Wright ZM, Basketter DA, Ryan CA, Gerberick GF, Kimber I (2001) The suitability of hexyl cinnamic aldehyde as a calibrant for the local lymph node assay. Contact Dermatitis 44:357–361 Dearman RJ, Hilton J, Evans P, Harvey P, Basketter DA, Kimber I (1998) Temporal stability of local lymph node assay responses to hexyl cinnamic aldehyde. J Appl Toxicol 18:281–284 Loveless SE, Ladics GL, Gerberick GF, Ryan CA, Basketter DA, Scholes EW, House RV, Hilton J, Dearman RJ, Kimber I (1996) Further evaluation of the local lymph node assay in the final phase of an international collaborative trial. Toxicology 108:141–152

Chapter 17 Use of Contact Hypersensitivity in Immunotoxicity Testing Jacques Descotes Abstract The histopathological examination of lymphoid organs together with a T-dependent antibody (TDAR) assay are the primary components of preclinical immunotoxicity assessment. Additional testing including measurement of cellular immunity may be considered. Besides ex vivo lymphocyte proliferation assays, either delayed or contact hypersensitivity models can be used. Contact hypersensitivity testing is typically performed either in mice or in guinea pigs and is directly derived from classical models used for the detection of contact sensitizing chemicals. Whatever the selected model, it is comprised of a sensitizing phase where the animals are applied a strong contact sensitizer topically, then a rest phase, and finally an eliciting phase where sensitized animals are challenged topically with the same contact sensitizer. In mice, the ear-swelling test is the reference procedure in which mice are sensitized to the ear or shaved abdominal skin and then challenged on the ear. Ear swelling usually measured from ear thickness reflects a cell-mediated immune response. In guinea pigs, a strong sensitizer is applied on the shaved skin of the abdomen or the interscapular area. The sensitized animals are challenged on another area of the shaved abdomen, and the cell-mediated response is assessed semiquantitatively from the magnitude of induced erythema inconsistently associated with edema. Treatment or exposure with immunosuppressive chemicals can result in a significantly decreased ear swelling or skin reaction. Contact hypersensitivity models are seldom used nowadays in preclinical immunotoxicity testing, most likely because of the lack of standardization and extensive validation as well as their use being restricted to mice or guinea pigs. Key words: Immunotoxicity testing, Contact hypersensitivity models, Ear-swelling test, Mice, Guinea pigs, Immunosuppression

1. Introduction The histopathological examination of main lymphoid organs, including the thymus, spleen, lymph nodes, and Peyer’s patches, and a T-dependent antibody response (TDAR) assay are the mainstay of the preclinical immunotoxicity assessment of medicines and industrial or environmental chemicals (1). When a cause for concern exists, additional immunotoxicity testing is required, R.R. Dietert (ed.), Immunotoxicity Testing: Methods and Protocols, Methods in Molecular Biology, vol. 598 DOI 10.1007/978-1-60761-401-2_17, © Humana Press, a part of Springer Science + Business Media, LLC 2010

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and an assay to measure cell-mediated immune responsiveness may be considered. Cell-mediated immunity assays can be performed ex vivo (i.e. lymphocyte proliferation assays). Alternatively, cell-mediated immunity testing in vivo may be the preferred option and includes either delayed hypersensitivity or contact hypersensitivity models. In vivo animal models to measure the adverse effects of medicines and chemicals on cellular immune responses are very close to models that can be used to predict the potential for inducing cell-mediated hypersensitivity reactions. One major aspect of cellmediated hypersensitivity to medicines and chemicals is contact hypersensitivity the most common clinical manifestation of which is contact dermatitis. Animal models to assess the potential of chemicals to induce contact sensitization have long since been in use (2). In the context of immunotoxicity testing, in vivo hypersensitivity responses to strong contact sensitizers can be measured to assess the possible immunosuppressive effects of medicines and chemicals.

1.1. Contact Hypersensitivity Contact hypersensitivity is a form of T cell-mediated immunity originally described by Bennacerraf and Gell (3), who showed that it is possible to reproduce the reaction seen to poison ivy and to various chemicals in humans experimentally by painting picryl chloride on the skin of guinea pigs. The critical events that must occur in generating a reaction are: (1) sensitization, (2) trafficking, and (3) elicitation, and these are reflected in the necessary three consecutive phases that characterize every contact hypersensitivity model used for immunotoxicity testing (4). In the sensitization phase, a naïve animal is exposed through the skin to a contact sensitizer that plays the role of a hapten. Usually no symptoms of exposure are seen. The hapten binds covalently to any protein, either a cell-associated or extracellular protein, which results in a chemical modification of the protein. The chemically modified protein can then be presented by antigen-presenting cells (APC) to T cells that recognize the modified protein as foreign. Haptens used experimentally, such as trinitrobenzene sulfonic acid and picryl chloride, primarily bind lysine and other nucleophilic side chains via an amide bond (5). The bulky benzene ring and the modified protein can be recognized by T cells. After phagocytosis of the hapten-protein, APC traffic back into the draining lymph node where the hapten-protein is presented to reactive T cells (6). Memory T cell clones are expanded in the lymph node where then they can generate the elicitation phase of the response.

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In the elicitation phase, local APCs present the hapten-protein to transiting memory T cells, and T cells recruit more inflammatory cells to the antigen deposition site. The Langerhans’ cells have been shown to play an important role by presenting hapten modified-self antigen in the context of MHC class II (7). Keratinocytes can also act as APCs by presenting hapten-modified self-proteins in the context of MHC class I molecules (8). APC, either Langerhans’ cells or another form of APC, must recognize not only the T cell receptor and CD8+ or CD4+ molecule, but also costimulatory molecules. The B7/CD28 ligand system has been shown to be important in regulating the T cell response to contact sensitizers. Finally, there is experimental evidence that CD4+ T cells regulate the magnitude and duration of the CD8+ T cell response. 1.2. General Aspects

Strong contact sensitizers are used to induce a specific cellular response that drugs and chemicals can inhibit if they do exert immunosuppressive effects. Whatever the selected model, the experimental protocol always comprises three consecutive phases. The first phase is the sensitization or induction phase during which the animals are given one or several topical applications of the reference contact sensitizer. The second phase is the rest period of variable duration that is required for the immune system to be prepared for mounting the expected response. The third phase is the eliciting or challenge phase where the same contact sensitizer is applied to the animals again. Overall, there are extremely few, if any, objective data to substantiate selected variations among experimental protocols, the concentration of the contact sensitizer and the number of topical applications during the sensitization phase, the duration of the rest period, and the concentration of the contact sensitizer during the induction phase. Limited efforts have been made to standardize these models. In fact, technical skill and acquired experience are essential so that results obtained with potent immunosuppressive chemicals are generally consistent despite obvious interlaboratory variations in the experimental protocol. These models are primarily used in mice and guinea pigs. Rats are normally not used, as contact sensitization is difficult to obtain in this species.

1.3. Contact Hypersensitivity Testing in Mice

Contact hypersensitivity testing for immunotoxicity assessment in mice typically utilizes ear swelling as the measured endpoint following sensitization by topical skin application(s). It is noteworthy, however, that in the mouse local lymph node assay, which is nowadays the first-line assay to assess the contact sensitizing potential of chemicals, thymidine incorporation in lymphocytes of the draining auricular lymph nodes is the measured endpoint instead of ear swelling. However, in this assay, sensitization still consists in topical applications of the test article on both the ears of mice (9).

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Asherson and Ptak (10) were the first authors to propose and substantiate the use of the ear-swelling test in mice to induce contact hypersensitivity to strong contact sensitizers, including picryl chloride, oxazolone, dinitrifluorobenzene (DNFB), and tetramethyl-p-phenylenediamine. It was later shown by others that earswelling response correlated with in vitro lymphocyte stimulation in mice sensitized with 2,4-dinitro-1-fluorobenzene (DNFB) (11), and that ear swelling induced by topical challenge on the ear of mice sensitized to picryl chloride (12) or DNFB (13) was associated with histological findings consistent with a cell-mediated reaction. A few attempts have been made to standardize the ear-swelling test in the mouse for immunotoxicity testing (14), and therefore, no widely accepted procedure is available. Inbred, as well as outbred, mice have been used, and although some strain-related differences have been shown in the susceptibility of mice to develop contact hypersensitivity, there is no conclusive evidence that ear response to a strong contact sensitizer actually differs sufficiently across mouse strains to induce a significant variability in the response to reference immunosuppressive compounds. In the sensitization (induction) phase, mice are given one or several topical applications. In fact, one topical application is most often sufficient, because a strong contact sensitizer is used. Topical application(s) can be directed on the ear skin (and then usually only one ear is used) or on the shaved skin of the abdomen. Again, no evidence supports the superiority of either procedure. The most commonly used contact sensitizers in the ear-swelling test are picryl chloride (10, 12, 14–19), DNFB (10, 11, 13) and oxazolone (10, 20–22). In contrast to the eliciting phase, an irritating concentration is typically used for the sensitization phase. The duration of the rest period is typically 7–9 days. Thereafter, sensitized animals are challenged (elicitation phase) with a topical application on both sides of one ear (the contralateral ear when topical sensitizing applications were to the skin of one ear) of a nonirritating concentration of the selected reference contact sensitizer. Ear swelling is commonly assessed from the thickness of the ear measured with a dial caliper immediately before challenge in conscious or mildly anesthetized mice and then after 24 h, 48 h, and/or 72 h. The peak of the reaction is normally 48 h. A radioisotopic assay has been proposed but results were very similar to those obtained with the use of a dial caliper (16). The ear-swelling test has been used extensively for pharmacological purposes to test the antiallergic and/or antiinflammatory effects of a number of candidate drugs. In the area of immunotoxicity testing, the ear-swelling test was used successfully to reproduce the immunosuppressive effects of a variety of reference compounds, including chlorpromazine, cyclophosphamide,

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diazepam, haloperidol, hydrocortisone, promethazine, lead, nickel, and selenium after systemic administration (14). The topical administration of the T-2 toxin (20), chlordane (21), and 2-butoxyethanol (22) was also shown to inhibit contact hypersensitivity to oxazolone in mice. Interestingly, the contact hypersensitivity response of mice sensitized to picryl chloride or 2,4,6-trinitro-1-chlorobenzene (TNCB) was shown to be enhanced by systemic treatment with thalidomide (19) or cimetidine (23), respectively, which suggests that contact hypersensitivity testing could be used in the assessment of immunostimulatory as well as immunosuppressive effects. 1.4. Contact Hypersensitivity Testing in the Guinea Pig

Contact hypersensitivity testing in the guinea pig is another possible model to investigate the immunosuppressive effects of medicines and chemicals. In fact, experimental contact dermatitis in guinea pigs sensitized to 1-chloro-2,4-dinitrobenzene (DNCB) was first described 14 years before ear swelling in mice (24), because at that time, mice were considered unable to mount a delayed hypersensitivity response (10). Shortly thereafter, several potent immunosuppressive agents including were shown to decrease contact hypersensitivity to DNCB in guinea pigs (25). Most often, outbred female albino (Dunkin–Hartley) guinea pigs are used. Although picryl chloride or oxazolone has been, but very rarely, used to induce contact hypersensitivity in guinea pigs, the majority of the published data were obtained using DNCB. Sensitization is performed topically on the shaved skin of one flank or the interscapular area. A wide range of concentrations from 7 to 50% has been used for sensitization, and the solvent may also differ, e.g., acetone or ethanol. The duration of the rest period is typically 14 days. Elicitation using a much smaller nonirritating concentration of DNCB (e.g. 0.05%) is applied on the shaved skin of one flank (the controlateral flank if sensitization was on one flank instead of the interscapular area). The intensity of the cutaneous reaction is read after 24 h and assessed semiquantitatively, e.g., 0 = no reaction, 0.5 = some red spots; I = red, confluent spots; and 2 = red, confluent and swollen spots. Groups of 10 animals can be used and an the arithmetic mean of the response can be calculated and compared across experimental treated and nontreated groups. In fact, contact hypersensitivity models in the guinea pig have very rarely been used for preclinical immunotoxicity testing. Chlorpromazine, a psychotropic drug the immunosuppressive effects of which have been described in other animal models and in humans, was shown to decrease contact hypersensitivity response to DNCB in guinea pigs (26). Topical cyclosporine was also shown to exert immunosuppressive effects in this model (27).

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Conclusion Despite species differences that impact on the method to be selected, contact hypersensitivity models are easy to perform and offer the advantage of measuring cellular immunity in vivo to take into account the influence of other systems, such as the nervous and endocrine systems the functions of which may be negatively affected and in turn impact on the cellular immune responses. Although in vivo cell-mediated immunity models have long since been proposed as useful tools for immunotoxicity assessment (28), they are seldom used nowadays. This is particularly true with regard to contact hypersensitivity models that present the major disadvantage of having been applied only to mice or guinea pigs that are not first-line animal species for immunotoxicity assessment. References 1. Descotes J (2006) Methods of evaluating immunotoxicity. Expert Opin Drug Metab Toxicol 2:249–259 2. Kimber I, Maurer T (1996) Toxicology of contact hypersensitivity. Taylor & Francis, London 3. Benacerraf B, Gell PG (1959) Studies on hypersensitivity. III. The relation between delayed reactivity to the picryl group of conjugates and contact sensitivity. Immunology 2:219–229 4. Krasteva M, Kehren J, Ducluzeau MT, Sayag M, Cacciapuoti M, Akiba H, Descotes J, Nicolas JF (1999) Contact dermatitis. I. Pathophysiology of contact sensitivity. Eur J Dermatol 9:65–77 5. Jacobsen C (1975) Trinitrophenylation of the bilirubin binding site of human serum albumin. Int J Pept Protein Res 7:161–165 6. Kripke ML, Munn CG, Jeevan A, Tang JM, Bucana C (1990) Evidence that cutaneous antigen-presenting cells migrate to regional lymph nodes during contact sensitization. J Immunol 145:2833–2838 7. Shelley WB, Juhlin L (1977) Selective uptake of contact allergens by the Langerhans cell. Arch Dermatol 113:187–192 8. Bacci S, Alard P, Dai R, Nakamura T, Streilein JW (1997) High and low doses of haptens dictate whether dermal or epidermal antigenpresenting cells promote contact hypersensitivity. Eur J Immunol 27:442–448 9. Gerberick GF, Ryan CA, Dearman RJ, Kimber I (2007) Local lymph node assay (LLNA) for

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17.

detection of sensitization capacity of chemicals. Methods 41:54–60 Asherson GL, Ptak W (1968) Contact and delayed hypersensitivity in the mouse. I. Active sensitization and passive transfer. Immunology 15:405–416 Phanuphak P, Moorhead JW, Claman HN (1974) Tolerance and contact sensitivity to DNFB in mice. I. In vivo detection by ear swelling and correlation with in vitro cell stimulation. J Immunol 112:115–123 Roupe G, Ridell B (1979) The cellular infiltrate in contact hypersensitivity to picryl chloride in the mouse. Acta Derm Venereol 59:191–195 Cho GY, Hough W (1986) Time course of contact hypersensitivity to DNFB and histologic findings in mice. J Korean Med Sci 1:31–36 Descotes J, Tedone R, Evreux JC (1985) Immunotoxicity screening of drugs and chemicals: value of contact hypersensitivity to picryl chloride in the mouse. Methods Find Exp Clin Pharmacol 7:303–305 Descotes J, Evreux JC (1981) Depressant effects of major tranquillizers on contact hypersensitivity to picryl chloride in the mouse. Experientia 37:1004–1005 Bäck O, Larsen A (1982) Contact sensitivity in mice evaluated by means of ear swelling and a radiometric test. J Invest Dermatol 78: 309–312 Goto Y, Inoue Y, Tsuchiya M, Isobe M, Ueno H, Uchi H, Furue M, Hayashi H (2000)

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19.

20.

21.

22.

Suppressive effect of topically applied CX-659S, a novel diaminouracil derivative, on the contact hypersensitivity reactionb in various animal models. Int Arch Allergy Immunol 123:341–348 Laschi-Loquerie A, Descotes J, Tachon P, Evreux JC (1984) Influence of lead acetate on hypersensitivity. Experimental study. J Immunopharmacol 6:87–93 Descotes J, Tedone R, Evreux JC (1988) Enhancement of antibody response and delayed-type hypersensitivity by thalidomide in mice. Fundam Clin Pharmacol 2:493–497 Blaylock BL, Kouchi Y, Comment CE, Pollock PL, Luster MI (1993) Topical application of T-2 toxin inhibits the contact hypersensitivity response in BALB/c mice. J Immunol 150:5135–5143 Blaylock BL, Newsom KK, Holladay SD, Shipp BK, Bartow TA, Mehendale HM (1995) Topical exposure to chlordane reduces the contact hypersensitivity response to oxazolone in BALB/c mice. Toxicol Lett 81:205–211 Singh P, Morris B, Zhao S, Blaylock BL (2002) Suppression of the contact hypersensitivity response following topical exposure to 2-butoxyethanol in female BALB/c mice. Int J Toxicol 21:107–114

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23. Belsito DV, Kerdel FA, Potozkin J, Soter NA (1990) Cimetidine-induced augmen tation of allergic contact hypersensitivity reactions in mice. J Invest Dermatol 94: 441–445 24. Zeligman I (1954) Experimental contact dermatitis. I. Dinitrochlorobenzene contact dermatitis in guinea pigs. J Invest Dermatol 22:109–120 25. Maguire HC Jr, Maibach HI (1961) Effect of cyclophosphoramide, 6-mercaptopurine, actinomycin D and vincaleukoblastine on the acquisition of delayed hypersensitivity (DNCB contact dermatitis) in the guinea-pig. J Invest Dermatol 37:427–431 26. Descotes J, Evreux JC (1982) Effect of chlorpromazine on contact hypersensitivity to DNCB in the guinea-pig. J Neuroimmunol 2(1):21–25 27. Nakagawa S, Oka D, Jinno Y, Takei Y, Bang D, Ueki H (1988) Topical application of cyclosporine on guinea pig allergic contact dermatitis. Arch Dermatol 124:907–910 28. Luster MI, Dean JH, Boorman GA (1982) Cell-mediated immunity and its application in toxicology. Environ Health Perspect 43:31–36

Chapter 18 Evaluation of Apoptosis in Immunotoxicity Testing Mitzi Nagarkatti, Sadiye Amcaoglu Rieder, Dilip Vakharia, and Prakash S. Nagarkatti Abstract Immunotoxicity testing is important in determining the toxic effects of chemical substances, medicinal products, airborne pollutants, cosmetics, medical devices, and food additives. The immune system of the host is a direct target of these toxicants, and the adverse effects include serious health complications such as susceptibility to infections, cancer, allergic reactions, and autoimmune diseases. One way to investigate the harmful effects of different chemicals is to study apoptosis in immune cell populations. Apoptosis is defined as the programmed cell death, and in general, this process helps in development and maintains homeostasis. However, in the case of an insult by a toxicant, apoptosis of the immune cells can lead to immunosuppression resulting in the development of cancer and the inability to fight infections. Apoptosis is characterized by cell shrinkage, nuclear condensation, changes in cell membrane and mitochondria, DNA fragmentation into 200 base oligomers, and protein degradation by caspases. Various methods are employed in order to investigate apoptosis. These methods include direct measurement of apoptotic cells with flow cytometry and in situ labeling, as well as RNA, DNA, and protein assays that are indicative of apoptotic molecules. Key words: Apoptosis, Immunotoxicity, Caspase, Bcl-2

1. Introduction Immunotoxicity is defined as the adverse effects to the immune response of the host caused by exposure to a toxicant (1). It is caused by chemical substances, medicinal products, airborne pollutants, cosmetics, medical devices, and food additives (2). Exposure to these substances results in many health complications such as susceptibility to cancer, autoimmune diseases, allergic reactions, and inability to fight infections (2). It is important to assess the toxic effects of environmental chemicals, medical device materials, and newly discovered drugs before they induce significant adverse effects on living organisms. Some of the adverse R.R. Dietert (ed.), Immunotoxicity Testing: Methods and Protocols, Methods in Molecular Biology, vol. 598 DOI 10.1007/978-1-60761-401-2_18, © Humana Press, a part of Springer Science + Business Media, LLC 2010

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effects on the immune system include: (1) reduction in antibody production, (2) reduction in cytokine secretion, (3) inability to distinguish between self and nonself, (4) hypersensitivity, and (5) induction of apoptosis in immune cell populations (3). Therefore, precise testing of immunotoxicity is required for the identification of hazardous materials and substances. One reliable way to measure immunotoxicity is to measure apoptosis in immune cell populations. Apoptosis is the process of programmed cell death, during which many changes occur such as membrane blebbing, cell shrinkage, mitochondria leakage, and DNA fragmentation into oligonucleosomal size of ~200 base pairs (4, 5). Apoptosis differs from another form of cell death, which is known as necrosis, in that the latter is characterized by random DNA fragmentation and cell swelling, resulting in lysis and release of cellular contents that induce an inflammatory response. The molecular changes of apoptosis are classified under two different pathways of apoptosis (4, 5). The intrinsic pathway, which occurs via the mitochondria, is initiated by an imbalance in antiapoptotic and proapoptotic members of the Bcl-2 family of proteins. Some of the proapoptotic molecules include Bad, Bid, Bax, Bim, and Bcl-xS while the antiapoptotic molecules are Bcl-2 and Bcl-xL. A shift toward proapoptotic factors results in the permeability of the mitochondrial membrane and cytochrome c leakage from the intermembrane space of the mitochondria into the cytosol. Cytochrome c then combines with pro-caspase 9 and apoptotic protease activating factor-1 (APAF-1) in the presence of adenosine triphosphate (ATP) to form the apoptosome. Active caspase 3 is produced in the apoptosome by cleavage of the pro-caspase 3, and this effector caspase activates endonucleases that cleave the DNA as well as the DNA repair enzyme, poly (ADP-ribose) polymerase (PARP) (6). Caspase 9 activity can be antagonized by the inhibitor of apoptosis proteins (IAPs). The antiapoptotic effect of IAPs is neutralized upon release of the mitochondrial protein known as the second mitochondrialderived activator of caspase or direct inhibitor of apoptosis protein (IAP)-binding protein with low pI (Smac/DIABLO) which acts by sequestering the IAPs (7). The extrinsic pathway is triggered by the ligation of death receptors belonging to the tumor necrosis factor (TNF) receptor family (e.g., TNF-R1 or CD95/Apo-1/Fas) with their respective ligands, TNF or FasL. This receptor–ligand interaction initiates the recruitment of a cascade of signaling molecules including the adaptor proteins TNF-R-associated death domain (TRAD) or Fasassociated death domain (FADD), which along with the recruitment of the procaspase 8 and 10 results in the formation of Death Inducing Signaling Complex (DISC). In the DISC, active caspase 8 and 10 are produced, which are the initiator caspases that activate caspase 3, resulting in apoptosis (4). The two apoptotic pathways

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are linked through the proapoptotic molecule, Bid, a member of the Bcl-2 family of proteins found in the cytosol. Bid is cleaved by caspase-8 to form a truncated protein tBid, which translocates to the mitochondria, where it induces the release of cytochrome c and causes mitochondrial dysfunction leading to activation of caspase 9. Signaling through the death receptors is inhibited by recruitment of an inhibitory protein, FLICE-like inhibitor protein (FLIP), to the DISC.

2. Materials 2.1. Annexin V/Propidium Iodide Staining

1. 12 × 75 mm round bottom culture tubes (VWR International, Westchester, PA). 2. Annexin-V-FLUOS Staining Kit (Roche Applied Science, Indianapolis, IN). 3. Phosphate Buffered Saline (PBS) (VWR International, Westchester, PA). 4. Annexin-V-FLUOS labeling solution: 20 mL Annexin-V-Fluos labeling reagent, 20 mL PI solution, 1 ml incubation buffer. All of these solutions are included in the kit.

2.2. TUNEL Assay

1. 12 × 75 mm round bottom culture tubes (VWR International, Westchester, PA) 2. In situ Cell Death Detection Kit (Roche Applied Science, Indianapolis, IN). 3. Staining Buffer: Phosphate Buffered Saline (PBS) with 1% FBS (Atlanta Biologicals, Lawrenceville, GA). 4. Fixation Solution: freshly prepared 4% paraformaldehyde (Sigma Aldrich, St. Louis, MO) in PBS. 5. Permeabilization Solution: freshly prepared 0.1% Triton-×-100, 0.1% sodium citrate in PBS (Sigma Aldrich, St. Louis, MO). 6. Labeling solution: 22.5 ml of labeling solution (nucleotide mix), 2.5 ml of enzyme (TdT). Both the labeling solution and the enzyme are the components of the kit.

2.3. In situ TUNEL Assay

1. Xylene (Sigma Aldrich, St. Louis, MO). 2. Ethanol (100%, 95%, 85%, 70%, and 50% diluted in deionized water). 3. PBS (VWR International, Westchester, PA). 4. 4% methanol-free formaldehyde (Polysciences, Warrington, PA) in PBS. 5. 0.85% NaCl solution.

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6. Proteinase K buffer (BD Biosciences, San Josa, CA). 7. DNase I (RNase-free) (Promega, Madison, WI). 8. DNase I buffer (Promega, Madison, WI). 9. DeadEnd Fluorometric TUNEL System including equilibration buffer, incubation buffer and 20× SSC (Promega, Madison, WI). 10. Propidium iodide (Sigma Aldrich, St. Louis, MO). 11. Anti-Fade Kit (Invitrogen, Carlsbad, CA). 2.4. Gel Electrophoresis for Detection of DNA Fragmentation

1. TE buffer pH 7.4 (Sigma Aldrich, St. Louis, MO). 2. TTE solution: TE buffer pH 7.4 with 0.2% Triton-×-100 (store at 4°C). 3. NaCl 5 M, ice cold. 4. Isopropanol, ice cold. 5. Ethanol at 70%, ice cold. 6. Bromophenol Blue (Sigma Aldrich, St. Louis, MO). 7. TBE buffer for electrophoresis (Sigma Aldrich, St. Louis, MO). 8. Ethidium bromide solution (Sigma Aldrich, St. Louis, MO). 9. Electrophoresis-grade agarose (Sigma Aldrich, St. Louis, MO). 10. DNA molecular weight markers (Invitrogen, Carlsbad, CA).

2.5. Mitochondrial Membrane Potential

1. DiOC6: 3,3-dihexyloxacarbocyanine iodide (Invitrogen, Carlsbad, CA).

2.6. Detection of Caspase 3/7 Activity

1. Apo-ONE homogenous caspase-3/7 assay kit including 100× substrate buffer (Promega, Madison, WI). 2. Blank: Apo-ONE caspase 3/7 reagent, cell culture medium only. 3. Negative control: Apo-ONE caspase 3/7 reagent, vehicletreated cells.

2.7. Protein Assay

1. Ammonium persulfate (APS) (Sigma Aldrich, St. Louis, MO). 2. Tetramethylethylenediamine (TEMED) (Sigma Aldrich, St. Louis, MO). 3. Sodium dodecyl sulfate (SDS) (Sigma Aldrich, St. Louis, MO). 4. Acrylamide (Sigma Aldrich, St. Louis, MO). 5. Tris-HCl: 0.5 M and 1.5 M (Sigma Aldrich, St. Louis, MO). 6. N¢N¢-bis-methylene-acrylamide (Sigma Aldrich, St. Louis, MO) 7. Glycerol (Sigma Aldrich, St. Louis, MO) 8. Glycine (Sigma Aldrich, St. Louis, MO) 9. Bromophenol blue (Sigma Aldrich, St. Louis, MO) 10. Methanol

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11. NaCl 12. Tween-20 (Sigma Aldrich, St. Louis, MO) 13. Nonfat dry milk 14. 2 beta-mercaptoethanol (Sigma Aldrich, St. Louis, MO) 15. BCA protein assay kit (Pierce, Rockford, IL) 16. Enhanced chemilluminescence (ECL) Reagent (GE HealthCare, Piscataway, NJ) 2.7.1.Western Blot Solutions

1. Transfer buffer: 3.03 g Tris base, 14.4 g glycine, 200 ml ethanol, adjust the volume to 1000 ml. 2. 10× TBS: 24.2 g Tris base, 80 g NaCl, adjust the volume to 1000 ml and pH to 7.6. 3. TBS-T 0.1%: 1 ml of 50% Tween-20, 500 ml 1× TBS 4. 5% Blocking solution: 15 ml 10× TBS, 135 ml deionized (DI) water, 7.5 g nonfat dry milk, mix well, and while stirring add 300 ml of 50% Tween-20. 5. APS: 100 mg ammonium persulfate, 1 ml DI water. 6. Stripping Solution: 3.125 ml 0.5 M Tris-HCl pH 6.8, 5 ml 10% SDS, 180 ml 2-betamercaptoethanol, 16.7 ml DI water. Incubate in 50°C water bath for 15 min. 7. 30% acrylamide (300 ml): 87.6 g acrylamide, 2.4 g N¢N¢bis-methylene-acrylamide, then complete it to 300 ml with DI water. 8. 10% SDS: Gently dissolve 10 g SDS in 90 ml of DI water and bring it up to 100 ml. 9. 1.5 M Tris-HCl ( pH 8.8): 27.23 Tris base, 80 ml DI water, adjust the pH to 8.8 and bring the volume up to 150 ml. 10. 0.5 M -HCl ( pH 6.8): 6 g Tris-base, 60 ml DI water, adjust pH to 6.8 and bring the volume up to 100 ml. 11. Sample Buffer: 0.6 ml DI water, 2 ml 0.5 M Tris-HCl pH 6.8, 5 ml 32% glycerol, 1.6 ml 20% SDS, 0.0002 g bromophenol blue. Prior to use, add 8% 2-mercaptoethanol. 12. 5× SDS-PAGE: 15 g Tris base, 72 g Glycine, 5 g SDS, adjust the volume to 1000 ml.

2.8. RT-PCR Analysis of Gene Expression 2.8.1. Extraction of Total RNA Using Trizol Reagent

1. Trizol Reagent (Invitrogen Corporation Carlsbad, California, Life Technologies, Frederick, MD) can be used to extract total RNA from cells or tissues as per the manufacturer’s instruction. 2. Chloroform (Sigma, St. Louis, MO). 3. Isopropanol (Sigma, St. Louis, MO). 4. Ethanol.

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5. TE buffer (Sigma Aldrich, St. Louis, MO) 6. Polypropylene microcentrifuge tubes. 2.8.2. Extraction of Total RNA Using RNeasy Mini Kit

1. RNeasy Mini kit (Qiagen, Valencia, CA). 2. RNase-free DNase(Qiagen Inc., Valencia, CA). 3. Ethanol (Sigma Aldrich, St. Louis, MO). 4. Polypropylene microcentrifuge tubes. 5. Sterile, RNase-free pipette tips. 6. Disposable gloves.

2.8.3. RT-PCR for Gene Expression

1. PCR tubes (Bioplastics, eEnzyme LLC, Gatthersburg, MD). 2. Mineral Oil (Sigma Aldrich, St. Louis, MO). 3. Bio-Rad iCycler PCR unit (BioRad Laboratories, Hercules, CA). 4. iScript cDNA synthesis kit (BioRad Laboratories, Hercules, CA). 5. PCR reagent kit, Epicenter Master mix (Epicenter, Madison, WI). 6. Moloney murine leukemia virus (Amersham, Arlington Heights, IL).

reverse

transcriptase

7. Oligonucleotide primers (Integrated DNA Technology, Coralville, IA). 8. AlphaImager™ (Alpha Innotech Corporation, San Leandro, CA).

3. Methods A variety of approaches in combinations is used to confirm apoptosis in immune cell populations. Using several methods will also help to determine which pathways are involved and which proteins play a role. For in vitro experiments, a time and a dose response should be performed in all of the experiments in order to determine the optimal time and dose for the induction of apoptosis. If primary immune cells are used directly from in vivo experiments following injection of an apoptosis-inducing chemical, it may be difficult to measure apoptotic cells because the phagocytes in vivo rapidly clear them. To overcome this, we have shown that in vitro culture of such cells for an additional 18–24 h followed by analysis of apoptosis can help to detect apoptotic cells because in such culture conditions, the cells undergoing apoptosis are less likely to come in contact with phagocytic cells (8, 9).

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3.1. Annexin V/Propidium Iodide Staining

As described previously, during apoptosis a cell undergoes many changes. One of the early characteristics of apoptosis is the translocation of phosphatidylserine (PS) from the inner leaflet of the plasma membrane to the outer one. Annexin V binds to PS with high affinity in the presence of Ca2+. Thus, fluorochrome-conjugated Annexin V can be used as a sensitive probe for the detection of PS on the cell surface. Annexin-V-Fluos stain identifies the apoptotic cells, while Propidium Iodide (PI) stains the DNA of necrotic cells.

3.1.1. Procedure

1. Collect the cells exposed to the agent either in vivo or in vitro (see Note 1) and place 106 cells per 12 × 75 mm round bottom culture tube. 2. Add 2 ml of phosphate buffered saline (PBS) per tube and wash the cells at 200×g for 10 min. 3. Discard supernatant and resuspend the cell pellet in 100 ml of Annexin-V-Fluos labeling solution. 4. Incubate for 10–15 min at room temperature. 5. In order to remove any excess fluorochrome, add 2 ml of phosphate buffered saline (PBS) per tube and wash the cells at 200×g for 10 min. 6. Determine fluorescence, using a flow cytometer. Cells that are Annexin V+ PI− are the early apoptotic cells, Annexin V+ PI+ are late apoptotic cells and Annexin V− PI+ are necrotic cells.

3.2. TUNEL to Detect DNA Fragmentation

The terminal deoxynucleotidyl transferase (TdT)-mediated dUTP nick end-labeling (TUNEL) kit is used to detect single- and doublestrand breaks in DNA in individual cells, which is a marker for early apoptotic cells. The kit employs the enzyme TdT, which catalyzes the addition of modified deoxyribonucleotides (dUTP) to the 3¢OH ends of single- and double-stranded nicks. The dUTP is fluorescently labeled and can therefore be detected with a flow cytometer.

3.2.1. Procedure

1. Collect treated cells and place 106 cells per 12 × 75 mm round bottom culture tube. 2. Add 2 ml of staining buffer per tube and wash the cells at 200×g for 10 min. 3. Discard supernatant and resuspend the cell pellet in 100 µl of fixation solution (4% paraformaldehyde in PBS) per tube. The fixation cross-links the DNA to the cellular constituents so it is not lost during the permeabilization step. 4. Incubate the cells at 4°C for 30 min. 5. Wash the cells twice by adding 2 ml of staining buffer per tube and by centrifugation at 200×g for 10 min.

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6. Add 100 ml of permeabilization solution per tube and incubate at room temperature for 5 min. This permits the enzymes to enter the cell through the permeable membrane. 7. Wash the cells twice by adding 2 ml of staining buffer per tube and by centrifugation at 200×g for 10 min. 8. Freshly prepare labeling solution, add 25 ml per tube, and incubate at 37°C for 1 h. 9. Wash the cells twice by adding 2 ml of staining buffer per tube and by centrifugation at 200×g for 10 min (see Note 2) 3.3. In Situ TUNEL Staining

The principle of this assay is identical to that described in TUNEL method, in which apoptotic cells can be visualized in frozen or paraffin-embedded tissue sections, smears, cytospin, or adherent cells. The assay can be fluorometric or colorimetric, depending on the incubation buffer mix. The following assay procedure is used for the fluorometric staining of paraffin-embedded sections.

3.3.1. Procedure

1. Wash paraffin-embedded sections twice by immersing in xylene in a Coplin jar for 5 min. This step deparaffinizes the tissue sections. 2. Immerse the slides in 100% ethanol containing Coplin jars for 5 min at room temperature. 3. Perform graded ethanol washes by immersing slides in 100%, 95%, 85%, 70%, and 50% ethanol respectively, for 3 min per wash. This step rehydrates the tissue sections. 4. Wash the slides first by immersing in PBS for 5 min at room temperature. 5. Place the slides in 4% methanol-free formaldehyde in Coplin jars for 15 min at room temperature. This step fixes the tissue sections. 6. Wash slides with PBS for 5 min. Repeat twice. 7. At this step, dry the slides and place them on a flat surface. 8. Prepare 20 mg/ml Proteinase K solution and add 100 ml to each slide in order to cover the tissue section. Incubate slides for 8–10 min at room temperature. 9. Wash slides with PBS for 5 min at room temperature. 10. Place the slides in 4% methanol-free formaldehyde in Coplin jars for 15 min at room temperature. 11. Once again, wash the slides with PBS for 5 min at room temperature in order to remove 4% methanol-free formaldehyde. 12. Tap the slides on a stack of paper towels in order to remove excess liquid. 13. Add 100 ml of equilibration buffer in order to cover the tissue sections.

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14. Incubate at room temperature for 5–10 min. 15. Prepare rTdT incubation buffer by mixing 45 ml of equilibration buffer, 5 ml of nucleotide mix, and 1 ml of rTdT enzyme per reaction. 50 ml is required per reaction. 16. Thaw the nucleotide mix on ice, always keep the incubation buffer on ice and protect it from light (see Note 3). 17. Using a paper towel, blot around the tissue in order to remove some of the equilibration buffer. 18. Add 50 ml of incubation buffer on each section and cover the tissue section with a plastic cover slip. The cover slip ensures even staining of the sections. 19. Place the slides in a humidified chamber, cover with foil and incubate at 37°C for 60 min. 20. Make up 2× SSC from stock by mixing 5 ml of 20× SSC with 45 ml of deionized water and fill a Coplin jar with the solution. 21. Remove the coverslips and immerse slides in 2× SSC for 15 min at room temperature. 22. Wash three times with fresh PBS for 5 min at room temperature. This step ensures the removal of unincorporated fluorescein-dUTP. 23. Prepare 40 ml of 1 mg/ml propidium iodide in PBS and fill a Coplin jar. 24. Place slides in PI solution for 15 min at room temperature in the dark. 25. Wash with deionized water for 5 min at room temperature. Repeat this step two more times. 26. Remove excess water from the slides. 27. Add one drop of antifade solution to the area. 28. Mount the slides using glass coverslips. 29. Seal the edges with clear nail polish and let them dry for 5–10 min. 30. Immediately, analyze slides under a fluorescent microscope. The green fluorescence of fluorescein can be viewed at 520 ± 20 nm and the red fluorescence of PI can be viewed at >620 nm. 3.4. Gel Electrophoresis for Detection of DNA Fragmentation

1. Dispense 1–5 × 106 cells in tubes. 2. Centrifuge the cells at 200×g at 4°C for 10 min. 3. Add 0.5 ml of TTE solution to the pellet and vortex vigorously. This procedure allows the release of fragmented chromatin from nuclei after cell lysis with Triton-×-100 and disruption

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of the nucleus following Mg++ chelation by EDTA in the TTE solution. 4. Centrifuge tubes at 20,000×g for 10 min at 4°C to separate fragmented DNA from intact chromatin. 5. Transfer supernatants to new tubes. 6. Add 0.5 ml of TTE solution to the pellet. 7. Add 0.1 ml of ice-cold 5 M NaCl to the 0.5 ml volume present in tubes having the supernatants or pellet and vortex vigorously. The addition of the salt removes histones from DNA. 8. Add 0.7 ml of ice-cold isopropanol to each tube and vortex vigorously. 9. Allow precipitation to proceed overnight at −20°C. 10. Pellet DNA by centrifugation for 10 min at 20,000×g at 4°C. 11. Discard supernatants by inverting the tubes. 12. Rinse the pellets by adding to each tube 0.5–0.7 ml of icecold 70% ethanol. 13. Centrifuge the tubes at 20,000×g for 10 min at 4°C. 14. Discard supernatants by rapidly inverting tubes. 15. Air dry the tubes for about 3 h. 16. Dissolve DNA by adding to each tube 20–50 ml of TE solution at 37°C for 1–3 days. 17. Mix DNA with 10× loading buffer to a final concentration of 1×. 18. Place the samples in a heating block at 65°C for 10 min. 19. Immediately load 10–20 ml of to each well of a standard 1% agarose gel containing ethidium bromide 0.5 mg/ml. 20. Appropriate DNA molecular weight markers should be included. Ethidium bromide is a potential carcinogen and should only be handled while wearing gloves. 21. Run the gel electrophoresis in TBE buffer at the desired voltage. The migration of samples is followed by the migration of bromophenol blue dye contained in the loading buffer. 22. Stop the electrophoresis when the dye reaches about 3 cm from the end of the gel. 23. To visualize DNA, place the gel on a UV transilluminator and take photos of the gel. Wear eye and skin protection when UV is on. DNA laddering is characteristic of apoptosis. 3.5. Mitochondrial Membrane Potential

During the earliest stages of apoptosis, even before the exposure of phosphatidylserine, there is loss in mitochondrial membrane

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potential and the mitochondria become leaky. This leads to translocation of cytochrome c into the cytosol and further advancement of the intrinsic apoptotic pathway. One easy way to measure mitochondrial membrane potential is with a potential-sensitive, membrane-permeable, lipophilic cation dye called 3,3-dihexyloxacarbocyanine iodide (DiOC6), which exhibits fluorescence after accumulation into energized systems. 3.5.1. Procedure

1. Culture immune cells at a density of 5 × 106 cells/ml with the toxicant. 2. Add 3,3-dihexyloxacarbocyanine iodide (DiOC6) at a final concentration of 40 nM 15 min prior to the end of incubation time. 3. After the incubation time, harvest the cells and place in 12 × 75 mm round bottom culture tubes. 4. Assess mitochondrial membrane potential (Dym) by using a flow cytometer with excitation and emission settings of 488 and 525 nM. A decrease in the fluorescence intensity is indicative of a loss of mitochondrial membrane potential (see Note 4).

3.6. Detection of Caspase Activity

The extrinsic apoptotic pathway is characterized by the activation of caspase 8, while the intrinsic pathway is characterized by the activation of caspase 9, and both of these pathways intersect at the activation of caspase 3. The existence of these proteins is directly proportional to the amount of apoptosis, and it can be measured with available reagent kits or with immunoblotting. Please refer to the next subheading (18.2.7 Protein assays) for immunoblotting protocols. The Caspase-Glo Assays from Promega use the luminogenic caspase-8 tetrapeptide substrate (Z-LETD-aminoluciferin), the caspase-9 tetrapeptide substrate (Z-LEHD-aminoluciferin), or the caspase-3/7 substrate (Z-DEVD-aminoluciferin), and a stable luciferase in buffer, which induces cell lysis and can detect luciferase activity. Upon cleavage of the substrate by the caspase, aminoluciferin is liberated and can contribute to the generation of light in a luminescence reaction. This section provides the assay protocol for the Apo-ONE homogenous caspase 3/7 assay kit.

3.6.1. Procedure

1. Incubate the blank, the negative control and the experimental samples in a 96 well-plates. 2. Thaw 100× substrate buffer to room temperature. 3. Prepare the Apo-ONE caspase 3/7 reagent by making a 1:100 dilution. 4. Add 100 ml to each well of the 96-well plate. 5. Mix the contents of the plate with a shaker at 300–500 rpm for 30 s.

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6. Incubate the plate at room temperature between 30 min to 18 h. 7. Perform a time response in order to determine the optimal time for caspase activity. 8. Measure the fluorescence of the wells with excitation wavelength at 499 nm and emission wavelength at 512 nm by using a plate reader. 3.7. Western Blotting for Protein Estimation

An increase in some important proapoptotic proteins and/or a decrease in the antiapoptotic molecules within a cell is an indicator of apoptosis. In the intrinsic pathway, the Bcl family proteins play a significant role as well as the leakage of cytochrome c from the mitochondria into the cytosol. Both in the intrinsic and extrinsic pathway of apoptosis, the effector caspase 3 plays a significant role, and thus, the cleavage of procaspase 3 to caspase 3 is assayed. PARP, an enzyme that is responsible for DNA repair, is also cleaved by caspase 3. Thus, caspase 3 activity is studied by estimating PARP cleavage. Furthermore, the cleavage of procaspase 8 and 10 is indicative of the involvement of the death receptor pathway, while cleavage of procaspase 9 denotes the induction of the mitochondrial pathway. Detection of the proapoptotic truncated Bid suggests cross-talk between the death receptor and mitochondrial pathways. All of these proteins can be quantitated by preparing lysates from cells as well as the cytosolic and mitochondrial fractions and immunoblotting for the protein of interest.

3.7.1. Procedure

1. Place the small plate onto a 1 mm spacer plate and slide the plates into the frame. 2. Tighten the clamps and put on the stand with the short plate facing you. 3. Insert a comb and make sure to leave 1 cm between the bottom of the loading lanes and the top of the resolving gel. 4. Prepare a resolving gel: 3.4 ml DI water, 2.5 ml 1.5 M Gel Buffer, 4 ml 30% acrylamide, 100 µl 10% SDS 5. In the last minute of gel preparation, add 50 ml 10% APS and 5 ml TEMED for the resolving gel. 6. Pour the resolving gel and add a few drops of 0.1% SDS in order to form a uniform-shaped gel. Cover the gel with water. 7. Once the gel polymerizes, drain the water and remove excess water with filter paper. 8. Prepare the stacking gel. 9. Stacking gel: 6.1 ml DI water , 2.5 ml 1.5 M Gel Buffer , 1.3 ml 30% acrylamide, 100 ml 10% SDS

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10. In the last minute of gel preparation, add 50 ml 10% APS and 10 ml TEMED for the stacking gel. 11. Pour the gel on top of the resolving gel and push the comb so that the top of the wells corresponds to the top of the glass. 12. Place the gel into the electrode assembly with the short plate on the inside. 13. Fill the center of chamber with buffer. 14. Denature samples as well as the ladder by boiling or using the PCR machine at 95°C for 3 min. 15. Load the ladder and the samples at the concentration of 15 mg of protein/lane. 16. Fill the empty wells as well as the outside part of the chamber with loading buffer and try to avoid air bubbles. 17. Run the samples on high at 200 V until the samples reach the buffer. 18. Saturate Whatman papers, the transfer membrane, and the brillo pads with transfer buffer. 19. In order to perform the gel transfer, disassemble the apparatus, and wash the chamber. 20. Place a Whatman paper on a brillo pad then place both on the black side of the sandwich device. 21. Separate the plate carefully, remove the gel and cut the bottom of the first lane. 22. Put the stacking gel on the Whatman paper with the cut on the bottom right corner. 23. Cut the right corner of the transfer membrane and place it onto the gel. 24. Place another wet Whatman paper on the gel, followed by the brillo pad, and close the chamber. 25. Put the black chamber into the transfer chamber containing an ice block and add enough transfer buffer until it reaches the top of the first row of holes. Run at 100 V for 1–2 h. 26. Separate the membrane from the gel, place it in the block solution protein side up and incubate on a shaker for 1 h. 27. Wash the membrane three times with 0.1% TBS-T by shaking at room temperature for 10 min each. 28. Prepare the primary antibody solution in 5% blocking solution, add it to the membrane, and shake at 4°C overnight. 29. Wash the membrane three times with 0.1% TBS-T by shaking at room temperature for 10 min.

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30. Prepare the secondary antibody solution in 5% blocking solution, add it to the membrane, and shake at room temperature for 1 h. 31. Wash the membrane three times with 0.1% TBS-T by shaking at room temperature for 10 min. 32. Freshly prepare enhanced chemilluminescence (ECL) solution according to the manufacturer’s directions. 33. Cover the blot with the solution, keeping the protein side up.v 34. Incubate for 1 min. 35. Remove air bubbles. 36. Develop the blot by using a fast film, usually XOMAT for 2–20 min. 3.8. RT-PCR Analysis of Gene Expression

Yet another way to detect the presence of the apoptotic molecules such as Fas and FasL is to use reverse transcriptase-polymerase chain reaction (RT-PCR). However, this is a semiquantitative assay and does not depict the protein expression. Earlier studies have used an extraction procedure that involved the partitioning of RNA into aqueous and organic phases for separation. In more recent studies, commercially available cartridges that use adsorption technology to purify RNA are used. Unlike the older techniques, the recent methods are time-saving and yield good quality total RNA without major concerns for its degradation. We describe both of the methods here.

3.8.1. RNA Extraction Using Trizol Reagent

1. Isolate total RNA from immune cells using Trizol Reagent. 2. Homogenize 100 mg of tissue in a glass homoginizer in 1 ml of Trizol reagent. Mix the homogenate with repetitive pipetting. 3. Transfer to a 2 ml polypropylene microfuge tube. 4. Pellet 5 million cells in a 2 ml microfuge tube and lyse in 1 ml of Trizol reagent with repetitive pipetting. 5. After five minutes at room temperature to allow complete dissociation of nucleoprotein complexes, add 0.2 ml of chloroform. 6. After the closing the lid, vigorously shake the tubes for 15 s. 7. After standing at room temperature for 15 min, spin tubes at 12,000×g for 15 min at 4°C to separate RNA containing aqueous phase (top) from DNA (interface) and protein containing organic phase (bottom). 8. Transfer the aqueous phase to a fresh tube and precipitate RNA by mixing with 0.5 ml of isopropanol.

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9. After standing at room temperature for 10 min, centrifuge at 12,000×g for10 min at 4°C to pellet RNA. 10. Wash RNA precipitate (not often visible) by suspending in 1 ml of 75% ethanol. 11. Vortex the tube and centrifuge at 7,500×g for 5 min at 4°C. 12. After removing ethanol, partially air dry the RNA pellet for 3–5 min and dissolve in 20–30 ml of double distilled water or TE buffer by warming at 55°C for 20 min. 13. The optical density at 260 nm of 1.0 corresponds to 0.04 mg/ml of RNA. The A260/280 ratio should be ³1.8 (see Note 5). 3.8.2. Extraction of Total RNA Using RNeasy Mini Kit

The commercially available RNAeasy Mini kit can isolate RNA longer than 200 bp within 30 min of cell lysis, without using alcohol precipitation or chloroform extraction steps. It uses a silicabased membrane and the speed of microspin technology to isolate highly pure RNA that is suitable for a wide range of applications.

Procedure

1. First lyse biological samples and homogenize in the presence of the denaturing reagent guanidine isothiocyanate containing buffer, which immediately inactivates RNases to ensure purification of intact RNA. 2. Add ethanol to provide appropriate binding conditions. 3. Apply the sample to an RNeasy Mini spin column, where the total RNA binds to the membrane and contaminants are efficiently washed away. 4. Elute high-quality RNA in 30–50 ml water. 5. Remove any residual DNA contamination that might affect PCR reaction by using a RNase-free DNase.

3.8.3. RT-PCR

The gene expression of proapoptotic and antiapoptotic genes can be measured by RT-PCR.

Procedure

1. Use 10 mg of RNA from each sample for reverse transcription to synthesize cDNA, using Moloney murine leukemia virus reverse transcriptase 2. Subject the cDNA samples to PCR amplification using synthetic oligonucleotide primers for proapoptotic genes such as Fas, FasL, or b-actin . Use b-actin as an internal control along with other genes of interest in PCR tubes together with PCR reagents supplied in Epicenter Master mix kit in amounts recommended by the manufacturer. The conditions of the PCR vary and may be as follows: denaturation at 94°C for 30 s, annealing at 56°C for 1 min, and extension at 72°C for 2 min. Totally, 30 cycles are performed. The following primers will be

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used: Fas sense primer, 5¢-GCACAGAAGGGAAGGAGTAC-3¢; Fas antisense primer, 5¢ GTCTTCAGCAATTC-TCGG GA-3¢ (amplified fragment, 455 bp); FasL sense primer, 5¢-GAGAAGGAAA-CCCTTTCCTG-3¢; FasL antisense primer, 5¢-ATATTCCTGGTGCCCATGAT-3¢ (amplified fragment, 940 bp); mouse b-actin sense primer, 5¢-ATCCTGACCCTGAACTACCCCATT-3¢; and b-actin antisense primer, 5¢-GCACTGTAGTTTCTCTTCGACACGA-3¢ (amplified fragment, 464 bp). 3. Electrophorese the PCR products through a 1.5% agarose gel containing ethidium bromide (Subheading 18.2.4). 4. Quantitatively measure the PCR products by taking a digital photograph of the gel exposed to UV light, using the imager that contains the UV source, a digital camera, and the software for computing the intensity of each band.

4. Notes 1. The immune cells may be cultured in RPMI 1640 supplemented with 10% FBS, 10 mM HEPES, 50 mM 2-mercaptoethanol, 100 U/ml penicillin, and 100 mg/ml streptomycin in the presence of vehicle or different concentrations of the toxicant for 24–48 h for the detection of apoptosis. 2. Positive control slides should be prepared for in situ TUNEL assay. After step 11 in subheading 18.2.3, add 100 ml of DNase I buffer to the slide and incubate at room temperature for 5 min. Then, remove the excess liquid, add 5–10 units/ ml of DNase I in DNase buffer to the slides, and incubate at room temperature for 10 min. Wash the slide at least three times with deionized water in a Coplin jar and continue with the staining steps. It is important to use separate Coplin jars for the positive control, since DNase contamination may affect the experimental slides. 3. The colorimetric DeadEnd TUNEL system kit may be obtained from a commercial source such as Promega (Cat. #G7130). The assay is very similar to the fluorometric assay, except that the rTdT reaction mix includes biotinylated nucleotide mix. Additional steps are necessary such as blocking the endogenous peroxidase, addition of peroxidase-labeled streptavidin followed by the substrate, hydrogen peroxide, and the chromogen, 3,3¢-diaminobenzidine (DAB), provided in the kit. 4. In the mitochondrial membrane potential detection assay (Subheading 18.2.5), expose the cells to 5 mmol/L carbamoyl cyanide m-chlorophenylhydrazone for 15 min at 37°C for a positive control.

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5. RNA is very sensitive to degradation by RNases and, therefore, precautions should to taken to prevent their introduction by wearing gloves, using autoclaved distilled water and buffers and dry baked plastic or glassware (Subheading 18.2.8). If necessary, prepare all the solutions with DEPC (diethylpyrocarbonate)-treated water, using 0.1% (v/v) DEPC in distilled water (dH2O). Allow the DEPC-treated water to incubate overnight at room temperature then autoclave the DEPCtreated water prior to use. Do not use DEPC-treated water to dissolve the final RNA pellet, as it interferes with optical density readings.

Acknowledgments This work was supported in part by NIH grants R01AI053703, R01ES09098, R01 AI058300, R01DA016545, R01HL058641, and P01AT00396. References 1. Vos J, Loveren V, Wester P, Vethaak D (1989) Toxic effects of environmental chemicals on the immune system. Trends Pharmacol Sci 10(7):289–292 2. Descotes J, Choquet-Kastylevsky G, Van Ganse E, Vial T (2000) Responses of the immune system to injury. Toxicol Pathol 28(3):479–481 3. Descotes J (2005) Immunotoxicology: role in the safety assesment of drugs. Drug Saf 28(2):127–136 4. Hengartner MO (2000) The biochemistry of apoptosis. Nature 407(6805):770–776 5. Igney FH, Krammer PH (2002) Death and anti-death: tumour resistance to apoptosis. Nat Rev Cancer 2(4):277–288

6. Kroemer G, Reed JC (2000) Mitochondrial control of cell death. Nat Med 6(5):513–519 7. Marsden VS, Strasser A (2003) Control of apoptosis in the immune system: Bcl-2, BH3only proteins and more. Annu Rev Immunol 21:71–105 8. Kamath AB, Nagarkatti PS, Nagarkatti M (1997) Evidence for the induction of apoptosis in thymocytes by 2, 3, 7, 8-tetrachlorodibenzo-p-dioxin in vivo. Toxicol Appl Pharmacol 142(2):367–377 9. Camacho IA, Hassuneh MR, Nagarkatti M, Nagarkatti PS (2001) Enhanced activationinduced cell death as a mechanism of 2, 3, 7, 8-tetrachlorodibenzo-p-dioxin(TCDD)induced immunotoxicity in peripheral T cells. Toxicology 165(1):51–63

Chapter 19 Dendritic Cells in Immunotoxicity Testing Donghong Gao and David A. Lawrence Abstract Dendritic cells (DCs) are now recognized to play the key role in the development of adaptive immunity by promoting activation of naïve T cells. Herein, we describe the methodologies to investigate how DCs can be modified by an environmental toxicant and subsequently influence immunity. The prototypic toxicant used as an example for altering DC development and functional influences on T cell development is lead (Pb). It has been reported that the environmental exposure to Pb enhances IgE production in children, which leads to an increase in the incidence of asthma. This effect has been suggested to be due to the preferential enhancement of helper T cell type 2 (Th2) cell responses by Pb. The predominant promotion of Th2 cell development is posited to be due to the altered characteristics of the bone marrow (BM)-DCs from Pb-treated mice (Pb-DCs) when compared to those of the BM-DCs that develop from progenitors in the absence of Pb. The Pb-DCs have a different immunophenotype as well as different cytokine expression after activation. In vitro and in vivo studies confirm that Pb-DCs have the ability to promote antigen-specific T cells to Th2 cells, favoring type-2-related humoral (HI) and cell mediated (CMI) immunity, which may be extracellular signal-regulated kinase (Erk)/mitogen-activated protein (MAP) kinase pathway dependent. Key words: Murine bone marrow derived dendritic cells, Pb, Cytokine, Phenotype, Th2

1. Introduction Dendritic cells (DCs) are the most important antigen-presenting cell (APC) for naïve T cells. It has been suggested that their phenotype, cytokine profiles, intracellular events, and costimulation signals affect their ability to direct T cell activation and differentiation (1–4). It is important to realize that like T cells, B cells, natural killers (NK) cells and macrophages, there are subsets of DCs. Based on expression of surface markers, there are at least three different types of DCs, and the differences among the DC subsets include differences in the expression of cytokines that

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influence the activation and expansion of different types of T cells. Additionally, the DC subsets of humans and mice have differences, and they likely develop from progenitors with different growth and differentiation factors. For example, in addition to granulocyte macrophage-colony stimulating factor (GM-CSF), TNF-a is required for the generation of DCs from human CD34+ bone marrow precursors (5) and human CD34+ hematopoietic progenitor cells (6). BM-DCs are not only sensitive to immunosuppressive and anti-inflammatory drugs, dietary products (7–11), but also sensitive to tick saliva (12), Thimerosal (13), the neurotransmitter norepinephrine (14) and environmental toxicants, such as Pb (15), and atrazine (16). It is not surprising that the chemicals that compromise cellular thiols, such as Pb alter DC structure/function since it is known that DCs are affected by oxidative stress and psychological stress, which can also induce oxidative products (17–24). In this chapter, we focus on mouse DCs and the environmental toxicant Pb is used to demonstrate the methods that can determine the association of changes to the characteristics of DCs from BM-DCs and the resultant influences on T cell activity. Pb preferentially promotes Th2 response in vivo and in vitro (25–27). Shown is how Pb modulates the development and APC function of BM-DCs (15). After Pb exposure during development stage, BM-DCs are phenotyped by using flow cytometry. Flow cytometers with simultaneous analysis of four or more cellular antigens is used to delineate the immunophenotype of DCs with fluorochrome conjugated antibodies that are specific for antigens known to be on DC subsets. Commercially available ELISA kits provide a useful tool to detect BM-DC cytokine production, T cell cytokine production, and serum antibody IgG1, IgG2a concentration. Th2 skewing is determined based on the T cell cytokine production, serum IgG1 and IgG2a concentration and ratio, as well as DTH responses.

2. Materials 2.1. Mice

1. BALB/c, C57BL/6, and BALB/c DO11.10 (OVA-reactive TCR transgenic mice, OVAtg) (2- to 4-month old) are obtained from the Wadsworth Center animal production unit. Mice are housed in a specified pathogen-free AAALAC-approved facility and maintained on mouse chow and acidified water ad libitum.

2.2. Generation of Bone Marrow DCs

1. Sterile 1× DPBS without Calcium or Magnesium (Sigma, St. Louis, MO). 2. Physiological saline (Baxter, Deerfield, IL).

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3. Supplement completed RPMI 1640 (Gibco, Grand Island, NY) with 1 mM nonessential amino acids, 1 mM sodium pyruvate, 1% sodium bicarbonate from BioWhittaker, 2 mM glutamine (Sigma, St. Louis, MO), 50 mM b-mercaptoethanol (Fluka, Ronkonkoma, NY), 25 mg/ml gentamicin (Sigma, St. Louis, MO), 1% penicillin–streptomycin–neomycin mixture (Gibco, Grand Island, NY), and 10% heat-inactivated fetal bovine serum (FBS, HyClone, Logan, UT). 4. 70% ethanol (v/v) 200 proof Ethyl Alcohol USP (Aaper Alcohol and Chemical CO., Shelbyville, Kentucky) and distilled deionized H2O (dd H2O) (Barnstead NANOpure Dlamond Life Science (UV/UF) ultrapure water system, Dubuque, Iowa). 5. Murine GM-CSF (Peprotech, Rocky Hill, NJ). 6. Prepare 10 mM PbCl2 (Fisher Scientific, Pittsburgh, PA) stock in physiological saline (Baxter, Deerfield, IL) and store at room temperature. 7. Lipopolysaccharide (Sigma, St. Louis, MO). 8. Dissolve chicken Egg Albumin (OVA) (Calbiochem, San Diego, CA) at 10 mg/ml in physiological saline (Baxter, Deerfield, IL) and store in single use aliquots at −80°C. 9. Limulus Amebocyte Lysate QCL-1000 (Biowhittaker, Walersville, MD) (see Note 1). 10. 60 × 15 mm tissue culture dishes (Falcon, Becton Dickinson, Franklin Lakes, NJ). 11. 100 × 15 mm Petri dishes (Falcon, Becton Dickinson, Franklin Lakes, NJ). 12. 100 × 20 mm tissue culture dishes (Falcon, Becton Dickinson, Franklin Lakes, NJ). 13. 15 ml polypropylene conical tubes (Falcon, Becton Dickinson, Franklin Lakes, NJ). 14. 50 ml polypropylene conical tubes (Falcon, Becton Dickinson, Franklin Lakes, NJ). 15. ½ CC Tuberculin Syringe with 27 G ½ diameter needle (Becton Dickinson, Franklin Lakes, NJ). 16. 5 ml Costar pipets (Corning Inc., Corning, NY). 17. 10 ml Falcon pipets (Becton Dickinson labware, Franklin Lakes, NJ). 18. CR-600 refrigerated centrifuge (International Equipment Company, Needham Heights, MA). 19. Coulter Particle counter (Becton Dickinson, Franklin Lakes, NJ). 20. Low O2 incubator (Forma Scientific Inc., Marjetta, OH).

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2.3. Immunization

1. Dissolve OVA (Calbiochem, San Diego, CA) at 10 mg/ml in physiological saline (Baxter, Deerfield, IL) and stored in single use aliquots at −80°C. 2. Physiological saline (Baxter, Deerfield, IL). 3. Plain capillary tubes (Fisher Ascientific Co., Pittsburgh, PA). 4. ½ CC Tuberculin Syringe with 27 G ½ diameter needle (Becton Dickinson, Franklin Lakes, NJ).

2.4. Blood Collection and Serum Preparation

1. Plain capillary tubes (Fisher Scientific Co., Pittsburgh, PA). 2. 1.7 ml and 0.6 ml micro-centrifuge tubes (Axygen Scientific, Inc., Union City, CA). 3. Eppendorf 5415C centrifuge (Brinkmann Instruments, Inc., Westbury, NY).

2.5. Measurement of Antigen Specific IgG Isotype in Serum by ELISA

1. Dissolve OVA (Calbiochem San Diego, CA) at 10 mg/ml in physiological saline (Baxter, Deerfield, IL) and stored in single use aliquots at −80°C. 2. Biotin-anti-mouse IgG1 and IgG2a (BD Bioscience, San Diego, CA). 3. Mouse IgG1 Anti-OVA monoclonal antibody (mAb) (Sigma, St., Louis, MO). 4. Coating buffer: 0.1 M NaHCO3, pH 9.5. Dissolve 0.1 M NaHCO3 (Baker, Inc., Phillipsburg, NJ) and 0.034 M NaCO3 (Baker, Inc.) in dd H2O (Barnstead NANOpure Dlamond Life Science (UV/UF) ultrapure water system, Dubuque, Iowa), adjust pH to 9.5. 5. Wash buffer: 0.05% Tween 20 (Sigma, St. Louis, MO) in 1× PBS, pH 7.2–7.4. 6. 1% BSA: dissolve (W/V) BSA (Sigma, St. Louis, MO) in 1× PBS, adjust pH to 7.2–7.4, sterilize with 0.22 mm filter (Corning Inc., Corning, NY). 7. 1× PBS: 137 mM NaCl, 2.7 mM KCl, 8.1 mM Na2HPO4, 1.5 mM KH2PO4, pH 7.2–7.4, 0.22 mm (Corning Inc., Corning, NY) filtered. 8. Substrate solution: 3, 3¢, 5, 5¢-Tetramethyl-benzidine Liquid Substrate (Sigma, Louis, MO). 9. Stop solution: 2 N H2SO4. Make from diluting 18 N H2SO4 (Baker Chemical Co., Phillipsburg, NJ) with dd H2O (Barnstead NANOpure Dlamond Life Science (UV/UF) ultrapure water system, Dubuque, Iowa). 10. ELISA plates (Corning Inc., Corning, NY). 11. ELISA reader (Bio-Tek, Burlington, VT).

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1. Mouse IgE ELISA Set (SB Biosciences, San Diego, CA) including Capture Antibody, Detection Antibody, StreptavidinHRP, and standards. 2. 1× PBS: 137 mM NaCl, 2.7 mM KCl, 8.1 mM Na2HPO4, 1.5 mM KH2PO4, pH 7.2–7.4, 0.22 mm (Corning Inc., Corning, NY) filtered. 3. Coating buffer: 0.1 M NaHCO3, pH 9.5. Dissolve 0.1 M NaHCO3 (Baker, Inc., Phillipsburg, NJ) and 0.034 M NaCO3 (Baker, Inc., Phillipsburg, NJ) in dd H2O (Barnstead NANOpure Dlamond Life Science (UV/UF) ultrapure water system, Dubuque, Iowa), adjust pH to 9.5. 4. Wash buffer: 0.05% Tween 20 (Sigma, Louis, MO) in 1× PBS, pH 7.2–7.4. 5. Assay diluent: 10% FBS (HyClone, Logan, UT) in 1× PBS. 6. Substrate solution: 3, 3¢, 5, 5¢-Tetramethyl-benzidine Liquid Substrate (Sigma, Louis, MO). 7. Stop solution: 2 N H2SO4. Make by diluting 18 N H2SO4 (Baker Chemical Co., Phillipsburg, NJ) with dd H2O (Barnstead NANOpure Dlamond Life Science (UV/UF) ultrapure water system, Dubuque, Iowa). 8. ELISA plates (Corning Inc., Corning, NY). 9. ELISA reader (Bio-Tek, Burlington, VT).

2.7. ELISA for Antigen Specific IgE in Serum

1. Mouse IgE ELISA Set (SB Biosciences, San Diego, CA) including Anti-mouse IgE (capture antibody), Biotinylated anti-mouse IgE (Detection antibody), Streptavidin-HRP, and recombinant mouse IgE (standard). 2. Dissolve OVA (Calbiochem, San Diego, CA) at 10 mg/ml in physiological saline (Baxter, Deerfield, IL) and store in single use aliquots at −80°C. 3. Make biotin-anti-OVA by labeling anti-OVA antibody (Sigma, St. Louis, MO) with 6-((biotinoyl)amino) hexanoic acid (Molecular Probes, Eugene, OR). 4. 1× PBS: 137 mM NaCl, 2.7 mM KCl, 8.1 mM Na2HPO4, 1.5 mM KH2PO4, pH 7.2–7.4, 0.22 mm (Corning Inc., Corning, NY) filtered. 5. Coating buffer: 0.1 M NaHCO3, pH 9.5. Dissolve 0.1 M NaHCO3 (Baker, Inc., Phillipsburg, NJ) and 0.034 M NaCO3 (Baker, Inc., Phillipsburg, NJ) in dd H2O (Barnstead NANOpure Dlamond Life Science (UV/UF) ultrapure water system, Dubuque, Iowa), adjust pH to 9.5. 6. Wash buffer: 0.05% Tween 20 (Sigma, St. Louis, MO) in 1× PBS, pH 7.2–7.4.

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7. Assay diluent: 10% FBS (HyClone, Logan, UT) in 1× PBS. 8. Substrate solution: 3, 3¢, 5, 5¢-Tetramethyl-benzidine Liquid Substrate (Sigma, St. Louis, MO). 9. Stop solution: 2 N H2SO4. Make by diluting 18 N H2SO4 (Baker Chemical Co., Phillipsburg, NJ) with dd H2O (Barnstead NANOpure Dlamond Life Science (UV/UF) ultrapure water system, Dubuque, Iowa). 10. ELISA plates (Corning Inc., Corning, NY). 11. ELISA reader (Bio-Tek, Burlington, VT). 2.8. Flow-Cytometric Analysis

1. 1× PBS: 137 mM NaCl (Mallinckrodt Baker, Inc., Phillipsburg, NJ), 2.7 mM KCl (Fisher Scientific, Pittsburgh, PA), 8.1 mM Na2HPO4 (Baker Chemical Co., Phillipsburg, NJ), 1.5 mM KH2PO4 (Fisher Scientific, Pittsburgh, PA), pH 7.2–7.4, 0.22 mm (Corning Inc., Corning, NY) filtered. 2. Staining buffer: 0.1% NaN3 (Fisher Scientific, Pittsburgh, PA) in 1× PBS. 3. Fixing buffer: 1% paraformaldehyde (Fisher Scientific, Pittsburgh, PA), 0.1% NaN3 (Fisher Scientific, Pittsburgh, PA) in 1× PBS. 4. Fc block (BD Bioscience, San Diego, CA). 5. Fluorescein-conjugated mAbs (BD Bioscience, San Diego, CA or Caltag Laboratories, Burlingame, CA). 6. Flow cytometry (Becton Dickinson & Co., Mountain View, CA).

2.9. Splenocyte Preparation

1. Sterile 1× DPBS without calcium and magnesium (Sigma, St. Louis, MO). 2. Make red blood cell lysing buffer from 0.017 M Tris (Sigma, St. Louis, MO) and 0.75% (w/v) NHCl4 (Sigma, St. Louis, MO), pH 7.4. Sterilize the buffer by passing through 0.22 mm filter system (Corning Inc., Corning, NY) and store at 4°C. 3. 60 × 15 mm tissue culture dish (Falcon, Becton Dickinson, Franklin Lakes, NJ). 4. Frosted microscope slides (Erie Scientific Co., Portsmouth, NH). 5. 15 ml polypropylene tube (Falcon, Becton Dickinson, Franklin Lakes, NJ). 6. Pasteur capillary pipets (Fisher Scientific Co., Pittsburgh, PA). 7. CR-600 refrigerated centrifuge (International Equipment Company, Needham Heights, MA). 8. Coulter Particle counter (Becton Dickinson, Franklin Lakes, NJ). 9. Low O2 incubator (Forma Scientific Inc., Marjetta, OH).

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1. Make 2% FBS RPMI medium from RPMI 1640 (Gibco, Grand Island, NY) supplemented with 1 mM nonessential amino acids, 1 mM sodium pyruvate, 1% sodium bicarbonate from BioWhittaker, 2 mM glutamine (Sigma, St. Louis, MO), 50 mM b-mercaptoethanol (Fluka, Ronkonkoma, NY), 25 mg/ml gentamicin (Sigma, St. Louis, MO), 1% penicillin– streptomycin–neomycin mixture (Gibco, Grand Island, NY), and 2% heat-inactivated fetal bovine serum (FBS, HyClone, Logan, UT). 2. Mouse CD4+ T cell enrichment kit (StemCell Technologies Inc, Vancouver, BC). 3. 15 ml polypropylene tube (Falcon, Becton Dickinson, Franklin Lakes, NJ).

2.11. Cell Culture

1. Make completed RPMI from RPMI 1640 (Gibco, Grand Island, NY) supplemented with 1 mM nonessential amino acids, 1 mM sodium pyruvate, 1% sodium bicarbonate from BioWhittaker, 2 mM glutamine (Sigma, St. Louis, MO), 50 mM b-mercaptoethanol (Fluka, Ronkonkoma, NY), 25 mg/ml gentamicin (Sigma, St. Louis, MO), 1% penicillin–streptomycin–neomycin mixture (Gibco, Grand Island, NY), and 10% heat-inactivated fetal bovine serum (FBS, HyClone, Logan, UT). 2. OVA peptide (OVAp; ISQAVHAAHAEINEAGR-339) solution (Peptide Synthesis core of Wadsworth Center, Albany, NY). 3. 24-well cell culture plate (Corning Inc., Corning, NY). 4. Coulter Particle counter (Becton Dickinson, Franklin Lakes, NY). 5. Low O2 incubator (Forma Scientific Inc., Marjetta, OH).

2.12. Cytokine Analysis

1. DuoSet ELISA Development System (R & D System, Minneapolis, MN) including Capture Antibody, Detection Antibody, Streptavidin-horseradish-peroxidase (StreptavidinHRP), and standard. 2. 1× PBS: 137 mM NaCl (Mallinckrodt Baker, Inc., Phillipsburg, NJ), 2.7 mM KCl (Fisher Scientific, Pittsburgh, PA), 8.1 mM Na2HPO4 (Baker Chemical Co., Phillipsburg, NJ), 1.5 mM KH2PO4 (Fisher Scientific, Pittsburgh, PA), pH 7.2–7.4, 0.22 mm (Corning Inc., Corning, NY) filtered. 3. Wash buffer: 0.05% Tween 20 (Sigma, St. Louis, MO) in 1× PBS, pH 7.2–7.4. 4. Block buffer: 1% BSA (Sigma, St. Louis, MO), 5% Sucrose (Fisher Scientific, Pittsburgh, PA) in 1× PBS with 0.05% NaN3 (Fisher Scientific, Pittsburgh, PA).

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5. Reagent diluent: 1% (W/V) BSA (Sigma, St. Louis, MO), in 1× PBS, pH 7.2–7.4, 0.22 mm (Corning Inc., Corning, NY) filtered. 6. Substrate solution: 3, 3¢, 5, 5¢-Tetramethyl-benzidine Liquid Substrate (Sigma, St. Louis, MO). 7. Stop solution: Make 2 N H2SO4 by diluting 18 N H2SO4 (Baker Chemical Co., Phillipsburg, NJ) with dd H2O (Barnstead NANOpure Dlamond Life Science (UV/UF) ultrapure water system, Dubuque, Iowa). 8. Variable volume pipette (Gilson, Midleton, WI) + tips (Axygen Scientific, Inc., Union City, CA). 9. ELISA plates (Corning Inc., Corning, NY). 10. ELISA reader (Bio-Tek, Burlington, VT). 2.13. Cell Proliferation

1. Make completed RPMI from RPMI 1640 (Gibco, Grand Island, NY) supplemented with 1 mM nonessential amino acids, 1 mM sodium pyruvate, 1% sodium bicarbonate from BioWhittaker (Lonza, Walkersville, MD), 2 mM glutamine (Sigma, St. Louis, MO), 50 mM b-mercaptoethanol (Fluka, Ronkonkoma, NY), 25 mg/ml gentamicin (Sigma, St. Louis, MO), 1% penicillin–streptomycin–neomycin mixture (Gibco, Grand Island, NY), and 10% heat-inactivated fetal bovine serum (FBS, HyClone, Logan, UT). 2. dd H2O (Barnstead NANOpure Dlamond Life Science (UV/ UF) ultrapure water system, Dubuque, Iowa). 3. OVA peptide (OVAp; ISQAVHAAHAEINEAGR-339) solution (Peptide Synthesis core of Wadsworth Center, Albany, NY). 4. [3H]-thymidine (Dupont-NEN, Wilmington, DE). 5. 96-well cell culture plate (Corning Inc., Corning, NY). 6. FilterMat filter paper (Skatron Instruments Inc., Sterling VA). 7. BAS cassette 2040 (Fuji Film, Stemford, CT). 8. Cell harvesters (Skatron Instruments Inc., Sterling VA). 9. Coulter Particle counter (Becton Dickinson, Franklin Lakes, NY). 10. Low O2 incubator (Forma Scientific Inc., Marjetta, OH). 11. BAS 2000 Fujix reader (Fuji Film, Stemford, CT). 12. TINA 2.0 program (Raytest, Stnaubenhardt, Germany).

2.14. DTH Assay

1. Dissolve OVA (Calbiochem, San Diego, CA) at 10 mg/ml in physiological saline (Baxter, Deerfield, IL) and stored in single use aliquots at −80°C.

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2. Physiological saline (Baxter, Deerfield, IL). 3. Sterile 1.7 ml micro-centrifuge tubes (Axygen Scientific, Inc., Union City, CA). 4. ½ CC Tuberculin Syringe with 27 G × ½ inch needle (Becton Dickinson, Franklin Lakes, NJ). 5. Spi Dial thickness gauge (Long Island Indicator Service Inc., Hauppauge, NY). 2.15. Protein Assay

1. BCA Protein Assay Kit (Pierce, Rockford, IL). Including BCA reagent A, BCA reagent B, and BSA. 2. 96-well cell culture plate (Corning Inc., Corning, NY). 3. 15 ml polypropylene conical tubes (Falcon, Becton Dickinson, Franklin Lakes, NJ). 4. ELISA reader (Bio-Tek, Burlington, VT).

2.16. Phosphoprotein Assay

1. Bio-Plex Phosphoprotein Assay kit (BIO-RAD, Hercules, CA), including coupled Beads, which are coated with antibodies of interest, detection abs, and lysates for control. 2. Bio-Plex Cell Lysis kit (BIO-RAD, Hercules, CA), including cell lysis buffer, cell wash buffer, factor 1, and factor 2. 3. Bio-Plex Phosphoprotein Detection Reagent kit (BIO-RAD, Hercules, CA), including assay buffer, wash buffer, antibody diluent, resuspension buffer, streptavidin-PE, 96-well filter plate, sealing tape, and instructions. 4. 500 mM Phenylmethylsulfonyl fluoride (PMSF): Dissolve 0.436 g PMSF (Sigma, St. Louis, MO) in 5 ml Dimethyl sulfoxide (DMSO) (Sigma, St. Louis, MO). Store the aliquots at −20°C. 5. Lysing solution: Make 10 ml of lysing solution by freshly of mixing 9.9 ml of cell lysis buffer, 40 ml of factor 1 and 20 ml of factor 2, then adding 40 ml of 500 mM PMSF. 6. Variable volume pipette (Gilson, Middleton, WI) + tips (Axygen Scientific, Inc., Union City, CA). 7. 1.7 ml micro-centrifuge tubes (Axygen Scientific, Inc., Union City, CA). 8. Vortex mixer (Barnstead/Thermolyne, Dubuque, Iowa) 9. Titer plate shaker (Lab-Line Instruments, Inc., Melrose Park, IL). 10. Timer (Fisher Scientific Co., Pittsburgh, PA). 11. Millipore multiscreen vacuum manifold (Millipore, Billerica, MA). 12. Eppendorf 5415C centrifuge (Brinkmann Instruments, Inc., Westbury, NY). 13. Luminex 100 or 200 LabMap System (Upstate, NY)

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3. Methods The bone marrow derived DCs are generated from the protocol published by Lutz et al. (28). Different doses of Pb are applied during BM-DC development so that Pb effects can be tested in a dose-dependent manner. Flow cytometry is utilized to test Pb effects on BM-DC phenotype. Then, equal amount of Pb-DCs or DCs are collected and cultured with LPS stimulation to test the cytokine production by ELISA. Pb-DC’s phenotyping and cytokine profile show Pb effects on BM-DC development, also indicating the potential of Pb effects on BM-DC APC function in terms of polarizing T cells. In vivo and in vitro studies of APC function of Pb-DCs are very critical. It confirms the prediction made based on the phenotyping and cytokine profile change. In vitro, testing cytokine production from supernatant (SN) of OVAtg CD4+ T cell cultured with Pb-DCs and antigen (OVAp) provides the information as to which antigen specific T cell (Th1 or Th2) is being activated. In vivo, naïve BALB/c mice are immunized with OVA pulsed Pb-DCs or DCs, and anti-OVA IgG1 and IgG2a production is monitored. Then, delayed type hypersensitivity (DTH) is tested. In vivo studies of HI (IgG1, IgG2a) and CMI (DTH) immunity provide further evidences of T cell polarization promoted by Pb-DCs. Mixed lymphocyte culture (MLC) presents the allogenecity of Pb-DCs or DCs, the other aspect of Pb-DCs. Finally, the purpose of testing several cell signaling pathways is to study the possible mechanisms contributed to Pb-DC induced T cell skewing. 3.1. Generation of BM-DCs

The protocol for generation of BM-DC is adapted from Lutz et al. (28). 1. Remove the femurs and tibias of BALB/c female 2–4-month old mice and place them in a 60 × 15 mm tissue culture dish containing 4 ml 1× DPBS. 2. Clear the surrounding muscles and put the intact bones into 70% ethanol for 2–3 min then wash them with 1× DPBS. 3. Cut both ends of the bones with scissors. 4. Flush the marrow with 1× DPBS by using a ½ CC Tuberculin Syringe with 27 G X ½ inch needle. 5. Further disintegrate the clusters within the marrow suspension by pipetting. 6. In order to remove the debris, transfer the marrow suspension to a 15 ml polypropylene tube and sit on ice for 2 min. 7. Remove the supernatant (SN) to a new 15 ml tube.

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8. Spin the tube at 150×g for 10 min and remove the SN. 9. Resuspend the marrow cell pellet in 10 ml 1× DPBS. 10. Measure the cell concentration using Coulter Particle Counter. 11. Spin again at 150×g then discard the SN. 12. Resuspend the marrow cell pellet in 10% FBS completed RPMI medium (see Note 2). 13. Culture 2 × 106 BM leukocytes in a 100 × 15 mm Petri dish with 10 ml completed RPMI medium containing 200 U/ml mGM-CSF with or without PbCl2 at 37ºC in a low O2 (5%) incubator. 14. After 3 days of incubation, additional 10 ml of complete RPMI containing the same amount of mGM-CSF with or without PbCl2 are added to the Petri dishes. 15. At day 6 and 8, collect and centrifuge half of the culture SN at 150×g for 10 min. 16. Put the marrow cell pellet back into its original dish with 10 ml fresh completed RPMI medium, containing mGMCSF, in the present or absent of Pb. 17. At day-10 (d10), collect nonadherent cells and SN are collected for analysis (see Note 3). 18. For generating mature BM-DC, reculture d10 nonadherent BM-DCs in 10 ml completed RPMI with 100 U/ml mGMCSF + 1 mg/ml lipopolysaccharide (LPS) ± Pb in 100 × 20 mm tissue culture dish for 2 days. 19. Collect nonadherent cells for flow analysis. 20. For cytokine analysis, culture 106 d10 nonadherent BM-DCs in 1 ml completed RPMI with 100 U/ml mGM-CSF + 1 mg/ ml LPS ± Pb in 24-well culture plate. 21. Collect SN after 2 days. 22. For a Pb dose-dependent study, use 1, 5, and 25 mM PbCl2. Otherwise, use just 25 mM PbCl2. 23. For generating OVA-pulsed d10 BM-DCs, at day 9 add OVA (100 mg/ml) to the culture. 24. One day later (d10), collect the nonadherent cells. 25. Wash two times with 1× DPBS at 150×g for 10 min. 3.2. Immunization

1. Inject BALB/c mice subcutaneously (subQ) in the footpad with 106 of overnight OVA-pulsed (100 mg/ml) d10 BM-DCs in 25 ml saline. 2. After one week, bleed and inject OVA (200 mg/100 ml) subQ to the mice.

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3. After another week, bleed these mice. 4. Store the serum at −20°C for IgG1, IgG2a and IgE analyses. 3.3. Blood Collection and Serum Preparation

3.4. Measurement of Antigen Specific IgG Isotype in Serum by ELISA

1. Obtain peripheral blood by retro-orbital phlebotomy into 1.7-ml Eppendorf tubes. 2. After clotting overnight at 4°C, collect the serum by centrifuging at 8,160×g for 5 min and store at −20°C for analysis (see Note 4). 1. Coat ELISA plate with 10 mg/ml OVA in Coating buffer, 50 ml per well for over night, at 4°C. 2. Wash plate three times with Wash buffer. 3. Block with 200 ml 1% BSA per well for 2 h at room temperature. 4. Repeat step 2. 5. Use 1% BSA to dilute the samples or internal control, add 50 ml serial diluted serum or internal control (see Note 5) per well. Incubate for 2 h at room temperature. 6. Wash six times with Wash buffer. 7. Add 0.5 mg/ml Biotin-anti-mouse IgG1 or IgG2a 50 ml per well. 8. Repeat step 6. 9. Add avidin-peroxidase and incubate at room temperature, dark, for 30 min. 10. Repeat step 6. 11. Stop the reaction with Stop solution. 12. Read the plate at 450 nm.

3.5. Total IgE Detection

1. Detect IgE production by using ELISA kit from BD Biosciences. Employ the assay protocol provided by the manufacturer. 2. Coat the plate with anti-mouse IgE mAb in Coating buffer for overnight. 3. Wash three times with Wash buffer. 4. Block the plate for 1 h at room temperature with Assay diluent. 5. Wash three times again. 6. Load serial diluted samples and standards on the plate. Dilute the samples or standards with Assay diluent. 7. Incubate for 2 h at room temperature or overnight at 4°C. 8. Wash six times with Wash buffer. 9. Add biotinylated anti-mouse IgE, incubate at room temperature for 1 h.

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10. Repeat step 8. 11. Add avidin-peroxidase and incubate at room temperature, dark, for 30 min. 12. Wash six times, add substrate. 13. Stop the reaction with Stop solution. 14. Read the plate at 450 nm. 3.6. ELISA for Antigen Specific IgE in Serum

1. Coat the plate with anti-mouse IgE mAb in Coating buffer for overnight. 2. Wash the plate three times with Wash buffer. 3. Block with 200 ml Assay diluent per well for 1 h at room temperature. 4. Repeat step 2. 5. Add serial diluted samples and standards to the plate. Incubate for 2 h at room temperature or overnight at 4°C. 6. Wash six times with Wash buffer. 7. Add 10 mg/ml OVA in 0.1 M NaHCO3, pH 9.5 to the sample wells. Incubate for 2 h at room temperature. 8. Wash six times with Wash buffer for sample wells only. 9. Add 2 mg/ml Biotin-anti-OVA to the sample wells and 0.5 mg/ml biotinylated anti-mouse IgE to the standard wells. Incubate for 1 h at room temperature. 10. Repeat step 6. 11. Add avidin-peroxidase and incubate at room temperature, dark, for 30 min. 12. Repeat step 6. 13. Stop the reaction with Stop solution. 14. Read the plate at 450 nm.

3.7. Flow-Cytometric Analysis

1. Analyze single-cell suspensions by multicolor flow cytometry. 2. 106 cells are suspended in 100 ml staining buffer. 3. Add 1 mg Fc block and 1 mg of fluorescein-conjugated mAbs (see Note 6) to the cells. 4. Incubate for 30 min on ice, dark. 5. Wash cells with 1 ml staining buffer. 6. Spin at 1,200 rpm (150×g) for 10 min. Discard the SN. 7. Resuspend the cells in fixing buffer. 8. Analyze using flow cytometer by gating out the majority of nonviable cells based on low forward angle light scatter (Fig. 19.1).

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CD11c Fig. 19.1. Dot plots of BM-DC development. BM cells were cultured with mGM-CSF ± 25 mM PbCl2 and collected and immunophenotyped on day-10. Cells were labeled with the designed mAbs and analyzed by flow cytometry. The quadrants shown were established based upon isotype control staining. Results are representative of nine independent experiments.

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3.8. Splenocyte Preparation

1. Euthanize mice with an overdose of CO2. 2. After 70% ethanol spray, open the left flank and remove the spleen. 3. Harvest each spleen into a sterile 60 × 15 mm tissue culture dish containing 2 ml of 1× DPBS without calcium and magnesium. 4. Transfer it into another 60 × 15 mm tissue culture dish containing the same amount of 1× DPBS inside a biosafety hood. 5. Homogenize the spleen with two frosted microscope slides. 6. Transfer the cell suspension into a 15 ml polypropylene tube by using a sterile pasteur capillary pipet. 7. After sitting for 2–3 min at room temperature to separate the cell debris, transfer the supernatant (the single-cell suspension) to a new tube. 8. Add 5 ml of red blood cell lysing buffer to the pellet after centrifugation at 150×g for 10 min. 9. After lysing for 5 min at room temperature, centrifuge the cell suspension again and resuspend in 10 ml of 1× DPBS. 10. Take 20 ml of this cell suspension to measure the cell concentration. 11. After centrifugation, resuspend the cell pellet in proper concentration in completed RPMI medium or further fractionated.

3.9. CD4+ T Cell Isolation

1. Isolate CD4+ T cells from spleens with negative-selection columns from StemCell Technologies Inc. Utilize the manufacturer protocol to perform the assay (see Note 7). 2. Incubate splenocytes with SpinSep enrichment cocktails in a 15 ml tube for 15 min at 4°C. 3. Wash with 10 ml 2% FBS RPMI medium. 4. Resuspend the antibody-treated cells in 2% FBS RPMI medium. 5. Incubate them with SpinSep Dense Particles on ice for 20 min. 6. Dilute the cell/particle suspension with 2% FBS RPMI medium. 7. Load the diluted cell/particle suspension on top of the SpinSep Density Medium in a new 15 ml tube. 8. Centrifuge for 10 min at 1,200×g at room temperature with the break off. 9. Separate CD4+ T cells from other cells by the density difference. The purity of CD4+ cells is 90 ± 3%.

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1. In order to evaluate d10 BM-DC APC function in vitro, cell culture technique is utilized. Culture 106 OVAtg CD4+ T cells with 105 of d10 Pb-DCs or DCs in present of 0.02 mg/ ml OVAp (see Note 8). 2. In order to exam the allogeneic stimulatory ability of d10 BM-DC, culture 106 single-cell suspension of C57BL/6 spleen cells with 105 Pb-DCs or DCs. 3. For testing the syngeneic stimulatory ability of d10 BM-DC, culture 106 BALB/c CD4+ T cells with 105 Pb-DCs or DCs. 4. Set up all culture with 1 ml completed RPMI in 24-well cell culture plate. 5. Incubate at 37ºC in a low O2 (5%) incubator for 4 days. 6. Collect all culture SN and store at −20ºC for cytokine analysis.

3.11. Cytokine Analysis

1. Detect cytokine production by using ELISA kits from R & D System. Use the manufacturer’s protocol to perform the assay. 2. Coat the plate with anti-mouse cytokine mAb in 1× PBS for overnight. 3. Wash three times with Wash buffer. 4. Block the plate for 2 h at room temperature with Block buffer. 5. Wash three times again. 6. Load serial diluted standards and samples on the plate. 7. Dilute the samples or standards with Reagent diluent. 8. Incubate for 2 h at room temperature or overnight at 4°C. 9. Wash six times with Wash buffer. 10. Add biotin-labeled anticytokine mAb. 11. Incubate at room temperature for 2 h. 12. Repeat step 8. 13. Add avidin-peroxidase and incubate at room temperature, dark, for 20 min. 14. Wash six times, add substrate. 15. Stop the reaction with Stop solution. 16. Read the plate at 450 nm (Fig. 19.2).

3.12. Cell Proliferation

1. The culture conditions are similar to those listed in the Cell Culture section, except that 2 × 105 of CD4+ T cells or SPLs and 2 × 104 of Pb-DCs or DCs (see Note 9) are set up with 0.2 ml completed RPMI in 96-well cell culture plate.

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Fig. 19.2. Pb effects LPS-induced cytokine production by d10 BM-DC cultures. BM cells were cultured with mGM-CSF + 0–25 mM PbCl2 for 10 days; then 106 cells were washed and recultured with mGM-CSF + LPS for 2 days (a). Additionally, BM cells were cultured with mGM-CSF ± 25 mM PbCl2 for 10 days; then 106 cells were washed and recultured with mGM-CSF + LPS ± PbCl2 for 2 days (b). SNs were collected for analysis of cytokine production. Results are presented as the mean ± SEM (N = 5, from 5 independent experiments); *, **, and *** indicate significant difference from control (0 mM Pb) cultures.

2. After 4 days, pulse each well with 0.5 mCi/well [3H]-thymidine for the last 6 h. 3. Harvest the cells on FilterMat filter paper by using a Cell harvester. 4. Then, expose the filter paper in a Fuji BAS cassette 2040 for 3 days. 5. Read the phosphostimulated luminesence image using a BAS 2000 Fujix reader and analyze via the TINA 2.0 program. 3.13. DTH Assay

1. After priming with OVA-pulsed d10 BM-DCs and immunization with OVA, mice are ready for DTH assay. 2. Inject 100 mg OVA in 25 ml saline subQ into one of the hind footpads of the OVA sensitized mice and inject 25 ml saline into another hind footpad of the same mouse.

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3. After 24 h, measure the footpad swelling response by using Spi Dial thickness gauge. DTH response is defined as the difference between the right and left footpad swelling (see Note 10). 4. Immunize BALB/c mice subQ twice with OVA served as positive controls. 3.14. Protein Assay

1. Test the protein concentration of cell lysates by using BCA Protein Assay Kit. Utilize the manufacturer protocol to perform the assay. 2. Mix BCA reagent A (containing mainly bicinchoninic acid) and BCA reagent B (containing cupric sulfate) as 50:1 ratio in a 15 ml tube. 3. Load 25 ml of samples and serial diluted BSA standards into the 96-well cell culture plate. The diluent used for the standard dilution should be the same as used for preparing samples. 4. Add 200 ml of BCA reagent A and B mixture to each well. 5. Incubate at 37°C for 30 min. 6. Cool the plate to room temperature. 7. Read the plate at 570 nm. Calculate the sample protein concentration based on the BSA standard.

3.15. Phosphoprotein Assay

Detect Phosphorylated proteins for cell signaling from d10 BM-DC lysate by using Bio-Plex Phosphoprotein Assay kit. Use the assay protocol provided by the manufacturer.

3.15.1. Cell Lysate Preparation

1. Wash d10 BM-DCs two times with cell wash buffer. 2. Resuspend the cells in lysing solution with the ratio at 107 cells in 2 ml lysing solution. 3. Agitate the cells. 4. Transfer the cell lysate to a tube and rotate for 20 min at 4°C. 5. Centrifuge the samples at 4,500×g for 20 min at 4°C. 6. Collect the SN. SN can be stored at −20°C, and undergo freezing/thaw cycles for up to five times. 7. Test fresh made or frozen cell lysate for protein concentration 8. Add an equal volume of assay buffer to the cell lysate before doing phosphoprotein detection.

3.15.2. Phosphoprotein Detection

1. Prewet filter plate with 50 ml assay buffer. 2. Remove the buffer by vacuum filtration over a vacuum manifold.

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3. Blot the bottom of the plate to a clean paper towel to dry excess liquid. 4. Dilute coupled beads to 1× with wash buffer, load 50 ml per well to the filter plate. 5. Wash the plate with 100 ml assay buffer per well for two times. 6. Repeat step 2. 7. Add 50 ml of cell lysate to each well. 8. Seal the plate and incubate overnight at 4°C in the dark on a plate shaker with 110 rpm. 9. Wash the plate with 100 ml assay buffer per well for three times. 10. Repeat step 2. 11. Dilute detection antibody to 1× with detection antibody diluent. 12. Load 25 ml per well to the plate. 13. Seal the plate and incubate for 30 min at room temperature in the dark on a plate shaker with 110 rpm. 14. Repeat step 7 and step 2. 15. Dilute streptavidin-PE to 1× with wash buffer. 16. Load 50 ml per well to the plate. 17. Seal the plate and incubate for 10 min at room temperature in the dark on a plate shaker with 110 rpm. 18. Repeat step 7 and step 2. 19. Add 125 ml resuspension buffer per well to the plate. 20. Seal the plate and incubate for 30 s at a room temperature in the dark on a plate shaker. 21. Use Luminex 100 or 200 system to collect and analysis data. 3.15.3. Statistical Analysis

Statistical analysis is performed by SigmaStat (Jandeel Scientific, San Rafael, CA) one-way ANOVA; p £ 0.05 was considered significant.

4. Notes 1. All solutions should be tested for endotoxin contamination. 2. All procedures should be done inside a sterile biosafety hood. 3. The red blood cells in BM should not be removed. BM leukocyte numbers are used to determine the amount of cells for initiating each culture, however, the red blood cells are also in the cell preparation. The average yield of BM-DCs is 11.7 ± 4.5 × 106 per dish in controls, and 8.4 ± 2.3 × 106 per

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dish in 25 mM PbCl2 treated cultures. If a small amount of BM-DCs is needed, then 24-well cell culture plates (7, 12), 12-well cell culture plates (29), or 6-well cell culture plates (9) can be used. 4. Step 2 may need to be repeated to get the pure serum. 5. For the internal control, use anti-OVA mAb starting with a concentration of 1,000 pg/ml. 6. The following fluorescein-conjugated mAbs are suggested for phenotyping of BM-DCs: anti-mouse CD11c, anti-mouse CD11b, anti-mouse Gr1, anti-mouse FceR1, anti-mouse CD3, anti-mouse CD8, anti-mouse CD19, anti-mouse CD45R/ B220, anti-mouse I-Ad, anti-mouse CD80, anti-mouse CD86, anti-mouse CD40, anti-mouse ICAM-1, anti-mouse F4/80, anti-mouse CD83, anti-CD31, anti-CD34, anti-mouse CD123. Based on the flow cytometry, 3-color (PerCP, phycoerythrin (PE), and Fluorescein isothiocyanate (FITC)) or 4-color (APC, PerCP, PE and FITC) antibody combination can be used. 7. The amount of spinSep enrichment cocktail, Density particles and Density medium is based on the purifying cell number. It is described in the manufacturer’s protocol provided with the kit. 8. The 10 to 1 ration of OVAtg CD4+ T cells to d10 BM-DCs and the concentration of OVAp are selected because these conditions have been shown to evoke a mixed Th1 and Th2 response (29, 30). 9. In this study, BM-DCs are not gamma-irradiated, because gamma-irradiated DCs may reduce T cell proliferation in mixed lymphocyte reaction (31). However, proper controls are needed. In our study, BM-DCs by themselves have a very low proliferation ability. 10. Since measuring DTH response means measuring a swelling response, the Dial thickness gauge should not stay on footpad too long. Gently put it against the footpad surface and read the result when it becomes stable. Measure two to three times for each footpad. Do not include big toe in gauge coverage surface. It is recommended that mice can be anesthetized before doing the footpad measurement.

Acknowledgments The authors thank the staff of the Immunology Core of Wadsworth Center for their assistance with the flow cytometry and phosphorimaging analyses. This work was supported, in part, by NIH grant ES11135.

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References 1. Adams S, O’Neill DW, Bhardwaj N (2005) Recent advances in dendritic cell biology. J Clin Immunol 25(3):177–188 2. Steinman RM (2003) Some interfaces of dendritic cell biology. APMIS 111:675–697 3. MacDonald AS, Straw AD, Dalton NM, Pearce EJ (2002) Cutting edge: Th2 response induction by dendritic cells: a role for CD40. J Immunol 168(2):537–540 4. Jenkins SJ, Mountford AP (2005) Dendritic cells activated with products released by schistosome larvae drive Th2-type immune responses, which can be inhibited by manipulation of CD40 costimulation. Infect Immun 73(1):395–402 5. Szabolcs P, Moore MA, Young JW (1995) Expansion of immunostimulatory dendritic cells among the myeloid progeny of human CD34+ bone marrow precursors cultured with c-kit ligand, granulocyte–macrophage colony-stimulating factor, and TNF-alpha. J Immunol 154:5851–5861 6. Caux C, Dezutter-Dambuyant C, Schmitt D, Banchereau J (1992) GM-CSF and TNF-alpha cooperate in the generation of dendritic Langerhans cells. Nature 360(6401):258–261 7. Mehling A, Grabbe S, Voskort M, Schwarz T, Luger TA, Beissert S (2000) Mycophenolate mofetil impairs the maturation and function of murine dendritic cells. J Immunol 165(5):2374–2381 8. Hackstein H, Morelli AE, Larregina AT, Ganster RW, Papworth GD, Logar AJ, Watkins SC, Falo LD, Thomson AW (2001) Aspirin inhibits in vitro maturation and in vivo immunostimulatory function of murine myeloid dendritic cells. J Immunol 166(12):7053–7062 9. Kim GY, Kim KH, Lee SH, Yoon MS, Lee HJ, Moon DO, Lee CM, Ahn SC, Park YC, Park YM (2005) Curcumin inhibits immunostimulatory function of dendritic cells: MAPKs and translocation of NF-kappa B as potential targets. J Immunol 174(12):8116–8124 10. Kim GY, Cho H, Ahn SC, Oh YH, Lee CM, Park YM (2004) Resveratrol inhibits phenotypic and functional maturation of murine bone marrow-derived dendritic cells. Int Immunopharmacol 4(2):245–253 11. Bronnum H, Seested T, Hellgren LI, Brix S, Frokiaer H (2005) Milk derived GM(3) and GD(3) differentially inhibit dendritic cell maturation and effector functionalities. Scand J Immunol 61(6):551–557 12. Cavassani KA, Aliberti JC, Dias AR, Silva JS, Ferreira BR (2005) Tick saliva inhibits

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Dendritic Cells in Immunotoxicity Testing 23. Rivollier A, Perrin-Cocon L, Luche S, Diemer H, Strub JM, Hanau D, van Dorsselaer A, Lotteau V, Rabourdin-Combe C, Rabilloud T, Servet-Delprat C (2006) High expression of antioxidant proteins in dendritic cells: possible implications in atherosclerosis. Mol Cell Proteomics 5:726–736 24. Kuppner MC, Scharner A, Milani V, Von Hesler C, Tschop KE, Heinz O, Issels RD (2003) Ifosfamide impairs the allostimulatory capacity of human dendritic cells by intracellular glutathione depletion. Blood 102:3668–3674 25. Heo Y, Parsons PJ, Lawrence DA (1996) Lead differentially modifies cytokine production in vitro and in vivo. Toxicol Appl Pharmacol 138(1):149–157 26. Heo Y, Lee WT, Lawrence DA (1997) In vivo the environmental pollutants lead and mercury induce oligoclonal T cell responses skewed toward type-2 reactivities. Cell Immunol 179(2):185–195 27. Heo Y, Lee WT, Lawrence DA (1998) Differential effects of lead and cAMP on development and activities of Th1- and Th2lymphocytes. Toxicol Sci 43(2):172–185

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Chapter 20 Evaluating Cytokines in Immunotoxicity Testing Emanuela Corsini and Robert V. House Abstract One of the most potentially useful tools in immunotoxicology is the assessment of cytokines, the proteins/ peptides that are responsible for regulating a variety of processes including immunity, inflammation, apoptosis, and hematopoiesis. Cytokine production measurements offer an outstanding promise and may eventually substitute for other more laborious procedures. The particular profile of cytokine production may provide an important information regarding the nature of many immunotoxic responses. Recent expansion in the knowledge of cytokine biology and the realization that cytokines play a role in human diseases have created a need for the precise assessment and accurate interpretation of their presence and activity in the body fluids, tissues and cells. Proper evaluation of cytokines requires attention to several technical details. Multi-cytokine analysis still needs to be standardized in terms of optimum source for analysis, protocols and quality control issues, such as the use of reference standards and the expression of results. Important practical details and considerations will be discussed in this chapter, including the source of the sample to be tested (circulating fluids, or ex vivo/in vitro isolated cells), the potential effects of collection, processing, and storage of the results of the assays, as well as potential variables associated with the source material (matrix effects, relevance, inhibitory substances), and factors influencing the choice of assay used (bioassay, immunoassay, molecular biology technique, flow cytometry). Key words: Bioassay, Cytokine, ELISpot, ELISA, Immunoassay, In vitro immunotoxicology, PCR

1. Introduction Cytokines play a key role in many manifestations of chemicallyinduced immunotoxicity, including both immunosuppression and immunoenhancement (1). Cytokines are small molecular weight proteins or peptides secreted by many cell types (particularly immune systems cells) that regulate the duration and intensity of the immune response. Type 1 cytokines (e.g., interferon-gamma (IFN-g), interleukin-12 (IL-12)) mediate the removal of malignant cells and virally infected cells, whereas Type 2 cytokines (e.g., IL-4, IL-5, IL-13) mediate the removal R.R. Dietert (ed.), Immunotoxicity Testing: Methods and Protocols, Methods in Molecular Biology, vol. 598 DOI 10.1007/978-1-60761-401-2_20, © Humana Press, a part of Springer Science + Business Media, LLC 2010

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of soluble bacterial antigens. Obviously, cytokines play key roles in all immune responses, and molecular immunotoxicology has indeed focused on the analysis of cytokine levels. Detection of specific cytokines reveals the state of immune response at any given time, such as when elevated tumor necrosis factor – alpha (TNF-a) levels reveal a state of inflammation. Cytokines offer important new avenues to explore both in terms of mechanistic understanding of immunotoxicity and of developing new assays to test the immunotoxic potential of the novel compounds. Effects on cytokines can be analyzed on two different levels: messenger ribonucleic acid (mRNA) and protein. The choice essentially depends on the aim of the study and on the equipment available. Proteins mediate biological activities directly, so they are a more direct measure of function than in mRNA. The enzyme-linked immunosorbent assay (ELISA) is by far the most common method used to assay the cytokines in biological samples; however, other cytokine assays are also available including multiplexed ELISAs and immunohistochemistry. Detection and quantitation of mRNA measures the presence of cytokines at a specific point in time within a tissue or cell, whereas protein is measured in a body fluid, possibly as a spill-over from the tissue, or in a supernatant following cell culture (2, 3). In the past, mRNA levels were assayed using Northern or dot blotting; nowadays, they are measured using reverse transcription-polymerase chain reaction (RT-PCR) (4, 5). More recently, analysis of cytokine production has been performed with intracellular staining followed by flow cytometry, a more informative and reliable approach since it confirms cytokine production at the single cell level, thus achieving higher specificity (6–8). 1.1. Sample Collection and Storage

Efforts to develop noninvasive collection methods include analysis of cytokines in saliva (9) or induced sputum (10) and exhaled breath condensate (11). However, these methods still require further optimization and validation before they are ready to be used widely. Most importantly, the sensitivity of the assay requires improvement, perhaps at the collection stage, to stabilize cytokines in sputum or saliva, since several cytokines are detected at levels that are too low to differentiate between control and disease states. Evaluation of cytokine levels in body fluids or ex vivo/in vitro cytokine production represents the testing future in immunotoxicology as their assessment becomes more and more convenient. Still, their analysis needs to be standardized in terms of optimum source for analysis and protocol.

1.1.1. Primary Sources

Cytokines can be measured in several body fluids, such as plasma, saliva, etc. or in vitro, in culture supernatants or in tissues. A common approach is to measure the relative concentration of

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different cytokines in peripheral blood following experimental treatment. This approach certainly has the advantage of simplicity, since it requires only limited experimental manipulation. Although it has been used successfully in many instances, there are problems with this method due to the biology of these molecules that needs to be considered: (a) The half-life of many cytokines in the circulation is very short (minutes), so that the timing of collection may be a significant factor. (b) Cytokines primarily act at a local level, resulting in a very low spill in plasma. Thus, detection of cytokines at a distant site may provide little information on the cytokine’s function in situ. (c) Cytokines are extremely potent mediators that are active at very low concentrations, and many cytokine assays are sensitive only for relatively high concentrations. (d) Baseline values for cytokines have not yet been reliably established in any species, humans included, making it difficult to interpret the biological significance of minor variations in cytokine levels. (e) When using blood as a source of cytokines an important consideration is whether whole blood, serum or plasma should be assayed. This is primarily a practical issue, but in general plasma (especially citrated plasma) has been found to clot in cytokine assays, resulting in high variability. Whenever possible, serum should be used as a more reproducible material for cytokine bioassays. Furthermore, complex sample matrices such as blood, serum or plasma may contain interfering factors such as rheumatoidfactors, heterophilic antibodies, binding proteins and complement components. This may affect the ability of the assay to quantify the analyte accurately (12). Recovery experiments are necessary to reveal if assays are affected in this way. 1.1.2. Secondary Sources

In most routine immunotoxicology studies using rodents, the spleen is the most common source of immune cells (T and B lymphocytes, macrophages, etc.). The spleen has the advantage of size, meaning a greater number of cells are available than using the alternative tissues. For assessment of mechanisms of hypersensitivity, the lymph nodes have proven to be a more relevant source of cells for evaluating cytokine production, particularly in the case of variations of the murine local lymph node assay (13). Following culture, the culture supernatant is collected and analyzed for the presence and relative concentration of the cytokine(s) of interest. It is important to remember that most cytokines are expressed only following the cellular activation.

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In the absence of an active in vivo immune response (such as following administration of an experimental antigen or infectious challenge), the basal levels of cytokines from cells cultured ex vivo are very low. Therefore, the cells need to be activated in vitro using a reagent appropriate for the cell population of interest (i.e., the mitogenic lectins concanavalin A [ConA] or phytohemagglutinin [PHA]), or a more physiologically-relevant stimulus such as an antibody specific for the CD3 receptor on T-cells (with or without antibodies directed against costimulatory receptors such as CD28) or lipolysaccharide (LPS) for monocytes/macrophages. In this type of experiment, it is important to titer the stimulus used to the lowest concentration that results in cellular activation. Otherwise, it is possible that the modest alterations in cytokine production may be masked. There are several advantages in using this approach: it minimizes many of the problems associated with in vivo cytokine assessment, the target population is better defined, positive controls can be included and human or nonhuman primate cells can be used. The primary disadvantage of this approach is that in vitro systems are not necessarily an accurate representation of an in vivo immune response. One must exercise care when attempting in vitro exposure systems since the results may be difficult to interpret (14). Regardless, it is often necessary to make extrapolations based on artificial systems and so this disadvantage is no greater than any other experimental situation. 1.1.3. Clean-up and Storage of Samples

Once collected, any “contaminating” cells or other debris should be rapidly removed by centrifugation (120–260 × g for 5 min) since they will continue to produce cytokines. Samples should be stored frozen, below −20°C. Cycles of thawing-freezing should be avoided to minimize degradation of the protein molecules. Sterile tubes are recommended, particularly if cytokines are going to be measured by bioassays.

1.1.4. Cryopreservation of Cells

It is often impractical to measure the cytokine production from cells that are stimulated ex vivo to produce cytokines, for example when analysis is to be peformed in a laboratory distant from the collection site (such as a clinical trial). In such cases, primary cells such as lymphocytes or monocytes may be carefully cryopreserved and either stored or shipped between laboratories. Published studies have shown that cytokine production is generally affected only marginally by this process, although some other measures of cell-mediated immunity may be slightly affected (15–17).

1.1.5. Immunoenzymatic Assay vs. Bioassay: Advantages and Disadvantages

Once the cytokine-containing material is obtained, the appropriate methodology must be utilized to accurately measure its concentration. The type of assay chosen will depend on the capabilities of the laboratory, as well as the type of information required. Methodology for assessing the production and action of cytokines

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Table 20.1 Advantages and disadvantages of cytokine assay techniques Assay technique

Advantages

Disadvantages

Bioassay

Detect only functional molecules. Exquisitely sensitive (less of pg/ml).

Often lack specificity. Requires cell culture. Time-, labor- and resource-intensive. Difficult to perform reproducibility

Immunoassays

Rapid and easy to perform. High specificity. No cell culture required. With multiplex ELISA several cytokines can be monitored simultaneously.

May not detect functional molecules. Less sensitive than bioassay. Reagents not available for all species.

Flow cytometry

Sensitive and extremely specific (single cell). Several cytokines can be monitored simultaneously. Other relevant molecules (i.e. surface markers) can be examined simultaneously.

Requires specialized equipment. Accurate and complete interpretation of results requires specialized skill.

mRNA expression

Most specific. Can detect changes at single cell level. Early detection of cytokines (transcription) High sensitivity

Generally expensive and time consuming. Requires specialized equipment and techniques. Message not necessarily translate into protein.

encompasses several technologies. The major types of cytokine assays currently in use include bioassays, immunoassays, molecular biology techniques, and flow cytometry. Each of these assay types exhibits advantages and disadvantages, and no one type of assay is best suited for all applications. A combination of techniques, and even combining the various techniques are usually used. Some of these techniques are described below. The advantages and disadvantages of the different cytokine assays are listed in Table 20.1. Bioassays are particularly important if a clinical use of the cytokine is expected, while in general immunoassays are better suited for immunotoxicology testing.

2. Materials 2.1 Immunoassays 2.1.1 Sandwich ELISA

1. Coating antibody and standard (R&D Systems, Minneapolis, MN). 2. 96 wells polystyrine plate (EIA/RIA plates, cat. n° 3369, Corning-Costar, Lowell, MA).

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3. Phosphate buffered saline (PBS): NaCl 0.14 M, KCl 2.7 mM, Na2HPO4.7H2O 8.1 mM, 1.9 mM. 4. Bovine serum albumin (BSA) (Sigma, St Louis, MO). 5. PBS/Tween 20: PBS containg 0.05% Tween 20 (Sigma, St Louis, MO). 6. HRP-conjugate antibody (R&D Systems, Minneapolis, MN). 7. Substrate solution, i.e. 3,3¢,5,5¢-tetramethyl-benzidine (TMB, Sigma Cat. T4444, the substrate is supplied as a one component ready to use solution). 8. Microplate reader (Emax from Molecular Devices, Sunnyvale, CA). 2.1.2 Competitive ELISA

1. Diluted primary antibody and standard (R&D Systems, Minneapolis, MN). 2. 96 wells polystyrine plate (EIA/RIA plates, cat. n° 3369, Corning-Costar, Lowell, MA). 3. Phosphate buffered saline (PBS): NaCl 0.14 M, KCl 2.7 mM, Na2HPO4.7H2O 8.1 mM, 1.9 mM. 4. Bovine serum albumin (BSA) (Sigma, St Louis, MO). 5. PBS/Tween 20: PBS containg 0.05% Tween 20 (Sigma, St Louis, MO). 6. Antigen-conjugate solution (R&D Systems, Minneapolis, MN). 7. ELISA reader (Emax from Molecular Devices, Sunnyvale, CA).

2.1.3 ELISpot

1. Anticytokine antibody (R&D Systems, Minneapolis, MN). 2. PBS: NaCl 0.14 M, KCl 2.7 mM, Na2HPO4.7H2O 8.1 mM, 1.9 mM. 3. Positive control (phytohaemagglutinin A, PHA, or phorbol ester such as Phorbol 12-Myristate 13-Acetate, PMA). 4. RPMI-1640 (Sigma, St Louis, MO). 5. Fetal bovine serum (Sigma, St Louis, MO). 6. Nonessential amino acids (Sigma, St Louis, MO). 7. Penicillin (Sigma, St Louis, MO). 8. Streptomycin (Sigma, St Louis, MO). 9. L-glutamine (Sigma, St Louis, MO). 10. PBS cointaing 0.01% Tween 20. 11. Biotinylated anticytokine Minneapolis, MN).

antibody

(R&D

Systems,

12. Streptavidin–alkaline phosphatase enzyme conjugate or streptavidin-HRP (R&D Systems, Minneapolis, MN).

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13. BCIP/NBT or AEC substrate (Sigma, St Louis, MO). 14. Imaging system (Cellular Technology Ltd, Shaker Heights, OH). 15. Polyvinylidene difluoride (PVDF)-backed microplate (Cellular Technology Ltd, Shaker Heights, OH). 2.2 FACS

1. Protein transport inhibitor (BD Biosciences, San Josè, CA). 2. Surface marker antibodies (BD Biosciences, San Jose, CA). 3. Paraformaldehyde or formaldehyde (Sigma, St Louis, MO). 4. PBS: NaCl 0.14 M, KCl 2.7 mM, Na2HPO4.7H2O 8.1 mM, 1.9 mM. 5. Fetal calf serum (FCS, Sigma, St Louis, MO). 6. Sodium azide (Sigma, St Louis, MO). 7. Cytokine antibodies (BD Biosciences, San Josè, CA). 8. Saponin (Sigma, St Louis, MO) 9. Fluorochrome-conjugated anticytokine Biosciences, San Josè, CA).

antibody

(BD

2.3 Bioassays

No materials to be included for this section.

2.4 Gene Expression

1. Reverse Transcription Kit (Cat. n°4368814 High capacity cDNA reverse transcription kit, Applied Biosystems). 2. Real-time PCR Mix (Cat. n°4324018 TaqMan Universal PCR Master Mix, Applied Biosystems). 3. PCR primers (Assays-on-Demand Gene Expression products). 4. TaqMan MGB probe (FAM dye-labeled). 5. ABI Prism SDS 7000 or equivalent instrument (Applied Biosystems, Foster City, CA). 6. Agarose (Sigma, St Louis, MO)

3. Methods 3.1. Immunoassays

Immunoassays are probably the most popular means of measuring cytokines. Several different formats exist, but the two most common used immunoassays are the enzyme linked immunosorbent assay (ELISA) and the radioimmunoassay (RIA); of these the ELISA is most often used. The ELISA format was developed for the measurement of small amounts of substances, typically picograms, in test samples. It is extremely useful in routine analytical determination. The ELISA format has two available techniques for antigen measurement: the sandwich technique and the competitive technique.

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The sandwich or double antibody technique begins with an antibody bound to a polystyrene well plus the antigen to be measured. For most applications, a polyvinylchloride (PVC) microtiter plate is best; however, manufacturers’ guidelines must be consulted to determine the most appropriate type of plate for protein binding. PVC will bind approximately 100 ng/well (300 ng/cm2). Following coating and sample/standard incubation, a second antibody specific for a different epitope is then added to the well; this second antibody is generally conjugated with an enzyme that converts a substrate to a colorimetric end product. Next, a substrate is added to the enzyme conjugate which is bound to the immune complex. If there are changes due to the presence of the enzyme conjugate bound to the immune complex, a positive test or color change will occur. The detection antibody can actually be detected by three methods: 1. The detection antibody is already labeled. 2. A labeled antibody recognizing the host of the detection antibody can be used if and only if the capture is from a different host than the detection. 3. A biotin-conjugated detection antibody which in a later incubation will bind streptavidin bound to some type of label. The label can be a fluorophor or an enzyme such as horseradish peroxidase (HRP) or alkaline phosphatase (AP). The type of label determines the sensitivity of the assay as well as the method of reading. Flurophor labeled assays are read with fluorometer, whereas enzymes can be read with spectrophotometers. Multiplexing in assays simply refers to the ability to output multiple readings from a single sample. For example, one well of a 96 well plate would react with the sample and would provide data for multiple assays. For example, using different commercially available cytokine array, with one sample and one well, one can measure several cytokines such as IL-1a, IL-1b, IL-2, IL-3, IL-4, IL-5, IL-6, IL-9, IL-10, IL-12, MCP-1, INF-g, TNF-a, MIP-1, and so forth. (see Note 1) 3.1.1. Typical ELISA (Sandwich Protocol)

1. Prepare coating antibody as directed. If maximal binding is required, use at least 1 mg/well. 2. Allow to incubate for 4 h at room temperature or 4°C overnight to allow complete binding. 3. Shake off coating solution and wash wells 3–4 times with phosphate-buffered saline (PBS). Remove the solutions or washes by flicking the plate over a sink. Remove the remaining drops by patting the plate on a paper towel. 4. Block unbound sites with 1–2% bovine serum albumin (BSA) in PBS (blocking solution) by adding 200 mL/well.

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5. Incubate plate for 1 hr at room temperature or, if more convenient, overnight at 4°C. 6. Shake off the blocking solution (without washing or drying). 7. Add 100 mL standard or sample to each well. 8. Incubate 1–2 h at room temperature. 9. Shake off sample and wash wells 3–4 times with PBS/Tween 20 (0.05%). 10. Add 100 mL HRP-conjugate antibody directed against the antigen to each well. 11. Incubate 1–2 h at room temperature. Primary and secondary antibody should be diluted in blocking solution to reduce nonspecific binding. 12. Shake off sample and wash wells 3–4 times with PBS/Tween 20 (0.05%). 13. Add 100 mL substrate solution to each well. Protect from light. 14. Incubate 10–30 min at room temperature. 15. Add 50 mL stop solution to each well. 16. Read optical density (OD) at 450 nm within 30 min in microplate reader. 17. Prepare a standard curve from the data produced from the serial dilutions with concentration on the X axis (log scale) vs. absorbance on the Y axis (linear). Interpolate the concentration of the sample from this standard curve. 3.1.2. Competitive ELISA

The antigen competitive ELISA begins with an antibody bound to a polystyrene well plus a test sample containing an antigen mixture to which an antigen-enzyme conjugate is added. At this point competitive inhibition occurs between the antigen-enzyme conjugate and an unlabeled antigen. Depending upon which antigen type is in excess, two different outcomes can follow when binding to a specific antibody occurs. After the formation of an immune complex from an antigen–antibody binding, the reagents are separated by a washing. Next a substrate is added to the immune complex. If the antigen-enzyme conjugate is the antigen in excess, a color change will occur indicating that the substrate has been chemically changed as a result of the enzyme conjugate being bound to the immune complex. If it is the unlabeled antigen that is in excess, there will be little to no change in color because the test sample contains antibody-type-specific antigen. 1. Add 100 mL of diluted primary antibody (capture) to each well. If maximal binding is required, use at least 1 mg/well. 2. Incubate for 4 h at room temperature or 4°C overnight to allow complete binding.

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3. Shake off coating solution and wash wells 3–4 times with PBS. Remove the solutions or washes by flicking the plate over a sink. Remove the remaining drops by patting the plate on a paper towel. 4. Block unbound sites with 1–2% BSA in PBS (blocking buffer) by adding 200 mL/well. 5. Incubate the plate for 1 h at room temperature or, if more convenient, overnight at 4°C. 6. Shake off blocking solution (without washing or drying). 7. Shake off sample and wash wells 3–4 times with PBS/Tween 20 (0.05%). 8. Add 50 mL of the standards or sample solution to the wells. 9. Add 50 mL of the antigen-conjugate solution to the wells (the antigen solution should be titrated). All dilutions should be done in the blocking buffer. 10. Incubate for at least 2 h at room temperature in a humidified atmosphere. 11. Shake off sample and wash wells 3–4 times with PBS/Tween 20 (0.05%). 12. Add substrate as indicated by the manufacturer. 13. After suggested incubation time has elapsed, measure optical densities at target wavelengths on an ELISA reader.(see Notes 2 and 3) 3.1.3. ELISpot

The ELISpot (Enzyme Linked Immuno-Spot) assay provides an effective method of measuring cytokine production of immune cells on the single cell level. The ELISpot has originally been developed for the detection of individual B-cells secreting antigen-specific antibodies. This method has since been adapted for the detection of individual cells secreting specific cytokines or other antigens. ELISpot assays employ the quantitative sandwich ELISA technique. A monoclonal antibody specific for a cytokine is pre-coated onto a polyvinylidene difluoride (PVDF)-backed microplate. Appropriately stimulated cells are pipetted into the wells and the microplate is placed into a humidified 37°C CO2 incubator for a specified period of time. During this incubation period, the immobilized antibody in the immediate vicinity of the secreting cells binds secreted cytokine. After washing away cells and any unbound substances, a biotinylated polyclonal antibody specific for the cytokine is added to the wells. Following a wash to remove any unbound biotinylated antibody, alkaline-phosphatase conjugated to streptavidin is added. Unbound enzyme is subsequently removed by washing and a substrate solution (BCIP/ NBT) is added. Blue-black colored precipitate forms at the sites of cytokine localization and appears as spots with each individual spot representing an individual cytokine-secreting cell. The spots

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can be counted with automated ELISpot reader systems or manually, using a stereomicroscope. The following protocol is an example of a typical ELISpot assay for quantifying the number cells producing interferon-g (IFN-g) in response to antigen or nonspecific activation using PHA. It may be optimized as necessary for the other applications. 1. On day one, coat plates with 100 mL (10 mg/mL) anticytokine antibody in sterile PBS. 2. Incubate overnight at 4°C. 3. Incorporate the following control wells into the assay: No cells; No primary antibody; No antigen stimulation; Positive control with PHA or phorbol ester such as PMA. 4. On day two, decant primary antibody solution. 5. Wash off twice unbound antibody with 150 mL sterile water per well. 6. Block membrane with 150 mL per well of culture medium (RPMI-1640, 10% fetal bovine serum, 1% nonessential amino acids, penicillin, streptomycin, glutamine) for at least 2 h at 37°C. 7. Purify human peripheral blood mononuclear cells (PBMC) using a Ficoll™ density gradient separation. 8. Wash cells in cold PBS. 9. Count and resuspend at a final concentration of 0.25–2 × 106 cells/mL in culture medium. If the expected response is not known, a serial dilution of cell concentrations is recommended. 10. Decant blocking medium. 11. Gently plate PBMC in 100 mL cell medium per well. 12. Incubate for 18–48 h at 37°C, 5% CO2, and 95% humidity. 13. On day three, decant cells. 14. Wash the plate six times with PBS/0.01% Tween 20. 15. Dilute biotinylated anticytokine antibody to 2 mg/mL in PBS/0.5% BSA. 16. Add 100 mL/well. 17. Incubate for 2 h at 37°C, 5% CO2, and 95% humidity. 18. Wash the plate six times with PBS/0.01% Tween 20. 19. Prepare streptavidin-alkaline phosphatase enzyme conjugate 1:1000 dilution in PBS. 20. Add 100 mL per well of streptavidin-alkaline phosphatase. 21. Incubate for 45 min at room temperature. 22. Decant streptavidin, wash 3 times with PBS/ 0.01% Tween 20, followed by three washes with PBS. 23. Add 100 mL/well BCIP/NBT substrate.

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24. Incubate for 5 min. 25. Stop spot development under running water and wash extensively. 26. Let the plate dry overnight in the dark. Spot intensity may decrease with exposure to light. 27. On day four, analyze the plate using imaging system. 3.2 FluoresceneActivated Cell Sorting (FACS)

Single-cell cytokine analysis techniques include ELISpot, in situ hybridization, immunohistochemistry, all of which have significant drawbacks requiring either high technical skill or tedious data collection and analysis. Flow cytometry is a powerful analytical technique in which individual cells can be simultaneously analyzed for several parameters including the expression of surface and intracellular markers defined by fluorescent antibodies. Recently, fluorescent anticytokine and antichemokine monoclonal antibodies have become very useful for the intracellular staining and multiparameter flow cytometric analysis of individual cytokine-producing cells within mixed cell populations, providing a high resolution method to identify the nature and frequency of cells which express a particular cytokine(s). In this technique, cells are stained with fluorescent anticytokine antibodies, which are subsequently visualized with the use of a flow cytometer. This method is very powerful for mechanistic-type immune function studies (18, 19) and is probably one of the most versatile from the standpoint of immunotoxicology. Flow cytometry assays for measuring cytokines have been standardized, adding to their value (20). The basic steps for intracellular staining of cytokine in cell culture include: 1. Cell stimulation: stimulate the cells in the presence of a protein transport inhibitor during activation. 2. Harvest the cells into conical tubes. 3. Pellet the cells (120–260 g for 5 min) and wash them once with PBS/0.5%BSA (blocking Fc receptors is recommended, since it may be useful for reducing nonspecific immunofluorescent staining). 4. Fix the cells in 4% paraformaldehyde in PBS or 2% formaldehyde in PBS and mix well. 5. Let the cells stand at RT in dark for 20 min 6. Pellet the cells (120–260 g for 5 min) 7. Wash the cells twice with ice-cold PBS/0.5%BSA/0.1mM sodium azide and resuspend cells in staining buffer (PBS/ 0.5%BSA/0.1mM sodium azide) for storing cells at 4°C or in 90% FCS/10% dimethylsulfoxice (DMSO) for storing at −80°C. Fixed cells can be stored to continue the intracellular staining at a later time.

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8. Stain the cells with surface marker antibodies. For frozen cells, wash twice to remove DMSO. Suspend the cells in icecold PBS/0.5%BSA/0.1 mM sodium azide (50 µl for each test = 1–2 × 106 cells). and 9. Transfer the cells to 96-well plate containing the surface antibodies. 10. Stain for 15 min on ice in dark. 11. Add 150 µl PBS and spin. 12. Wash once with 200 µl PBS and spin (120–260 × g for 5 min). 13. Resuspend in 100 µl PBS (containing no protein). 14. Pellet the cells. 15. Resuspend in 150 µl of permeabilization buffer (0.5% saponin in PBS/0.5%BSA/0.1 mM sodium azide). 16. Mix gently with multichannel pipette. 17. Incubate the cells at room temperature for 10 min 18. Pellet the cells (flick out supernatant) 19. Intracellular staining: thoroughly resuspend the fixed/permeabilized cells in 25 ml per well of permeabilization buffer containing a pre-determined optimal concentration of a fluorochrome-conjugated anticytokine antibody or appropriate negative control. 20. Incubate the cells at RT for 30 min in the dark 21. Wash twice with 150–200 ml of permeabilization buffer. (Note that one wash may be sufficient, but more washes may decrease the background). 2 2. Wash twice with 200 ml of PBS/BSA/Azide buffer (no Saponin) 23. Suspend the cells in 200 ml of PBS/BSA/Azide buffer (no Saponin). 24. Transfer to FACS tubes. For most cytokines, cells can be left in the staining buffer and can be analyzed the next day. Extended incubation prior to analysis may result in reduced fluorescent signals. 25. Run FACS analysis. (For an example, see Chapter 21) (see Note 4) 3.3 Bioassays

While immunoassays maybe a more convenient method for quantization of cytokines, they only measure the immunological reactive material. They may or may not detect the biologically inactive material, such as cytokines bound to soluble receptors or degraded cytokine molecules. Bioassays, however, detect biologically active cytokines and can be as accurate and precise as immunoassays (21).

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A bioassay (biological assay) is one in which living material (cells, isolated tissues or whole animals) serves as the indicator system for the cytokine. Although for cytokine research this generally refers to an in vitro system utilizing defined cell lines, in fact the cytokine bioassay has evolved from whole animal models through isolated primary cells (or tissues), and then into immortalized cell lines and genetically engineered cells. Examples of bioassay types include: induction of proliferation (IL-1), maintenance of viability (IL-2), stimulation of migration (various chemokines), induction of secondary function (IL-6), and inhibition of function (IL-10). A description of the specific techniques is beyond the scope of this chapter; rather, the interested reader is directed elsewhere (22, 23). Bioassays provide valuable information concerning the potency of cytokine products. This is essential for evaluating batch-to-batch consistency, appropriate formulations and stability. Bioassay data are crucial at all stages in the development of cytokine products, from early research to final quality control of finished product. Over the years, World Health Organization international standards have been used to reduce the variation in the estimates of cytokine preparations within and between the laboratories for both immunoassays and bioassays. (see Note 5) 3.4 Gene Expression (RT-PCR Protocol)

Many cellular functions are regulated by changes in gene expression. Thus, quantification of transcription levels of genes plays a central role in the understanding of gene function and of abnormal alterations in regulation that may result in an immunotoxic effect. By looking at the genetic expression for cytokines, activational events may be evaluated at very early time points. Several molecular biology assays have been used for cytokine analysis, including Northern blot, dot or slot blot, Rnase protection assay, and so forth. But real-time reverse transcriptase polymerase chain reaction (RT-PCR) is by far the most widely used method to quantify cytokines from cells, tissues, or tissue biopsies. Real-time PCR, also known as kinetic PCR, qPCR, qRT-PCR and RT-qPCR, is quantitative PCR method for the determination of cope number of PCR templates such as DNA or cDNA in a PCR reaction. The method allows for the direct detection of PCR product during the exponential phase of the reaction, combining amplification and detection in a single step without the need for postPCR processing. There are two main methods of real-time PCR: TaqMan and intercalator-based. Both methods require a special thermocycler equipped with a sensitive camera that monitors the fluorescence in each well at frequent intervals during the PCR reaction. TaqMan PCR requires a pair of PCR primers, an additional fluorogenic probe, which is an oligonucleotide with both a reporter fluorescent dye and a quencher dye attached. Intercalator-based method, also known as SYBR Green method,

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requires a double-stranded DNA dye which binds to newly synthesized double-stranded DNA and gives fluorescence. TaqMan method is more accurate and reliable than the SYBR green method but also more expensive. The TaqMan assay was the first real-time PCR assay that has been developed. The amount of fluorescence released is directly proportional to the amount of product generated in each PCR cycle and thus can be applied as a quantitative measure of PCR product formation. One can choose among a diversity of competing instrumentations present in the market. All of them run the PCR reaction as a closed tube and measure product accumulation in real time during the course of PCR amplification. Differences between the instrumentations are the sample format (tubes, microplates, strip tubes, capillaries, etc.), the maximum sample number (ranging from 16 to 384), the length of a run (ranging from 30 min to 2 h), the light source (halogen or laser), the fluorescence wavelength detection, the possibility of performing single or multiplex (i.e., measuring different fluorescence emissions simultaneously) PCR reactions, the availability of melting curve analysis, and finally the price. In all cases, a software package is provided that measures the increase in fluorescence emission in real time, during the course of the reaction. Although the method allows fast, sensitive, and accurate quantification, different control assays are necessary for the method to be reliable. By construction of complementary DNA (cDNA) plasmid clones, standard curves are generated that allow direct quantification of every unknown sample. Furthermore, the choice of a reliable housekeeping gene is very important. Co-amplification of contaminating genomic DNA is avoided by the designing sets of primers that are located in different exons or on intron–exon junctions. A large number of cytokines and their receptors have been cloned at both cDNA and genomic level thus facilitating the exploitation of molecular biology within immunotoxicology. Sequences from databases can be easly obtained from EMBL (http://www.ebi.ac.uk:queries:queries.html) and NCBI (http:// www.ncbi.nlm.nih.gov). The isolation of RNA is the most critical step in the analysis of mRNA expression levels. Isolated RNA molecules are highly susceptible to degradation via the activity of ribonuclease, an enzyme which contaminates most of the laboratory apparatus and must be excluded from the test RNA. Some simple rules for creating such a laboratory environment have been laid down by Blumberg (24). Besides quality, the reproducibility in RNA yield is also important since poor reproducibility may result in a larger variability between the samples within an experimental group. Several RNA isolation kits employing columns have been marketed and are suitable in routine RNA analysis. Column-based

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methods have the advantage that they do not require a phase separation step, in which pipetting off the upper phase (that contains the RNA) may introduce additional variability. Furthermore, they do not require hazardous compounds such as phenol and chloroform. Isolated RNA are best stored in the short-term as an alcoholic precipitate at −20°C, at 0–80°C for longer period. 1. Reverse transcription Total RNA

2.0 mg

RT Buffer 10X

0.8 ml

Random primers 10X

2.0 ml

dNTP mix 25X

1.0 ml

Multi Scribe

1.0 ml

2. Incubate the tubes at 25°C for 10 min and then at 37°C for 2 h. 3. Store the 1st strand cDNA at −20°C until use for real-time PCR. 4. Real-time PCR . For a 50 ml reaction (96-well plate), use Assays-on-demand Gene Expression products, which consist of a 20X mix of the unlabeled PCR primers and TaqMan MGB probe (FAM dye-labeled): TaqMan mix

25.0 ml

20X Assays-on-demand

2.5 ml

Diluted cDNA in RNase free water

22.5 ml

If different reaction volumes are used, amounts should be adjusted accordingly. 5. Set up the experiment and the following PCR program on ABI Prism SDS 7000. 50°C 2 min, 1 cycle 95°C 10 min, 1 cycle 95°C 15 s (denature)-> 60°C 30 s (anneal/extend), 40 cycles 6. Prepare the mixture in each optical tube. 7. Analyze the real-time PCR result with the SDS 7000 software. 8. Examine the PCR specificity by using 5 ml from each reaction on a 3% agarose gel. (see Note 6) 4. Concluding Comments

A variety of methodologies are available to assess the function of T-cells (25), and the choice of which assays to employ can significantly affect the quality of the data produced. One of the most potentially powerful methods is the quantification of cytokines, which are the key signaling mediators in all innate and adaptive

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immune responses (26). In this chapter, we have provided details on the “how” of cytokine assessment, and many more techniques are available. However, one must also consider the “why” of cytokine assessment. An understanding of how these mediators function in both normal and pathological conditions will provide a key discriminator in determining which cytokines should be evaluated, at what time they should be measured (both postxenobiotic exposure as well as what stage within the physiological production cycle to examine), and how these data fit into a broader understanding of immunotoxic insults. Cytokines should not be “shotgunned”, that is, one should not measure them solely for the sake of measurement since the resulting data will be incomplete at the best and misleading at the worst. Rather, the data from these assays should be incorporated into an overall understanding of immunotoxicity from a mechanistic standpoint.

5. Notes 1. The multiplex ELISA has several advantages: (a) Less sample is required: In a 16 cytokine array, 50 ml of sample is required to detect all 16 cytokines, whereas running individual ELISAs would require almost 1 ml of sample. (b) Time: In approximately 3 h, 16 cytokines can be measured, whereas running 16 individual ELISAs would require 48 h. (c) Cost: A 16-plex array costs approximately $1,000, whereas purchasing 16 individual ELISAs would cost approximately $6,400 (in 2009 dollars). 2. Competitive ELISAs yield an inverse curve, where higher values of antigen in the samples or standards yield a lower amount of color change. 3. Although immunoassays have many desirable features, there are some problems to be aware of: (a) Several intrinsic soluble factors, such as heterophilic antibodies, may bind the animal antibodies that are used in an immunoassay producing an erroneous result. Naturally occurring anticytokine antibodies and soluble receptors can also interfere with the results of immunoassays. (b) Depending on the specificity of the antibodies used, the concentration of the cytokine may be seriously under estimated. (c) Another important consideration in the performance of both bioassays and immunoassays is the use of appropriate reference standards. These standards allow one to quantitate

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the amount of cytokine present, as well as providing continuity between the assays and over time. Not surprisingly, but often overlooked or disregarded, is the fact that not all commercially available recombinant cytokines, immunoassay kits, or antibody reagents are equivalent. This is not to imply that any of these reagents are necessarily “correct” or “better.” Rather, recombinant cytokines may differ somewhat in conformation, sequence, or post-translational processing, and thus yield different results in a bioassay or in-house developed immunoassays. In other words, different kits can give different results in terms of absolute values. 4. Critical parameters for cytokine staining include: cell type and activation protocol, the time of cell harvest following activation, the inclusion of a protein transport inhibitor during cell activation, and the choice of anticytokine antibody. Use of monensin or brefeldin A blocks intracellular transport processes and results in the accumulation of most cytokine proteins in the rough endoplasmic reticulum or Golgi complex. This leads to an enhanced ability to detect the cytokine-producing cells. Since these agents have a dose- and time-dependent cytotoxic effect, exposure must be limited. Furthermore, one should be aware of the possible effects of transport inhibitors on the expression levels of cell surface markers. 5. Two major disadvantages of bioassays include: (a) Alack of specificity of the indicator cells. A defining feature of cytokines is their pleiotropism and redundancy; most cytokines described to date can elicit several different responses in target cells. In addition, most cytokineresponsive cells respond to a number of different cytokines. The result is that there are very few available cell lines that respond only to a single specific cytokine. (b) Interference by extraneous factors. The ability of various factors to interfere with the assay is hard to anticipate or control. A number of intrinsic factors such as b2macroglobulin, endogenous soluble cytokine receptors, rheumatoid factors, heterophilic antibodies, and natural anticytokine antibodies are found in the body fluids, and all can interfere with the results (27). 6. In interpreting the results, one should always take into account that a discrepancy may exist between mRNA and protein levels. Furthermore, it is the protein that comprises the biological activity. On the other hand, a drawback of protein assessment over analysis of mRNA levels is that cytokines can only be measured in body fluids or cell supernatants but not in (intact) tissue. Cytokines may, in some cases, exert their effects only within a certain organ or tissue with a limited spill-over effect into the peripheral blood. Furthermore, the measurement of

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cytokine release into cell culture supernatant is essentially a cumulative measure and, unless time points are chosen very carefully, will not yield information on the way in which cells are responding to various stimuli and how a cascade of events is progressing. Thus, molecular techniques may exhibit their greatest utility when combined with the other assays.

References 1. House RV, Descotes J (2007) Cytokines in human health: immunotoxicology, pathology and clinical applications. Humana, Totawa, NJ 2. House RV (2001) Cytokine measurement techniques for assessing hypersensitivity. Toxicology 158:51–58 3. Whiteside TL (2002) Cytokine assays. Biotechniques 10:12–15 4. Vandebriel RJ, Van Loveren H, Meredith C (1998) Altered cytokine (receptor) mRNA expression as a tool in immunotoxicology. Toxicology 130:43–67 5. Overbergh L, Giulietti A, Valckx D, Decallonne R, Bouillon R, Mathieu C (2003) The use of real-time reverse transcriptase PCR for the quantification of cytokine gene expression. J Biomol Tech 14:33–43 6. Jason J, Larned J (1997) Single-cell cytokine profiles in normal humans: comparison of flow cytometric reagents and stimulation protocols. J Immunol Methods 207:3–22 7. Pala P, Hussell T, Openshaw PJ (2000) Flow cytometric measurement of intracellular cytokines. J Immunol Methods 243:107–124 8. Letsch A, Scheibenbogen C (2003) Quantification and characterization of specific T-cells by antigen-specific cytokine production using ELISPOT assay or intracellular cytokine staining. Methods 31(2):143–149 9. Robroeks CM, Jöbsis Q, Damoiseaux JG, Heijmans PH, Rosias PP, Hendriks HJ, Dompeling E (2006) Cytokines in exhaled breath condensate of children with asthma and cystic fibrosis. Ann Allergy Asthma Immunol 96(2):349–355 10. Winkler O, Hadnagy W, Idel H (2001) Cytokines detectable in saliva of children as appropriate markers of local immunity of the oral cavity – an approach for the use in air pollution studies. Int J Hyg Environ Health 204(2–3):181–184 11. Simpson JL, Wood LG, Gibson PG (2005) Inflammatory mediators in exhaled breath, induced sputum and saliva. Clin Exp Allergy 35(9):1180–1185

12. Whicher J, Ingham E (1990) Cytokine measurements in body fluids. Eur Cytokine Netw 1(4):239–243 13. Ryan CA, Gerberick GF, Gildea LA, Hulette BC, Betts CJ, Cumberbatch M, Dearman RJ, Kimber I (2005) Interactions of contact allergens with dendritic cells: opportunities and challenges for the development of novel approaches to hazard assessment. Toxicol Sci 88(1):4–11 14. Mitschik S, Schierl R, Nowak D, Jörres RA (2008) Effects of particulate matter on cytokine production in vitro: a comparative analysis of published studies. Inhal Toxicol 20(4):399–414 15. Jeurink PV, Vissers YM, Rappard B, Savelkoul HF (2008) T cell responses in fresh and cryopreserved peripheral blood mononuclear cells: kinetics of cell viability, cellular subsets, proliferation, and cytokine production. Cryobiology 57(2):91–103 16. Kreher CR, Dittrich MT, Guerkov R, Boehm BO, Tary-Lehmann M (2003) CD4+ and CD8+ cells in cryopreserved human PBMC maintain full functionality in cytokine ELISPOT assays. J Immunol Methods 278(1–2):79–93 17. Maecker HT, Moon J, Bhatia S, Ghanekar SA, Maino VC, Payne JK, Kuus-Reichel K, Chang JC, Summers A, Clay TM, Morse MA, Lyerly HK, DeLaRosa C, Ankerst DP, Disis ML (2005) Impact of cryopreservation on tetramer, cytokine flow cytometry, and ELISPOT. BMC Immunol 6:17 18. Arora SK (2002) Analysis of intracellular cytokines using flowcytometry. Methods Cell Sci 24:37–40 19. Ghanekar SA, Maecker HT (2003) Cytokine flow cytometry: multiparametric approach to immune function analysis. Cytotherapy 5(1):1–6 20. Maecker HT, Rinfret A, D’Souza P, Darden J, Roig E, Landry C, Hayes P, Birungi J, Anzala O, Garcia M, Harari A, Frank I, Baydo R, Baker M, Holbrook J, Ottinger J, Lamoreaux L, Epling CL, Sinclair E, Suni MA, Punt K,

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Calarota S, El-Bahi S, Alter G, Maila H, Kuta E, Cox J, Gray C, Altfeld M, Nougarede N, Boyer J, Tussey L, Tobery T, Bredt B, Roederer M, Koup R, Maino VC, Weinhold K, Pantaleo G, Gilmour J, Horton H, Sekaly RP (2005) Standardization of cytokine flow cytometry assays. BMC Immunol 6:13 21. Meager A (2006) Measurement of cytokines by bioassays: theory and application. Methods 38:237–252 22. Mire-Sluis AR, Page L, Thorpe R (1995) Quantitative cell line based bioassays for human cytokines. J Immunol Methods 187:191–199 23. House RV (1999) Cytokine bioassays: an overview. Dev Biol Stand 97:13–19

24. Blumberg DD (1987) Creating a ribonuclease-free environment. Methods Enzymol 152:20–24 25. Suni MA, Maina VC, Maecker HT (2005) Ex vivo analysis of T-cell function. Curr Opin Immunol 17:434–440 26. House RV (2002) Preclincial immunotoxicity assessment of cytokine therapeutics. In: Thomas JA, Fuchs RL (eds) Biotechnology and Safety Assessment, 3rd edn. Academic, San Diego, pp 191–231 27. Heney D, Whicher JT (1995) Factors affecting the measurement of cytokines in biological fluids: implications for their clinical measurement. Ann Clin Biochem 32: 358–368

Chapter 21 Flow Cytometry in Preclinical Drug Development Patrick B. Lappin Abstract Flow cytometry has many applications in clinical medicine allowing rapid and highly specific characterization of cells in solution (e.g., peripheral blood) or from dissociated tissues. The data generated from these analyses may be used to diagnose and monitor progression of disease as well as aid in prognostication of selected pathologic processes. In recent years, flow cytometric techniques have established a foothold in preclinical drug development providing an ability to identify and characterize both cell morphology and function, as well as to more clearly assign observed alterations in one or more cell attributes as intended or unexpected effects of new biopharmaceutical entities. The inclusion of flow cytometric evaluation assays (some described in this chapter) during preclinical drug development has increased and enhanced the detail of data generated to support the safety and efficacy of new biopharmaceuticals. Flow cytometry analyses used in preclinical drug development that are described in this chapter include immunophenotyping, peripheral blood cross-reactivity, binding activity and stability and cell receptor dynamics. Key words: Preclinical drug development, Flow cytometry, Cross-reactivity, Monoclonal antibody, Immunophenotyping, Receptor dynamics

1.Introduction Flow cytometry has long been utilized as a means to evaluate the properties of individual cells collected from biological systems. Characterization of surface or internal antigen expression, subcellular components (e.g., mitochondria) and/or activity attributes (e.g., protein expression, calcium flux, cell proliferation) using flow cytometry has increased the ability to specifically define simple and complex structural/functional changes in diverse cell populations that may occur following exposure to biopharmaceuticals. The examination of these changes with consideration for normal interspecimen variability can aid in the identification and characterization of adverse or toxic effects. R.R. Dietert (ed.), Immunotoxicity Testing: Methods and Protocols, Methods in Molecular Biology, vol. 598 DOI 10.1007/978-1-60761-401-2_21, © Humana Press, a part of Springer Science + Business Media, LLC 2010

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Flow cytometry technology involves the examination of individual cells based on a physical interaction of whole cells, subcellular components or labeled cell proteins with focused light of one or more specific wavelengths and the conversion of light to electronic signals subsequently displayed as event data. Laser scanning cytometry (LSC) uses laser-based optics and microscopy coupled with complex analysis software and fluorescence immunohistochemistry to characterize cell morphologic and biologic attributes within adherent cultured cells and tissue sections. LSC evaluation of intact monolayers or tissue sections allows the examination of individual cell characteristics while preserving the tissue/monolayer morphology; potentially, LSC can be applied to any tissue or cell monolayer type. With LSC, individual cell characteristics collected during analysis are traditionally directed by the instrument operator, thus requiring considerable hands-on time and extended data acquisition time for a large number of specimens with multiple measurable cell attributes. Traditional flow cytometry utilizes similar mechanics to evaluate individual suspended cells (disaggregated tissues, body fluids). All cells present in a sample suspension are available for analysis, and typically representative portions of the sample containing all cell types are processed through the cytometer. These diverse cell populations may additionally be subdivided into discrete cell types for evaluation post sample processing (subpopulation examination by electronic gating strategies on whole populations collected). As such, large numbers of cells from mixed populations can be subdivided and rapidly analyzed for a multitude of cell attributes, limited only by the ability to provide intact cells in suspension and the availability of suitable detector molecules for targets of interest. While both LSC and traditional flow cytometry have applications in biomedical research, traditional flow cytometric analysis has been more rapidly and widely assimilated into drug discovery, preclinical development and clinical safety evaluations. Preclinical and clinical flow cytometry applications are the focus of this chapter. 1.1. Regulatory Oversight

Despite the impressive technological power associated with flow cytometry, clearly defined guidance is required to ensure the quality and consistency in the data generated, particularly when the data is to be utilized in a regulatory submission package for drug approval. Regulatory guidance ranges from site-specific standard operating procedures (SOPs) defined by Good Laboratory Practices (1) to national and international guidance documents that define standards for the conduct of all aspects of drug development (2–4). Assays developed and used to characterize drug candidates may be otherwise regulated only by the limits of scientific acumen, the capabilities of flow cytometry equipment and staff, and available financial resources. Flow cytometric analyses

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directed toward unique properties of novel pharmaceuticals may not only yield data that are valuable in the characterization of these compounds, but may provide a basis of information or methodology for adaptation to flow cytometry-based assays that can be used in preclinical drug development or clinical research. Some of the flow cytometric assays used in drug discovery may be classified as “one-offs” specific for the compound and desired endpoint. Others may provide the basis for modification to other test systems and other classes of compounds, and they may be utilized as biomarkers or surrogate endpoints for monitoring once a candidate drug is in the marketplace. Preclinical drug development typically comprises multiple experimental designs (pharmacodynamic/pharmacokinetic, safety pharmacology, toxicology) utilizing the same test compound in one or more laboratory species. The most robust drug safety packages include measurable endpoints that are comparable across multiple studies, and when possible, across multiple species. For multiple candidates from a single drug class, cross-drug/study assay adaptability provides significant power to study conclusions, enhancing data acceptance by national and international regulatory agencies. The examination of peripheral blood and tissue mononuclear cells using traditional flow cytometry has become a standard for the detection and or monitoring of immunomodulation in both preclinical and clinical arenas. The increasing number of very specific markers for cellular proteins or chemical targets has encouraged the elaboration of testing and the complexity of data generated. 1.2. General Uses of Flow Cytometry in Discovery, Development and Clinical Research 1.2.1. Discovery

1.2.2. Development

Uses of flow cytometry technology in drug discovery include genomic and proteomic screening (bead technology), cell target detection and specificity, drug hit-to-lead generation and lead optimization (5). These applications are discussed elsewhere and are beyond the scope of this review. Suffice it to say that flow cytometry has the potential to provide high throughput screening of a large number of compounds resulting in expedited discovery timelines. The primary applications of flow cytometry to drug development involve assays to detect morphologic and/or functional alterations to one or more cell populations (usually immune cells) in association with the administration of biopharmaceutical agents. Assays may be designed or utilized to measure/monitor the anticipated pharmacologic effect and/or to identify the target/ off-target toxicity. Flow cytometric evaluation may be as simple as determining the presence/absence, magnitude and duration of a single morphologic effect, or as complex as identifying functional interactions within or between the cells that may impact a critical downstream process (e.g., drug binding to platelets resulting in

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decreased platelet aggregation and increased bleeding time, in which receptor saturation data provide a pharmacodynamic basis for dose selection in the transition from nonclinical to clinical testing (6)). The bulk of this chapter details some of the applications of flow cytometry in preclinical drug development. 1.2.3. Clinical

Most of the basic flow cytometry procedural steps used in discovery and preclinical stages of biopharmaceutical evaluation are applicable to the clinical phase of drug development. For example, applications for clinical flow cytometry evaluation include measurement of CD4+ T-lymphocytes to diagnose and monitor the progression of acquired immunodeficiency syndrome (AIDS), lymphocyte immunophenotyping to detect, characterize and evaluate the impact of treatment on immunosuppressive disease, or diagnosis of and therapeutic planning for lymphoid neoplasia (7, 8). Interestingly, the character and dynamics of some clinical effects observed with immunomodulatory therapeutics and measured using flow cytometry may be identical to the alterations tracked in preclinical drug development. Some clinical flow cytometric parameters may serve as biomarkers that are routinely evaluated to provide a measure of therapeutic efficacy and/or treatment optimization (e.g., peripheral blood B-lymphocyte monitoring with anti-CD20 therapeutics).

1.2.4. Flow Cytometry Method Qualification/ Validation

Procedures (assays) designed to generate biologic data must exhibit the following experimental characteristics: (1) precision (high level of sensitivity and accuracy), (2) specificity (predictable yet limited target), (3) consistency (similar result expectation across samples), (4) reproducibility (similar/same results expected within subdivided individual samples), and (5) robustness (experimental process and resulting data that are not unduly impacted by minor differences in experimental methodology). Validation of these methods requires the ability to consistently generate data that is within the published limits of regulated control reagents specific for the method/assay to be validated. Validation of general methods used for flow cytometric analysis is a relatively straight-forward process following the standard practices described elsewhere (9). Complete validation of specific flow cytometric assays, however, can present significant challenges based on the species for which the assay is being conducted. Appropriate flow cytometry assay control reagents exist for human peripheral blood mononuclear cells and for similar cells in a limited number of laboratory animal species. Similar control reagents are generally lacking for rats and nonhuman primates (two common species utilized for preclinical safety and pharmacodynamic studies) resulting in the inability to “officially” validate these assays. However, under the guidance of testing facility SOPs, close concordance with regulatory guidances and using

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sound scientific practices, flow cytometry assays can be qualified (incomplete validation) using some published literature and historical database ranges (if available), and site experience to enable utilization in regulated preclinical studies. Additionally, biological samples utilized in the conduct of these assays must be consistent (sample content/quality based on the collection and/ or processing methods) and stable (not subject to degradation within the limits of the assay). 1.2.5. Specific Flow Cytometry Applications

The following sections provide examples of flow cytometry-based assays which are currently in use and in association with preclinical drug development. Additional description of the data generated, applications of data to preclinical toxicology and general procedural rules for flow cytometry assays are provided.

2. Materials (see Note 1) 2.1. Immunophenotyping (Single Cell Suspensions from Body Fluids, Bone Marrow or Lymphoid Tissues): Non-human Primate and Rat

1. Control cells (e.g., lyophilized Coulter Cyto-Trol™ cells – Beckman Coulter, Miami, FL) reconstituted with provided diluent. 2. 1% Neutral Buffered Formalin (NBF) – 10% NBF diluted 1:10 in 1× PBS. 3. 1× Phosphate-buffered Saline (1× PBS). 4. Roswell Park Memorial Institute (RPMI) 1640 – Bone marrow only. 5. Fetal Bovine Serum (FBS) – Bone marrow only. Note: 1 mL FBS to 9 mL RPMI 1640 for 10% solution. 6. Fluorochrome-conjugated cell target antibodies and corresponding isotype control antibodies, as appropriate (order species specific). 7. Red blood cell lysing reagents (based sample type and on manual or automated lysis). (a) Manual lysis, used as directed (e.g., BD FACS™ Lysing Solution, Becton Dickinson (BD Biosciences), San Jose, CA). (b) Automated lysis (e.g., Coulter TQ-Prep Workstation and ImmunoPrep® Reagent System (Beckman Coulter, San Jose, CA)). 8. Optional: Quantitation beads (e.g., Flow-Count™ Fluorospheres, Beckman Coulter, San Jose, CA).

2.2. Peripheral Blood Cross-reactivity Evaluation: Multiple Species

1. Pure test antibody, biotinylated test antibody or test antibody– fluorochrome conjugate to be evaluated for cross-reactivity (may include all forms AND may be multiple forms of one or more test antibodies). Usually provided by the investigator.

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2. Pure isotype antibody, biotinylated isotype antibody or isotype–fluorochrome conjugate corresponding to the 1 or more test antibodies. May be provided by the investigator or available from a commercial provider (see Note 2). 3. Fluorochrome-conjugated target cell antibodies (if available) for use in confirmation of appropriate test antibody binding. 4. Streptavidin–fluorochrome secondary reagent if any biotinylated antibody is utilized. 5. Fluorochrome-conjugated isotype anti-test antibody if any pure test antibody is utilized. 6. 1% NBF: 10% NBF diluted 1:10 in 1× PBS. 7. 1× phosphate-buffered saline (1× PBS). 8. Wash buffer (1× PBS or other buffer defined by test/isotype antibody specifications). 9. Red blood cell lysing reagents (based on sample type and on manual or automated lysis (see Subheading 21.2.1, item 7)). 10. Additional reagents based on requirement of the specific test/ isotype antibody or analysis procedure may include different buffer solutions (e.g., 1× PBS with 10% BSA) or cell preservative/antimicrobial (e.g., sodium azide). 11. Additionally, positive/negative control cell lines or transfected cells (known to express target antigen) may be used for experimental quality control in conjunction with or in place of cell target antibodies. 2.3. Binding Activity and Stability Testing for Novel Test Antibody: Cultured Cell Lines (see Note 3)

1. Appropriate positive and negative control cell lines (ATCC, Manassas, VA). Positive control cells must express detectable target antigen specific for the test antibody; negative cells must be free of target antigen. Control populations must be individual cell suspensions OR be amenable to disaggregation without loss of target antigen. 2. Appropriate media for maintenance and/or growth of positive and negative cell lines (see Note 4). 3. One or more test antibodies (pure and biotin-labeled are forms most often tested; usually provided by the investigator). The corresponding isotype control antibodies are also required to test and ensure the specificity of labeling. 4. Labeled antibodies may be provided or pure antibody may be biotinylated following receipt. 5. Streptavidin–fluorochrome conjugate – labeling reagent for biotinylated test/isotype antibodies. 6. Goat (or other species) anti-human IgG–fluorochrome conjugate – labeling secondary antibody for pure test/isotype antibodies (see Note 5).

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7. 10% BSA in 1× PBS. 8. 1% BSA/0.1% sodium azide in 1× PBS (staining buffer). 9. 1% BSA/0.1% sodium azide/1% NBF. 2.4. Receptor Occupancy Assay: Multiple Species (see Note 6)

1. Pure test antibody (unlabeled) – usually provided by the investigator. 2. Fluorochrome-conjugated test antibody – may be labeled and provided by the investigator or provided as pure test antibody to be subsequently labeled on-site or by a subcontractor. 3. Fluorochrome-conjugated cell target antibodies if appropriate (can only be used if there is no binding interference with the test antibody). 4. Red blood cell lysing reagents based on sample types and on manual or automated lysis (see Subheading 21.2.1, item 7).

3. Methods Flow cytometric analysis of suspended, unfixed cells may present several unique challenges to the investigator. Considerations for potential impacts of sample collection, handling, processing and even sample evaluation on the accuracy, quality and consistency of flow cytometry data must be included during the experimental design phase. Consistent sample collection is imperative to the generation of quality data. Liquid samples containing suspended cells (whole tissue aspirates/disaggregations, bone marrow, peripheral blood) must be collected using the same collection process (e.g., syringe vs. evacuated sample container), with the same anticoagulant (concentration and volume/sample) and subsequently held under the same conditions (temperature, humidity, time) prior to processing. While the collection procedure and anticoagulant are unlikely to waver during the course of an experiment, storage conditions may be inconsistent and result in considerable variability in the results. Some surface antigens on tissue and peripheral blood cells are extremely labile and greatly affected by the temperature and storage time. Sample handling during collection and storage may also impact the quality and consistency of the data generated by flow cytometric analysis. Rough handling and excessive or delayed processing of samples to be used for platelet activation assays would clearly result in suboptimal results (if any were to be obtained at all). Rough handling or excessive/delayed processing of peripheral blood cells may result in ex vivo cell activation (important if an end point is the measurement of effects of a

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biopharmaceutical agent on cell activation in vivo) or loss of surface antigens (not all cells of a subpopulation will be affected uniformly). Excessive processing (multiple sample washes) of peripheral blood or disaggregated tissue lymphocytes may result in the loss of selected cell subtypes (B-lymphocyte specific reductions – personal observation). The use of monoclonal or polyclonal antibodies for phenotyping of immune cells by flow cytometry also requires the consideration of antibody structure and antibody–antigen interaction properties. For any experiment in which data will be compared between individuals or test (dose) groups, a specific antibody clone (from a specific manufacturer) that is known to be crossreactive with the test species and, if possible, the use of a specific lot of that clone should be utilized throughout the experiment to ensure consistent results. Most antibodies derived from large commercial manufacturers exhibit highly consistent labeling patterns/proportions from lot to lot (this character, however, rarely holds true when comparing the same antibody clone from two different manufacturers). Inter-lot comparability is often antibody specific (e.g., high level of lot comparability with anti-CD4 clones but poor consistency between lots from some anti-CD14 clones). Antibody lot comparability evaluations must be completed prior to placing a new lot of antibody into an established project to avoid data interpretation errors. Antigen sequences are not uniformly conserved across species, and as such, a single antibody type/clone will not always cross react with the same antigenic target in multiple species. Additionally, the same antigenic target in multiple species may be associated with different biologic pathways making the use of a common antibody to target a specific cell population completely appropriate in one species and inaccurate in another (10). The application of cross-reactivity method development experiments to a more comprehensive study design may be required to confirm an appropriate antigen–antibody binding. Most commercially manufactured antibodies specific for individual species are titred for optimal performance during the quality control process performed by the manufacturer. In late stage preclinical safety assessment, it is not uncommon for human cell-directed antibodies to be utilized for non-human primate studies. Following the confirmation of cross-reactivity and antigen specificity, these anti-human antibodies should be titred to determine the optimal concentrations for cell detection in a non-target species, to ensure appropriate and consistent performance. With the advancement of flow cytometry systems and increases in the number of detection channels available on newer instruments, care must also be taken in the selection of target label (fluorochrome conjugated antibody or histochemical agent)

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utilized for multi-parameter evaluation. The excitation and emission spectra for a small number of fluorochromes can be optimized for maximal separation and minimal overlap, but the expansion of analyses into six or more fluorochromes results in more complex spectral separation issues, potentially confounding the ability to detect rare events. Additionally, some fluorochromes are large molecules that may introduce stearic hindrance complications where two or more antigen targets are proximate to one another. Appropriate assignment of antibody–fluorochrome conjugates to specific cell populations based on the expected cell density, antigen characteristics (if known) and fluorochrome properties will enhance the detection and quantitation of some cell subtypes. Note: Instrument initialization and set-up procedures vary between flow cytometers. These procedures are included in specific instrument operation manuals and will not be addressed here. The data generated in the course of flow cytometry analysis is formatted by the user prior to and during analysis; specific steps associated with gating and data analysis/interpretation are beyond the scope of this chapter. Tube or multiwell plate formats may be used for sample preparation and analysis. All methods descriptions below are confined to assays done with 12 × 75 mm polypropylene tubes. Additionally, all methods described below encompass the use of monoclonal or polyclonal antibodies; flow cytometric analyses using the histochemical markers (e.g., myeloperoxidase for bone marrow analysis) are described elsewhere (11) but are not included in this chapter. The following materials and methods apply to experimental procedures involving non-human primate samples (unless otherwise specified). 3.1. Immunophenotyping: Non-human Primate and Rat

This assay is used to detect changes in cell counts or ratios and may be used to monitor progression of and/or recovery from those changes. Similar experimental methodology may be used to detect markers of cell activation, cell maturity and some markers involved with intra- or intercellular processes. The availability of appropriately cross-reactive antibodies suitable for flow cytometry may limit the ability to measure some antigen expression by this method, requiring the use of other technologies (e.g., ELISA, Western Blot). 1. Prepare human control cells per manufacturer’s directions. 2. Centrifuge prepared control cells at room temperature (RT) for approximately 5 min at 300 × g. 3. Remove the supernatant and wash the cell pellet once with 1× PBS. 4. Resuspend the cells and centrifuge at RT for approximately 5 min at 300 × g. 5. Remove the wash supernatant.

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6. Resuspend the control cells in 0.5 mL of 1% NBF. 7. Retain at RT until analysis (see Note 7). 8. Acquire the appropriate cell suspension sample(s): (a) Peripheral blood or body fluid (cerebrospinal fluid, tissue cysts) aspiration in anticoagulant (no further preparation). (b) Bone marrow – samples collected in anticoagulant from appropriate marrow cavity (humerus/pelvis in monkey; femoral shaft in rat). i. Immediately resuspend samples in 5–15 mL ice cold RPMI/10% FBS. ii. Centrifuge for approximately 5 min at 4°C at 380 × g. iii. Resuspend the pellet in 5 mL of ice cold RPMI/10% FBS. (c) Tissue aspirate – mechanical disaggregation to conserve surface proteins (repeated pipetting). Resuspend cells in 1X PBS or other appropriate media (d) Lymphoid tissue – mechanical disaggregation [manual mashing/scraping; automated Medimachine™ (Becton Dickinson)]. i. Filter and resuspend cells in 1× PBS or other appropriate media. Note: Proteinase will disrupt/destroy some proteins and should not be used for cell surface labeling experiments. 9. Resuspend the cells uniformly by gently mixing control and test sample(s) for 3–5 min. 10. For all samples other than peripheral blood, measure the number of nucleated cells in an aliquot of resuspended cells (to allow standardization of cellular density for staining). 11. Place 20–100 mL aliquots of appropriately diluted (if required) test sample or undiluted control sample in the bottom of each tube (sample on the side of the tube may not be exposed to labeling agent) (see Note 8). 12. Add the appropriate volume of labeling antibody (single or mixture) to control and test sample aliquots. 13. Gently mix, and incubate for 30 min at room temperature with samples protected from bright light (labeling interval, temperature may vary with the antibody). 14. Lysis, fixation and wash steps (a) Peripheral blood, bone marrow or body fluid aspirates: i. Lyse/fix test samples (per manufacturer’s instructions).

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ii. Immediately analyze all samples OR refrigerate lysed/fixed samples until analysis (4°C for no longer than 172 h). (b) Tissue aspirates or disaggregated lymphoid tissues: i. Lyse test samples. ii. Centrifuge lysed samples for approximately 5 min at RT at 350 × g. iii. Remove the supernatant and resuspend the cell pellet in 0.5 mL of 1% NBF (see Note 9). 15. Add 100 mL of Flow-Count™ Fluorospheres and briefly vortex mix (optional) (see Note 10). 16. Analyze the prepared control and test samples for specific cell subpopulations. 3.2. Cross-reactivity: Multiple Species (see Note 11)

1. Procure anticoagulated blood samples from species to be tested. Note: If sample procurement requires using an off-site vendor, ensure that detailed collection, handling and shipment instructions are provided to the appropriate individuals. 2. Dilute test and isotype antibodies to the appropriate concentrations using 1× PBS or another specified diluent. Dilutions should be made fresh daily for multi-day experiments. 3. Prepare any other secondary antibody or labeling reagents as needed for the study design. 4. Gently mix all blood samples for 3–5 min to uniformly resuspend cells. 5. Place 20–100 mL aliquots of blood for each individual into the bottom of the appropriate tube (see Note 12). 6. Add the appropriate volume of test antibody/isotype antibody (based on concentrations to be tested) to the sample aliquots. 7. Gently mix and incubate for 30 min at room temperature with samples protected from bright light (labeling interval, temperature may vary with the antibody). If using test/isotype antibodies conjugated directly to a fluorochrome label, cell target antibodies (if available) can also be added at this step. If test antibody and corresponding isotype antibody require a secondary and/or tertiary labeling step, one or more wash steps between labeling may be required. 8. Wash and secondary/tertiary labeling steps (if necessary) (a) If using pure (unlabeled) test/isotype antibody, follow the initial incubation by the addition of 1 mL wash buffer (1× PBS or other buffer).

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(b) Resuspend the cells. (c) Centrifuge for 5 min at RT at 350 × g. (d) Remove the supernatant. (e) Resuspend the resulting cell pellet in the appropriate volume of secondary antibody (conjugated to biotin or to a fluorochrome). (f) Gently mix and incubate for 30 min at room temperature protected from bright light. If using a biotinylated secondary antibody, the second incubation step is followed with: i. Resuspend the cells in wash buffer. ii. Recentrifuge and remove the wash supernatant. iii. Resuspend the cell pellet in the conjugated streptavidin–fluorochrome. Additional cell target antibodies can also be added to the sample at this step. (g) If using biotinylated test/isotype antibody secondary labeling reagent (streptavidin–fluorochrome conjugate): i. Resuspend the cells in wash buffer. ii. Centrifuge for approximately 5 min at RT at 350 × g. iii. Remove the wash supernatant. iv. Incubate the cells incubated with streptavidin–fluorochrome for 30 min at room temperature protected from bright light. Additional cell target antibodies can also be added to the sample at this step. 9. Complete the red blood cell lysis after the last labeling step (based on test/isotype antibody format described above). 10. Lyse and fix samples (per manufacturer’s instructions). 11. Immediately analyze all samples OR refrigerate the lysed samples until analysis (4°C for no longer than 172 h). 12. Vortex mix briefly prior to analysis (see Note 13). 13. Analyze samples for test antibody binding. If cell target antibodies are utilized, test antibody binding to specific subpopulations can also be determined. Identification of non-specific binding of the isotype antibody is used to determine the specificity of test antibody bindings. 3.3. Binding Activity and Stability Testing: Cultured Cell Lines (see Note 14)

1. Prepare dilutions of the test/isotype antibodies in an appropriate buffer or staining buffer to provide 5–10 concentrations that span the range from undetectable to at or above the expected therapeutic concentration. 2. Prepare dilutions fresh daily for multi-day experiments. 3. Aliquot an appropriate volume of positive or negative control cells (approximately 106 cells per mL) in labeled tubes.

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4. Centrifuge tubes for approximately 5 min at 4°C at 380 × g. 5. After centrifugation, remove the supernatant and resuspend each cell pellet in the appropriate volume of pure or biotinylated test or isotype antibody. 6. Vortex mix for approximately 5 s. 7. Incubate at 2–8°C for 30 min. 8. Following incubation, centrifuge cells for 5 min at approximately 4°C at 380 × g. 9. Remove the supernatant and resuspend the cells in 3.0 mL of cold staining buffer. 10. Vortex mix for approximately 5 s. 11. Centrifuge for 5 min at approximately 4°C at 380 × g. 12. Remove supernatant and resuspend cell pellets in an appropriate volume of streptavidin–fluorochrome (for biotinylated test/isotype antibodies) or anti-human IgG–fluorochrome (for pure test/isotype antibodies). 13. Vortex mix for approximately 5 s. 14. Incubate at 2–8°C for 30 min. 15. Following incubation, centrifuge cells for 5 min at approximately 4°C at 380 × g. 16. Remove the supernatant and resuspend the cells in 3.0 mL of cold staining buffer. 17. Vortex mix for approximately 5 s. 18. Centrifuge cells for 5 min at approximately 4°C at 380 × g. 19. Remove the supernatant and resuspend the cells in 0.5 mL of cold fixation buffer. 20. Vortex mix for approximately 5 s. 21. Analyze positive and negative control cells labeled with test or isotype antibodies to determine the optimal concentration, labeling specificity and binding stability (if measured). 3.4. Receptor Occupancy (see Note 14)

1. Preparation of control samples for generation of negative– positive antigen availability range. Note: this preparation can be done concurrent with test subject sample preparation. (a) Aliquot 100 mL of blood into two clean labeled tubes. (b) To one tube, add a saturating concentration of unlabeled test antibody (“spiked sample”) and incubate at RT for 10 min. (c) After incubation with unlabeled test antibody, add 10 mg (or another concentration based on antibody titration – if available) of fluorochrome-conjugated test antibody and one or more cell target antibodies (if appropriate).

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(d) Incubate at RT for an additional 15 min. (e) To the second sample tube (“unspiked sample”), add 10 mg fluorochrome-conjugated test antibody and one or more target cell antibodies (if appropriate). (f) Incubate for 15 min. (g) Lyse the red blood cells in both samples per the manufacturer’s instructions. (h) Vortex mix for approximately 5 s. (i) Immediately analyze all samples (see Note 16). 2. Preparation of test subject samples (requires two aliquots of 100 mL of anticoagulated blood placed into clean labeled tubes). (a) To one tube, add a saturating concentration of unlabeled test antibody (“spiked”). i. Incubate at RT for 10 min. ii. After incubation with unlabeled test antibody, add 10 mg of fluorochrome-conjugated test antibody and one or more cell target antibodies (if appropriate). iii. Incubate at RT for an additional 15 min. (b) To the second sample tube (“unspiked”), add 10 mg fluorochrome-conjugated test antibody and one or more target cell antibodies (if appropriate). i. Incubate for 15 min. (c) Lyse the red blood cells in both samples per the manufacturer’s instructions. (d) Briefly vortex mix. (e) Immediately analyze all samples. 3. Sample analysis (a) Analyze “spiked” and “unspiked” control samples prepared on each experimental day to determine the range of negative to positive for that day. Figure 21.1 shows typical labeling patterns for “spiked” and “unspiked” samples, as well as an example of partial labeling indicative of incomplete receptor availability. (b) Positive, negative or partial labeling for control and test subject “spiked” and “unspiked” samples are defined in Table 21.1 (see Note 17). (c) Use additional measurements over time to determine persistence and/or recovery of target antigens.

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Fig. 21.1. Histogram patterns of “Unspiked” and “Spiked” peripheral blood samples subsequently stained with fluorochrome-conjugated test antibody. Histogram (a) displays a typical pattern for “Unspiked” control cells exhibiting an abundance of available antigen (a single peak in region D). Histogram (b) shows the expected pattern for a “Spiked” control sample (a single peak of unlabeled cells in region C). Histogram (c) displays predominantly negative cells with a small shoulder of positive cells in region F, indicating limited antigen availability in this sample. The patterns exhibited in these representative samples could indicate complete receptor availability due to lack of test antibody binding or full post-occupancy recovery (Histogram (a)), complete test antibody binding or lack of receptor recovery (Histogram (b)), or partial test antibody binding/recovery (Histogram (c)). Note: differences in the widths of regions C, D and F or Y-axis scale on these representative histograms reflect minor daily variation in positive/negative control standards used to generate these data. These differences highlight the importance of establishing controls on each analysis day. Y-axis represents the number of events. X-axis indicates the relative fluorescence intensity of all events evaluated.

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Table 21.1 Expected receptor occupancy labeling pattern In vivo test antibody binding pattern

Pure test antibody “spiked” samplea “Unspiked” sample

Saturation

No fluorescent signal

No fluorescent signal

No binding

No fluorescent signal

Maximal fluorescent signal

Intermediate binding

No fluorescent signal

Partial florescent signal

Lack of antibody specificity or absence of No fluorescent signal target antigen

No fluorescent signal

“Spiked” sample exhibiting positive signal may reflect absence of pure test antibody receptor saturation

a

4. Notes 1. Unless and otherwise specified, multiple manufacturers exist for many of the common reagents used in the described assays. Selection of a specific provider for common reagents is at the user’s discretion. Additionally, reagents required for instrument operation, instrument QC, compensation, etc. may be specific to the brand and model of instrument (e.g., flow cytometer) utilized in the analysis. These reagents are described in the instrument operations manuals and are not discussed here. 2. Pure test antibody may be used “as is” – binding is identified with a fluorochrome or biotin-conjugated secondary antibody directed at the isotype of test antibody. If a biotin-conjugated anti-test antibody is used, a streptavidin–fluorochrome reagent combination is subsequently applied for signal detection. 3. This type of assay may be done to characterize one or more test antibodies for the subsequent use in other experimental procedures (e.g., stability of biotinylated antibody for tissue cross-reactivity testing). Binding activity may involve the analysis for the percentage of cell labeling over a wide range of antibody concentrations, as well as the evaluation of pure antibody vs. labeled antibody (e.g., biotin) binding efficiency. Stability evaluation may involve the effects of binding by temperature (e.g., one or multiple freeze-thaw cycles), storage time or other variable experimental conditions. 4. Cell culture capability (maintenance, storage – liquid nitrogen, growth) may be required based on the cell population and requirements of the experimental procedure. 5. Secondary antibody must be derived from a species different from the primary (test/isotype) antibody, but with specificity against the subtype of the primary antibody.

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6. This assay is utilized to determine receptor dynamics following in vivo exposure of a target antigen on a peripheral blood cell to a test antibody. Receptor dynamics may include: (1) the percentage of antigens bound by test antibody based on the concentration of test antibody administered, (2) the duration of test antibody–antigen binding following exposure (single or multiple doses), and (3) the presence/absence of antigen recovery and the time interval required for partial/complete recovery of the unbound antigen. Data derived from this procedure may further define pharmacodynamic properties of the test antibody to help optimize optimizing dosing intervals. This assay may also be used to characterize the differences in receptor dynamics between similar cells in human and appropriate laboratory species. 7. For non-human primate immunophenotyping by flow cytometry, control cells (e.g., CytoTrol™) are used to confirm the antibody reactivity and cytometer performance on each analysis day. These cells are not non-human primate controls and cannot be utilized to validate non-human primate assays. 8. The ideal cell concentration (for most commercially available flow cytometry immunophenotyping antibodies) is 1–2 × 106 nucleated cells/mL for peripheral blood or other body fluids and 5–10 × 106 nucleated cells/mL for tissue cells or bone marrow. The low volume stated in the “per aliquot” range is based on the availability of sample from the rodents and the number of markers required to be analyzed (more markers per animal may require less sample per aliquot). 9. All samples should be briefly vortex mixed prior to analysis. 10. Quantitation fluorospheres may be used to provide real-time subset counts; alternatively, percentages derived from the cytometer analysis can be multiplied by absolute lymphocyte counts (from hematologic analysis) to provide absolute subset counts. 11. Each experiment will vary based on the species evaluation requirements, availability of one or more test antibodies and specific properties of the test antibodies. In most instances, three or more concentrations of test antibody spanning a range of anticipated therapeutic blood/tissue concentrations are evaluated in at least three unrelated individuals from each test species. Data generated in cross-reactivity studies performed using flow cytometry may be used for lead selection (multiple test antibodies evaluated) or species selection (based on presence and robustness of cross-reactivity). 12. The low volume stated in the “per aliquot” range is based on the availability of sample from rodents and the number of

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markers required to be analyzed (more markers per animal may require less sample per aliquot). 13. Red blood cell lysis may be done prior to labeling for some test antibodies if the presence of red cells interferes with labeling or interpretation. However, for prelabeling lysis to be successful, the target antigen must be unaffected by the lysing process. 14. The following description is based on the procedural steps performed at refrigerated temperatures. The temperature required for centrifugation/incubation may vary based on the character of the test/isotype antibodies and the experience of the laboratory. 15. Anticoagulated blood samples will be collected from test subjects prior to initial dosing with test antibody to establish a baseline for each test subject. Subsequent sample collections will be scheduled over the proposed course of the experiment. On each experimental day, a single sample is collected from an untreated test subject to be used to establish a negative– positive antigen-availability control range. 16. The “spiked” sample is used to measure the receptor availability after complete saturation of the antigen (theoretically, should exhibit no labeling because pure test antibody should bind to all available receptors and block binding of fluorochrome-conjugated test antibody). Conversely, the “unspiked” sample is used to measure the receptor availability after no exposure to test antibody (theoretically, should exhibit a measure of 100% antigen availability). 17. For samples in which in vivo test antibody dosing results in saturation of binding sites, both the “spiked” and “unspiked” samples will exhibit minimal to no binding of the fluorochrome-conjugated test antibody (i.e., no fluorescent signal on analysis). For samples in which in vivo test antibody exposure results in no or little receptor binding, the “spiked” sample will be fluorescence-negative while the “unspiked” sample should exhibit maximal fluorescence positivity. Samples for which a portion of available receptors are bound during in vivo dosing will exhibit no fluorescent signal in “spiked” samples and a partial (percentage of maximum) fluorescent signal in the “unspiked” sample. If cell target antibodies are also utilized, receptor occupancy can be measured on specific cell subpopulations.

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Acknowledgments The author would like to acknowledge Charles River Laboratories for providing the opportunity to build on a long term interest in flow cytometry through supported development of a laboratory and experimental assays that clearly enhance preclinical safety assessment. The author would like to thank Dr. Keith Reimann for development and optimization of the receptor occupancy assay, and for his advice and encouragement in general flow cytometry analyses. The author would also like to acknowledge Dr. Rafael Ponce for editorial review and content advice, and Dr. Stephanie Fraser for technical and editorial contributions to this work. References 1. Good Laboratory Practice Regulations, 21 CFR 58, Department of Health and Human Services, Food and Drug Administration, U.S. Government Printing Office, Washington, DC. Effective 1979 2. International Conference on Harmonization (ICH) Harmonized Tripartite Guideline “S6” Note for Guidance on Preclinical Safety Evaluation of Biotechnology-derived Pharmaceuticals, (62 FR 61515, November 18, 1997; CPMP (Committee for Proprietary Medicinal Products)/ICH/302/95, September 1997) 3. International Conference on Harmonization (ICH) Harmonized Tripartite Guideline “S8” Note for Guidance on Immunotoxicity Studies for Human Pharmaceuticals,” (70 FR 61133-61134, October 20, 2005; CPMP (Committee for Proprietary Medicinal Products)/ICH/423/02, November 2005) 4. US Food and Drug Administration (2001) Guidance for industry immunotoxicology evaluation of investigational new drugs. US Food and Drug Administration; Center for Drug Evaluation and Research 5. Sklar LA, Carter MB, Edwards BS (2007) Flow cytometry for drug discovery, receptor pharmacology and high-throughput screening. Curr Opin Chem Biol 5:527–534

6. Visich J, Ponce R (2008) Science and judgment in establishing a safe starting dose for first-in-human trials of biopharmaceuticals. In: Cavagnaro J (ed) Preclincal safety evaluation of biopharmaceuticals: a science-based approach to facilitating clinical trials. Wiley, Hoboken, NJ, pp 971–984 7. O’Gorman MR, Zijenah LS (2008) CD4 T cell measurements in the management of antiretroviral therapy – a review with an emphasis on pediatric HIV-infected patients. Cytometry B Clin Cytom 74(Suppl 1):S19–S26 8. Hedley DW, Chow S, Goolsby C, Shankey TV (2008) Pharmacodynamic monitoring of molecular-targeted agents in the peripheral blood of leukemia patients using flow cytometry. Toxicol Pathol 36(1):133–139 9. Owens MA, Vall HG, Hurley AA, Wormsley SB (2000) Validation and quality control of immunophenotyping in clinical flow cytometry. J Immunol Methods 243:33–50 10. Ianelli CJ, Edson CM, Thorley-Lawson DA (1997) A ligand for human CD48 on epithelial cells. J Immunol 159(8):3910–3920 11. Criswell KA, Bleavins MR, Zielinski D, Zandee JC, Walsh KM (1998) Flow cytometric evaluation of bone marrow differentials in rats with pharmacologically induced hematologic abnormalities. Cytometry 32(1):18–27

Chapter 22 Enhanced Histopathology Evaluation of Lymphoid Organs Susan A. Elmore Abstract Enhanced histopathology is a tool that the pathologist can use as a screening test to identify immunomodulatory compounds. This assessment is based on the assumption that chemically induced alterations may result in qualitative or quantitative changes in the histology of the lymphoid organs. It involves the histological evaluation of various lymphoid organs and their respective tissue compartments to identify specific cellular and architectural changes. Although this methodology cannot directly measure immune function, it does have the potential to determine whether or not a specific chemical causes suppression or enhancement of the immune system. As with all screening tests, evaluation of, and comparison with, control tissues are crucial in order to establish the range of normal tissue changes for a particular group of animals. Laboratory animals include species other than rat and mouse; therefore, recognition of species differences in the structure and function of the immune system should be noted as well as identification of which differences are biologically relevant for the endpoint being considered. Consideration should also be given to the nutritional status, antigen load, age, spontaneous lesions, steroid hormone status, and stress for each strain and group of animals. General guidelines for the examination of each of the lymphoid organs are provided in this chapter. Key words: Enhanced histopathology, Thymus, Spleen, Lymph nodes, MALT, Bone marrow

1. Introduction Immunomodulatory compounds may produce changes in cell production and cell death as well as cellular trafficking and recirculation. Such changes may be expressed as alterations in cell type, cell density or compartment sizes within lymphoid organs (1–4). Due to the recent increased focus on ensuring consistency in the evaluation of xenobiotics for immunotoxicity, The Society of Toxicologic Pathology (STP) authorized the formation of the STP Immunotoxicity Screening Working Group to publish a “best practice” concept for the proper examination and reporting of lymphoid organs for enhanced histopathology (5). Subsequently, a series of articles was published in a special edition of the Toxicologic R.R. Dietert (ed.), Immunotoxicity Testing: Methods and Protocols, Methods in Molecular Biology, vol. 598 DOI 10.1007/978-1-60761-401-2_22, © Humana Press, a part of Springer Science + Business Media, LLC 2010

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Pathology Journal describing the methodology of enhanced histopathology as well as providing specific examples to illustrate this methodology (6–10). One component of enhanced histopathology is that the separate compartments of each lymphoid organ should be evaluated individually. This specialized type of evaluation has been shown to increase the sensitivity and specificity of immuno-histopathology (11, 12). Each lymphoid organ has separate anatomic compartments that support specific immune functions, thus the rationale for evaluating each compartment separately. Another component of enhanced histopathology involves the reporting of any identified tissue changes in a semiquantitative descriptive, rather than an interpretive fashion. Semiquantitative descriptive terms include “reduced numbers of lymphocytes” and “increased numbers of lymphocytes” whereas interpretive terms include “atrophy” and “hyperplasia.” Reporting tissue changes in descriptive terms avoids the misinterpretation of tissue changes, recognizing that cell trafficking and recirculation rather than an increase or decrease in resident cell populations may account for the changes in cell numbers. Any identified changes should be put into the appropriate toxicologic and pathologic perspective. The final interpretations and conclusions should be discussed within the pathology narrative.

2. Basic Approach to Enhanced Histopathology 2.1. Required Information

Before microscopic evaluation of the immune system is performed, information on dosing regimen, clinical signs, organ weights (spleen and thymus), body weights, hematology and clinical chemistry parameters and any gross lesions should be reviewed. Organ weights should be interpreted in the context of all other clinical, histopathology and clinical pathology data from the study. The age, nutritional status, spontaneous lesions, steroid hormone status, stress and overall health for each strain and group of animals should be considered when interpreting microscopic changes in lymphoid organs (13–15). Various species, such as rats, mice, rabbits, guinea pigs, hamsters, dogs, and non-human primates may be used to address the immunotoxic potential of xenobiotics and there are numerous and significant species differences that should be considered. Understanding these species differences would allow selection of the appropriate species for immunological analysis and welldefined differences may be used to delineate immunologic mechanisms. The various species differences in the structure and function of the immune system and identification of those differences that may be important in the conduct of immunotoxicity testing has been reviewed (16).

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2.2. Tissues to Evaluate

The lymphoid tissues that should be evaluated include the thymus, spleen, bone marrow, and lymph nodes and depending on route of application, mucosa associated lymphoid tissues (MALT). Although a standard list of lymph nodes may be routinely evaluated, the most proximal regional lymph nodes draining the site of xenobiotic application should always be included on that list. Sainte-Marie et al. (17) and Tilney (18) provide detailed descriptions of lymphatic drainage patterns in the rat and the lymph nodes involved. For orally administered compounds, cervical and mesenteric lymph nodes and Peyer’s patches (gut associated lymphoid tissue, GALT) should be examined. The nasal associated lymphoid tissue (NALT), bronchial associated lymphoid tissue (BALT) and bronchial lymph nodes should be examined for inhalation studies. Peripheral lymph nodes (i.e., popliteal, axillary, etc.) that do not drain the site of application may be useful for evaluating the systemic, rather than regional, immunomodulatory effects of compound administration. However, it should be noted that there might be variability of normal histology in these nodes due to collection, sectioning and embedding techniques.

2.3. Determining Range of Normal

Evaluation of concurrent control tissues is crucial in order to establish the range of normal tissue changes for a particular group of animals. However, it is important to understand that for some lymphoid organs, such as those exposed to dietary antigens (submandibular lymph node, mesenteric lymph node, Peyer’s patches), the range of normal appearance may be broad. Therefore, for each group of lymphoid organs in control animals, it is prudent to determine what morphologic spectrum of tissue changes will be considered “within normal limits.” There may be an occasional animal within a group of control animals that has tissue changes considered outside the range of normal. In this case, it is recommended that the same grading criteria applied to the test animals also be applied to these control animals. Criteria used to define “normal” may be detailed in the Materials and Methods section or in the pathology narrative.

2.4. Preventing Diagnostic Drift

To prevent diagnostic drift it is best to evaluate one group of lymphoid organs at a time. As an example, one would evaluate all the thymuses from the negative control group first, establish the “range of normal” for that group of animals, and consider if any of the control animals fall outside of this range. Then choose a few slides that are representative of the normal range and keep them available as reference slides. The thymuses from the positive control group would be evaluated next and graded with labspecific established grading criteria. The next step would be to evaluate the thymuses from the high dose group, recording any changes considered outside the range of normal when compared

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to negative controls. The representative control slides may be referred to when necessary to prevent diagnostic drift. The thymuses from the remaining test groups would be evaluated next and graded accordingly. Once all the thymuses have been evaluated, the next lymphoid organ would be evaluated in a similar fashion. 2.5. When to Use Blind Scoring

The need for determining the “range of normal” within the control tissues before evaluating tissues from treatment groups indicates that “blind evaluation” or “blind scoring” cannot be done for the first pathology evaluation. However, evaluating a group of tissues without knowledge of treatment group or dosing regimen as a way to identify subtle tissue changes between groups may be helpful after the initial read has been performed.

2.6. Knowledge of Normal Structure, Function and Histology

Lastly, it is critical to understand the normal structure, function and histology of each lymphoid organ and its potential interactions with other organs before performing enhanced histopathology evaluation. A series of articles describing the normal structure and histology of lymphoid organs has been published as a reference guide for pathologists and is available in A Monograph on Histomorphologic Evaluation of Lymphoid Organs (19–23). A thorough knowledge of normal cell populations and cell trafficking is required in order to identify cellular and structural abnormalities. It is also important to understand how lymphoid interact with one another and how the immune system interacts with other organ systems. Changes within one lymphoid organ may be reflected in another lymphoid organ. For example, changes within the bone marrow may be reflected in the thymus since it requires bone marrow progenitor cells. Moreover, changes within the immune system may be reflected in other organ systems and vice versa. For example, destruction of red blood cells in the peripheral blood may result in increased numbers of erythroid progenitor cells in the bone marrow and increased extramedullary hematopoiesis (EMH) in the red pulp of the spleen. Therefore, any observed change within a lymphoid organ should be interpreted in the context of the complex interactions of the different lymphoid organs and/ or other organ systems as well as all available study data such as dosing regimen, body weight, organ weight, clinical pathology, nutritional status, antigen load, age, spontaneous lesions, stress, etc.

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Fig. 22.1. Section of thymus from a control 90-day-old male Sprague–Dawley rat. The thymus is divided into a morphologically distinct cortex (C) and medulla (M) separated by a vascular corticomedullary zone. The cortex stains more darkly and contains predominately densely packed, small immature T lymphocytes amid a sparse epithelial cell population. Compared to the cortex, the medulla is paler staining with fewer T lymphocytes and a more robust epithelial cell population.

3. How to Evaluate Lymphoid Organs Using Enhanced Histopathology 3.1. Enhanced Histopathology of the Thymus

The thymus contains two structurally and functionally distinct compartments. First there is the cortex, which contains predominately mature double positive (CD4+CD8+) T cells, and secondly the medulla, which contains predominately single positive (CD4+CD8- and CD4-CD8+) T cells (Fig. 22.1). Enhanced histopathology of the thymus involves the separate evaluation of the size and cellularity of these two compartments. Increased numbers of apoptotic lymphocytes can be the result of immunotoxicity and should also be noted (Fig. 22.2). Endogenous glucocorticoid release in response to stress and debilitation can occur within a group of animals and this can also result in increased numbers of apoptotic lymphocytes and tingible body macrophages. However, lymphocytes in the cortex normally undergo numerous cell divisions and negative selection before entering the medulla, so apoptosis is a normal finding in this population of rapidly dividing cells. Therefore, an increase in the number of apoptotic cells should be recorded only after careful comparison with controls. Other cellular changes should also be noted such as necrosis, inflammation, pigment and EMH. An increase or decrease in the cortex:medulla ratio is another parameter that should be determined. However, within each lobule the plane of section results in variation of this ratio when measured at multiple points. Therefore, a subjective qualitative

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Fig. 22.2. Section of thymus from a 90-day-old male Sprague–Dawley rat treated with 1 mg/kg bodyweight dexamethasone 3 h before tissue collection. This section of thymus illustrates a treatment-related increase in cortical apoptotic lymphocytes with scattered tingible body macrophages (arrows) containing engulfed apoptotic bodies. Grading criteria would be lab or study-specific and should be used for each lesion.

assessment can be done from a low to medium microscopic magnification by determining if there is an overall increase or decrease in the cortex:medulla ratio in the section of thymus analyzed when compared to controls. A standardized trimming procedure should be followed to ensure proper tissue orientation and accurate comparisons (21). An increase in the numbers of epithelial cords and tubules within the medulla is another feature to evaluate. The loss of medullary lymphocytes can result in the epithelial component of the medulla appearing more prominent, but not necessarily hyperplastic or hypertrophied. Since prominent and hyperplastic medullary epithelial cells are also a common age-related change found in association with thymus involution, consideration of the animals age and comparison with controls should help to determine if this histological change is test article or age-related. The epithelium-free areas (EFAs) in the thymus are lymphocyte-rich regions, devoid of stromal elements, non-vascularized, and with unknown function (Fig. 22.3). The occurrence and extent of EFAs varies between rat strains. It is postulated that they may be lymphocyte reservoirs (24) or proliferation sites of lymphocytes (25, 26). EFAs are located in the subcapsular region and serial sections show that they run from the subcapsular area to deep in the cortex, often bordering the medulla (27). To thoroughly evaluate this compartment of the thymus, special stains would be useful (CD4/CD8+, keratin-, laminin-). Therefore, the evaluation of EFAs should be performed during the initial hematoxylin and eosin (H&E) screen and evaluated with special stains if

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Fig. 22.3. Section of thymus from a control Wistar rat. The epithelium free areas (EFAs) (arrows) in the thymus are lightly encapsulated, lymphocyte-rich areas that run from the subcapsular region to deep in the cortex, occasionally bordering the medullary areas. These structures are found in the rat and are considered to be strain-dependent. Due to the lack of stromal elements, special stains such as keratin and laminin, may be helpful to more closely evaluate these regions. Reprinted from Toxicol. Pathol. Ref. 8 with permission from Sage Publications.

the initial screen indicated that this compartment may have been affected. A checklist for the changes that can be observed in the thymus for enhanced histopathology is given in Table 22.1. This table is intended to be an example of a guideline that the pathologist can use during histological evaluation. 3.2. Enhanced Histopathology of the Spleen

Compartments to evaluate in the spleen are the hematogenous red pulp and the lymphoid white pulp. The white pulp is further separated into the periarteriolar lymphoid sheaths (PALS, T cellrich areas), lymphoid follicles (B cell-rich areas) and marginal zones (B cell and macrophage-rich areas) (Fig. 22.4). These compartments should be evaluated separately for changes in size and cellularity. The presence, severity grade and location of plasma cells, apoptotic cells, tingible body macrophages, pigmented macrophages, granulocytes and hematopoietic cells should be noted (Fig. 22.5). A careful evaluation of the amount of EMH should be noted in the red pulp so that subtle lesions can be detected. Other lesions such as granulomas, macrophage aggregates, fibrosis and necrosis should also be diagnosed during the evaluation. An example of a checklist for the changes to be noted in the spleen for enhanced histopathology is given in Table 22.2. This table is intended to be an example of a guideline that the pathologist can use during histological evaluation.

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Table 22.1 Thymus Enhanced Histopathology Checklist Chemical/Animal # Yes/Severity grade

No

Cortex Increased/Decreased size Increased/Decreased number of lymphocytes Increased numbers of apoptotic cells Increased numbers of tingible body macrophages Necrosis Medulla Increased/Decreased size Increased/Decreased number of lymphocytes Increased numbers of apoptotic cells Increased numbers of tingible body macrophages Increased numbers of Hassall’s corpuscles Necrosis Prominent epithelial cords and tubules Cortex/medulla ratio Increased/Decreased Epithelium free areas (EFAs): rats only Not evaluated Not present Increased/Decreased size Increased/Decreased number of lymphocytes Increased numbers of apoptotic cells Increased numbers of tingible body macrophages Necrosis Other (give location) Inflammation Cysts Pigment Extramedullary hematopoiesis (EMH) Other Comments

The red pulp contains macrophages and white blood cells including lymphocytes, but it is predominately composed of red blood cells. For this reason, there can be variation in splenic size and weight, as well as histological variation in red pulp size and erythrocyte cellularity depending upon the method of euthanasia and the efficacy of exsanguination at necropsy. The potential variations in splenic weight are more likely to be encountered in dogs and non-human primates than in rodents due to its storage function in these species (28). Changes in spleen weight that are dose-related and confirmed with histopathological evaluation may be considered toxicologically significant.

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Fig. 22.4. Section of spleen from a control 90-day-old male Sprague–Dawley rat. The white pulp of the spleen is subdivided into the T cell-rich periarteriolar sheath (PALS), the B cell-rich follicle (F) and the B cell and macrophage-rich marginal zone (MZ). The arrow indicates a cross section through the central artery.

Fig. 22.5. Section of spleen from a 90-day-old male Sprague–Dawley rat treated with 1 mg/kg bodyweight dexamethasone 6 h before tissue collection. Treatment-related lesions include increased lymphocyte apoptosis in the PALS region and marginal zone as well as a decrease in the follicle area. Grading criteria would be lab or study-specific and should be used for each lesion.

3.3. Enhanced Histopathology of the Lymph Nodes

The lymph node consists of multiple lymphoid lobules, each containing three major functional compartments that support specific immune functions. These are (1) the cortical area composed of predominately B cell lymphoid follicles and interfollicular region (2) the T cell-rich paracortical area (also called deep

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Table 22.2 Spleen Enhanced Histopathology Checklist Chemical/Animal # Yes/Severity grade

No

Periarteriolar lymphoid sheath Increased/decreased size Increased/decreased number Increased/decreased lymphocytes Marginal zone Increased/decreased size Increased/decreased lymphocytes Increased/decreased macrophages Follicles Increased/decreased numbers Increased/decreased lymphocytes Increased/decreased germinal centers Increased apoptosis Increased Tingible body macrophages Red pulp Increased/decreased size Increased/decreased lymphocytes Increased/decreased hematopoietic cells Increased numbers Plasma cells Apoptotic cells Tingible body macrophages Pigmented macrophages Dendritic cells Granulocytes/mast cells Granuloma/macrophage aggregates

Location

Fibrosis Necrosis Other Comments

c ortical unit) and (3) the medulla with sinusoids and medullary cords composed of predominately plasma cells and macrophages (Fig. 22.6). Each of these compartments should be evaluated separately for changes in area, cell type and cell density. The presence, location, and severity grade of apoptotic cells, tingible body macrophages, necrosis, pigmented macrophages, granulocytes, granuloma/macrophage aggregates, prominent high endothelial venules (HEV), erythrocyte rosette formation, etc. should be noted. The various sinuses (subcapsular, transverse, paracortical and medullary), although not lymphoid compartments, are

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Fig. 22.6. Section of mesenteric lymph node from a control 90-day-old male Sprague– Dawley rat. The lymphoid lobule is the basic anatomical and functional unit of the lymph node. The three major functional compartments of the lymphoid lobule are (1) the cortical area containing B cell-rich follicles (F) with or without germinal centers (G) and the interfollicular region (2) the T cell-rich paracortex (P) with paracortical sinuses and (3) the medulla (M) with sinusoids and medullary cords. In this image, the dense cellularity of the medullary cords contrasts sharply against the sparse cellularity of the surrounding medullary sinuses.

important areas of cell trafficking and should also be evaluated for changes in area, cell type and cell density. The checklist presented in Table 22.3 can be used to aid the pathologist in the evaluation of the various lymph node compartments. The plane of section is an important variable to consider when evaluating lymph nodes. The plane of section (transverse, longitudinal or tangential) can affect the relative size of the cortex, paracortex and medulla. It is important that a section be taken through the middle of the longitudinal axis of the lymph node in order to be able to examine all compartments (6). But even with longitudinal sections of smaller nodes, such as the bronchial, axillary or popliteal lymph nodes, only portions of cortex and paracortex may be present (Fig. 22.7). The entire mesenteric lymph node chain should be collected and sectioned longitudinally in order to avoid cross-section variability. In some species such as the rat, multiple slides may be needed in order to evaluate the entire chain. 3.4. Enhanced Histopathology of the MALT

The mucosa associated lymphoid tissues are scattered aggregates of nonencapsulated organized secondary lymphoid tissue within the mucosa that play an important role in local immune responses (23). These lymphoid aggregates are located along the surfaces of all mucosal tissues such as the gut (GALT), nasopharynx (NALT), bronchus (BALT), conjunctiva (CALT), lacrimal duct (LDALT), larynx (LALT) and salivary duct (DALT). However,

Table 22.3 Lymph Node Enhanced Histopathology Checklist (Indicate lymph node location) Chemical/Animal # Yes/Severity grade Cortex Increased/Decreased area Number of follicles Increased/Decreased Germinal center development Increased/Decreased Increased/Decreased numbers Lymphocytes Increased numbers Apoptotic cells Tingible body macrophages Plasma cells Pigmented macrophages Granulocytes (indicate type) Necrosis Granuloma/macrophage aggregates Interfollicular area (note changes) Paracortex Increased/Decreased area Prominent HEV Increased/Decreased numbers Lymphocytes Increased numbers Apoptotic cells Tingible body macrophages Plasma cells Pigmented macrophages Granulocytes (indicate type) Necrosis Granuloma/macrophage aggregates Medullary cords Increased/Decreased area Increased/Decreased numbers Lymphocytes Macrophages Plasma cells Increased numbers Apoptotic cells Tingible body macrophages Pigmented macrophages Granulocytes (indicate type) Necrosis Granuloma/macrophage aggregates Subcapsular/transverse, medullary sinuses Increased numbers Lymphocytes Macrophages Plasma cells Pigmented macrophages Granulocytes (indicate type) Other/Comments

No

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Fig. 22.7. Section of superficial cervical lymph node from a control 90-day-old Sprague– Dawley rat. Only portions of cortex and paracortex may be present in longitudinal sections of smaller lymph nodes, as seen in this example.

the three main regions of MALT that are most frequently evaluated in toxicity studies are the GALT, NALT and BALT (Fig. 22.8a–d). The functional compartments of MALT are the lymphoid follicles, the interfollicular region, the subepithelial dome region, and the overlying follicle-associated epithelium (FAE). The evaluation of MALT for enhanced histopathology would include changes in the number and size of follicles and germinal centers and changes in the size and density of the interfollicular area. Other changes to note would include the presence, severity and location of apoptotic cells, tingible body macrophages, necrosis, plasma cells, granulocytes, pigment and macrophages. An example of a checklist is given in Table 22.4. 3.5. Enhanced Histopathology of the Bone Marrow

The bone marrow is the largest primary lymphoid organ and should be included in the battery of lymphoid tissues examined for enhanced histopathology. While enhanced histopathology involves evaluation of the separate compartments in each lymphoid organ, bone marrow is unique in that it lacks specific anatomic compartments. Evaluation should include an estimate of cellular density and a myeloid/erythroid (M:E) ratio. An increase or decrease in the numbers of megakaryocytes, adipocytes, stromal cells and amount of hemosiderin should also be noted. Other lesions to note include necrosis, hemorrhage, fibrosis, granulomas and neoplasia. An example of a checklist is given in Table 22.5. Although changes in the lymphoid lineage would be the best indicator of immunomodulation, conclusive identification of lymphoid lineage cells is typically not accomplished with H&E evaluation of rodent bone marrow. Lymphoid lineage cells are difficult to

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Fig. 22.8. Sections of mucosa associated lymphoid tissues (MALT) from control 90-day-old Sprague–Dawley rats. The MALTs are scattered aggregates of nonencapsulated organized secondary lymphoid tissues that respond to specific antigens encountered along all mucosal surfaces. (a) Example of a Peyer’s patch from the small intestine, which is one type of gut associated lymphoid tissue (GALT). The general structure of a Peyer’s patch at this magnification includes centrally located follicles flanked by parafollicular or interfollicular regions. The follicles may or may not have germinal centers, depending on degree of dietary antigenic stimulation. (b) Example of two aggregates of nasopharyngeal associated lymphoid tissue (NALT) (arrows) on the lateroventral floor of the proximal nasopharyngeal duct. (c) higher magnification of NALT with a quiescent follicle with subjacent interfollicular region and overlying follicle-associated epithelium (FAE). The FAE is attenuated, contains no goblet cells and fewer cilia relative to the respiratory epithelium. (d) Example of bronchial associated lymphoid tissue (BALT) within the wall of a bronchiole. The distribution of BALT is always between an artery and a primary bronchiole and there is a predilection for branching points. The presence and amount of BALT in rodents is strain and species specific.

distinguish from many of the other nucleated cells in the H&Estained bone marrow. This limits the extent of enhanced histopathology of the bone marrow. However, H&E evaluation of the marrow is an excellent screening tool. When potential changes in bone marrow cellularity are identified in tissue sections, cytology should be performed for a more comprehensive assessment. If the M:E ratio appears to be altered, comparison with a complete blood count may provide useful information for distinguishing which cell

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Table 22.4 MALT Enhanced Histopathology Checklist (specify type: GALT, BALT) Chemical/Animal # Yes/Severity grade

No

Follicles Increased/Decreased size or number Increased/Decreased numbers of lymphocytes Increased/Decreased germinal centers Interfollicular area Increased/Decreased size Increased/Decreased numbers of lymphocytes FAE ulceration Prominent HEV Increased numbers of Apoptotic cells Tingible body macrophages Plasma cells Pigmented macrophages Granulocytes (type)

Give Location

Granuloma/macrophage aggregates Necrosis Other Comments

type is increased or decreased. For a more thorough evaluation, a differential count on a bone marrow smear could be used to determine which cell line was altered and a quantitative assessment of that alteration could be determined. Flow cytometry could also be done, providing quantitative and immunophenotyping information about hematopoietic and lymphopoietic cell populations. Cytology would also be needed to confirm any alteration in the maturation index, which is the ratio between the number of proliferative phase cells to the number of maturation phase cells. Changes in bone marrow cellularity can be an indicator of systemic toxicity. However, the majority of changes in the bone marrow that are observed in toxicity studies are a response to hematological changes or lesions elsewhere in the body. For this reason, a consideration of the overall health of the animal, and all tissue changes in the body, are required in order to differentiate primary (direct toxic effect) from secondary (physiological response) effects on the bone marrow. For example, pyometra or suppurative skin ulcerations may impose an increased demand for neutrophils and result in an increase in the myeloid cell lineage in the bone marrow. Clinical chemistry is another tool that can provide important information regarding alteration of organ function, such as the liver or kidney that may have an effect on bone marrow cellularity.

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Table 22.5 Bone Marrow Enhanced Histopathology Checklist Chemical/Animal# Yes/Severity Grade Increased/Decreased number of cells Myeloid Erythroid Myeloid/Erythroid ratio Megakaryocytes Adipocytes Reticular adventitial cells Macrophages Granulocytes (type)

No

Ratio =

Maturation index abnormal? Myeloid Erythroid Hemosiderin Increased Decreased Necrosis Hemorrhage Fibrosis Granuloma Neoplasia Other Comments

References 1. Ruehl-Fehlert C, Bradley A, George C, Germann PG, Bolliger AP, Schultee A (2005) Harmonization of immunotoxicity guidelines in the ICH process – pathology considerations from the guideline Committee of the European Society of Toxicological Pathology (ESTP). Exp Toxicol Pathol 57:1–5 2. Basketter DA, Bremmer JN, Buckley P, Kammuller ME, Kawabata T, Kimber I, Loveless SE, Magda S, Stringer DA, Vohr HW (1995) Pathology considerations for, and subsequent risk assessment of, chemicals identified as immunosuppressive in routine toxicology. Food Chem Toxicol 33:239–43 3. Schuurman HJ, Kuper CF, Vos JG (1994) Histopathology of the immune system as a tool to assess immunotoxicity. Toxicology 86:187–212 4. Vos JG (1980) Immunotoxicity assessment: screening and function studies. Arch Toxicol Suppl 4:95–108

5. Haley P, Perry R, Ennulat D, Frame S, Johnson C, Lapointe JM, Nyska A, Snyder P, Walker D, Walter G (2005) STP position paper: best practice guideline for the routine pathology evaluation of the immune system. Toxicol Pathol 33:404–7 6. Elmore SA (2006) Enhanced histopathology of the lymph nodes. Toxicol Pathol 34: 634–47 7. Elmore SA (2006) Enhanced histopathology of the spleen. Toxicol Pathol 34:648–55 8. Elmore SA (2006) Enhanced histopathology of the thymus. Toxicol Pathol 34:656–65 9. Elmore SA (2006) Enhanced histopathology of the bone marrow. Toxicol Pathol 34:666–86 10. Elmore SA (2006) Enhanced histopathology of mucosa-associated lymphoid tissue. Toxicol Pathol 34:687–96 11. Harleman JH (2000) Approaches to the identification and recording of findings in the lymphoreticular organs indicative for immunotoxicity in

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regulatory type toxicity studies. Toxicology 142:213–9 Kuper CF, Harleman JH, Richter-Reichelm HB, Vos JG (2000) Histopathologic approaches to detect changes indicative of immunotoxicity. Toxicol Pathol 28:454–66 Gopinath C (1996) Pathology of toxic effects on the immune system. Inflamm Res 45(Suppl):S74–S78 Levin S, Semler D, Ruben Z (1993) Effects of two weeks of feed restriction on some common toxicologic parameters in Sprague– Dawley rats. Toxicol Pathol 21:1–14 Odio M, Brodish A, Ricardo MJ Jr (1987) Effects on immune responses by chronic stress are modulated by aging. Brain Behav Immun 1:204–15 Haley PJ (2003) Species differences in the structure and function of the immune system. Toxicology 188:49–71 Sainte-Marie G, Peng FS, Belisle C (1982) Overall architecture and pattern of lymph flow in the rat lymph node. Am J Anat 164:275–309 Tilney NL (1971) Patterns of lymphatic drainage in the adult laboratory rat. J Anat 109:369–83 Willard-Mack CL (2006) Normal structure, function, and histology of lymph nodes. Toxicol Pathol 34:409–24 Cesta MF (2006) Normal structure, function, and histology of the spleen. Toxicol Pathol 34:455–65

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21. Pearse G (2006) Normal structure, function and histology of the thymus. Toxicol Pathol 34:504–14 22. Travlos GS (2006) Normal structure, function, and histology of the bone marrow. Toxicol Pathol 34:548–65 23. Cesta MF (2006) Normal structure, function, and histology of mucosa-associated lymphoid tissue. Toxicol Pathol 34:599–608 24. Van Ewijk W (1984) Immunohistology of lymphoid and non-lymphoid cells in the thymus in relation to T lymphocyte differentiation. Am J Anat 170:311–30 25. Duijvestijn AM, Sminia T, Kohler YG, Janse EM, Hoefsmit EC (1982) Rat thymus micro-environment: an ultrastructural and functional characterization. Adv Exp Med Biol 149:441–6 26. Godfrey DI, Izon DJ, Tucek CL, Wilson TJ, Boyd RL (1990) The phenotypic heterogeneity of mouse thymic stromal cells. Immunology 70:66–74 27. Bruijntjes JP, Kuper CF, Robinson JE, Schuurman HJ (1993) Epithelium-free area in the thymic cortex of rats. Dev Immunol 3:113–22 28. Greaves P (2007) Hematopoietic and lymphatic systems. In: Greaves P (ed) Histopathology of preclinical toxicity studies: interpretation and relevance in drug safety evaluation, 3rd edn. Elsevier, Amsterdam, pp 116–125

Chapter 23 Immunotoxicity Testing in Nonhuman Primates Stephanie Grote-Wessels, Werner Frings, Clifford A. Smith, and Gerhard F. Weinbauer Abstract Biological relevance is generally the major justification for using nonhuman primates (NHP) during preclinical safety assessment. This holds particularly true for the evaluation of biopharmaceuticals with NHP often being the species of choice. For safety assessment of small molecules, NHP are used in case of a higher degree of metabolic similarity, to detect the highly specific immunotoxic side effects and to discriminate toxicity from efficacy of immunomodulatory drugs. Unlike for rodent immunotoxicity studies, standardized tests and protocols are generally less available for NHP. The immunotoxicity testing protocols described in the present chapter have been adapted for application to NHP samples. In principle, rodent protocols can be transferred to NHP. Fortunately, most of the immunotoxicity parameters delineated in the ICH S8 guideline can be applied to NHP specimens. Exceptions are the host resistance assay and the delayed type hypersensitivity test. Owing to the close structural and physiological similarity between NHP and human, human test kits or reagents are often well suited for application to NHP samples. For data evaluation it should be noted that no inbred strains of NHP are available, resulting in a large inter-animal variability for most immunotoxicity assay results. The experimental protocols and reagents described in this chapter were developed specifically for the cynomolgus monkey (Macaca fascicularis), currently the most commonly used NHP species in toxicology. In many instances, these protocols will also be applicable to rhesus monkeys (M. mulatta) and potentially to other Old World macaques. For the marmoset (Callithrix jacchus), a New World monkey also used in toxicology, the choice of available immunotoxicity testing protocols is much reduced when compared to macaques. Key words: Nonhuman primate, Macaque, Immunotoxicity evaluation, Immunophenotyping, Cytokines, Immunoglobulins, Lymphocyte proliferation, Bone marrow examination, Marmoset, Cynomolgus

1. Introduction Biological relevance is generally the major justification for using nonhuman primates (NHP) during preclinical safety assessment. This holds particularly true for the evaluation of R.R. Dietert (ed.), Immunotoxicity Testing: Methods and Protocols, Methods in Molecular Biology, vol. 598 DOI 10.1007/978-1-60761-401-2_23, © Humana Press, a part of Springer Science + Business Media, LLC 2010

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biopharmaceuticals with NHP being often the species of choice (1). This species-choice is typically dictated by similarities of the pharmacological and pharmacodynamic activity and a lesser likelihood of anti-drug antibody formation (2, 3). For safety assessment of small molecules, NHP are used in case of a higher degree of metabolic similarity, to detect the highly specific immunotoxic side effects and to discriminate toxicity from efficacy of immunomodulatory drugs (4). For the evaluation of reproductive or developmental immunotoxicity, macaques are often the species of choice since there are close physiologic similarities to humans with regard to gonadal functions, pregnancy, and pre- and postnatal development (5). The latter includes but is not limited to the levels of immunoglobulins and maturation of the immune system prior to and after birth (6, 7). General and comprehensive overviews for (immunotoxicity) assessment of (bio)pharmaceuticals have recently been published (8, 9). Unlike the rodent immunotoxicity studies, standardized tests and protocols are generally less available for NHP (e.g. species comparison is provided in reference (10)). The immunotoxicity testing protocols described in the present chapter have been adapted for application to NHP material. In principle, rodent protocols can be transferred to NHP. Fortunately, most of the immunotoxicity parameters delineated in the ICH S8 guideline (11) can be applied to NHP specimens. Exceptions are the host resistance assay and the delayed type hypersensitivity (DTH) test. The host resistance assay is usually more demanding for primates since (1) special facilities are needed to maintain infected animals and separate them from the main colony, (2) large group sizes would be needed to yield meaningful results and (3) using death as an endpoint in an experiment with NHP is obviated by ethical considerations. DTH reactions with protein immunization and challenge have been reported for mice and rats, but NHP DTH reactions are very difficult to reproduce consistently(11). In addition, one should make sure that the DTH response is not mistaken for an antibody and complement-mediated Arthus reaction (11). Intradermal antigen challenge after previous immunization for determination of DTH may not always predict classical DTH, although an extended method with microscopic biopsy evaluations has been described (11–13). In contrast to rodents, few test reagents have been developed for NHP. Owing to the close structural and physiological similarity between NHP and humans, human test kits or reagents are often well suited for application to NHP samples. Nevertheless, a well planned method transfer and/or validation should be performed before using human reagents for NHP. For example, it was shown that the use of the common combination for immunophenotyping of human natural killer cells (CD16 in combination with CD56) is not appropriate for macaques (14). For data

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evaluation, it should be noted that no inbred strains of NHP are available, resulting in large inter-animal variability for most immunotoxicity assay results. This fact, in combination with comparatively smaller group sizes of NHP studies, results in reduced statistical power to detect significant effects based upon group comparisons. It is, therefore, recommended that predose assessment of immunotoxicity parameters be included in order to enable longitudinal evaluation of every individual animal in a NHP toxicity study. The experimental protocols and reagents described below were tested and developed specifically for the cynomolgus monkey (Macaca fascicularis). To date, this is the most commonly used NHP species in toxicology (15). In many instances, these protocols will also be applicable to rhesus monkeys (M. mulatta) and potentially to other Old World macaques. For the marmoset (Callithrix jacchus), a New World monkey also used in toxicology, the choice of available immunotoxicity testing protocols is much reduced when compared to macaques. Reference to protocols applicable to marmoset specimen is made wherever appropriate.

2. Materials 2.1. Immuno phenotyping

1. FACSTM Lysing solution 10× (Becton Dickinson Immuno cytochemistry systems, San Jose, USA), stored at room temperature, ready to use after dilution with ultrapure water (Millipore). Dilution is stable at room temperature for up to 2 weeks. 2. Cell Wash (Becton Dickinson, Heidelberg, Germany). 3. The following antibodies are suitable for blood analysis of cynomolgus monkeys: anti-CD3-FITC (clone SP34), antiCD4-PE (clone M-T477), anti-CD8-APC (clone RPA-T8), anti-CD16-PE (clone 3G8), anti-CD20-APC (clone L27), anti-CD45-PerCP (clone TÜ116), Isotypes IgG3-FITC (clone J606), IgG1-PE (clone MOPC-21), IgG2a-PE (clone G155178), IgG1-APC; (Becton Dickinson, Heidelberg, Germany or Beckman-Coulter, Krefeld, Germany), stored at 5 ± 3°C. 4. The following antibodies are suitable for blood analysis of marmoset monkeys: anti-CD2-PE (39C1.5), anti-CD4-PC7 (SFCI12T4D11), anti-CD8-ECD (clone SFCI21Thy2D3), anti-CD16-PC5 (3G8), anti-CD20-FITC (H299), antiCD56-PC5 (N901), Isotypes IgG1-FITC, IgG2A-PE (all Becton Dickinson, Heidelberg, Germany or Beckman-Coulter, Krefeld, Germany), stored at 5 ± 3°C. 5. 1 mL whole blood in EDTA anticoagulant, stored at room temperature.

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6. For collection of lymphocytes from lymphatic tissue: Medimachine, Cat.No. 340588; Medicons 50 mm, non sterile, Cat.No. 340592; Filcons 50 mm, non sterile, Cat.No. 340603, (all Becton Dickinson, Heidelberg, Germany); Staining buffer (HBSS, 1%BSA, 0.1% NaN3, no special provider recommendations). 2.2. Determination of Cytokines

1. Human Th1/Th2 Cytokine FlowCytomix (BMS710FF, Bender MedSystems GmbH, Vienna, Austria). 2. Assay buffer (1×): dilute stock solution 1:10 with Aq. bidest and store at 5 ± 3°C. 3. Biotin conjugate Mix: mix 600 mL of each biotin-conjugate with up to 6 mL of reagent dilution buffer as is necessary (depending on number of cytokines determined, see test kit protocol). 4. Bead Mix: mix 300 mL of each bead with up to 3 mL of reagent dilution buffer. 5. Reconstitute Th1/Th2 standard with distilled water according to the manufacturer`s instructions. 10 mL of each reconstituted standard is added to a vial and filled up to the final volume of 200 mL with assay buffer (1×). A serial dilution is prepared of the standard mixture (assay buffer: standard mixture = 2:1, seven serial dilutions) 6. Streptavidin–PE solution: For 96 reactions, mix 176 mL of stock solution with 5,324 mL assay buffer. 7. Test serum or plasma.

2.3. Functional Natural Killer (NK) Cell Testing

1. NK cell Testkit (Orpegen Pharma, Heidelberg, Germany) containing target cells, interleukin-2, complete medium, DNA-staining solution. 2. Isopaque-Ficoll tubes: Histopaque-1077, Sigma Aldrich, Taufkirchen, Germany. 3. Sterile phosphate buffered saline (#70011-036, Invitrogen, Carlsbad, CA, USA) single-use tubes (#352052, Becton Dickinson, Heidelberg, Germany). 4. 5 mL of monkey blood in heparin anticoagulant.

2.4. T-Cell Dependent Antibody Response

1. KLH = keyhole limpet haemocyanin (#H-7017, Sigma Aldrich, Taufkirchen, Germany). 2. Coating buffer: Borate buffered saline, pH 8.4, 0.1 M boric acid; prepare 75 mM NaCl with MilliQ-water, stored at 5 ± 3°C. 3. Tween-20, 10% in A. bidest, stored at 5 ± 3°C. 4. Wash buffer: Borate buffered saline with 0.01% Tween-20, pH 7.8, store at 5 ± 3°C.

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5. Blocking buffer: Borate buffered saline with 3% BSA (Sigma A7888) pH 8.3–8.5, stored at 5 ± 3°C. 6. Assay diluent buffer: wash buffer with 1% BSA, pH 8.3–8.5, stored at 5 ± 3°C 7. KLH coating solution: reconstitute KLH according to manufacturing instructions; aliquots can be stored at −20 ± 4°C for at least 3 months. Coat polypropylene tubes with blocking buffer. Dilute KLH stock solution to 5 mg/mL with coating buffer. Use 100 mL KLH coating solution/well to coat the ELISA plate. 8. Cynomolgus monkey IgG solution: reconstitute IgG (Cappel # 55937, MP Biomedicals, Solon, OH, USA) according to manufacturer’ instructions and store aliquots at −80 ± 4°C. Coat polypropylene tubes with blocking buffer. Dilute IgG stock solution to 1 mg/mL with coating buffer. Use 100 mL IgG coating solution/well to coat ELISA plates. 9. Rabbit-anti-cynomolgus monkey IgG–HRP conjugate: reconstitute IgG–HRP conjugate (Cappel # 55433, MP Biomedicals, Solon, OH, USA) according to the manufacturer’s instructions; aliquots can be stored at −20 ± 4°C for at least 12 months. Test the dilution of IgG-HRP conjugate stock solution for each preparation to yield an optical density of 0.4–1.5. Use 100 mL/well to coat the ELISA plates. 10. TMB substrate (#T-0440, Sigma Aldrich, Taufkirchen, Germany), storage at 5 ± 3°C. 11. Stop solution: Phosphoric acid, 1 M in distilled water. 2.5. Bone Marrow Examination

1. Bovine serum albumin or foetal bovine serum (1–10%). 2. Methanol. 3. Modified Wright’s stain. 4. Immersion oil. 5. Standard light microscope; ×10 eye pieces; ×20, ×50 and ×100 objective lenses.

2.6. Immunhisto chemical Staining of Tissue

1. The following antibodies are suitable for staining cynomolgus monkeys tissues: anti-CD3 (clone A0452), anti-CD20 (clone L26), anti-CD57 (clone TB01, all Dako, Hamburg, Germany), anti-CD2 (clone 2CO2) and anti-CD4 (clone 4B12, Medac, Wedel, Germany), anti-CD8 (clone BC/1A5, Biocarta, San Diego, USA) (5, 16). 2. Pre-treatment module buffer 1 (citrate buffer, Medac, Wedel, Germany). 3. TBS-Buffer (tris buffered saline, #T5912, Sigma Aldrich, Germany).

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4. Hydrogen peroxide block reagent (#TA-125-HP, Medac, Wedel, Germany). 5. EnVision + DualLink reagent (#K4061, Dako, Germany). 6. DAB (Diaminobenzidine) Plus Substrat (#TA-004-HDX, Medac, Wedel, Germany). 7. Mayers Hämalaun. 8. 99% Ethanol. 9. XEM-200 (Vogel GmbH, Gießen, Germany) and CV Mount (Leica, Wetzlar, Germany). 2.7. Lymphocyte Proliferation Assay

1. Blood samples with heparin. 2. Dulbecco’s modified Eagle’s medium (DMEM, high glucose (4.5 g/L) with Penicillin and Streptomycin 1×, (PAA, Pasching, Austria); Glutamine 1×,; non-essential amino acids 1×, (Invitrogen, Carlsbad, California) and 5 × 105 M 2-mercapto-ethanol, Sigma Aldrich, Taufkirchen, Germany). 3. Ficoll-Paque Plus solution (Amersham, Uppsala, Sweden). 4. Human AB serum (PAA, Pasching, Austria). 5. PHA stock solution (Phytohaemagglutinin from Phaseolus vulgaris, 1 mg/mL, Sigma Aldrich, Taufkirchen, Germany). 6. IL-2 stock solution (Proleukin®, 10,000 U/mL, Chiron, Ratingen, Germany). 7. 6-Methyl-3H-thymidine solution 20 Ci/mmol (740 GBq/ mmol; Hartmann Analytic, Braunschweig, Germany). 8. EDTA solution: 0.547 M NaCl, 0.0134 M KCl, 0.031 M Na2HPO4, 0.0059 M KH2PO4, 0.0107 M EDTA, 0.0323 M NaHCO3, 0.001% phenol red. 9. Scintillation cocktail Rotiszint (Roth, Karlsruhe, Germany).

3. Methods 3.1. Immuno phenotyping

Immunophenotyping by flow cytometry is a useful technique for the analysis of heterogeneous leukocyte populations in peripheral blood that can also be applied to the cell suspensions that have been derived from bone marrow or other tissues. The cells are incubated with fluorochrome-labelled antibodies that are specific for cell surface antigens (e.g. CD4 for T-helper cells). When analyzed on a flow cytometer, cells that have bound the labelled antibody can be quantified on the basis of their fluorescence emission. Simultaneously, cell populations are distinguished from each other based on size and granularity using light scatter properties. Anti-human antibodies show a high degree of cross-reactivity with cynomolgus monkey cells, whereas for marmosets crossreactivity might be more frequently assay-limiting (see Note 1).

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1. Setup antibody mixture for isotype controls with 20 mL of each antibody: anti-IgG3-FITC, anti-IgG1-PE, anti-IgG1APC, anti-IgG2-PE, anti-CD45-PerCP. 2. Setup T-cell antibody mixture with 20 mL of each antibody: anti-CD3-FITC, anti-CD4-PE, anti-CD45-PerCP, antiCD8-APC. 3. Setup NK-/B-cell antibody mixture with 20 mL of each antibody: anti-CD3-FITC, anti-CD16-PE, anti-CD45-PerCP, and 5 mL of anti-CD20-APC.

3.1.2. Preparation of Samples

1. For each blood sample, three vials are needed: Vial 1 with 100 mL isotype antibody mixture, vial 2 with 80 mL T-cell antibody mixture and vial 3 with 65 mL NK-/B-cell antibody mixture. 2. Invert the tube with EDTA whole blood 2–3 times and add 100 mL blood/vial. 3. Incubate for 15 min in the dark at room temperature. 4. Add 2 mL FACS lysing solution/vial. 5. Incubate for 10 min in the dark at room temperature. 6. Centrifuge samples at 300–500 × g for 5 min at room temperature. 7. Decant and vortex the supernatant. 8. Add 2 mL cell wash/vial. 9. Centrifuge samples at 300–500 × g for 5 min at room temperature. Decant the supernatant and add 500 mL of cell wash.

B & W IN PRINT

10. Analyze samples using the flow cytometer (see Fig. 23.1).

Fig. 23.1. Typical plots for cynomolgus monkey blood cells. Frequently, distinction by forward scatter (FSC) and sideward scatter (SSC) alone (left plot, proposed lymphocyte population gated) do not yield clear discrimination of cell populations. Additional gating by CD45 vs. SSC allows for good discrimination of lymphocytes from monocytes (right plot).

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3.1.3. Preparation of Cells from Lymphatic Tissue

1. Fill a 50 mm medicon (Becton Dickinson, Heidelberg, Germany) with 1 mL PBS and 0.9 g of tissue. 2. Fix medicon in Medimachine and run for 2 min. 3. Filter the meshed solution from the medicon through a 50 mm filcon. 4. Flush the medicon with 1 mL of PBS and filter the solution as above. 5. Filter the whole solution through another 50 mm filcon. 6. Dilute the cells 1:10 in staining buffer. 7. Use 0.1 mL for staining as above.

3.2. Determination of Cytokines

For cytokine determination in serum of cynomolgus monkeys a commercially available multiplex assay can be used. Fluorescent polystyrol beads are coupled with antibodies specific to the cytokines to be detected. A mix of coupled beads is incubated with the serum to be tested and cytokines in the sample bind to the antibodies coupled to the beads. A biotin conjugated antibody mix is added, which binds to the cytokines bound to the capture antibodies and Streptavidin–Phycoerythrin (PE) is added which binds to the biotin conjugates. Beads are differentiated by their sizes and distinct spectral signature by flow cytometry which enables the calculation of cytokine concentration. 1. Mix a 25 mL sample with 25 mL Beadmix. 2. Add 50 mL of biotin-conjugate, protect from light with an aluminium foil. 3. Incubate at room temperature (18–25°C) for 2 h on a shaker at 500 rpm. 4. Prepare Streptavidin-PE solution. 5. Add 1 mL assay buffer (1×), and centrifuge samples at 200 × g for 5 min at room temperature. Repeat this step one more time. 6. Add 50 mL of Streptavidin–PE solution to each vial, protect from light with aluminium foil. 7. Incubate at room temperature (18–25°C) for 1 h on a shaker at 500 rpm. 8. Add 1 mL assay buffer (1×). 9. Centrifuge the samples at 200 × g for 5 min at room temperature. Repeat steps 8 and 9 one more time. 10. Add 500 mL assay buffer (1×) to each vial. 11. Flow cytometer setup: adjust voltage of FL-2 so that the bead populations are positioned in the left part of the dot plot. This ensures that the bead populations of standard 1 will be visible on the screen. Define the number of events

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Table 23.1 Final flow cytometer setup for Beckman Coulter FC500 Detectors

Gain

FSC

200–400

20

SSC

200–400

5–10

FL1

200–400

1

FL2

500

1

FL3

250

1

FL4

580

1

FL5

250

1

Compensation

FL1

FL2

FL1

–

–

FL2

–

–

FL3

–

–

FL4

–

10

so that 300 events per analyte are measured. A final setup (e.g., for Beckman Coulter FC500) may look like that shown in Table 23.1. 12. Adjust parameters of FL-2 so that the bead population with the highest PE (FL-2) signal remains close to the right axis (in order to guarantee that bead populations with low PE/ low concentrated analytes are detectable) while the whole population is visible. 13. Analyze samples starting with the standard curve (S1–S7 and blank), followed by the samples. 3.3. Functional NK Cell Testing

This method allows the quantitative determination of the cytotoxic activity of NK cells. The systems comprise: cryopreserved prestained human K562 target cells, complete medium and a DNA-staining reagent. The K562 target cells are labelled with a lipophilic green fluorescent membrane dye in order to discriminate the effector and target cells. After the incubation period in the cytotoxicity assay, killed target cells are identified by a DNAstain, which penetrates the dead cells and specifically stains the nuclei. In this manner, the percentage of target cells killed by effector NK cells can be determined (see Note 2). 1. Prepare mononuclear cells by Ficoll-Hypaque separation. 2. Wash with PBS (1×).

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3. Resuspend in complete medium. 4. Thaw one vial of cryopreserved target cells by rapid agitation in a 37°C water bath. 5. Transfer the cell suspension to the 50 mL centrifuge tube containing 50 mL of complete medium. 6. Centrifuge cells at 120 × g for 5 min at room temperature. 7. Resuspend in 1 mL of complete medium. 8. Mix mononuclear cells with K562 target cells at the desired ratios ranging from 50:1 to 12.5:1. 9. Incubate in a final volume of 100 mL. 10. Add 30 mL Interleukin-2 (200 U/mL) to the selected cell aliquots (“high control” samples) and vortex. 11. Centrifuge the tubes for 2–3 min at 120 × g. 12. Incubate the tubes for 120 min or 240 min in a humidified CO2 incubator. At the end of the incubation, place the tubes on ice until flow cytometric analysis. 13. Add 50 mL DNA staining solution per tube. 14. Vortex and incubate for 5 min at 0°C (light protected in ice bath). Measure the cell suspensions within 30 min. 15. Analyze cells on a flow cytometer using the blue-green excitation light (488 nm argon-ion laser, e.g. FACScan, LYSIS II software or FACScalibur, CELLQuest software). During data acquisition, set a “live” gate in the FL1 histogram on the green fluorescent target cells in order to discriminate effector and target cells. Use a target control tube for this purpose. Collect at least 2,500 events per sample (see Figs. 23.2 and 23.3). 3.4. T-Cell Dependent Antibody Reaction (TDAR)

A T-cell dependent antibody response requires functional antigen presenting cells, T-helper cells, and antibody-producing B-cells and is, therefore, an appropriate screening for an alteration of large parts of the immune system. Test animals are immunized with a new antigen, and blood samples taken thereafter are tested for antibodies (both IgM and IgG for cynomolgus monkeys) against this antigen by ELISA or other techniques. For marmosets, determination of specific IgM is not possible due to a lack of a specific detection antibody. When the antigen is given several weeks before the start of the test item dosing and memory responses can be tested by an antigenic challenge during dosing.

3.4.1. Immunization of Cynomolgus Monkeys/ Marmoset Monkeys

Immunize cynomolgus monkeys with 100 mg KLH plus Freund’s incomplete adjuvant (i.d. or s.c.) or tetanus toxoid (TT, Infanrix® i.m., dose as for human use). Immunize marmoset monkeys with 50 mg KLH plus Freund´s incomplete adjuvant (i.d. or s.c.) or tetanus toxoid (TT, Infanrix® i.m., 50% of dose as for human use).

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specific activity (%)

Fig. 23.2. Typical plots for determination of NK cell function. Left plot: The slightly larger target cells are gated. Right plot : Gated cells are transferred, and green (FL1) fluorescent cells are investigated (FL1 low cells represent gated effector cells). The cells in the upper right quadrant are dead target cells which have taken up propidium iodide.

mean - IL2

40

mean + IL2

20 0 0h

4h

24 h

Timepoint

48 h

Fig. 23.3. NK cell activity before and after intravenous application of 0.25 mg/kg dexamethasone to three cynomolgus monkeys (mean ± SD). In vitro stimulation generally results in a higher NK cell activity, except shortly after in vivo suppression when the killing activity is completely blocked.

3.4.2. Determination of Anti-KLH-IgG/IgM in Serum of Cynomolgus/ Marmoset Monkeys

1. Coat the plates with 100 mL coating buffer/well (four plates per well monkey IgG containing as positive controls) and block with 275 mL blocking buffer/well. 2. Prepare a dilution of samples and control samples with assay diluent (ratio 1:50). 3. Dilute all samples again 1:27 with assay diluent. 4. Cover the plates and shake at room temperature for approximately 3 h. 5. Wash the wells with washing buffer for four times. 6. Add 100 mL rabbit anti-cyno/marmoset IgG-HRP or rabbit anti-cyno IgM-HRP. 7. Cover the plates and shake at room temperature for approximately 2 h.

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8. Wash the wells with washing buffer for four times. 9. Add 100 mL TMB per well. 10. Cover the plates and shake at room temperature for approximately 30 min. 11. Add 100 mL stop solution. 12. Analyze the samples on an ELISA reader. 13. Calculate titres using the following formula: Units/mL (titre): [(OD sample – OD blank)/(OD positive control – OD blank)] × dilution (see Fig. 23.4 and Note 3). 3.5. Bone Marrow Examination

Bone marrow is not only the site of haematopoietic cell production in normal, healthy adult mammals, but is a primary or central lymphoid organ of the immune system. Immunotoxicity studies should include examination of a bone marrow smear, which provides a direct visual assessment of cellular morphology and a good overview of activity of the tissue from which cells of the haematopoietic and immune systems are produced. Bone marrow examination is a sensitive tool to detect an impaired cell replication and/or development. Samples are usually collected from sternum, femur, iliac crest or humerus at post mortem, and are normally obtained by removing the complete bone, opened by splitting or cutting, and extracting a small quantity of marrow with forceps.

3.5.1. Preparation of Smears

1. Since the marrow of the most healthy animals is viscous and in order to spread successfully in the manner of a blood smear mix the marrow in approximately equal proportions with autologous serum or plasma, bovine serum albumin or foetal bovine serum. 25000

Titer

20000 15000 IgM

10000

IgG

5000 0 pre-KLH d8

d11

d15

d22

d25

d29

Immunization day Fig. 23.4. A typical development of anti-KLH antibodies in cynomolgus monkeys. In this case one immunization was performed on day 1 and a booster immunization was given on day 15. One week after immunization higher titers for IgM than for IgG are observed. The booster vaccination induces a strong elevation of IgG titers, but no further elevation of IgM titers.

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3.5.2. Staining of Smears

Fix the smears in methanol for 20 min before staining with a Romanowsky dye, such as modified Wright’s stain, for approximately 10 min.

3.5.3. Microscopical Examination

1. Make an initial examination of a stained smear at a magnification of approximately ×20 to enable an assessment of cellularity, and to gauge the adequacy of megakaryocyte numbers. Unusual features such as agglutination, rosetting, excessive rouleaux of mature cells, and infiltrations of malignant cells can also be identified at this stage. 2. Identify suitable areas for closer examination (×50 and ×100, see Fig. 23.5). On a spread preparation a monolayer of evenly spread, separated and easily identified nucleated cells will generally be in areas trailing behind the marrow fragments and will consist of cells released from those fragments. Choose a variety of areas across the smear for examination to even out the concentrations of specific cell types such as erythroblast islands, and to take account of any possible uneven distribution of the various cell types: It may be possible to identify areas of potential blood contamination at this time. 3. Perform three levels of analysis: Visual appraisal, M:E ratio, or a full differential (myelogram). 4. Visual appraisal: Include an assessment of cellularity, distri bution, and morphology in the examination. Examine smears for:

Fig. 23.5. Photograph of cynomolgus monkey whole bone marrow cells. Modified Wright staining, magnification × 100.

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Micronuclei.

●●

Abnormal cell replication (mitosis).

●●

●●

bnormal cells e.g. lymphoblast’s, hairy cells, histiocytes, A foamy macrophages.

●●

Abnormal inclusion bodies e.g. Aűer rods.

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Platelet budding.

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Normal cell maturation.

●●

Parasites.

●●

●●

●●

●●

3.6. Immunohisto chemical Staining

n approximate normal numerical relationship between A erythroid and myeloid compartments.

ecognize the unusual findings in haematopoietic cell lines R and other (plasma cell, megakaryocyte, macrophage, etc) cell lines, morphology, maturation, or the presence of abnormal cells e.g., lymphoblasts, carcinoma cells, etc, must be noted. or cells normally found in the bone marrow which are not F a part of the haemopoeitic system (e.g., osteoclasts and osteoblasts, contaminating epithelial cells and so on), describe only if abnormal. :E ratio: Classify cells according to their morphology into M erythroid, myeloid, and ‘other’ cell types: A total of at least 200 cells must be counted. Divide the myeloid cell total (M) by the erythroid cell total (E) to obtain the M:E ratio. For most primate species this is around 1.0 (equal numbers of developing myeloid cells to developing erythroid cells), although `normal´ values should be determined for each laboratory. yelogram: Classify cells according to their morphology M into erythroid, myeloid, and “other” cell types: Erythroid and myeloid cells are further classified according to their stage of maturation. It is important at this point to recognize the differences in cell morphology, the appearance of unusual cell patterns or abnormalities which could indicate bone marrow toxicity. 500 nucleated cells should be evaluated to provide increased confidence in the results (see Note 4).

Use antibodies directed against cell specific antigens for immunohistochemical staining of tissue sections. Evaluate the tissue distribution of these antigens among the sections. Using this approach and the specificity of the antigens, different cell types can be identified by their morphological or immunohistochemical properties (see Note 5). Staining of paraffin embedded spleen tissue with an anti-CD3 antibody

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1. Collect sections on superfrost slides (Merck, Braunschweig, Germany) and fix in 4% paraformaldehyde. 2. Incubate sections in pre-treatment module buffer 1 at 98°C for 35 min. 3. Rinse with TBS-buffer for several times. 4. Incubate for 30 min at room temperature in diluted antiCD3 antibody (1:75 in TBS buffer). 5. Rinse with TBS buffer for several times. 6. Quench endogeneous peroxidase activity by incubation in Hydrogen Peroxide Block reagent for 15 min the rinse with TBS buffer several times. 7. Incubate in EnVision + DualLink reagent for 30 min. 8. Rinse with TBS buffer several times. 9. Add DAB plus substrate for 10 min then rinse the sections with distilled water several times. 10. Perform counterstaining with Mayers Hämalaun for approximately 4 s. 11. Clear with distilled waterand rinse with tap water for 5 min. 12. Dehydrate by concentrations.

using

sequentially

increasing

ethanol

13. Incubate sections for 2 × 5 min in XEM-200. 14. Mount slides with coverslips using CV Mount. 3.7. Lymphocyte Proliferation Assay 3.7.1. Preparation of PBMCs from Blood Samples

1. Dilute blood samples with an equal volume of PBS. 2. Underlay blood/PBS mixtures with 3–5 mL Ficoll-Paque Plus solution (20°C). 3. Centrifuge underlayered cells at 1,000 × g and 20°C for 20 min. 4. Transfer the interphase containing the PBMCs into a new tube. 5. After washing with HBSS, resuspend the cells in medium (DMEM containing 10% human AB serum). 6. Adjust PBMC cell titres to 1 × 106 cells/mL with medium.

3.7.2. Stimulation of PBMCs with Mitogens

1. Working concentrations of mitogens: PHA: 20 mg/mL, 4 mg/ mL, 0.8 mg/mL, IL-2:100 U/mL, 20 U/mL, 4 U/mL. 2. Dispense PBMCs on the 96 well plates containing mitogens (1 × 105 cell per well). 3. Shake the plates by using a microplate shaker. 4. Centrifuge at 80 × g and 20°C for 10 min. 5. Incubate the cell culture plates for 72 h at 37°C.

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3.7.3. Measurement of Proliferative Response

1. Dilute the 3H-thymidine solution to a concentration of 8 mCi/ mL (296 kBq/mL). 2. Add 50 mL of the 3H-thymidine solution to each microculture. 3. Subsequently, incubate the culture plates for 8 h at 37°C. 4. After the pulse, discard 100 mL of the supernatants and replace it with EDTA solution. 5. Incubate the plates for an additional 10 min at room temperature. 6. Harvest the cells and transfer them onto glass fibre filters. 7. After 2 h of air drying, transfer the filters into scintillation vials and cover with 2 mL scintillation cocktail. 8. Finally, measure the incorporated radioactivity using a scintillation counter (see Note 6).

4. Notes 1. Immunophenotyping: All anti-CD antibodies should be validated before use in GLP-studies. Since anti-human antibodies are used for monkey samples (see Fig. 23.6), cross-reactivity and plausibility of staining (e.g. co-staining with CD4 if the 80 70

% of Lymphocytes

60

human♂, n =10 human♀, n =10

50

cyno♂, n =75 cyno♀, n =75

40 30 20 10 0 CD3+

CD4+

CD8+

CD16+

CD20+

Fig. 23.6. Comparison of human and cynomolgus monkey cell populations (CD3 + , T-Lymphocytes; CD4 + , T-helper cells; CD8, cytotoxic T-cells; CD16 + , NK-cells, CD20 + , B-cells) showed no significant differences between species.

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new marker is known to be expressed on T-helper cells only) should be confirmed. If blood samples are to be stored before measurement, the stability of the marker on the target cells should be shown (see Fig. 23.7), and a titration of different concentrations is crucial if fewer antibodies than recommended by the provider are to be used. Other validation parameters, such as intra-run precision or staff-robustness, are method and not antibody dependent. 2. NK cell function testing: Normally, different effector target cell ratios (12.5:1, 25:1, 50:1) are used. Limitations in blood volumes might not allow for triplicates in all ratios. Testing only one ratio (25:1) in triplicates might result in more reliable data than using singletons of all ratios. This should be tested during method validation. 3. TDAR: Note that the titre calculation is one of many different suitable methods, which is not NHP specific. For anti-KLH ELISA, it is recommended that one batch of KLH can be used for all samples from one study, since there is a high batch-tobatch variability. As a consequence, titres from different studies determined with different KLH batches are not entirely comparable. 4. Bone marrow examination: As with any clinical pathology parameter, interpretation of bone marrow data should never be made in isolation. Clinical signs, the results of haematological, biochemical and immunological assays, post mortem findings and, where appropriate, immunohistochemistry and 100 0h

24 h

48 h

% of Lymphocytes

80

60

40

20

0 CD3+

CD4+

CD8+

CD16+

CD20+

Fig. 23.7. Stability of lymphocyte populations in blood (CD3 + , T-Lymphocytes; CD4 + , T-helper cells; CD8, cytotoxic T-cells; CD16 + , NK-cells, CD20 + , B-cells), stored on a roller at room temperature (mean ± SD, n = 3). Within 48 hour no shift of populations is seen. The standard immunophenotyping protocol had been used.

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histopathological findings, should all be taken into account while assessing any given therapeutic response. For example, peripheral reticulocyte counts show the functional effectiveness of the erythroid cell component of bone marrow, whereas peripheral neutrophil “band” cell (metamyelocyte) counts indicate effective myeloid cell production. The individual cell compartment counts demonstrate the maturation of the individual cell lines. The M:E ratio demonstrates the relationship between these two cell compartments in the marrow. Alterations to this parameter may indicate a reduction in one of the cell compartments (myeloid or erythroid), or alternatively, an increase in the other: This effect can be clarified by having a consideration of the peripheral cell counts and by reference to a histological section of a bone marrow section in determining the overall cellularity of the marrow. 5. Immunohistochemical staining: The tissue distribution of each antibody should be determined immunhistochemically for the assessment of antibody sensitivity and specificity. All anti-CD antibodies should be validated before use in GLP-studies. In parallel to each staining, a positive and negative control staining should be performed. 6. Lymphocyte proliferation assay: for the prepared PBMCs, a viability higher than 85% and less than 20% erythrocytes is recommended. Despite the comparable cell numbers, absolute PBMC proliferation rates between individual donors may vary by a factor of 3. The stimulation index is defined as the quotient of the proliferative response in presence of the mitogens and the proliferative response without mitogens. PBMCs of cynomolgus monkeys show some differences with regard to mitogen responsiveness as compared to human PBMCs. The use of LPS (lipopolysaccahride) or PMA (phorbol myristyl acetate) is not recommended because for this species LPS is not a good inductor of proliferation. Stimulation with PMA yields in a massive proliferation in the presence of the highest concentration of PMA but not at lower concentrations. Thus, for the assessment of lymphocyte proliferation PHA and/or IL-2 should be used.

Acknowledgments The authors are indebted to Prof. Eberhard Buse, Pathology Department of Covance Laboratories GmbH Münster, for developing and providing the immunohistochemistry staining protocol. The authors wish to specifically acknowledge the dedicated and competent work of our laboratory technical staff.

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References 1. Chapman K, Pullen N, Graham M, Ragan I (2007) Preclinical safety testing of monoclonal antibodies: the significance of species relevance. Nat Rev Drug Disc 6:120–126 2. Cavagnaro JA (2002) Preclinical safety evaluation of biotechnology-derived pharmaceuticals. Nat Rev Drug Discov 1:469–475 3. Frings W, Cavagnaro JA (2005) Predicted clinical immunogenicity: intended or unintended. In: Weinbauer GF, Buse E, Müller W, Vogel F (eds) New developments and challenges in primate toxicology. Waxmann Verlag, New York, pp 9–21 4. Thomas PT (2002) Non-clinical evaluation of therapeutic cytokines: Immunotoxicologic issues. Toxicology 174:27–35 5. Buse E, Habermann G, Osterburg I, Korte R, Weinbauer GF (2003) Reproductive/developmental toxicity and immunotoxicity assessment in the non-human primate model. Toxicology 185:221–227 6. Hendrickx AG, Makori N, Peterson P (2000) Non-human primates: their role in assessing development effects of immunomodulatory agents. Hum Exp Toxicol 19:219–225 7. Buse E (2005) Development of the immune system in the cynomolgus monkey: the appropriate model in human targeted toxicology. J Immunotoxicol 2:211–216 8. Herzyk DL, Bussiere JL (eds) (2008) Immunotoxicology strategies for pharmaceutical safety assessment. Wiley, New Jersey 9. Cavagnaro JA (ed) (2008) Preclinical safety evaluation of biopharmaceuticals. Wiley, New Jersey

10. House RV, Thomas PT (2002) Immuno toxicology: fundamentals of preclinical assessment. In: Derelanke MJ, Hollinger MA (eds) Handbook of toxicology, 2nd edn. CRC, Boca Raton, FL 11. International Committee on Harmonisation: Immunotoxicity studies for human pharmaceuticals. (2006) ICH Topic S8 (ICH guidelines are available at http://www.ich. org) 12. Price KD, Mezza L, Diters R, Wells S, Devona D, Tzogas Z, Haggerty H (2004) Development and immunomodulation of delayed type hypersensitivity (DTH) in cynomolgus monkeys. Toxicologist 78:431 13. International Committee on Harmonisation: Preclinical safety evaluation of biotechnologically derived pharmaceuticals. ICH Topic S6 (ICH guidelines are available at http://www. ich.org) 14. Carter DL, Shieh TM, Blosser RL, Chadwick KR, Margolick JB, Hildreth JE, Clements JE, Zink MC (1999) CD56 identifies monocytes and not natural killer cells in rhesus macaques. Cytometry 37:41–50 15. Weinbauer GF, Niehoff M, Niehaus M, Srivastav S, Fuchs A, van Esch E, Cline JM (2008) Physiology and endocrinology of the ovarian cycle in macaques. Toxicol Pathol 36 (suppl):7S–23S 16. Buse E, Habermann G, Vogel F (2006) Thymus development in Macaca fascicularis (Cynomolgus monkey): an approach for toxicology and embryology. J Mol Histol 37:161–70

Part V Evaluation in Humans

Chapter 24 Fundamentals of Clinical Immunotoxicology Robert V. House Abstract Whereas animal studies are invaluable for screening various chemical and drugs for immunotoxic potential, such systems are necessarily limited in their predictive value for humans given the differences in physiology, immune system structure and function, and various other parameters between humans and nonhuman animals. However, prospective experimental studies in humans are not always practical or ethical. What is needed is an approach for combining animal data, human data collected in the course of clinical studies, and modern tools of bioinformatics and systems biology. In this chapter, we will explore current assays and methodologies for assessing immunotoxic potential in humans using this multi-parameter approach. Key words: Clinical trials, Cytokine storm, Human immunotoxicology, Immunomodulator, Immunotherapy, Methods, Vaccines

1. Introduction: Basic Terminology and Concepts

The subject of this review is clinical immunotoxicology: de facto, the study of immunotoxicity in humans. We should preface our discussions by specifying what we mean by the term immunotoxicity/immunotoxicology. There are undoubtedly as many definitions of the term as there are investigators including some “official” definitions. For this review, I submit my own version: “Pathological deviation from a homeostatic norm for host resistance.” This definition presumes some generalizations and potential heresies; allow me to explain. Except for true toxicity (that is, poisoning) that directly affects immune constituents, what we sometimes call immunotoxicity may in some cases actually be a biologically appropriate response. For example, cyclosporin acts via normal receptor pathways, although the ultimate response is not necessarily “healthy.” In this same context, the unintended consequences of biopharmaceuticals

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(discussed later) usually make sense in retrospect. The key differentiator is that the response is pathological in context, and that it affects host defense in some measurable way. The secondary concern of this definition is that any immunotoxic reaction causes a deviation from homeostasis. A transient deviation from the norm is acceptable if it is truly transient since the immune system is highly adaptable and self-regulating. It is established dogma that the immune system tends to correct itself, so subtle damage may not even be evident. The multiple redundancies in mechanisms and rapid replenishment of immune system constituents are necessary for survival in a threat-rich environment. Deviations in isolated immune response parameters are straightforward to detect in highly controlled animal models; however, in humans such subtle changes might not be so easily observed. Figure 24.1 illustrates the myriad ways that immune homeostasis might be perturbed, and some of the probable pathways by which the immune system would restore the organism to a state of immune homeostasis. From this simple illustration, it is clear that “immunotoxicity” is not simply down or up (suppressed or augmented), and that any mode of modulation may have consequences on host defense. Most of these interactions have already been described in some form in animal studies, and many have been shown to be true in clinical studies as well (although often not in the strict context of immunotoxicology).

Fig. 24.1. Spectrum of immunomodulation.

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2. Why Not Depend on Animal Studies?

2.1. When Immunomodulation Is a Good Thing: The Example of Vaccines

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Biomedical research has an ultimate goal of delivering appropriate health care to humans. It would seem axiomatic then that we should use the shortest path available to gaining data representative of human disease. Yet, the vast bulk of biomedical data have been generated in animal studies, both small (rodents and lagomorphs) and large (primarily canines and nonhuman primates). Why is this so? First, humans are not always “good” experimental animals (although they are always the relevant model). A prospective study in humans would yield only a fragmentary, incomplete picture of reality, at least from a mechanistic standpoint, since humans are highly outbred (that is, genetically dissimilar), and so each human in effect is its own experiment “group.” Perhaps as important, humans are highly heterogeneous from an environmental control standpoint; the diversity of human lifestyle can never fully be accounted for in experimental designs (diet, preexisting health status, medications, stress levels, past exposure to infectious or carcinogenic insults, and so on). Obviously, the single most important reason that humans are by and large impractical for prospective studies to identify potential immunotoxicants is that it is unethical to test potential toxicants in humans without any anticipated benefits to the subjects. While we must, therefore, often default back to nonhuman animal studies to gain knowledge applicable to humans, it will be increasingly important to generate first-level data from humans or human-like surrogates. As the field of immunotoxicology has matured, a greater realization of the need to assess immune stimulation (as opposed to only immune suppression) has emerged. However, the adverse consequences of inadvertent immune stimulation remain largely undefined at present (1). Indeed, current regulatory guidance documents for immunotoxicology suggest the need to consider inadvertent immune stimulation, but offer no advice on how such data would be used in safety evaluation. On the other hand, certain types of medical products are specifically designed to induce or modulate an immune response. Safety assessment of this type of product raises an entirely different question, namely, how can one differentiate the intended immunomodulation from the unintended immunomodulation? The introduction of preventive vaccines into the medical armamentarium has arguably had a greater effect on overall human health than any other medical development. Rather than treating morbidity, vaccines prevent infection and, thus, prevent much suffering and expenditure of resources. Perhaps paradoxically, this very decrease in human morbidity has led first to complacency,

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and now a backlash in the form of an increasingly vigorous anti-vaccine lobby that not only argues against the need for vaccines, but raises unfounded fears about their safety as well. As with other human medicines, safety evaluation for vaccines begins long before human clinical trials are allowed to take place. However, for preventive vaccines the use of healthy volunteers significantly limits the tolerance for risk; therefore, the acceptance criteria for volunteers, and the parameters for evaluation of health during the clinical trial portion of the development life-cycle, are tightly controlled. For vaccines, this emphasis on safety continues for the entire development lifecycle, including postlicensure. Indeed, certain vaccines may require very large Phase 3 studies given the importance of ensuring safety. Naturally, as Paracelsus noted so long ago, poison is in everything; vaccines, like any other medicine, carry an inherent risk and no vaccine is ever completely safe. However, there is an important difference; unlike almost all other medicines, vaccines are routinely administered to large numbers of otherwise healthy individuals. Thus, the bar for safety on vaccines is nearly always much higher than therapeutics. Couple this with the fact that certain vaccines under development (most notably biodefenserelated vaccines and pandemic influenza vaccines) may never be used, and the risk/benefit ratio is difficult to estimate. Before we discuss how vaccine safety can be maximized, let’s explore some of the more controversial aspects of vaccine safety – or at least controversial in the eyes of anti-vaccine activists. 2.1.1. Do Vaccines Overload the Immune System?

An interesting argument making the rounds is that the multiplicity of vaccines currently in use somehow quantitatively “overloads” the immune system causing it to be unable to respond to infection (2). Some simple calculations reveal that the potential immune repertoire of the human neonate is approximately 109– 1011 antigens. Therefore, the capacity to respond to antigenic stimuli is approximately 105 vaccines simultaneously, assuming an antigenic load of 100 antigens/vaccine. Based on the standard 11 vaccine regimen to American children, children are now routinely exposed to 125 vaccine antigens (not, however, simultaneously). Clearly, this is far below the level that theoretically it would overload a neonatal immune response (3). Consider that in 1960, there were over 3,200 antigens per five vaccines total; it becomes obvious that the actual antigenic burden has decreased. Vaccination can result in short-term hyporesponsiveness to in vitro immune tests but not to infection. The majority of studies have demonstrated that immunization with multiple vaccines does not predispose one to infection with unrelated organisms; in fact, vaccinated individuals appear to be less susceptible to nonvaccine related infections (3).

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2.1.2. Does Vaccination Lead to Autoimmunity?

The induction of autoimmune-type conditions has been associated in the literature with various vaccines. The primary proposed mechanism for this is a real (or imagined) similarity between the vaccine components (generally for viral vaccines) and self proteins. For example, both Hepatitis B and influenza vaccines have been suspected of causing multiple sclerosis, BCG and Hib vaccines have been associated in the literature with type 1 diabetes, and Lyme disease vaccine with chronic arthritis. Although such association may be theoretically plausible, there have to date been no clear-cut causal associations determined for any link between vaccination (for any disease) and the induction of autoimmunity in humans (4–7).

2.1.3. Does Vaccination Lead to Increases in Asthma or Allergy?

One argument against vaccination is based on the empirical observation that concomitant with the increase in vaccinations and decrease in so-called childhood infections has been a rise in the incidence and prevalence of allergic reactions, including asthma. The so-called “Hygiene Hypothesis” was proposed in which the prevention of childhood infections results in a stronger Th2 response, predisposing an individual to allergic reactions (8). At first blush, the hypothesis appears to make sense, however, it may be based on what can be thought of as a number of immunological fallacies (4), including: (a) Vaccines do not prevent most “childhood infections,” which are predominantly viral infections of the upper respiratory or intestinal tracts; (b) Other strong Th2 conditions, such as pregnancy or advanced melanoma, do not predispose toward allergies; (c) Helminthic infections decrease, rather than increase, allergies. Several major, well-controlled clinical trials have failed to detect a correlation between allergy and vaccination. While this remains a controversial area, many investigators agree that the hygiene hypothesis is unlikely to play a significant role, if any, in the increasing incidence and prevalence of allergy in the developed world (4, 9, 10). This is not to say that allergic reactions to vaccine don’t occur; as previously noted, vaccines have safety concerns like any other medicine, and allergic reactions are common in all medicines. Allergic reactions to vaccines are rare (a few cases per every 10,000 vaccinations), and nearly always occur in response to the presence of various vaccine components such as animal proteins (ovalbumin, chicken proteins, gelatin, calf lymph), yeast proteins (recombinant products), preservatives (thimerosal), adjuvants (aluminum), and antibiotics. Reactions can also occur to packaging (latex in stoppers). Overall, however, allergic reactions to the active vaccine components, or hypersensitivity reactions to the process of immunization per se, generally are not a concern.

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2.1.4. Does Vaccination Lead to Autism?

The preceding discussions of vaccine safety all have direct relevance to immunotoxicology in humans (tolerance, hypersensitivity and autoimmunity). However, although no immunological correlation has been associated with it, one additional and major vaccine safety must be considered here since no review is complete without it: autism. Autism is a collection of related syndromes, all with some degree of learning disorder. The incidence and prevalence of autism appears to be on the increase, and through a series of associations too numerous to detail here, this condition has been associated with vaccinations, particularly measles/mumps/rubella (MMR) vaccination (11). Ultimately, autism became linked in the public eye with the presence of the preservative thimerosal in vaccines; however, multiple clinical and epidemiological studies have discounted any such association (12–14). An interesting recent paper emphasizes that the environmental risk factors for autism are more likely to exert their effects prenatally, rather than during childhood (15). Regardless of the evidence, this will likely continue to be a topic of heated controversy for the foreseeable future.

2.2. When Immunomodulation Is Not a Good Thing: The Example of Immunomodulatory Antibodies

As suggested earlier, the term immunomodulation may as easily be applied to immunostimulation as to immunosuppression. The great majority of research over the past 30 years has focused on immunosuppression (16). While this focus has produced much important data, it can be argued that this tells only part of the story since adverse immunostimulation has the potential to be as deleterious to the host (17). Until recently, however, this risk has been largely theoretical since few natural toxicants or therapeutics have been shown to have even moderate stimulatory effects on the resting immune response. This situation changed when therapeutics began to be designed with the express purpose of stimulating the immune system. The “test case” for how such therapeutics could affect the immune system is clearly the antiCD28 monoclonal antibody TGN1412. TGN1412 is a superagonist intended to stimulate T-cells irrespective of activation status, and to preferentially stimulate Th 2 cells. Preclinical studies were performed according to the accepted toxicology standards, and no untoward effects were observed and no adverse effects were expected in humans according to the standards in place at the time of the study. However, once the antibody was infused into human volunteers in the phase 1 study, several individuals responded with a massive release of inflammatory cytokines leading to shock and multi-organ failure; this syndrome is referred to as a “cytokine storm” and is an extreme medical emergency (18, 19). In the intervening years since this tragedy, much has been written about how this could have been avoided; it is clear in hindsight that the clinical trials should have been designed and conducted differently, and it is hoped that these lessons will not be

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soon forgotten (20, 21). The more important point here is that animal data will never fully replace human data, and so the importance of true clinical immunotoxicology will become ever more important as more of these types of therapeutics are developed. Testing strategies have been suggested, and some basic foundations have been established (22). 2.3. Clinical Considerations for Developmental Immunotoxicology

3. General Testing Considerations and Approach

A topic of increasing focus and significance within immunotoxicology is the effect of xenobiotic exposure on the developing immune system, so-called developmental immunotoxicology (DIT). As initially suspected and now well-documented by experimental evidence in animal studies, the developing immune system is sensitive to a wide range of toxicants (23, 24). Perhaps more than any other area of immunotoxicology, DIT appears to have made the most advanced transition to date between nonclinical and clinical studies; this would be due at least in part to the relatively naive (antigenically speaking) immune system in children. Several excellent reviews have been written on how DIT studies can be applied to children (25–27), and a number of publications describe experimental studies in this area (28–31). In this author’s opinion, this particular branch of immunotoxicology promises to be one of the most rapidly developing in terms of practical data for human safety assessment.

As described above, the concept of prospective, controlled experimentation on humans is usually infeasible from an ethical consideration. However, this is not to say that human studies per se are totally unallowable. True human (clinical) studies range from highly controlled clinical trials to large, population-based observational studies. Clinical studies, specifically those done in the course of licensing therapeutic or prophylactic medicines, resemble animal studies to the degree that exposure parameters of interest can often be controlled and outcomes can be monitored. However, such studies (particularly when extensive biological monitoring and functional immune tests are to be performed) can be expensive, and exposures as well as outcomes of interest may be difficult to study in the available time frame since volunteers are not always available for long-term exposures or extended follow-up. Some examples of clinical studies of special relevance to immunotoxicity assessment include vaccine studies or studies of patients administered immunosuppressive therapy such as transplant patients or those with chronic inflammatory diseases such as arthritis. Observational or epidemiological studies are likewise valuable, can be of varying size, and can be cross-sectional, retrospective,

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or prospective in nature. In addition to the high costs associated with such studies, observational studies are challenging for many reasons, including potential confounding by host factors (age, gender, lifestyle, medical history) and environmental factors (frequency of exposure to chemicals and infectious agents). Rigor in observational studies is ensured by the use of multivariable analysis techniques such as regression modeling, providing there is sufficient sample size and information on potential confounders. Well-designed epidemiological studies can contribute valuable information to the assessment of risk due to immunotoxic exposures. Because of having a concern about the individual variation, a confirmatory evaluation and a cross-sectional or longitudinal study design can be employed using randomized normal, nonexposed individuals. It is also important to obtain a complete medical history including any potential immune dysfunction and accurate exposure assessments (32). Regardless of the potential value of observational/epidemiological studies, there is no substitute for hard data, and it is the collection of such data that will concern us for the remainder of this review. Table 24.1 below shows a sample of some of the clinical observations that would lead one to suspect immunotoxicity in humans; in the following sections, we will describe the various tests available for obtaining such data for immunotoxicity assessment in a clinical setting. 3.1. Conducting Research in Humans

With that brief background, we next turn to understanding some of the assays and approaches germane to clinical evaluation of immunotoxicity. We must first begin by understanding the special

Table 24.1 Findings indicative of immunotoxicity in human studies Finding

Indicative of

Relative common Decreased white cell counts

Immune suppression

“Flu-like” symptoms

Immune stimulation

Allergy/pseudoallergy • Frequent and mild (rashes) • Rare and severe (anaphylaxis)

Immune stimulation (hypersensitivity)

Relative rare Increased incidence of lymphoma Immune suppression Infectious complications

Immune suppression

Autoimmunity

Immune stimulation? (autoimmunity)

Cytokine storm

Immune stimulation

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considerations involved in human research. Medical research involving human subjects is done to improve diagnosis, prophylaxis and therapy. In addition, it is done to further our understanding of the etiology and pathogenesis of disease, which is information often impossible to obtain in any way except by going directly into humans. It follows that a human subject can be defined as a living individual about whom an investigator conducting research obtains data through intervention or interaction with the individual, or identifiable private information. Such information may include bodily materials, diagnostic specimens (including residuals), and even private information that a subject provides to investigators. It is paramount in doing human research that ethical considerations remain the highest priority. The Hippocratic Oath states “I will apply dietetic measures for the benefit of the sick according to my ability and judgment; I will keep them from harm and injustice…I will neither give a deadly drug to anybody if asked for it, nor will I make a suggestion to this effect.” The overarching principal can best be stated “Primum non nocere” (first, do no harm). Basic ethical principles include respect for the person (acknowledge personal dignity and autonomy and protect those with diminished autonomy), beneficence (do no harm, maximize potential benefits and minimize potential harm), and justice (fairness in the distribution of research benefits and burdens). 3.2. Specific Tests Available to Assess Immunotoxicity in Humans

Having now established that human “experimentation” can be performed under certain tightly controlled conditions, we turn our attention to the “how” of such research. Basing our approach on the seminal work of Luster et al., we can likewise divide human immunotoxicology testing in a tiered approach. Table 24.2 illustrates the variety of tests that can be performed, including those that might be considered routine (analogous to Tier 1 tests), and those that are more mechanistic/investigatory in nature (similar to Tier 2). In the subsequent sections, we will cover each of these approaches in greater detail.

3.2.1.Complete Blood Count and Differential

Complete white blood cell counts and differentials on all individuals whose immune status is being evaluated are easily incorporated into a Tier 1-type panel of immune assays. These data are expressed as absolute lymphocyte count for each cell type. Higher absolute lymphocytes counts should be expected in children than adults and in certain ethnic groups. Lymphocyte counts consistently below 1,500/mm3 are termed lymphocytopenia and may signal a defect in the T-cell compartment. Lymphocytopenia can be associated with primary immune deficiency disease, but can also occur secondary to conditions such viral infections, malnutrition, severe stress, autoimmune diseases, and hematopoietic malignancy. When lymphocytopenia is repeatedly observed, a bone marrow

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Table 24.2 Immune assessment tests for humans I. Primary screening tests General immune indicators

Screening procedures

These tests are usually included in general health panels

• These assay methods have been extensively standardized among laboratories • Results for these parameters are clinically interpretable • For these parameters, reference ranges are well established across age groups

1. Complete blood count and differential counts 2. Serum immunoglobulin levels; less often including quantification of different immunoglobulin classes 3. Delayed type hypersensitivity (DTH) test

II. Additional confirmatory tests

More detailed tests for the immune system

Immune parameters

These tests should be • The assays are often less included when indicated standardized by clinical findings • Some date may be difficult to interpret • Reference ranges are not as well established particularly across all age groups

1. Phenotyping for major lymphocyte subsets 2. Primary antibody response to immunogen (vaccination) 3. Assessment of cytokine production 4. Nonspecific/innate immune assessment: NK cell function; autoantibodies; granulocyte function

Modified from (32)

biopsy is sometimes recommended as an aid for ruling out other diseases and for the identification of normal plasma cells, pre-B-cells, or diagnosis of bone marrow depression or dysplasia. Individuals with lymphocytopenia should be reevaluated and further assessed for changes in cell-mediated immune function. Lymphocytosis (higher than expected lymphocyte counts) can be caused by chronic infections or allergic reactions. Abnormalities in granulocyte counts also occur include monocytosis (associated with stress, infections and hematologic disorders) and eosinophilia (caused by allergic reactions, parasitic disease, neoplasia and adrenocortical dysfunction) (32). It is important to note that any of these observations must be seen as an isolated event in time, and readings can be skewed be nonimmunological events (such as physiologic lymphocytosis resulting from epinephrine). Conclusions regarding these data thus must be made in conjunction with the other supporting data (33). 3.2.2. Immunoglobulin Concentrations

Perhaps the most relevant measure of immune status that can be learned from serum chemistry tests is the measure of globulins; although this assessment includes lipoproteins and acute phase proteins, the most relevant for our purposes is the measurement

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of immunoglobulin (Ig) concentrations in serum (33). This is a straightforward test that can be easily included in retrospective epidemiological studies. Methods for quantifying Igs include single-radial diffusion, double diffusion in agar gel, immunoelectrodiffusion, radioimmunoassay, and enzyme-linked immunosorbent assay (ELISA), among others. Electrophoresis may be used to further separate Igs into various fractions, adding granularity to the results. The adverse health effects associated with small changes in immunoglobulin levels is unclear, although large decreases may suggest lymphotoxicity. Conversely, globulins can increase in certain pathological conditions, including infection or other types of immune stimulation. 3.2.3. Delayed-Type Skin Testing

Skin testing is sometimes used to assess cellular immune competence since delayed hypersensitivity, a localized cutaneous response, is mediated by T-cells and the production of inflammatory cytokines. Antigens commonly used to elicit a positive skin response include purified protein derivative of agents such as mumps, trichophyton, candida, tetanus, or diphtheria (all agents that many individuals are expected to exhibit a recall response to due to prior exposure either environmentally or via immunization). These antigens usually are administered by intradermal injection and are grouped in a panel for greater coverage. Skin responses are read at 48 and 72 h for maximal diameter of erythema and induration. Unfortunately, the test is not considered very sensitive unless severe immunosuppression is suspected, which is unlikely to occur. In general, DTH response would not be a routine screening option for human immunotoxicity.

3.2.4. Phenotypic Analysis by Flow Cytometry

Evaluation of surface markers (the so-called cluster of differentiation [CD] molecules) on immune system cells by flow cytometry has provided considerable information on the development and activation state of the human immune system in children and adults, as well as assisting in the clinical diagnosis for immunological disorders. CD markers have been identified for many relevant cell subpopulations, providing a powerful tool for tracking not only the identity and location of various immune cell populations but for assessing activity as well. Flow cytometry is now well established as a vital technique in nonclinical immunotoxicology testing (34). For clinical immunotoxicology, the obvious choice of experimental sample would be peripheral blood; some more involved clinical studies might provide access to the other samples such as lung lavages, but these would be relatively rare. The most commonly used procedure for processing experimental samples for immunofluorescence is the first to stain an aliquot of whole blood with fluorescent-conjugated antibodies, followed by differential

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analysis on a flow cytometer. The relative proportion of each cell type in a sample is then determined. The technique is well developed and reference values for many cell populations across multiple species have been established (35). Flow cytometric analysis of cell populations (and activity), while listed as a second-order test in Table 24.2, can easily be performed as either a mechanistic study, or as a more routine screen (although the “Tier 1” tests listed in Table 24.2 would generally be performed in many clinical evaluations when immunotoxicity might be suspected and so would not generally be included as a routine screen due to the costs). Perhaps the greatest future potential value of flow cytometry lies in safety evaluation of biopharmaceuticals; with these agents, modulation of the immune response is an intended (or at least strongly suspected) consequence. Flow cytometry is able to provide at least a secondary read on immune structure (that is, mobilization of various immune cell populations) and function, and should be included in any such evaluation (36, 37). 3.2.5. Specific Antibody Assessment

The T-dependent antibody response (TDAR) has become the de facto primary ex vivo assay for evaluation of immunotoxicity in animal studies (38). The benefit of the TDAR, as fully documented in the literature, is that elucidation of this response requires the concerted interaction of T-cells, B-cells (plasma cells), antigen-presenting cells, complement, and cytokines; thus, a single assay encompasses essentially all of the components of an adaptive immune response. By extension, a xenobiotic-induced deficit of any single component should be manifested as an immunotoxic event. The TDAR was found to be one of the most predictive of all immunotoxicity assays in animals and is widely used as an initial screening assay (39). Of course, the TDAR requires immunization with foreign antigens (generally sheep erythrocytes or keyhole limpet hemocyanin), and the effector cells are taken from the spleen; neither parameter is practical for human testing. However, an increasing initiative in clinical immunotoxicology is to quantify the response to vaccines, usually by measurement of antibody titers following vaccination. This approach obviates any concern with administration of foreign antigens since vaccines go through rigorous safety testing, and the conditions for administration to induce an optimum immune reaction are well known in advance. In addition, no tissue collection is required since antibody titers in the serum are all that is required. The use of vaccine studies to evaluate immunotoxicity in humans was first proposed by van Loveren et al. (40). Some of the first work in this area was the demonstration of a decreased response to hepatitis vaccination in students exposed to ultraviolet radiation in association with polymorphisms in genes controlling

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inflammatory cytokines (41). More recently, vaccination has been used to evaluate the immunotoxic effect of the exposure to various pesticides (42–44). While assessing vaccination responses for clinical immunotoxicology has not been validated, results to date appear to support the ability to detect changes in populations with moderate degrees of immunosuppression; this suggest a level of sensitivity unlikely be achieved with more commonly employed clinical tests (immunophenotyping, immunoglobulin levels, etc.) 3.2.6. Cytokine Measurement

Cytokines are protein/peptide molecules that act as chemical messengers in many physiological systems within the body. Although originally discovered in the context of the immune response, they are now known to mediate many different processes (often in the context of immune or immune-related physiological processes). Due to their central role in controlling the complex interactions in both innate and adaptive immunity, as well as the more recent understanding of the role they play in transitioning between innate and adaptive immunity, cytokines are an obvious target for research in clinical immunotoxicology (45, 46). The various methodologies for assessing cytokines (bioassays, immunoassays, and molecular techniques), and the technical considerations associated with their performance, have been covered in detail elsewhere (47, 48). However, there are important strategic considerations that are often overlooked yet which are vitally important, particularly for clinical immunotoxicology: (a) Cytokine analysis by itself will practically never give a complete story. Cytokines are rarely the ultimate effector mechanism for anything; rather, they are mediators between other systems. Thus, cytokine data should be part of the strategy, not the answer; (b) The list of potential cytokine targets is immense, and increasing all the time (see especially http://www.copewithcytokines. de). Study designs that simply shotgun cytokine targets based on assay availability are wasteful of resources and may unnecessarily complicate the issue; rather, cytokine analysis should be hypothesis-driven and the number of candidates should be minimized to answer the question at hand. For example, is the response expected to be mediated via a differential T-helper cell response (Th1, Th2, Th3, Th17)? Does the response have a significant inflammatory component? And so on. (c) Cytokines generally act at a local level but rarely systemically (in fact, global release of cytokines, particularly in the form of a cytokine storm, is often pathological). Detection of cytokines systemically reveals little about what is happening at the intended target. This is especially a concern in clinical immunotoxicology, since access to tissues other than

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serum or plasma is generally difficult outside the confines of a clinical trial. Given the latter consideration, care should always be exercised when evaluating immunotoxicity in humans by means of circulating cytokines. Cytokine levels in the circulation may be modulated by a variety of factors such as stress, exercise, low-level infections and so forth. Therefore, a “snapshot” approach is unlikely to provide a clear picture of transient effects. (As mentioned in the introductory remarks, we should be wary of assigning the term “immunotoxic” to transient observations.) One approach is to supplement a systemic cytokine evaluation with an assessment of ex vivo cytokine production by immune cells; this is particularly well suited to clinical evaluation since it produces a “cleaner” result (that is, one that is relatively free of physiological interference). A recent approach (49, 50) for evaluating cytokine production is the use of the Cell Chip, a T-cell-based fluorescent chip utilizing reporter cell lines. This system purportedly is able to determine the T-cell immunotoxicity in vitro; while this system has not been fully developed, it is reasonable to postulate that modifications of such a system to include various human immune cells would be a powerful adjunct to in vivo clinical immunotoxicology assessment. 3.2.7. Natural Killer (NK) Cells

Assessment of NK cell activity has a long association with the evaluation of immunotoxicity in nonhuman models. For most studies in animals, NK cell activity has been used most often as a measure of nonspecific host defense, particularly against viruses and tumor cells. However, recent work with NK cells has revealed them to be a key nexus of innate and adaptive immunity, serving both effector (killing) function as well as crucial immunoregulatory activities (51, 52). The gold standard assay for assessing NK cell function is the chromium-release assay in which immune cells (in the case of humans, generally peripheral blood leukocytes) are co-cultured with NK-sensitive tumor cells (the K562 cell line in most cases) that have been radiolabeled with 51Cr (53). Generally, several effector target cell ratios are employed so that a response curve can be constructed. Following a 4-h incubation period, the released radiolabel is collected and the amount of radiolabel released (as the percentage of total releasable radiolabel, corrected for spontaneous release) is determined. This assay is relatively simple to perform and enjoys good reproducibility between laboratories (54). More recently, several variations of the basic assay have been developed that obviate the need for radiolabels, rendering the assay safer and more accessible to laboratories that cannot use radiolabeled materials (55).

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3.2.8. Neutrophil Function

Neutrophil function has generally been seen as a mechanistic/ confirmatory measure of potential immunotoxicity, at least in nonhuman studies. This may represent a bias toward the assessment of adaptive immunity rather than innate immune defense, or it may reflect the difficulty in obtaining sufficient cells from rodents. Regardless, for human studies the neutrophil represents an important target of opportunity to understand how exogenous factors might affect this important mediator of host defense. Numerous techniques are available for measuring neutrophil function including phagocytosis, elucidation of cytokines, quantitation of reactive oxygen species and reactive nitrogen species and microbial killing; an excellent review has been recently published (56). In addition, enumeration of neutrophils from peripheral blood is an easy assay that can yield much information (33).

3.2.9. Autoantibodies

A number of autoimmune-type diseases may manifest as loss of tolerance to nonself antigens and can be detected by the presence of autoantibodies in the circulation. Antibodies to cellular components and nuclear antigens (ANA, DNA, mitochondria) and to rheumatoid factor (RA) and their frequency in a population may reflect an immune alteration. Standardized diagnostic kits are available to detect the presence of these autoantibodies in sera. Although these tests have great potential value in understanding human autoimmune conditions, these tests (and the resulting data) are often complex and great care must be exercised in interpretation (57, 58).

4. The Future of Clinical Immunology? 4.1. The Promise of Synthetic Immune Systems

One frontier approach for understanding the effect of xenobiotics on the human immune response will be the use of what I term a “synthetic immune system.” (For the sake of clarity in definition, I use the term synthetic to differentiate this concept from “artificial” immune systems, which are computer architecture/software systems that purportedly model an organic immune response to make complex decisions…in essence a form of artificial intelligence.) For the current discussion, a synthetic immune system would be an in vitro cum ex vivo construct in which human-derived tissues are maintained in an in vitro environment engineered to contain an appropriate microenvironment supporting all of the necessary interactions for developing an adaptive immune response. For example, in such a system a biocompatible matrix would be seeded with endothelial cells and antigen-presenting cells such as dendritic cells (or other supportive elements) and peripheral blood leukocytes (PBL) from individual donors. These systems would

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develop an appropriate three-dimensional structure, including demarcation of B- and T-cell areas. This, of course, would be a significant enhancement over crude systems such as the MishellDutton culture system in that the milieu can be specifically tailored to mimic the human environment (59, 60). In addition, the donor population can be carefully controlled to allow for variability in human immune response, and if a long-term donor population can be maintained, a high degree of consistency and reproducibility can result. At present (2008), this technology is already being developed commercially to study in vitro immunization in the context of vaccine development by VaxDesign (http://www.vaxdesign.com). However, there is no obvious reason why this concept could not be adapted to assess the immunotoxicity in vitro as well. As with more traditional in vitro toxicology studies, certain variables will require development (metabolism of compounds, effects of vehicle, etc.) However, the potential benefits of this approach should more than offset any of the disadvantages posed by these relatively simple technical tweaks. 4.2. Not-Quite-Human Human Models

Since it is axiomatic that normal prospective (controlled) immunotoxicology studies cannot be ethically performed in humans (except, arguably, for vaccines which are designed to be immunomodulatory), and likewise accepted that nonhuman animals will only ever be a poor substitute for collection of human data, one option might be the creation of animals that mimic humans as closely as possible in terms of structure and function of the immune system. This can be accomplished by either “replacing” the immune system of rodents with a human immune system or by selectively altering the rodent’s immune system by genetic manipulation. These disparate approaches are described below. Regardless of the approach taken, these animals can be used much as their “normal” counterparts in controlled studies.

4.3. Replacement Strategy

The strategy for replacing a rodent immune system with a human one is relatively straightforward, although not necessarily simple. The framework is to use a mutant mouse strain lacking a functional immune system. There are at least two good candidates for use as this “backbone” including the severe combined immunodeficient (SCID) mouse (and the Nonobese diabetic (NOD) variant), which lacks functional T- and B-cells, and the beige/nude/ xid (x-linked immunodeficient) mouse (61, 62). These animals lack T- and B-cell function, thus forming a tabula rasa for the development of an adaptive immune response. Creation of such chimeric animals is an expensive proposition, and thus use of these models will probably always be reserved for highly specific mechanistic studies. Moreover, the very limited immunotoxicology studies that have been done in this model demonstrated a high

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variability, suggesting that much basic development remains to be done. Still, the ability to directly challenge a “human” immune system that would respond in an immunologically appropriate manner would be of great benefit to understand the potential clinical outcomes of immunotoxic exposure, and this model should be evaluated further. 4.4. Modification Strategy

Another strategy for packaging a humanized immune response in a rodent body is the use of transgenic animals (63). Put simply, transgenic technology allows for the insertion of specific, functioning genetic elements from one organism into another. Two essential technologies may be used: nonhomologous recombination, and homologous recombination. Of the two, homologous recombination is the most specific and will be covered here. The specifics of homologous recombination have been covered elsewhere (64) and will not be reiterated here in detail. In summary, murine embryonic stem cell lines either have specific genetic sequences eliminated so that they no longer express certain phenotypes (knockout mice), or have foreign genetic elements inserted so that they express novel proteins or other phenotypes (knockin). Once the embryonic stem cells are so modified, genetic chimeras are produced and founder lines of animals are bred from the chimeras. By a combination of knockout and knockin, mice can express various elements of a human immune response although the animal’s immune system is not totally replaced by a human counterpart. While these animals lack the entirety of the human immune response offered by reconstituted immunodeficient mice, they are considerably easier to create and maintain, and are much less expensive. The concept of using transgenic mice in immunotoxicology studies was first proposed by Løvik (65). Since that time, these animals have been used in a variety of mechanistic-type immunotoxicology studies, most often in the evaluation of autoimmunity, hypersensitivity and safety assessment of human biopharmaceuticals (64). The cost and technical resources that are necessary to create transgenic animals have so far prohibited a more widespread acceptance of this model in immunotoxicology screening; however, generation of a sufficient corpus of information using this model would help bridge the gap between animals and humans.

5. Putting All the Pieces Together In this review we have discussed both standard and “frontier” types of clinical immunotoxicology testing assays and systems. Clearly, we are still only in the early stages of understanding how best to determine whether and how xenobiotic exposure – whether

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Fig. 24.2. Strategies for turning clinical data into immunotoxicology knowledge.

inadvertent such as environmental or industrial, or deliberate such as therapeutics – affects the human immune system. Keeping in mind some of the considerations covered in the early part of this review regarding how we can assess whether or not such exposure is truly “immunotoxic,” and likewise keeping in mind the sheer volume and diversity of data as would be collected using this variety of tests, we need a better frame work for turning these data into true knowledge. This will be our focus for the remainder of the review. In Fig. 24.2, I have suggested one approach that builds on present understanding based on existing preclinical immunotoxicology data as well as data collected from various human studies such as clinical trials and epidemiological testing (clinical data trending and data mining from human studies). This is how we currently gain most of our understanding of potential human immunotoxicity, and it will continue to be a powerful (if not completely adequate) tool on an ongoing basis. To take this approach to the next level, it will be necessary to use the computational power of next-generation concepts such as bioinformatics, systems biology and the so-called converging technologies such as nanotechnology/biotechnology/informatics/cognitive science (NBIC). This approach already shows great promise in vaccine development (66), and should provide much better predictability for the studies of human immunotoxicology. Finally, Fig. 24.3 presents a more comprehensive (and admittedly more speculative) strategy which overlays the various approaches described in this review in a temporal/logical manner, and how these studies might be used to form a bridge between current existing data generated in small animal (primarily rodent) models, through nonhuman primates (assumed to be more closely

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Fig. 24.3. Meta-analysis for bridging nonclinical and clinical data.

relevant to human data) and finally to humans. In this paradigm, one can more fully comprehend the wide variety of data streams leading into the considerable systems biology computational/ modeling tools that would be required to form predictive models for understanding human immune effects. These tools are only now being developed, so for the near (and perhaps intermediate) term we must still depend on the incomplete (and, as we have seen, sometimes wholly inadequate) extrapolation from animal data to human situations.

6. Summary and Conclusions So the question remains: Are we closer to true clinical immunotoxicology? Clearly, given the scientific advances that have taken place over the 30-year history of immunotoxicology as a discipline, we are far closer to understanding retrospectively – and perhaps predicting – the toxic effect of xenobiotics on the human immune response than ever before. Given the ethical constraints against human experimentation, of course, prospective studies are unlikely to ever be conducted (and rightly so) with the exception of highly controlled clinical trials of immunomodulatory therapeutics and prophylactics (such as vaccines). In addition, retrospective studies are useful only in highly specific situations. For the near term

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then, our best current approach is extrapolation from animal studies to humans based on weight-of-evidence, as well as careful study of data resulting from clinical trials of human therapeutics, particularly those intended (or at least expected) to have direct or indirect effects on the immune response. As more sophisticated experimental and information systems evolve (bioinformatics, synthetic immune systems, and so forth), a systems biology approach to clinical immunotoxicology will serve to combine multiple streams of data (in vitro, ex vivo, and in silico) into a more comprehensive approach that will greatly improve not only the design of novel therapeutics but our prediction of potential environmental and industrial hazards as well.

References 1. House RV, Hastings KL (2004) Multidimensional immunomodulation. J Immunotoxicol 1: 123–129 2. Gregson AL, Edelman R (2003) Does antigenic overload exist? The role of multiple immunizations in infants. Immunol Allergy Clin North Am 23(4):649–664 3. Offit PA, Quarles J, Gerber MA, Hackett CJ, Marcuse EK, Kollman TR, Gellin BG, Landry S (2002) Addressing parents’ concerns: do multiple vaccines overwhelm or weaken the infant’s immune system? Pediatrics 109(1):124–129 4. Offit PA, Hackett CJ (2003) Addressing parents’ concerns: do vaccines cause allergic or autoimmune diseases? Pediatrics 111(3): 653–659 5. Schattner A (2005) Consequence or coincidence? The occurrence, pathogenesis and significance of autoimmune manifestations after viral vaccines. Vaccine 23(30):3876–3886 6. Goriely S, Goldman M (2007) From tolerance to autoimmunity: is there a risk in early life vaccination? J Comp Pathol 137(Suppl 1):S57–S61 7. Siegrist CA (2007) Mechanisms underlying adverse reactions to vaccines. J Comp Pathol 137(Suppl 1):S46–S50 8. Garn H, Renz H (2007) Epidemiological and immunological evidence for the hygiene hypothesis. Immunobiology 212(6):441–452 9. Ramsey CD, Celedón JC (2005) The hygiene hypothesis and asthma. Curr Opin Pulm Med 11(1):14–20 10. Sánchez-Solis M, García-Marcos L (2006) Do vaccines modify the prevalence of asthma and allergies? Expert Rev Vaccines 5(5):631–640

11. Taylor B (2006) Vaccines and the changing epidemiology of autism. Child Care Health Dev 32(5):511–519 12. Offit PA, Coffin SE (2003) Communicating science to the public: MMR vaccine and autism. Vaccine 22(1):1–6 13. Parker SK, Schwartz B, Todd J, Pickering LK (2004) Thimerosal-containing vaccines and autistic spectrum disorder: a critical review of published original data. Pediatrics 114(3): 793–804 14. DeStefano F (2007) Vaccines and autism: evidence does not support a causal association. Clin Pharmacol Ther 82(6):756–759 15. Dietert RR, Dietert JM (2008) Potential for early-life immune insult including developmental immunotoxicity in autism and autism spectrum disorders: focus on critical windows of immune vulnerability. J Toxicol Environ Health B Crit Rev 11(8):660–680 16. House RV, Luebke RW (2006) Immunotoxicology: thirty years and counting. In: Luebke R, House R, Kimber I (eds) Immunopharmacology and immunotoxicology, 3rd edn. CRC, Boca Raton, pp 3–20 17. Ponce R (2008) Adverse consequences of immunostimulation. J Immunotoxicol 5(1):33–41 18. Suntharalingam G, Perry MR, Ward S, Brett SJ, Castello-Cortes A, Brunner MD, Panoskaltsis N (2006) Cytokine storm in a phase 1 trial of the anti-CD28 monoclonal antibody TGN1412. N Engl J Med 355:1018–1028 19. Wang H, Ma S (2008) The cytokine storm and factors determining the sequence and severity of organ dysfunction in multiple organ dysfunction syndrome. Am J Emerg Med 26(6):711–715

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Part VI Wildlife Testing

Chapter 25 In Vivo Functional Tests for Assessing Immunotoxicity in Birds Keith A. Grasman Abstract Various methods have been adapted for assessing the effects of environmental contaminants on the structure and function of the immune system in wild and captive birds. This chapter describes two integrative functional assays that have been adapted to a variety of avian species and have proven to be sensitive biomarkers for immunotoxicological effects. The phytohemagglutinin (PHA) skin test measures T cellmediated immunity. PHA is injected intra- or sub-dermally into the wing web of the elbow joint (or interdigitary skin or wattle). The PHA stimulates T lymphocytes to release cytokines that cause an inflammatory influx of leukocytes and fluid. The thickness of the wing web is measured before and 24 h after injection. A stimulation index, which reflects T cell function, is calculated as the increase in skin thickness caused by the PHA minus the increase caused by an injection of phosphate buffered saline (PBS) in the other wing web. In addition to its sensitivity to contaminants, ecological studies have shown that the PHA skin response is positively associated with rates of survival and colonization of new areas (i.e., ability to found new local populations) in wild birds. The sheep red blood cell (SRBC) hemagglutination assay measures the antibody response to immunization with SRBC antigens, integrating the functions of B lymphocytes, helper T lymphocytes, and macrophages. A SRBC suspension is injected i.v., and a blood sample is collected approximately 6 days later. Plasma (or serum) from the blood sample is serially diluted in a microtiter plate, and SRBCs are added. The magnitude of the antibody response is defined as the titer – the highest dilution of plasma in which the concentration of antibody is sufficient to agglutinate the SRBCs. Both IgM and IgG titers can be measured. This avian test is very similar in principle to the anti-SRBC ELISA and splenic plaque forming assays used for immunotoxicological testing in rodents. However, this avian hemagglutination assay does not require a species-specific secondary antibody (as does the ELISA), and this minimally invasive, nonlethal procedure is amenable to studies of protected species, as opposed to the splenic assay. The PHA and SRBC assays have been employed successfully in both the laboratory and field. In ecological studies birds must be recaptured 24 h or 6 days after the initial injections, limiting their use in some species. However, their sensitivity to a variety of contaminants and their ease of adaptability to a variety of species have made the PHA and SRBC tests some of the most commonly used assays for screening and monitoring immunotoxicity in birds. Key words: Antibody, Avian, Birds, Hemagglutination, Immunosuppression, Immunotoxicology, Phytohemagglutinin, T cell-mediated immunity

R.R. Dietert (ed.), Immunotoxicity Testing: Methods and Protocols, Methods in Molecular Biology, vol. 598 DOI 10.1007/978-1-60761-401-2_25, © Humana Press, a part of Springer Science + Business Media, LLC 2010

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1. Introduction In one of the first studies in the field of immunotoxicology, Friend and Trainer (1) found that exposure to PCBs increased the mortality of mallard ducklings (Anas platyrhynchos) subsequently challenged with duck hepatitis virus. Since that time, many additional studies have documented the effects of various environmental contaminants on the structure and function of the immune system in birds (see (2) for a review). Contaminants that affect the avian immune system include heavy metals, pesticides, and organic industrial pollutants. Some of these immunotoxicological investigations have studied wild birds as indicators of the environmental health effects of contaminants. Laboratory studies, primarily of chickens, quail, and mallards, have allowed investigations under more controlled conditions intended to parallel field studies and provide more detailed information on immunotoxicological mechanisms. Immunotoxicology testing in birds often presents a number of unique challenges compared to similar studies in laboratory rodents. A large body of immunological literature for chickens, driven by a concern for infectious diseases in the poultry industry, provides a general understanding of avian immunology and immunological methods that can be used to study other species. However, relatively little is known about the basic immunology of wild or exotic species, and few species-specific reagents are available (e.g., monoclonal antibodies against immunoglobulins or cell surface proteins). Protocols for state-of-the-art immunological methods (e.g., involving cell culture, flow cytometry, ELISA) often require much experimentation and optimization to adapt to exotic species. In many ecological studies, the distance between field study sites and the laboratory may limit the usefulness of state-of-the-art immunological methods. Difficulty in capturing certain species may limit sample sizes. Certain immunological methods may be inappropriate because of difficulty in recapturing individuals (e.g., blood sampling for antibody titers a week after immunization). Choice of methods may be constrained by small body sizes, which limit blood and tissue samples, and (or) the need for nonlethal, minimally invasive tests when studying protected species (e.g., threatened or endangered). Despite these challenges, a wide variety of methods have been developed to screen for immunotoxicity and investigate immuntoxicological mechanisms in both wild and captive birds (see (2, 3) for reviews). These methods range from simple to state-of-the-art; some assess the structure of the immune system while others measure specific immunological functions or processes. White blood cell hematology may indicate deficiencies in specific leukocyte types or elevated leukocyte numbers resulting from active infections. The masses, cellularity, and histological

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characteristics of primary immune organs such as the bursa of Fabricius (the site of B lymphocyte maturation in birds) and thymus (the site of T lymphocyte maturation) give important data regarding immunological development. Similar endpoints for the spleen, a secondary immune organ, indicate infection status. Separation and quantification of plasma or serum proteins by electrophoresis can be used to measure overall concentrations of immunoglobulins (gamma-globulins) and acute phase proteins (alpha- and beta-globulins) that change in response to infection and inflammation. Methods have been developed to isolate and cryopreserve avian mononuclear cells (lymphocytes and monocytes) for in vitro tests such as mitogen-induced proliferation and phagocytosis assays (4, 5). These methods allow cells collected in the field to be transported to the laboratory and archived until time of analysis. The methods described in this chapter are in vivo functional tests that have been employed in a large number of laboratory and field studies in a diversity of avian species. These integrative assays can be employed individually, but when used together they screen or assess a range of immunological functions. The phytohemagglutinin (PHA) and SRBC tests are nonlethal, minimally invasive methods that are amenable to studies of many different avian species, including those that are classified as threatened or endangered. They require no species-specific reagents or advanced laboratory equipment and, hence, have been used in many studies of wild birds. However, both assays do require the recapture of test subjects, and the SRBC assay is limited to species with body masses of approximately 100–150 g in order to provide sufficient volumes of blood. 1.1. Phytohemagglutinin Skin Test for T Cell-Mediated Immunity

The PHA skin response test is an in vivo method of measuring T lymphocyte-mediated immunity. It measures the swelling caused by inflammatory leukocyte and fluid infiltration after an intradermal injection of PHA. Mitogens such as PHA are derived from lectins, which are plant or bacterial proteins that bind to specific sugar components of glycoproteins attached to the surface of cells. PHA specifically binds to the surface of T lymphocytes. In the skin response test, PHA stimulates T lymphocytes to release lymphokines, resulting in an increase in vascular permeability and the influx of a variety of leukocytes. Hence, this assay tests certain T cell functions. A large increase in skin thickness indicates a strong T cell-mediated immune response. Treatment of birds in the laboratory with irradiation or immunosuppressive drugs to eliminate T lymphocyte function decreases the PHA skin response 50–60% (6–8). The protocol described in this chapter and previously published by Grasman et al. (9) and Grasman and Fox (10) is a modification of the procedures of Fairbrother and Fowles (11) and Schrank et al. (7).

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The PHA skin test has been employed in a large number of studies in the field of “immuno-ecology” – investigations of the interactions between immune function and ecological variables in wild birds. Meta-analyses of these studies have demonstrated the ecological relevance of T cell function as assessed by the PHA skin response (12, 13). Wild birds with lower PHA responses demonstrate reduced abilities to survive and colonize new areas (i.e., found new local populations). Hence, associations between environmental contaminants and reduced PHA responses in wild birds likely are connected to population-level consequences. 1.2. Hemagglutination Assay for Antibody Response to Immunizations with Sheep Red Blood Cells

Agglutination tests measure the production of specific antibodies after the injection of foreign antigens. Blood plasma collected 6–7 days after immunization is serially diluted and incubated with a fixed amount of antigen. Antibodies can form visible complexes with large, insoluble antigens. The reciprocal of the greatest dilution of antibody at which there is a visible complex between antigen and antibody is defined as the titer. A high titer indicates a strong antibody response to the original immunization. Hemagglutination tests use red blood cells from another species, usually sheep, as the antigen. The hemagglutination protocol in this chapter is a modification of the microtiter procedure of Gross and Siegel (14, 15) as previously described by Grasman et al. (9) and Grasman and Fox (10). In laboratory studies, a second SRBC immunization can be administered at some point following the initial blood sampling to stimulate a secondary or memory antibody response. Some species may respond poorly to RBC antigens from sheep, and RBCs from other species may stimulate higher antibody titers. For instance, chukar partridge (Alectoris graeca) RBCs stimulate strong titers in Japanese quail and are more preferable than sheep RBCs for this species (Coturnix coturnix; (8)). The avian anti-SRBC hemagglutination test is very similar in principle to the anti-SRBC splenic plaque-forming assay and the anti-SRBC ELISA used in laboratory rodents, which have been recognized as extremely sensitive immunotoxicity screening assays (16, 17) and have been incorporated into regulatory toxicology programs in the US and Europe (18, 19).

2. Materials 2.1. Phytohemagglutinin Skin Test for T Cell-Mediated Immunity

1. Phytohemagglutinin-P (Sigma L-1668) with 5 mg of active ingredient stored in manufacturer’s bottle with septum at 2–8°C or on wet ice until use (Sigma Chemical Co., St. Louis, MO) (see Note 1). 2. Sterile phosphate buffered saline (PBS). Store in 20 ml aliquots in bottles with septa. Autoclave to sterilize the PBS before sealing the septa.

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3. Disposable syringes and needles (Becton-Dickinson, Franklin Lakes, NJ). One 5 ml syringe with 20–22 gauge needle for preparing PHA. Two 1 ml syringes with 26 gauge intradermal or subcutaneous needles (1/2 or 5/8″) per bird for PHA and PBS injections. 4. Pressure sensitive calipers (micrometer) with a low-tension spring and 5–6 mm diameter contacts (The Dyer Company, Lancaster, PA). 2.2. Hemagglutination Assay for Antibody Response to Immunizations with Sheep Red Blood Cells 2.2.1. Preparation of SRBC Suspensions

1. SRBC from an individual sheep preserved in Alsever’s solution (Colorado Serum Company, Denver, CO) (see Note 2). 2. Stock solution of 0.85% saline for washing SRBC. 3. Disposable supplies for washing SRBCs: two 10 ml disposable syringes with 20–22 gauge needles, one 0.45 mm syringe filter, one sterile 15 ml centrifuge tube, one sterile disposable dropper for removing supernatant. 4. Clinical centrifuge and rotor for 15 ml centrifuge tube.

2.2.2. Immunization Injections

1. Disposable 1 ml syringes. One with a 20–22 gauge needle for preparing final SRBC suspension. One per bird with a 30 gauge needle (1/2″) for SRBC injection. 2. Sterile 0.85% saline. Store in 50 ml aliquots in bottles with septa. Autoclave to sterilize the saline before sealing the septa.

2.2.3. Blood Collection and Processing

1. Sterile Vacutainer tubes or syringes of appropriate size for the bird species being studied (see Note 3). 2. Disposable supplies for preserving plasma: one disposable dropper per bird for removing plasma, cryovials for storing plasma, and centrifuge tubes (only if syringes are used for collecting blood). 3. Clinical centrifuge and rotors for blood tubes. 4. Liquid nitrogen, dry ice, or −80°C freezer for storing blood plasma.

2.2.4. Hemagglutination Microtiter Assay

1. 0.85% saline. 2. 2-mercaptoethanol (Sigma Chemical Co., St. Louis, MO). 3. 96-well microtiter plates with round-bottomed wells.

3.Methods 3.1. Phytohemagglutinin Skin Test for T Cell-Mediated Immunity

1. Prepare a 1 mg/ml solution by adding exactly 5 ml of sterile phosphate buffer saline (PBS) into the PHA vial. 2. Use the 5 ml syringe to remove 5 ml from the PBS bottle. 3. Add the PBS to the PHA bottle through the septum.

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3.1.1. Preparation of PHA and PBS for Injection

4. Thoroughly mix the PHA solution. 5. Option: Syringes for injection can be preloaded to simplify the injection procedure and reduce handling time for birds. For each bird, draw 0.1 ml of PHA through the septum of the bottle into a 1 ml syringe fitted with a 26 gauge intradermal needle. 6. Fill a second syringe with sterile PBS. 7. Flush any air bubbles out of both syringes as they are filled. 8. Store the PHA and PBS solutions and (or) syringes in the dark at 2–8°C or on wet ice until use. 9. Freshly mix PHA-P on each day of injections.

3.1.2. Injections

1. Pluck the feathers from both sides of each wing web, clearing an area minimally large enough to accommodate the contacts of the calipers. 2. Measure and record to the nearest 0.05 mm the thickness of each wing web using the pressure-sensitive calipers. Take two measurements for each wing web. The mean will be used for analysis. 3. Inject 0.1 ml of PHA intra- or sub-dermally into one wing web and 0.1 ml of PBS into the other wing web (see Notes 4 and 5). When injecting a group of birds, alternate between right and left wings for the PHA injection. 4. Record the time of injection, band or identification number, and PHA-injected wing for each bird (see Notes 6–8 for comments on sample sizes, study design, and age of birds).

3.1.3. Measurement and Calculation of Response

1. Twenty-four hours (±3–4 h) after injections, measure and record the thickness of each wing web (see Notes 9–11). 2. Record the time of the measurement. 3. Calculate the mitogen stimulation index (SI) as the increase in wing web thickness caused by PHA minus the increase caused by PBS (see Note 12).

3.2. Hemagglutination Assay for Antibody Response to Immunizations with Sheep Red Cells 3.2.1. Preparation of SRBC Suspensions

1. Using a sterile 10 ml disposable syringe and a 22 gauge or larger needle, draw up approximately 10–11 ml of the 50% SRBC-Alsevers suspension and deliver into a sterile centrifuge tube (see Note 13). 2. Centrifuge the suspension for 5 min at 2,575 × g or until no cells remain in the supernatant (see Note 14). 3. Discard the supernatant using a sterile syringe or disposable pipette. 4. Fill a sterile 10 ml disposable syringe with normal saline, then attach a 0.45 mm syringe filter and 22 gauge or larger needle.

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5. Add saline to the SRBC to the original volume. 6. Re-suspend the cells by gentle mixing. 7. Repeat steps 2–6 two more times, with the exception that at the end of the last wash, add saline to make a 50% SRBC suspension (see Note 15). 3.2.2. Immunization Injections

1. Use a sterile 1% suspension of SRBC in normal saline. Prepare it soon before injections are begun (see Note 16). 2. Thoroughly mix the 50% SRBC suspension. 3. Then use a sterile 1 ml disposable syringe and a 22 gauge or larger bore needle to transfer 1 ml of the SRBC suspension through the septum to the bottle containing 50 ml of sterile normal saline. 4. Keep this suspension on wet ice throughout the injection period. 5. After thoroughly mixing the 1% SRBC suspension, fill a 1 ml disposable syringe with a 30 gauge needle with 0.1 ml of the suspension. 6. Flush any air bubbles out of the syringe. 7. Inject 0.1 ml of the SRBC suspension into the wing (or jugular vein) of the chick (see Notes 5–7 for comments on sample sizes, study design, and age of birds, and Notes 17 and 18 for comments on injections).

3.2.3. Blood Collection and Processing

1. Collect blood for antibody analysis 6 days post injection (if possible). Do not collect blood prior to day 6 (see Notes 19 and 20). 2. Collect a blood sample of at least 1 ml, but more is preferable dependent upon the size of the bird. 3. Use heparin or EDTA as an anticoagulant. 4. Keep blood on ice and in the dark till use. 5. Centrifuge the blood for 5 min at 2,570 × g. 6. Store plasma in 1 ml cryovials in liquid nitrogen or at −80°C.

3.2.4. Hemagglutination Microtiter Assay

1. Dilute the 50% suspension of washed SRBCs (as described above) to prepare a 0.25% suspension with enough volume to fill the total number of plates to be run that day. 2. Add 50 ml of normal saline are added to each well of a 96 well microtiter plate with round-bottomed wells. 3. Add 25 ml of plasma are added to the first well of each row. Each plasma sample should be run in duplicate on different microtiter plates. 4. Conduct serial twofold dilutions of each plasma sample across its row (12 dilutions or wells per sample).

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5. Add 25 ml of a 0.25% SRBC suspension in normal saline is added to each well. 6. After gentle agitation, incubate the plates for 3 h at 37°C. 7. Determine the titers as the log2 of the reciprocal of the highest dilution showing agglutination or spreading of the RBCs. If the amount of antibody is insufficient to cause agglutination, the RBCs form a small, dense, round “button” at the bottom of the well (see Note 21). 8. (see Note 22) To measure IgG activity, add 25 ml of 0.20 M 2-ME in normal saline to the first well of each row of the microtiter plate. 9. Add 25 ml of normal saline to the rest of the wells. 10. Add 25 ml of plasma to the wells containing 2-ME. 11. After 1 h of incubation at 37°C, follow steps 4–7 above to complete the assay. 12. Determine IgG titers after incubation as described above. 13. Calculate IgM titers by subtracting the IgG titers from the total hemagglutination titers.

4. Notes 1. Note that this is the protein form of the PHA extract. Sigma Chemical Co. reports a shelf life of 6 months for the product (personal communication from technical support), although experience suggests that the product maintains its activity for several years if stored properly. 2. The SRBCs are preserved as an approximately 20–25% (by volume) suspension (equal volumes of whole blood and Alsevers). Store at 2–7°C or on wet ice until use for up to 21 days after blood sampling. 3. Generally, maximum volumes of blood samples should not exceed 1% of body mass. Needles should also be of appropriate gauge but should not have a diameter smaller than 26 gauge in order to prevent lysis of avian RBCs. Heparin or EDTA are both appropriate anticoagulants for this assay. Alternatively, serum from clotted blood also is appropriate. 4. If the surface area and (or) thickness of the skin in the wing web is limited, the injection volume can be decreased (e.g., to 0.03 ml), although the mass of PHA (0.1 mg) should be kept the same unless smaller amounts are demonstrated to be sufficient to induce a strong stimulation index in control animals. 5. If the surface area and (or) thickness of the skin in the wing web is limited and reducing injection volume does not work

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well, alternate injection sites include the wattle (in chickens) or interdigitary skin (webbed or nonwebbed feet). Further more, different injection sites can be employed if the test is to be repeated during longer studies. 6. Optimal sample sizes for the PHA skin test and SRBC test vary depending on study design and variability within treatment groups or study sites, but previous studies have shown that n = 20–25/group or site yields excellent statistical power and small confidence intervals. Some groups may be smaller (n = 10) if other groups are larger. 7. As with many immunotoxicity tests, especially those conducted on captive animals, a positive control group treated with an immunosuppressive drug (e.g., corticosterone) can be useful for verifying the sensitivity of the assay in a particular experimental setting. 8. While precocial or semiprecocial birds usually respond to PHA and SRBCs during the first few weeks after hatch, altricial species may respond poorly until further immunological development occurs. 9. The most common time for measuring inflammation is 24 h after injection of PHA. If logistics necessitate earlier of later times, preliminary tests should be conducted to ensure that the response is of sufficient magnitude at that time point. In work with quail, SIs were comparable at 12 and 24 h post injection but decreased by 48 h (Grasman, unpublished data). 10. In some birds, significant edema (fluid) may cause the inflamed site to compress significantly under the tips of the calipers. The use of an engineering-grade calipers with broad (5–6 mm) tips and a low-tension spring can alleviate some of the difficulty of measurement. Furthermore, consistent readings can be facilitated by lightly tapping on the arms of the calipers until compression of the inflamed area stops. Averaging two independent measurements enhances accuracy and reduces variability. 11. Consistency also is promoted by having a single investigator make all measurements. However, multiple investigators can train together to use a similar measurement technique. 12. The placebo injection of PBS in the alternate wing web is important because some individuals display greater swelling in both wing webs compared to other individuals. Subtraction of the swelling caused by PBS alone accounts for this tendency. 13. Because SRBC antigens can vary from sheep to sheep, the protocol recommends using SRBCs from the same individual sheep or both immunization and the hemagglutination plate assay. However, SRBCs from different sheep or composite samples from multiple sheep also can be used.

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14. SRBCs should be in good condition for both immunizations and the hemagglutination plate assay. Lysis of SRBCs can be caused by prolonged storage (more than 21 days in Alsevers solution), improper storage temperatures, rough handling during processing or transport, and improper saline concentrations. SRBC condition can be assessed by examining the color of the supernatant following washing. A clear red or pink supernatant suggests that lysis has occurred. An opaque red or pink supernatant may reflect suspended intact cells or lysis, and should be centrifuged further. If no lysis has occurred, the supernatant should be pale yellow if it contains Alsevers and plasma following the first wash, or colorless if it contains saline solution following later washes. 15. The same syringe and syringe filter can be used each time, but make sure that saline is not drawn into the syringe through the filter. The saline should be filtered only as it is pushed out of the syringe. The final 50% suspension can be stored at 2–7°C or on wet ice for several hours until used for immunizations or the hemagglutination plate assay. 16. While some avian protocols may use larger numbers of SRBCs for immunization, the smaller numbers recommended here have been shown to induce a strong response and, more importantly, a greater difference between controls and immunosuppressed birds. 17. I.V. injection is recommended over i.p. because the latter route might damage abdominal air sacs in birds. 18. If injected properly, the dark blood in the vein often will be pushed aside momentarily by the lighter SRBC suspension. If a light pink bubble forms under the skin, then the injection probably missed the vein. Record the band or identification number of the injected chick. Be sure that the SRBC suspension in the syringe does not begin to settle if there is a delay between loading the syringe and injecting the bird. If so, gently rock the syringe to mix the suspension. 19. The timing of blood sampling following immunization is critical, especially for primary responses that are often of lower magnitude and shorter duration than secondary or memory responses. Like in rodents, a 6–7 day interval between immunization and blood sampling works well for primary responses in many bird species. Sampling earlier or later may miss the peak titer. 20. Peak antibody activity usually occurs 6 days after the primary antigen injection (see Note 19). Blood for antibody analysis should be collected 6 days post injection if possible, but antibody activity may remain high for several days if poor weather or logistics prevent collection on day 6. Blood should not be

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collected before 6 days post injection because antibody titers will not yet be at their peak. 21. The titer, as measured by the hemagglutination plate assay, depends not only on the concentration of antibody in the plasma sample but also on the concentration of SRBCs added to the plate. If initial testing shows titers to be too low, a smaller number of SRBCs can be used to increase the titers. Likewise, if titers are too high, smaller numbers of SRBCs can be used. Note that this means that titer values cannot be compared between studies unless the same SRBC concentrations were used. 22. The above procedures describe the basic hemagglutination test, which measures the activity of total hemagglutinating antibodies. Incubation of plasma in 2-mercaptoethanol (2-ME) dissociates IgM and leaves IgG intact. The 2-MEresistant antibodies are equivalent to IgG while the 2-ME-sensitive antibodies are equivalent to IgM.

References 1. Friend M, Trainer DO (1970) Polychlorinated biphenyl: Interaction with duck hepatitis virus. Science 170:1314–1316 2. Fairbrother A, Smits J, Grasman KA (2004) Avian immunotoxicology. J Toxicol Environ Health B 7:1–33 3. Grasman KA (2002) Assessing immunological function in toxicological studies of avian wildlife. Integrative Comp Biol 42:34–42 4. Finkelstein M, Grasman KA, Croll DA, Tershy B, Smith DR (2003) Immune function of cryopreserved avian peripheral white blood cells: Potential biomarkers of contaminant effects in wild birds. Arch Environ Contam Toxicol 44:502–509 5. Lavoie ET, Grasman KA (2005) Isolation, cryopreservation, and mitogenesis of peripheral blood lymphocytes from chickens (Gallus domesticus) and wild herring gulls (Larus argentatus). Arch Environ Contam Toxicol 48:552–558 6. Edelman AS, Sanchez PL, Robinson ME, Hochwald GM, Throbecke GJ (1986) Primary and secondary wattle swelling response to phytohemagglutinin as a measure of immunocompetence in chicks. Avian Dis 30:105–111 7. Schrank CS, Cook ME, Hansen WR (1990) Immune response of mallard ducks treated with immunosuppressive agents: Antibody response to erythrocytes and in vivo response to phytohemagglutinin-P. J Wildl Dis 26:307–315

8. Grasman KA, Scanlon PF (1995) Effects of acute lead ingestion on antibody- and T cellmediated immune responses in Japanese quail. Arch Environ Contam Toxicol 28:161–167 9. Grasman KA, Fox GA, Scanlon PF, Ludwig JP (1996) Organochlorine-associated immunosuppression in prefledgling Caspian terns and herring gulls from the Great Lakes: an ecoepidemiological study. Environ Health Perspect 104 (Supplement):829–842 10. Grasman KA, Fox GA (2001) Associations between altered immune function and organochlorine contamination in young Caspian terns (Sterna caspia) of the Great Lakes, 1997–99. Ecotoxicology 10:101–114 11. Fairbrother A, Fowles J (1990) Subchronic effects of sodium selenite and selenomethionine on several immune functions in mallards. Arch Environ Contam Toxicol 19:836–844 12. Moeller AP, Cassey P (2004) On the relationship between T-cell mediated immunity in bird species and the establishment success of introduced populations. J Animal Ecol 73: 1035–1042 13. Moeller AP, Saino N (2004) Immune response and survival. Oikos 104:299–304 14. Gross WB, Siegel PB (1980) Effects of early environmental stresses on chicken body weight, antibody responses to RBC antigens, feeding efficiency, and response to fasting. Avian Dis 24:569–579

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15. Gross WB, Siegel PB (1981) Long-term exposure of chickens to three levels of social stress. Avian Dis 25:312–325 16. Luster MI, Portier C, Pait DG, White KL Jr, Gennings C, Munson AE, Rosenthal GJ (1992) Risk assessment in immunotoxicology. I. Sensitivity and predictability of immune tests. Fund Appl Toxicol 18:200–210 17. Luster MI, Portier C, Pait DG, Rosenthal GJ, Germolec DR, Corsini E, Blaylock BL, Pollock P, Kouchi Y, Craig W, White KL,

Munson AE, Comment CE (1993) Risk assessment in immunotoxicology. II. Relationships between immune and host resistance tests. Fund Appl Toxicol 221:71–82 18. ICICIS Group Investigators (1998) Report of validation of assessment of direct immunotoxicity in the rat. Toxicology 125:183–201 19. US Environmental Protection Agency (1998) Health Effects Test Guidelines OPPTS 870.7800 Immunotoxicity. USEPA Washington, D.C

Part VII In Vitro Alternatives

Chapter 26 In Vitro Testing for Direct Immunotoxicity: State of the Art D.P.K. Lankveld, H. Van Loveren, K.A. Baken, and R.J. Vandebriel Abstract Immunotoxicity is defined as the toxicological effects of xenobiotics including pharmaceuticals on the functioning of the immune system and can be induced in either direct or indirect ways. Direct immunotoxicity is caused by the effects of chemicals on the immune system, leading to immunosuppression and subsequently to reduced resistance to infectious diseases or certain forms of nongenotoxic carcinogenicity. In vitro testing has several advantages over in vivo testing, such as detailed mechanistic understanding, species extrapolation (parallelogram approach), and reduction, refinement, and replacement of animal experiments. In vitro testing for direct immunotoxicity can be done in a two-tiered approach, the first tier measuring myelotoxicity. If this type of toxicity is apparent, the compound can be designated immunotoxic. If not, the compound is tested for lymphotoxicity (second tier). Several in vitro assays for lymphotoxicity exist, each comprising specific functions of the immune system (cytokine production, cell proliferation, cytotoxic T-cell activity, natural killer cell activity, antibody production, and dendritic cell maturation). A brief description of each assay is provided. Only one assay, the human whole blood cytokine release assay, has undergone formal prevalidation, while another one, the lymphocyte proliferation assay, is progressing towards that phase. Progress in in vitro testing for direct immunotoxicity includes prevalidation of existing assays and selection of the assay (or combination of assays) that performs best. To avoid inter-species extrapolation, assays should preferably use human cells. Furthermore, the use of whole blood has the advantage of comprising multiple cell types in their natural proportion and environment. The so-called “omics” techniques provide additional mechanistic understanding and hold promise for the characterization of classes of compounds and prediction of specific toxic effects. Technical innovations such as high-content screening and high-throughput analysis will greatly expand the opportunities for in vitro testing. Key words: Immunotoxicity, In vitro, Mechanism, Parallelogram, 3R, Myelotoxicity, Lymphotoxicity, Whole blood, T cell, Proliferation, Cytokine, Cytotoxic T cell, Natural killer cell, Antibody, Dendritic cell, Genomics, Proteomics

1. Introduction Immunotoxicity is defined as the toxicological effects of xenobiotics, including pharmaceuticals, on the functioning of the immune system and can be induced in either direct or indirect ways. R.R. Dietert (ed.), Immunotoxicity Testing: Methods and Protocols, Methods in Molecular Biology, vol. 598 DOI 10.1007/978-1-60761-401-2_26, © Humana Press, a part of Springer Science + Business Media, LLC 2010

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Direct immunotoxicity is caused by the effect of chemicals on the immune system leading to immunosuppression and subsequently to reduced resistance to infectious diseases or certain forms of nongenotoxic carcinogenicity when the suppressed immune system is challenged. Indirect immunotoxicity is caused by specific immune responses to the compounds themselves or to self-antigens altered by these compounds and is mainly manifested as allergy or autoimmunity (1). Since many compounds induce immunotoxicity and, hence, adversely affect the health of exposed individuals, the evaluation of immunotoxicity is nowadays incorporated into the overall toxicological investigation of compounds. So far immunotoxicity is mainly investigated in animal models. However, these in vivo studies are expensive, require a considerable number of animals, and raise important ethical concerns. For this reason European policy is promoting alternative testing methods and assessment strategies to the use of laboratory animals in order to reduce and, whenever possible, replace animals employed for scientific studies (2). A workshop, hosted by the European Centre for the Validation of Alternative Methods (ECVAM) in 2003 (3), reported on the state-of-the-art of in vitro systems for evaluating immunotoxicity. Based on recommendations from this workshop, prevalidation studies for detection of immunosuppressive activity in vitro were initiated and revealed promising results (4). Although in vitro testing is more straightforward for direct immunotoxicants, at present the development and validation of in vitro immunotoxicity assays focuses primarily on chemical sensitization, since this field of toxicity testing uses a high number of animals when compared to the other fields of toxicity testing. With respect to the other types of immunotoxicity, which are immunostimulation, autoimmunity, and developmental immunotoxicity, little or no efforts in the area of in vitro testing have been made as yet. These processes are not only complex involving delicately balanced interactions encompassing many tissues, they are often badly understood, and cell systems that may mimic some of the processes involved are difficult to devise. In this report an overview of the presently available in vitro assays for direct immunotoxicity testing will be given. In addition, the limitations of the development and use of in vitro testing systems in general and for testing direct immunotoxicity per se will be discussed.

2. In Vitro Testing for Direct Immunotoxicity

In animal models, the direct immunotoxic potential of compounds is investigated according to a tiered approach. In the first tier, general immunotoxicity of the test chemical is assessed by

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integrated analyses of changes in lymphoid tissue and immune cell populations in combination with the standard toxicity endpoints (differential cell counting, lymphoid organ weights, histology, and bone-marrow cellularity) and knowledge of the health status of the animals. When this initial screening reveals any signals for immunotoxicity, a second tier study is performed in which the functionality of the immune system is studied in more depth (5). In line with this approach, the ECVAM workshop has recommended the use of a flow chart/decision tree approach for the in vitro detection of immunotoxicants. Initially, a set of in vitro tests should evaluate whether or not a compound is immunotoxic. When a potential immunotoxicant has been detected, the pre-screening assays should be followed by more detailed in vitro mechanistic assays (3). A proposal for such a decision tree is shown in Fig. 26.1. 2.1. Myelotoxicity

The immune system is a complex, multicomponent system that comprises lymphoid organs and interacting, specialized cells located all over the body. The major lymphoid organs are the bone marrow, spleen, thymus, lymph nodes, and localized areas of lymphoid tissue in the respiratory and intestinal tract, the bronchusassociated lymphoid tissue (BALT) and Peyer’s patches, respectively. The major cell type of the immune system is the lymphocyte, although accessory immune cells like macrophages, natural killer (NK) cells, neutrophils, eosinophils, basophils, dendritic cells (DC), keratinocytes, and epithelial cells play pivotal roles in the immune response as well. Compound Myelotoxicity CFU-GM assay

YES Immunotoxic

NO

Lymphotoxicity

Human whole-blood cytokine release assay Lymphocyte proliferation assay Mixed lymphocyte reaction Cytotoxic T-lymphocyte assay Natural Killer cell assay T cell-dependent antibody response Dendritic Cell maturation Fluorescent Cell Chip

Fig. 26.1. Proposed decision tree for in vitro testing for direct immunotoxicity.

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Since all immune cells arise from pluripotent hematopoietic stem cells in the bone marrow, the pre-screening for immunotoxicity should be started with the evaluation of myelotoxicity (Fig. 26.1). Compounds which are capable of damaging or destroying the bone marrow will most likely express an immunotoxic effect, since the effectors of the immune system itself will no longer be available. Thus, if a compound is myelotoxic, it will be de facto an immunotoxicant and in that case no additional evaluation of other endpoints is required (3). The classical bioassay for hematopoietic stem cell activity consists of measurement of colony-forming units in the spleen of irradiated mice after injection of bone marrow cells (5). At present, a scientifically validated human and murine in vitro colony forming units-granulocyte/macrophage (CFU-GM) assay is available for evaluating the potential myelotoxicity of xenobiotics, quantified by the number of surviving bone marrow progenitors as a function of exposure level (6, 7). Next to granulocytes/macrophages, the adverse effects of xenobiotics can be studied on other specific hematopoietic cell types of interest, such as erythrocytes and thrombocytes by using the burst forming units-erythroid (BFU-E) assay and the colony forming units-megakaryocyte (CFU-MK) assay. Nowadays, several in vitro bone marrow systems are commercially available, although they have the disadvantage of being rather expensive, especially when human cells are used. However, prices may decline as, with respect to human cells, not only bone marrow cells but also umbilical cord blood cells have proven to be a reliable source of human myeloid progenitor cells. In a study specifically directed to compare these cell types, results demonstrated that the source of progenitors did not affect determination of the inhibitory concentration (IC) value of six drugs tested (6). Future refinement and optimization of the model might result in application of the assay in 96-well plates, so that high throughput screening (HTS) of myelotoxic compounds can be performed (6, 7). 2.2. Lymphotoxicity

Compounds that do not possess any myelotoxic potential may still be lymphotoxic and thus contribute to immunosuppression. Hence, according to the decision tree approach (Fig. 26.1), the second step in the evaluation for direct immunotoxicity should be testing for lymphotoxicity. As part of this, a set of functional in vitro assays are available. Initially, viability of lymphocytes has to be determined since noncytotoxic concentrations of the test compound have to be used for evaluating the basic functionality of the lymphocytes. Viability can be assessed by various validated tests, from which the 3-(4, 5-dimethylthiazol-2-yl)-2, 5 diphenyltetrazolium bromide (MTT), lactate dehydrogenase (LDH) and trypan blue dye exclusion assays are most frequently used. While the MTT assay measures metabolic activity, the other two

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measure membrane integrity. When viability is higher than 80%, basic functionality can be determined by measuring cytokine production, T and B cell proliferation, cytotoxic T lymphocyte activity, NK cell activity, antibody production, and DC maturation. In the following section, the ins-and-outs of the assays measuring this functionality will be discussed separately with respect to their value in evaluating direct immunotoxicity in vitro (8). 2.2.1. Human Whole-Blood Cytokine Release Assay

Cytokines are produced in the first steps of the immune response, indicating that quantitative alterations in cytokine levels can be used as a measure of immunomodulation. Of the cytokine assessment designs currently available, the whole blood cytokine release assay is the only assay that has been prevalidated for use in in vitro immunotoxicology. The principle of the assay, described by Langezaal et al. (9), is based on the well-known human whole-blood method for pyrogen testing (10). In brief, human blood is treated with lipopolysaccharide (LPS) or staphylococcal enterotoxin B (SEB) following which monocytes and Th2-lymphocytes produce interleukin- (IL-) 1-b and IL-4, respectively. After incubation for 40 h in the presence or absence of immunotoxic and non immunotoxic test compounds, IL-1b and IL-4 production is quantified in the supernatant, and IC50 (i.e., the 50% inhibitory concentration) and SC4 (i.e., the fourfold stimulating concentration) values are calculated to establish immunotoxic potency (9). The human whole-blood cytokine release assay offers the major advantage that the species differences between humans and animals are avoided. Moreover, the culture technique of the assay is extremely simple. It avoids preparation and culture artifacts since human primary cells are employed in their physiological proportions and environment. The outcome of the whole blood cytokine release assay was found to be more homogenous between different donors than the respective models using isolated cells. It is also advantageous that the same model can be used to monitor immune functions both ex vivo and in vitro. The recent development of cryo-preserved human whole blood overcomes limitations such as blood availability, risks of infections, or abnormal responses and the pooling of donors offers the opportunity of standardized materials. Furthermore, effects of different blood cell populations can be tested in the whole blood cytokine release assay, since depending upon the type of stimulus, human blood cells release different cytokine patterns originating from several blood cell populations. Stimulation with LPS results in the release of IL-1b, IL-6, and tumor necrosis factor- (TNF-) a by monocytes, whereas stimulation with SEB and prolonging the incubation period from 48 to 72 h results in the release of various cytokines such as IL-2, IL-4, IL-13, and interferon- (IFN-) g.

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IL-4 is chosen as the lymphokine read-out, since it is not produced by monocytes and reflects activation of B cells by Th2 cells, thus allowing assessment of the interplay of two major lymphocyte populations. IL-1b and IL-4 are suitable endpoints for toxicity measurements against monocytes and lymphocytes, respectively (9). Combination of the endpoints resulted in a sensitivity of 67% and a specificity of 100%. However, due to the highly pleiotropic and redundant nature of cytokines, in which a single function may be affected by multiple cytokines simultaneously, the ECVAM Workshop now recommends that the measurement of cytokine levels should not be restricted to IL-1b and IL-4, but should be extended to a broad panel of cytokines. Next to this, the impact of the assay will also increase when it is extended to other cell types and to the measurement of more general mediators such as eicosanoids, nitric oxide, or degranulation products (3). The assay is transferable to another laboratory, since a high correlation of IC50 values, established in two laboratories, was obtained for several test compounds (9). The differences found in the amounts of cytokine production between donors, were ascribed to factors such as the number of whole blood cells, age, number of cytokine receptors, and polymorphisms. However, these inter-donor differences had a minimal influence on the calculated IC50 values and did not impair testing, indicating that the assay is robust. The interindividual variation in leukocyte numbers is a major concern of the whole blood cytokine release assay. However, in healthy donors the normal range of leukocyte numbers is fairly narrow, and by using a differential blood cell count, responses can be normalized to the number of a given leukocyte population in the experiment (3). In the test described by Langezaal et al. (9), lymphocytes were generally more sensitive for immunosuppression than were the monocytes. This is probably because lymphocytes produce and release their cytokines later than do monocytes, and because lymphocytes proliferate in vitro after stimulation with SEB giving toxicants more time to exert their toxicity against the lymphocytes. However, Langezaal et al. (9) also found that some compounds suppressed IL-1b release more than they suppressed IL-4 release, showing that they possess a greater specificity towards monocytes. By calculating IC50 values against IL-1b and IL-4 release, information can be obtained about the specificity of compounds in suppressing monocytes or lymphocytes and, thus, about the potency of the compound in suppressing the specific immune function. To examine the in vivo relevance of the outcome of the whole blood cytokine release assay, it was assumed that compounds clinically known to be immunosuppressive should exhibit IC50 values below human therapeutic plasma concentrations. It was found

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that the plasma concentrations of all nonimmunotoxic compounds tested were lower than the corresponding IC50 value in vitro, indicating that the assay brings only a few false positive results. Of the immunosuppressive compounds, three out of six indeed showed an IC50 value below the human plasma concentration, whereas the reverse was true for the other three compounds. The latter discrepancy might be due to the fact that these compounds were either alkylating agents or compounds that require metabolism to be immunotoxic. It has been shown that these kinds of agents might be identified as being nonimmunosuppressive when using the whole blood cytokine release assay. Alkylating agents in vivo mainly act on proliferating T- and B-lymphocytes and the formation of antibodies. Since these processes are no endpoints in the whole blood cytokine release assay, immunotoxicity of alkylating agents cannot be identified by using this assay. Similarly, compounds that rely on metabolic activation before they are immunotoxic cannot be identified by the whole blood cytokine release assay, since metabolism is predominantly lacking in this assay. Preliminary experiments revealed that introducing metabolism into the assay by adding microsomes showed no interference with the blood incubation. On the other hand, co-cultures of genetically engineered cells expressing cytochrome P450 enzymes interfered with cytokine release by monocytes and lymphocytes (9). For measuring cytokines and their receptors a wide range of assay systems are available, such as Luminex, ELISA, flow cytometry, and molecular biology techniques such as RT-PCR (3). 2.2.2. Lymphocyte Proliferation Assay

The adaptive immune response requires the proliferation of lymphocytes, so inadequate proliferation may be an indicator of immunosuppression. Lymphocytes proliferate in response to mitogens or antigens. For T cells, the mitogens concanavalin A (Con A) or phytohaemagglutinin (PHA) or the combination of anti-CD3 (stimulation) and anti-CD28 (co-stimulation) can be used. While anti-CD3 mimics T cell receptor activation, anti-CD28 enhances proliferation, cytokine production, and survival of T cells. B cell proliferation is stimulated by the mitogens LPS or S. typhimurium mitogen (STM) or the combination of anti-CD40 and IL-4 (5). In some assays, pokeweed mitogen is used for the induction of lymphocyte proliferation, but this mitogen lacks specificity since the proliferation of both T and B cells is stimulated (5). Cell proliferation is most widely measured by the incorporation of radiolabeled thymidine (3H-thymidine) into cellular DNA. In rodent studies, the proliferation assays are conducted on isolated splenocytes, thymocytes and lymph node cells, while human assays are conducted on lymphocytes isolated from peripheral blood. Using a human assay offers the advantage that species differences between humans and animals are avoided. However, in

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contrast to lymphocytes obtained from uniform rodent strains, human lymphocytes lack uniformity, resulting in reduced reproducibility. In general, the lymphocyte proliferation assay has distinct advantages including relatively low cost, readily available source material and high feasibility as the method has been extensively reported in scientific literature (3). Recently, the lymphocyte proliferation assay has confirmed the immunotoxic effect of six compounds that have known immunosuppressive activity in vivo, in human, rat, and murine cells (4). Unfortunately, in a follow-up study (Vandebriel et al., in preparation), human peripheral blood mononuclear cells (PBMC) and murine splenocytes showed a high inter- and intra-laboratory variability resulting in a rather poor predictivity. This disappointing outcome may partially be explained by the fact that the dataset was incomplete as the participating laboratories decided not to repeat flawed or suboptimal experiments in order to evaluate the robustness of the test. A follow-up is needed and pre-optimization of the test in each laboratory is required to increase the predictivity and robustness of the assay. For mouse and rat splenocytes, the T cell mitogen Con A was found superior over the B cell mitogen LPS. This may in part be due to the fact that immunosuppression mainly affects T cells, suggesting a clearer effect on these cells than on B cells. The most obvious disadvantage of the lymphocyte proliferation assay is the requirement of radioactive isotopes. In recent years, several nonradioactive alternative test methods have been developed including flow cytometric assays measuring BrdU incorporation or ATP content and colorimetric assays using MTT, trypan blue or Alamar Blue. The alternatives have proven to be sensitive, rapid, easy to perform, and applicable to HTS. However, their dynamic range is much smaller than that of 3H-thymidine, leaving the latter the technique of choice for measuring lymphocyte proliferation. 2.2.3. Mixed Lymphocyte Reaction

The mixed lymphocyte reaction (MLR) is, next to the lymphocyte proliferation assay, an in vitro assay that can be used for the evaluation of immunotoxicity. The principle of the test is based on the ability of T lymphocytes to proliferate in response to allogeneic cells. In short, suspensions of lymphocytes from spleen or lymph nodes are co-cultured with allogeneic stimulator cells for 4 days, after which radiolabeled thymidine (3H- thymidine) is added and radioactivity is counted within 24 h. The net MLR response is the difference in cell proliferation between lymphocytes that were co-cultured with the allogeneic cells and those co-cultured with syngeneic cells. For the murine MLR, splenocytes are used as a source for both responder and stimulator cells, whereas in the rat MLR, lymph node lymphocytes are used as responder and stimulator cells,

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since the use of rat splenocytes has resulted in poor lymphocyte proliferation (11). In inbred populations, the activating capacity of the stimulator cells is mediated by the foreign major histocompatibility complex (MHC) class I or class II molecules expressed on these cells, while in noninbred populations, a pool of allogeneic cells can be used as stimulators. In order to inhibit the proliferation of stimulator cells, these cells are g-irradiated or treated with mitomycin C before incubation with the responder cells. Priming of the responder cells is not necessary, since a sufficiently high number of T cells respond to the stimulator cells (5). Although the MLR is potentially useful for predicting chemically induced adverse immune effects, the assay has not been standardized and validated across laboratories and its concordance with host resistance assays or its predictive value for immune effects is presently not known (3). 2.2.4. Cytotoxic T-Lymphocyte Assay

The cytotoxic T-lymphocyte (CTL) assay is a continuation of the MLR response in which the T lymphocytes further differentiate into cytotoxic effector cells. CTL are a population of CD8+ lymphocytes characterized by specific cytotoxicity for target cells harboring intracellular antigens, such as tumor cells and virally infected cells. Measuring CTL function contributes to the evaluation of the effect of various test compounds on cell-mediated immunity, as well as serving as the basis of more detailed mechanistic immunotoxicology studies (12). Naïve CTL’s require a sensitization period in which CTL precursors undergo proliferation and differentiation into cytotoxic effector cells. Since many cell types and mediators are involved in this sensitization process, assessment of CTL function may not only reveal deficits in the effector phase of the immune response, but also in cellular activation and regulatory pathways (3, 12). In the conventional assay, splenocytes are removed from rodents exposed to the test compound and co-cultured with P815 mastocytoma cells. After 5 days, splenocytes are harvested and added to fresh 51Cr-radiolabeled P815 cells following which cytotoxicity is determined by measuring 51Cr release into the supernatant. P815 cells are used as sensitizer and target cells, since this cell line has proven to be allogeneic and more convenient than primary allogeneic cells such as lymphocytes. A culture period of 5 days is necessary for the T cells to differentiate into cytotoxic effector cells. For in vivo testing, this extended period of culture enables the spleen to recover from any adverse effect mediated by the test compound and thus makes the assay less useful in assessing immunotoxicity (5, 12). Following this conventional assay, House and Thomas (12) developed a more simplified version in which, not the animals, but their isolated splenocytes were exposed to the test compound in vitro. Evaluation of the splenocyte CTL assay revealed consistent

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control values, limited assay-to-assay variability, low background toxicity, and relatively high sensitivity to modulation following exposure to xenobiotics. Next to splenocytes, peripheral blood lymphocytes from animals can be used for the in vitro testing of CTL function. By using peripheral blood, the number of animals can be reduced and repeated testing of the same animal is enabled. Moreover, this method offers the ability for testing CTL function in humans, thereby avoiding species differences. For assessing human CTL function, the Jurkat leukemic cell line can be used as stimulating and target cell line. 2.2.5. Natural Killer Cell Assay

Natural Killer (NK) cells are involved in the nonspecific defense system against tumors, microbe-infected cells, and MHC Class I mismatched cells. For this reason, NK cell activity is recommended as a critical endpoint to measure the immunotoxicity of xenobiotics including environmental chemicals and pharmaceuticals (13). In humans and rodents, cells with NK activity can be identified by morphological techniques and functional assays. Most of the cells that show NK activity are nonadherent, nonphagocytic lymphocytes that are morphologically associated with large granular lymphocytes and can be enumerated by measuring the expression of surface markers (mainly CD56) by flow cytometry (3, 5). The classical assay to measure NK cell activity is the chromium-51 (51Cr) release assay. In brief, effector cells are collected from the spleen (spleen cells depleted of red blood cells) or peripheral blood (mononuclear cells) and mixed with 51Cr-labeled target cells, such as YAC-1 lymphoma cells for rodents or K562 erythroleukemia cells for humans. Following 4 h of incubation, cytolytic activity of the effector cells is determined by measuring 51Cr release into the culture supernatant as a result of lysis of the target cells (3, 5, 13). Although this assay is regarded as the “gold standard” cytotoxicity assay, it has the disadvantage that 51Cr is a volatile health hazard, which must continuously be replaced because of its short half-life. Additionally, there is a high spontaneous release of 51Cr, and radioisotopes per se are known to have detrimental effects upon cell function. Moreover, the inter-laboratory variability of the assay is also of potential concern (13, 14). To overcome some of these drawbacks, new flow cytometrybased, nonradioactive methods for measuring NK cell activity have been developed (14, 15). Briefly, effector cells are mixed with fluorochrome-labeled target cells to enable the discrimination of target from effector cells. After 18 h of incubation, a DNA-labeling fluorochrome is added to estimate the proportion of dead target cells. Subsequently, both fluorochromes can be distinguished by flow cytometry, and NK cell activity can be calculated (13, 15). Hence, the flow cytometric assay avoids the

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problems associated with the use of radioactivity and is less time consuming in comparison to the 51Cr-release assay, less expensive, and more amenable to standardization, since it is included in the proficiency testing profile of many laboratories (13, 14). In addition, the flow cytometric assay requires only a minimal number of effector cells (less than 0.5 ml peripheral blood), thus enabling repeated testing of the same animal or human being (15). The relatively high spontaneous release of fluorescent dye from the target cells can be omitted by using carboxy-fluorescein succinimidyl ester (CFSE) as primary fluorochrome, since CFSE has proven to bind covalently to cell proteins (13). Many cells with high NK activity are found in spleen and peripheral blood, whereas lymph nodes have less NK activity and thymus and bone marrow show only marginal activity (5). For the in vitro testing of NK cell activity, both spleen and peripheral blood cells can be used, although blood cells are indicated to be more sensitive than spleen cells (15). Using peripheral blood will limit the use of experimental animals and offers the additional advantage of repeated testing of the same animal. By using human peripheral blood, animals can even be replaced and species differences can be avoided. Since peripheral blood cells do not fully retain their activity after cryo-preservation, the use of fresh peripheral blood is recommended (13). In human peripheral blood, NK cells account for 7–41% of the circulating effector lymphocytes (3). There is not a unique phenotype characteristic of NK cells. So far, at least 48 different subsets have been identified and no definite correlation between immunophenotype and NK cell activity has been found. The NK cell activity is exquisitely sensitive to modulation by toxic substances, but changes in NK activity are rather nonspecific and not often related to pathological conditions of the immune system. In contrast, they have been associated with various conditions of unknown etiology (16). The NK assay has been used thoroughly to monitor in vivo exposure. These days, the predictive potential of the assay after in vitro exposure is under intensive investigation (3). Besides measuring NK activity, NK cells can also be used for determining the cytokine expression following the exposure to test compounds. This is because NK cells produce a wide range of cytokines, such as TNF-a, TNF-b, IFN-a, IFN-b, IFN-g, granulocytemacrophage colony-stimulating factor, and IL-3 (3, 17). 2.2.6. T Cell-Dependent Antibody Response

In animals, production of a T cell-dependent antibody response is considered to be the “gold standard.” However, there are currently no good systems for the in vitro T cell-dependent antibody production using human cells. Development of human in vitro systems will require optimization of the antigen, culture conditions, and assay endpoints. In addition, there is some concern whether

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a primary immune response can actually be induced in human peripheral blood lymphocytes. One potential starting point would be an in vitro immunization culture system based on the MishellDutton assay, an in vitro model for evaluating the humoral immune response of mouse splenocytes to sheep red blood cells. However, in itself the assay is still not optimal for this use due to significant variability in the results found between laboratories. Due to the high predictivity of human immunotoxicants provided by the in vivo antibody induction assay, as well as the possibility of bridging between in vivo and in vitro potentially afforded by this system, it is recommended that development of this in vitro system is given the highest priority (3). In fact, using the Mishell-Dutton assay a correct prediction of six out of seven immunosuppressive compounds and all four nonimmunotoxic compounds was made (18). 2.2.7. Dendritic Cell Maturation

Dendritic Cells (DC) have a pivotal role in the immune response in that they are the only cell type which are able to activate naïve T cells, thereby inducing adaptive immune responses. In general, they can be found in two maturation stages, the immature stage in which they are exquisitely capable of antigen uptake, and the mature state in which they are extremely good at antigen presentation (19). This maturation process also occurs in case of contact sensitization where Langerhans cells, a specialized type of DC, after uptake of haptenized proteins (as immature DC) migrate from the skin to the regional lymph node where they present the hapten-protein complex to T cells (as mature DC). Thus, in vitro models for identifying skin sensitizers have used the capacity to induce DC maturation as a means to identify sensitizers. Immature DC are cultured from human PBMC or DC-like cell lines, and DC maturation is established on the basis of expression of surface markers (such as CD54 and CD86) and cytokine production. In contrast to the plethora of studies which identify sensitizers on the basis of DC maturation, very few studies addressed the direct immunotoxic effects of compounds on DC. Recently, suppressive effects of anti-inflammatory agents on nickel-induced DC maturation were shown providing a means to establish immunosuppressive activity of compounds on DC maturation (20).

2.2.8.Fluorescent Cell Chip

A new in vitro system for measuring cytokine expression has been developed. This system named “fluorescent cell chip” is based on a number of cell lines derived from different origins (T cells, mast cells, and macrophage-monocytes) that each are transfected with various cytokine reporter cell constructs and that regulate the expression of a fluorescent protein in a similar way as they regulate expression of cytokines (21, 22). In the prototype “cell chip,” only one cell type, the murine thymoma cell line EL-4,

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was transfected with the regulatory sequences of IL-2, IL-4, IL-10, IFN-g, and b-actin fused to the gene for enhanced green fluorescent protein (EGFP) with one cell line per cytokine (22). A T cell line was chosen since the exposure effects of immunotoxic compounds often comprise alterations in T cell activity. The four cytokines were selected based on the fact that together they indicate effects on the size of T cell response and the balance between Th1 and Th2 response: IL-2 plays a pivotal role in the growth and function of T cells, IFN-g is produced by Th1 only, IL-4 is produced by Th2 only and IL-10 suppresses T cell function and plays a key role in the differentiation and function of regulatory T cells that limit immune responses. The T cell line transfected with b-actin is used as a control for specificity (22). It was chosen to set up the Fluorescent Cell Chip using murine cells in order to compare the results to the extensive in vivo mouse database. Further development will include the use of human cells. Following the exposure to several compounds, EGFP-mediated fluorescence can be measured by flow cytometry or in a 96-well format and changes in fluorescence intensity represent changes in cytokine expression and correlate to cytokine levels. When the cells are simultaneously stained for viability, fluorescence can be measured in viable cells only (21, 22). This prototype in vitro immunotoxicity screening has several important advantages. The major advantage is the possibility to apply this test in HTS. Other methods to measure cytokine expression, such as RT-PCR, Luminex and ELISA are more timeconsuming. Fluorescence can be detected by several methods such as Fluostar® plate reader, fluorescence microscopy, and flow cytometry. The latter has the advantage of concurrent assessment of viability, the possibility to measure fluorescence intensity per cell, and possible effects on cell shape and size (22). The fluorescence microscopy-based assay could be particularly useful when the amount of tested compound is limited because the readout can be obtained using small numbers of cells and a small volume of incubation media (23). The Fluostar® plate reader has the advantage of testing a large number of samples in a limited amount of time, thus enabling HTS. One limitation of the prototype “cell chip” is the absence of antigen presenting cells. For a substance to give an effect in this test system, it must interact directly with the cells. Immunosuppressive compounds often exert their effects directly on T cells, suggesting that the prototype “cell chip,” which consists of only T cells, is a suitable system to evaluate the immunosuppressive potential. Using the prototype “cell chip,” the immunomodulatory potential could be detected for twelve out of thirteen model substances (22). Subsequently, the experimental protocol was modified to a 96 well-plate format, and the assay was employed for testing 46 substances (24). The tested compounds reproducibly

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generated compound-specific patterns of changes in fluorescence that correlated with the available in vivo immunotoxicity data and allowed for hierarchical clustering of their expected activities based on pattern similarity analysis. In addition to cytokines, the expression of other mediators may be used as an endpoint for in vitro immunotoxicity screening. As part of this, BW5147.3 murine thymoma cells expressing enhanced cyan fluorescent protein (ECFP; an enhanced cyan variant of GFP) under the control of the murine c-fos promoter was tested for determining the level of cellular stress induced by several substances (23). The level of ECFP fluorescence was in good agreement with the level of c-fos expression and could be used as an indicator of c-fos induction. When this cell line was exposed to a series of substances of unknown action, different and substance-specific changes in fluorescence level were observed. These results suggest that inclusion of additional control cell lines may allow better distinction between direct toxic insult and more specific action on gene expression mediated by test substances in reporter cell lines (23). In conclusion, the fluorescent “cell chip” is a new and promising approach for in vitro screening of chemicals for their immunotoxicity, although further refinement of the system by the expansion with other cell types and cytokines is required to enhance the precision and sensitivity of the test system for testing overall immunotoxicity (22). 2.3. Additional Assay Considerations

2.3.1. Toxicogenomics

In addition to the previously described functional assays, mechanistic assays can be used as potential screening tools for immunotoxicity and as methods to identify the mechanism of action of immunotoxicants in an attempt to expand and better understand the endpoints measured by the functional assays. In the field of direct immunotoxicity testing, “omics” approaches such as genomics and proteomics appear to be valuable and are increasingly used for this purpose. Until now, most toxicogenomics studies were in vivo experiments, but this technique also represents a major advancement for in vitro testing. Toxicogenomics studies the adverse effects of xenobiotics by means of gene expression profiling. Microarray analysis, which allows simultaneous measurement of the expression of thousands of genes in a given sample, is nowadays a widely applied technique. In short, total RNA is isolated from control and compoundexposed samples, each labeled with a different fluorescent dye, and hybridized onto microarray slides comprising multiple copies of DNA segments representing specific genes. Scanning of the slides yields intensity values for all genes evaluated. After processing and statistics, a set of differentially expressed genes can be derived. Clustering of genes showing similar expression patterns and

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pathway analysis are then applied to evaluate effects of toxicant exposure (25). Since the function of many gene products is known, and the expression patterns presumably correlate with the amount of active product produced, gene expression profiling provides insight into the mechanisms of action of xenobiotics. In addition, gene expression profiling may aid in the characterization of classes of compounds and identification of biomarkers for prediction of specific toxic effects (26–28). This approach is based on the assumption that exposures leading to the same endpoint will share changes in gene expression and is supported by several proof-of-principle studies with well-characterized chemicals (29–32). In order to identify biomarkers for immunotoxicity, overlapping transcriptional effects of model compounds were studied by Baken et al. (33). Microarray analysis was performed in murine spleens after in vivo exposure to bis(tri-n-butyl)tinoxide (TBTO), cyclosporine A, benzo(a)pyrene, and acetaminophen. The process that was most significantly affected by all toxicants was cell division, and it was concluded that the immunosuppressive properties of the model compounds appeared to be mediated by cell cycle arrest. Since highly proliferating immune cells will be particularly sensitive to the effects on cell division, evaluation of cell proliferation thus remains a valuable tool to assess immunosuppression. Patterson and Germolec (34) examined gene expression changes induced by the prototype immunosuppressive agents 2,3,7,8-tetrachlorodibenzo-p-dioxin, cyclophosphamide, diethylstilbestrol, and dexamethasone in murine thymus and spleen. Preliminary data showed that, although most transcriptional effects were compound-specific, some genes were regulated by all compounds. These genes were mainly involved in apoptosis, immune cell activation, antigen presentation and processing, and again cell proliferation. Although the specificity and predictivity of inhibition of cell division for immunotoxicity in general should be confirmed by testing a larger range of compounds, both studies show that microarray analysis offers opportunities to discover gene expression changes that may be indicative of immunosuppression. Regarding in vitro toxicogenomics, in a current research project a range of immunotoxic compounds is analyzed in in vitro exposure systems based on mouse primary thymocytes, a mouse thymoma cell line (EL-4), human PBMC and a human T cell line (Jurkat cells) to identify more specific biomarkers and to gain more insight into underlying mechanisms. By following this parallelogram approach, in which in vitro/in vivo rodent and in vitro human gene expression profiles are compared, the relevance of the outcome for humans in vivo can be assessed. Preliminary results indicate that the TBTO-specific gene expression profile in these cell cultures is highly comparable to the previously found

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expression profile of TBTO in murine spleens and primary rat thymocytes (35). Pathway analysis revealed that TBTO mainly induces DNA damage, DNA repair, and apoptosis. In addition, proteomic analysis of EL-4 cells exposed to TBTO confirmed this outcome. Microarray analysis is able to detect known and novel effects of a wide range of immunomodulating agents. However, this approach has also several experimental pitfalls (36, 37). The impact of exposure duration and dose level on the outcome of microarray analysis was illustrated by a series of experiments on TBTO. Induction of thymocyte apoptosis by TBTO appeared to precede inhibition of cell proliferation, since the former was found after short exposure times in vitro, whereas the latter was the main finding at later time points in in vitro and in vivo studies (38). Administration of a high dose of TBTO to mice resulted in significant regulation of gene expression in the thymus, whereas absence of overt gene expression changes was found in rat thymus after exposure to a somewhat lower dose, even though immunotoxic effects were observed as indicated by the involution of this organ (35). The use of both low and high doses in a study on hexachlorobenzene (HCB) (39) revealed the complexity of cells and mediators that participated in the response to this compound. Such approaches may provide valuable insight into gene expression changes in the presence and absence of pathological or cellular effects. Correct interpretation of gene expression profiles in terms of functional effects is often challenging in toxicogenomics. Changes in expression of genes mediating a certain process do not always point all to the same direction, for example, and not all genes taking part in a certain pathway will necessarily be regulated. Furthermore, induction of an immune response may be required for immunomodulators to exert their effects, which may therefore be more easily detected after stimulation by antigens or mitogens. The interpretation of in vivo microarray results may also be complicated by the effect of changes in cell populations on gene expression profiles. When assessing effects in spleen, influx of cells via the blood (possibly as a result of xenobiotic exposure) may cause altered abundance of certain mRNAs and thus altered gene expression profiles, as was seen after the exposure to a high dose of HCB (39). Furthermore, effects of xenobiotics may differ per cell type, and when effects of several xenobiotics are compared in the same organ, different compounds may affect different cell types. For a correct interpretation of genomic results, anchoring of gene expression profiles to pathological and functional endpoints is important (36, 40). It is equally important to establish correlation of absence of changes in gene expression with functional effects, since effects may only be observable in specific experimental settings or at other levels than the transcriptome, such as

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post-transcriptional or post-translational. Results of in vitro approaches should most ideally be confirmed with in vivo effects, since functional differences may exist between the cells in culture or in vivo, and in vitro designs lack interaction of various different cell types (25).

3. Discussion of In vitro Assays 3.1. The Roles of In Vitro Assays 3.1.1. Obtain Mechanistic Understanding of Immunotoxic Effects

A first role for in vitro assays is in obtaining a mechanistic understanding of immunotoxic effects. Especially when gaining knowledge at the molecular level, it is very useful to perform at least part of the investigation in vitro since confounding factors such as contributions by other cells than the ones of interest, or even more complicating the influx or efflux of cells are prevented. Examples include gene expression profiling (38) and signal transduction.

3.1.2. Parallelogram Approach

A second role for in vitro assays is in the so-called parallelogram approach. In this parallelogram there are four cornerstones, one of which is the health effect of exposure to a chemical, assessed as an endpoint (e.g., infection model) in experimental animals, and another the quantitative prediction of this endpoint in humans (Fig. 26.2). The other cornerstones are the assays of parameters that are relevant to the mechanism of the adverse effect in experimental animals and humans and are used for species comparison (41). For direct immunotoxicity, these in vitro assays often employ human PBMC or human T cell lines (e.g., Jurkat) and mouse splenocytes or mouse T cell lines (e.g., CTLL2). Any assay can be used, provided that they can be performed using similar cell types from human and laboratory animal origin. This parallelogram approach can be used to provide information on the dose– response relationship in humans. An example is the dose of TBTO that affected resistance in humans (42). In concert with information on actual exposure, dose–response data can be used for the characterization of risk for adverse health effects in humans.

3.1.3. Replace, Reduce and Refine the Use of Laboratory Animals

A third role of in vitro assays is to replace, reduce, and refine the use of laboratory animals. One application is the use of in vitro methods as a pre-screen for animal studies, thereby reducing the

Fig. 26.2. Parallelogram approach.

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number of animals. Another application is the complete replacement of animal studies by in vitro assays. In comparison with in vivo assays, in vitro assays provide a more controlled and easily manipulated cellular environment and have fewer potential concerns, such as access of the test compound to the cells of interest and their distribution. Furthermore, in vitro exposure facilitates a greater control over the timing of exposure and dose of the test compound. Overall, in vitro studies are relatively inexpensive and less time-consuming. 3.2. General Limitations of In Vitro Methods

Some general limitations for in vitro methods exist, however, and they also apply to in vitro immunotoxicology testing. By using in vitro systems, it is difficult to reproduce the integrity of the immune system. Most in vitro models lack organ architecture, thereby diminishing the possibilities for cell–cell interactions and interactions between the different components of the immune system. In addition, most in vitro systems lack the possibility for metabolism, indicating that compounds that require biotransformation to exert immunotoxic effects would require special culture systems, such as the presence of supersomes, microsomes, S9 fractions, or the use of transgenic cell lines (43). Moreover, physiochemical properties of the test compound, such as solubility, the need for serum, the effect of vehicle on cells, and chemical binding to cells, may interfere with the in vitro system. Finally, in vitro assays lack neuro–endocrine interactions, memory responses, and recovery effects. The resolution of these deficiencies will require the development of novel culture systems (3).

3.3. Prevalidation Studies

Despite these limitations, several prevalidation studies for testing direct immunotoxicity have been initiated. They have shown promising results in detecting immunosuppression in vitro. In line with the tiered approach recommended by ECVAM, pre-screening for direct immunotoxicity should start with the evaluation of myelotoxicity, since all immune cells arise from hematopoietic stem cells in the bone marrow. When compounds do not possess any myelotoxic potential, the process of evaluating direct immunotoxicity should be continued by lymphotoxicity testing (Fig. 26.1). With respect to the latter, the whole blood cytokine release assay and the lymphocyte proliferation test have proven to be the most reliable functional assays. Following myelotoxicity testing, the whole blood assay is the only one that has been prevalidated for testing direct immunotoxicity in vitro. As suspected by their names, the lymphocyte proliferation assay focuses on lymphocytes only, whereas the whole blood assay focuses on the various blood cell populations present in whole blood. Currently, the whole blood assay is used only for measuring cytokine release by monocytes and lymphocytes. Investigating additional cell types such as NK cells as well as additional endpoints

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should improve the value of the whole blood assay. The inclusion of new endpoints should not be restricted to additional cytokines or to the more general mediators such as eicosanoids, nitric oxide, or degranulation production, but should also include more immune-specific mediators such as antibodies. 3.4. Advantages of In Vitro Assays

Not only the whole blood cytokine release assay and the lymphocyte proliferation assay, but in fact all assays aimed to evaluate direct immunotoxicity in vitro can be performed by using peripheral blood. Peripheral blood has the advantage that it can easily be obtained and that the same individual (or animal) can be repeatedly tested. Moreover, peripheral blood, especially whole blood, has the advantage that primary cells are employed in their physiological proportions and environment. By using human peripheral blood, the number of laboratory animals can significantly be reduced and species differences are avoided. The “cell chip” has proven potential for in vitro testing of direct immunotoxicity. The value of this assay will likely improve when other cell types and endpoints are included. The prototype “cell chip” consists of T cells, since they are generally the most sensitive target cells in case of direct immunotoxicity. However, next to T cells, other cell types also play pivotal roles in the immune response. Mast cells, DC, keratinocytes, and lung epithelial cells, among others, are target cells of indirect immunotoxicity and are therefore used for in vitro testing for this type of immunotoxicity. Inclusion of these cell types may be of additional value for testing direct immunotoxicity.

3.5. Comparison of Whole Blood Cytokine Release Assay and Fluorescent Cell Chip

Although the whole blood cytokine release assay and the fluorescent cell chip rely on a similar endpoint, cytokine expression, the outcome of both assays might differ as different cell cultures are used. In the whole blood assay primary cells are used, whereas in the “cell chip” cell lines are employed. Cell lines provide a more homogeneous source than do the primary cultures and can be grown in essentially unlimited quantities without (in case of animal cells) the need to sacrifice animals. However, the fact that a cell can grow indefinitely in culture implies that it has lost some of the differentiation properties of a primary cell, and this might affect the outcome of the assay. There is a debate on whether primary cells or cell lines are preferred in direct immunotoxicity testing. This issue is addressed in ongoing studies that compare gene expression profiles of primary thymocytes and the thymoma cell line EL-4 after exposure to TBTO with the gene expression profile of the thymus from mice exposed to the same compound (Janssen et al., in preparation). In fact, for TBTO exposure, gene expression profiles are or will be generated for rat and mouse thymus in vivo (35), rat and mouse thymocytes in vitro ((38); Janssen et al., in preparation),

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and human PBMC in vitro (Madhu et al., in preparation). So, not only at the phenotypic level (42) but also at the genomic level dose–response relationships in humans will be available. The phenotypic inter-species extrapolation may in this respect be regarded as a “phenotypic anchor.” 3.6. Using In Vitro Assay for InterSpecies Extrapolation

Another example of using in vitro assays for inter-species extrapolation is the prediction of the human maximum tolerated dose (MTD) of myelo-suppressive xenobiotics. Besides detecting myelo-suppressive agents, the (CFU-GM) assay can also correctly predict (87%) of the human MTD of myelo-suppressive xenobiotics by adjusting mice-derived MTD for the differential sensitivity between CFU-GM from mice and humans. The inhibition of CFU-GM in vitro correlates with the absolute neutrophil count decrease in vivo. The model offers the opportunity for studying onset and recovery of neutropenia. Using this assay in the preclinical testing phase of drug development will substantially decrease the risk of a lethal overdose to patients in phase I clinical trials, and the trial will be completed more quickly.

3.7. “Omics” Approaches to In Vitro Testing

In addition to the functional in vitro assays, “omics” approaches such as genomics and proteomics are promising for in vitro testing of direct immunotoxicity. Besides elucidating the mechanisms of action, they offer the opportunity for characterization of classes of compounds and identification of biomarkers for prediction of specific toxic effects. Incorporation of these results into functional assays will substantially lead to the improvement of sensitivity and specificity of testing.

3.8. The Future of In Vitro Testing

The future of in vitro testing for direct immunotoxicity will see a number of simultaneous, independent developments (Table 26.1). Not only will assay development be continued, it is imperative

Table 26.1 Developments in in vitro testing for direct immunotoxicity Assay set-up: Choice for species Choice between primary cells and cell lines Assay development: Knowledge which assays have the best predictivity (Pre) validation of these assays Inclusion of “omics” techniques: Functional genomics Proteomics Technical improvements: High-content screening High-throughput screening

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that, on the basis of the predictivity of an assay or the combined predictivity of two (or more) assays, several in vitro assays should be prevalidated. They should preferably be of human origin, as inter-species extrapolation is avoided. Assay development is supported by novel techniques such as functional genomics (microarray analysis, siRNA gene knockdown) and proteomics. In addition, there is a tremendous technical innovation in the field of high-content screening (e.g., computer-automated fluorescence microscopy) and HTS (e.g., using microfluidics).

4. Conclusion In conclusion, major opportunities exist to advance in vitro testing for direct immunotoxicity. This should include prevalidation of existing assays and selection of the assay (or combination of assays) that performs best. To avoid inter-species extrapolation, assays should preferably use human cells. Furthermore, the use of whole blood has the advantage of comprising multiple cell types in their natural proportion and environment. The so-called “omics” techniques provide additional mechanistic understanding and hold promise for the characterization of classes of compounds and prediction of specific toxic effects. Phenotypic anchoring and functional genomics are key factors to fulfill this promise. Technical innovations such as high-content screening and high-throughput analysis will greatly expand the opportunities for in vitro testing. As improved mechanistic understanding, technical innovations, and an open mind to in vitro alternatives by legislative bodies are all required, advancement of in vitro immunotoxicity testing requires efforts by research institutions, industry, and government.

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Index A Acquired immunodeficiency syndrome (AIDS)................................................... 85, 306 Acute phase protein Complement C3......................................................... 69 C-reactive protein................................................. 66, 67 Plasminogen............................................................... 69 Serum amyloid A........................................................ 69 Adhesion molecules.................................10, 58, 61–70, 104 Adjuvant.............................43, 162, 188–191, 350, 367, 810 Adult trigger..................................................................... 21 Agarose gel......................................................250, 256, 298 Alkaline phosphatase................... 30, 34, 288, 290, 292, 293 Allergen............................7, 9, 221, 222, 224, 226, 227, 230 Allergy............................ 4, 9, 21, 54, 78, 230, 367, 370, 402 Alternative methods................................................8, 9, 402 Anaphylaxis................................................................ 9, 370 Annexin V............................................................8, 243, 247 Antibiotics................................................................ 81, 367 Antibody production........................................41, 103, 106, 160, 161, 242, 405, 411 Antigen presentation................. 83, 113, 114, 196, 412, 415 Antigen-presenting cells (APC)...............................83, 144, 234, 259, 374, 377 Apoptosis.................................................... 63, 88, 144, 197, 207, 241–257, 327, 331, 332, 415, 416 Arachidonic acid....................................................10, 56, 83 Arthritis............................41, 43, 54, 58, 103, 105, 367, 369 Arthus reaction............................................................... 342 Assay validation...................................................... 307, 420 Asthma.................................................... 10, 23, 54, 57, 367 Astrocyte.......................................................................... 66 Atrazine.................................................................. 187, 260 Autism...................................................................... 23, 368 Autoimmunity............................................ 4, 10, 18, 21, 23, 41–48, 367, 368, 370, 379, 402 Autophagy.................................................................. 82–83 Avian.......................................................388–390, 394, 396

B BALB/c mice.................................. 133, 202, 268, 269, 277 Basophil.........................................................55–57, 65, 403 Bioassay.............285–287, 289, 295–296, 299, 300, 375, 404 Biopharmaceuticals.......................... 303, 342, 363, 374, 379

Blastogenesis.................................................................. 138 B lymphocyte.......................................................23, 63, 64, 285, 306, 310, 389, 407 Body weight............................................180, 288, 324, 326 Bone marrow................................ 54–60, 63, 64, 78, 80, 82, 83, 85, 88, 130, 199, 260–261, 268, 307, 309, 311, 312, 319, 325, 326, 335–338, 345, 346, 352–354, 357, 358, 371, 372, 403, 404, 411, 418 Bovine serum albumin (BSA)...............................30, 34–36, 87, 188–191, 262, 265–267, 270, 288, 290, 292–295, 308, 309, 344, 345 Breast cancer................................................................... 186 Bronchoalveolar lavage........................................... 112, 113 Bronchus-associated lymphoid tissue (BALT).............23, 198, 325, 333, 335–337, 403 Brown Norway rats......................................................... 133

C Calcium flux................................................................... 303 Candida...........................................................100, 101, 373 Caspase....................................................242–244, 351–352 Cell culture....................................... 36, 145–148, 152, 199, 208, 244, 265–268, 275, 277, 279, 284, 287, 294, 301, 318, 355, 388, 415, 419 Cell line........................................................ 47, 81, 82, 113, 120, 150, 208, 214, 215, 296, 300, 308–309, 314–315, 337, 354, 358, 376, 379, 409, 410, 412–415, 417–420 Cell mediated immunity (CMI)...............................99, 101, 105, 106, 185, 186, 195, 198, 268 Cell proliferation............................................8, 64, 66, 160, 266, 275–276, 279, 405, 407, 408, 415, 416 Chelation........................................................................ 250 Chemilluminescence.............................................. 245, 254 Chemoattractant.................................................. 64–66, 69 Chemokine.......................63–66, 70, 71, 144, 187, 294, 296 Chemotaxis..........................................................63, 67, 187 Chicken....................................................46, 174, 187–189, 191, 261, 267, 388, 395 Childhood leukemia................................................... 23, 44 Children’s health............................................................... 19 Chimeric animals........................................................... 378 Cigarette smoking.............................................42, 151, 197 Clinical pathology......................... 54, 55, 70, 324, 326, 357

425

Immunotoxicity Testing 426 Index

Cluster of differentiation (CD) CD3.............. 36, 279, 286, 343, 345, 347, 354–357, 407 CD4...................................................... 36, 41, 122, 187, 195–197, 235, 265, 268, 274, 275, 279, 306, 310, 327, 328, 343, 345–347, 356, 357 CD8............................................ 36, 110, 113, 187, 196, 235, 279, 327, 328, 343, 345, 347, 356, 357, 409 CD11...................................................88, 104, 105, 279 CD16........................................................207, 209, 213, 214, 342, 343, 347, 356, 357 CD19........................................................................ 279 CD20................................. 306, 343, 345, 347, 356, 357 CD28........................................... 36, 235, 286, 368, 407 CD34...........................................................88, 260, 279 CD40...........................................................88, 279, 407 CD45................................................................ 343, 347 CD49.................................................207, 208, 213, 217 CD56................................. 207, 209, 215, 342, 343, 410 CD80.................................................................. 88, 279 CD83........................................................................ 279 CD86...........................................................88, 279, 412 CD95................................................................ 207, 242 CD123...................................................................... 279 Colony forming assays................................................ 85–86 Committee for Proprietary Medicinal Products (CPMP)............................................................ 6 Complement................................44, 55, 57, 62, 66–71, 104, 174–176, 179, 180, 182, 185, 285, 297, 342, 374 Confocal microscopy.................................................. 86–87 Contact hypersensitivity......................................... 233–238 Converging technologies................................................ 380 Cortex:medulla ratio............................................... 327, 328 Coulter Counter......................................175, 177, 179, 201 Cyclooxygenase................................................................ 83 Cyclophosphamide.........................................102, 111, 124, 129, 130, 188, 236, 415 Cyclosporine A..........................................42, 161–163, 415 Cynomolgus monkey......................................343, 345–348, 350–353, 356, 358 Cytokine......................................... 9, 10, 28, 32, 53, 58–67, 69–71, 78, 90, 98, 99, 101–104, 112–115, 122, 133, 144, 159, 160, 197, 242, 259, 260, 265–266, 268, 269, 275, 276, 283–301, 344, 348–349, 368, 370, 372–377, 405–407, 411–414, 418–420 Cytokine storm..........................................98, 368, 370, 375 Cytomegalovirus......................................100, 101, 105, 114 Cytotoxic T lymphocyte (CTL).............................5, 61, 64, 99, 109, 110, 113, 114, 133, 144, 186, 195–202, 405, 409–410

D Delayed type hypersensitivity response (DTH response)....................... 7, 19, 28, 33, 37, 47, 186–188, 190, 196, 260, 276, 277, 342, 373

Dendritic cell............................................ 9, 36, 64, 83, 144, 159, 186, 187, 195, 197, 222, 259–279, 332, 377, 403, 412 Developmental immunotoxicity (DIT)...........................................17–24, 43, 369 Dexamethasone..............................................102, 111, 120, 124, 161, 183, 187, 328, 331, 351, 415 Diethylstilbestrol...............................................19, 121, 415 9,10-Dimethyl-1,2-benzanthracene (DMBA)............... 147 Dinitrifluorobenzene (DNFB)....................................... 236 2,4-Dinitrochlorobenzene (DNCB)............................... 229 Dioxins, 2,3,7,8-tetrachlorodibenzop-dioxin...................................42, 121, 187, 415 DNA fragmentation...............................................242, 244, 247–250 DNA microarray.............................................................. 70 Dosing regimen...................................................... 324, 326

E Ear challenge.................................................................. 236 Effector cells.....................................................63, 187, 196, 199, 201, 202, 211, 212, 351, 374, 409–411 Effector/Target ratio................................ 58, 63, 65, 66, 69, 120, 122, 124, 144, 187, 196, 199, 201, 202, 209–212, 242, 252, 349–351, 357, 374–376, 404, 409–411 Elicitation phase..................................................... 234–236 Environmental Protection Agency (EPA).................. 6, 188 Enzyme linked immunosorbent assay, ELISpot..............................30–32, 34, 35, 37, 288–289, 292–294 Eosinophil..................................... 55, 57, 65, 123, 372, 403 Epidemiology................................................................. 4, 7 Epitopes.................................................................... 43, 175 Epstein-Barr virus.................................................. 101, 114 Erythema...............................................................8, 40, 373 Estimated concentration................................................. 227 Extended histopathology.................................................... 6

F Fas ligand................................................................ 197, 207 Fc receptor (FcR)........................................................... 294 Fibrosis................................................................... 332, 338 Flow cytometry............................................36, 87–89, 144, 208–209, 213–214, 264, 268, 287, 294, 303–320, 337, 373–374 Fluorescein isothiocynate (FITC)............................. 87, 88, 209, 213, 215, 216, 279, 343, 347 Fluorescence-actived cell sorting (FACS)............ 213, 289, 294–295, 307, 343, 347 Fluorescent cell chip................................412–414, 419–420 Fluorochrome.................................. 289, 307–309, 316, 320 Fluorophor..................................................................... 290 Foot pad challenge.......................................................... 187 Foxp3........................................................................ 41, 197

Immunotoxicity Testing 427 Index

G

I

Gastrointestinal-associated lymphoid tissue (GALT)........................... 23, 325, 333, 335–337 Gavage.......................................................... 29, 32, 37, 114, 124, 126, 131, 163 Gel electrophoresis..........................................244, 249, 250 Gender......................................... 41, 44, 145, 162, 188, 370 Gene expression..............................................9, 10, 91, 197, 245–246, 254, 255, 289, 296, 298, 414–417, 419 Gestation.......................................................................... 19 Good laboratory practices (GLP)........................... 356, 358 Granulysin.......................................................207–209, 214 Granzyme........................................ 196, 197, 207–209, 214 Guinea pig................................................ 4, 7–10, 174, 175, 182, 221, 234, 235, 237, 238, 324

ICH S8 guidance............................................................ 162 Immune complex............................. 47, 48, 68, 78, 290, 291 Immune dysfunction..............18, 19, 22, 23, 40, 41, 44, 370 Immune enhancement.............................................. 45, 176 Immune surveillance............................................... 143, 144 Immunization............................................. 47, 98, 196, 262, 269, 342, 350, 352, 366, 367, 374, 378, 388, 390, 391, 393, 395, 396, 412 Immunobiology........................................................ 67, 221 Immunocompetence.............................. 18, 97, 98, 110, 124 Immunofluorescence.......................................... 86–87, 373 Immunoglobulin (Ig) IgA........................................................7, 114, 123, 139 IgE..............................................................7, 9, 10, 122, 123, 138, 139, 263, 270, 271 IgG IgG1............................................. 10, 28, 30, 34, 35, 99, 209, 214, 215, 260, 262, 268, 270, 343, 347 IgG3........................................................... 343, 347 IgG2a.................................................28, 30, 34, 35, 99, 208, 260, 262, 268, 270, 343 IgM................................................. 7, 99, 100, 114, 123, 129, 137–139, 160–171, 174–176, 179, 180, 208, 350–352, 394, 397 Immunohistochemistry..................... 62, 284, 294, 357, 358 Immunological priming.................................................. 222 Immunomodulation.......................... 55, 335, 364–368, 405 Immunophenotyping..............................................7, 10, 99, 104, 306, 319, 337, 346, 356, 357 Immunostimulation............................... 10, 28, 98, 368, 402 Immunosuppression............................................. 4, 6, 9, 11, 21, 97, 98, 101, 103, 111, 114, 115, 121, 144, 160, 162, 283, 368, 373, 375, 402, 404, 406, 408, 415, 418 Immunotoxicant...............................................7, 18, 21, 45, 109, 198, 365, 402–404, 412, 414 Induction phase...................................................... 235, 236 Industrial chemicals.......................................................... 10 Infection....................................... 18, 43–45, 47, 55–59, 65, 67, 81, 82, 98, 99, 101–106, 110–115, 120–126, 129–133, 135, 178, 181, 186, 196, 197, 199, 200, 230, 241, 365–367, 371–373, 376, 388, 389, 405, 417 Infectious disease..................................... 6, 18, 98, 388, 402 Inflammation............................................ 18, 47, 53–71, 78, 122, 123, 187, 284, 327, 330, 389, 395 Inflammatory misregulation............................................. 48 Influenza......................................................................... 111 Informatics..................................................................... 380 Inhalation..........................................................10, 198, 325 Innate immunity........................................99, 100, 102, 105 Integrin............................................................... 61–62, 104

H Haemophilus.................................................................. 103 Hapten.....................................................222, 234, 235, 412 Hazard identification................. 46, 160, 222, 224, 228, 230 Hemagglutination assay.......................................... 390–394 Hematopoietic stem cells (HSC)............................ 75, 76, 85, 88, 404, 418 Hepatitis B..................................................................... 367 Herpes virus................................................................... 105 Hexyl cinnamic aldehyde................................................ 227 Histamine..............................................................56, 57, 64 Histiocyte....................................................................... 354 Homeostasis..................................................18, 66, 76, 364 Horseradish peroxidase........................................... 265, 290 Host resistance bacteria............................................................. 101–105 fungus....................................................................... 101 parasite......................................................101, 119–139 protozoan.................................................................. 120 tumor.........................................................101, 143–154 virus...........................................................101, 109–115 Human................................................. 3–11, 18, 22, 23, 28, 39, 42, 45, 47, 67, 69, 80, 82, 88, 91, 98, 101, 103, 104, 111, 114, 115, 120, 121, 123, 125, 130, 186, 187, 207–215, 234, 237, 260, 285, 286, 293, 306, 310, 311, 319, 342, 344, 346, 349, 350, 355, 356, 358, 363–382, 404–408, 410–413, 415, 417, 419–421 Humoral immunity (HI).............................5, 110, 112, 121 Hygiene hypothesis........................................................ 367 Hypercontractility.......................................................... 122 Hypersensitivity.......................................... 7, 18, 27–29, 33, 57, 185–192, 195, 233–238, 242, 268, 285, 342, 367, 370, 372, 373, 379 Hyporesponsiveness........................................................ 366

Immunotoxicity Testing 428 Index

Interferon Interferon-a............................................................. 411 Interferon-b.............................................................. 411 Interferon-g..............................................28, 36, 60, 63, 64, 78, 81, 90, 405, 411, 413 Interlaboratory reproducibility........................................... 5 Interleukin (IL) Interleukin-1............................................60, 63, 64, 67, 112, 290, 296, 405, 406 Interleukin-2............................................57, 63, 64, 83, 214, 290, 296, 346, 355, 358, 405, 413 Interleukin-3...............................................86, 290, 411 Interleukin-4............................................28, 36, 78, 91, 122, 197, 276, 283, 290, 405–407, 413 Interleukin-5...............................................57, 283, 290 Interleukin-6............................................60, 63, 64, 67, 112, 290, 296, 405 Interleukin-8........................................................ 64, 65 Interleukin-9............................................................ 290 Interleukin-10............................. 78, 197, 290, 296, 413 Interleukin-12.....................................78, 197, 283, 290 Interleukin-13............................. 78, 122, 197, 283, 405 Interleukin-18.......................................................... 197 Interleukin-21.......................................................... 197 Interleukin-23.......................................................... 197 Interleukin-27.......................................................... 197 International Conference on Harmonization of Technical Requirements for Registration of Pharmaceuticals for Humans Use (ICH).......................................... 6 Intracellular adhesion molecule.................................. 61, 62 Intranasal exposure.......................... 102, 103, 111, 112, 199 Intravenous injection...............................177, 222, 224, 230 In vitro....................... 9, 28, 47, 68, 123, 145, 186, 209, 212, 216, 236, 246, 247, 260, 268, 275, 284, 286, 296, 351, 366, 376–378, 382, 389, 401–421 In vivo................................................. 27–37, 111, 144–146, 148–149, 165, 168, 186, 216, 234, 238, 246, 247, 260, 268, 286, 310, 318–320, 351, 376, 387–397, 402, 406–409, 411–420

J Juvenile..................................................... 19, 22, 24, 41, 43, 111, 151, 188, 191, 192, 198

K Keyhole limpet hemocyanin (KLH)...................... 160–166, 168–171, 174, 188–190, 344, 345, 350–352, 357 Kupffer cell................................................................. 78, 83

L Langerhans cell................................................222, 235, 412 Larvae..............................................121–132, 134–136, 138

Lead......................................19, 42, 103, 187, 208, 237, 319 Leukotrienes..........................................................55, 56, 66 Lipopolysaccharide (LPS).................. 36, 61, 67, 81, 84, 87, 90, 261, 268, 269, 276, 286, 358, 405, 407, 408 Listeria.................................................................... 104, 105 Local lymph node assay (LLNA)...........................8–10, 47, 221–231, 235, 285 Lupus.......................................................................... 40, 42 Lyme disease.................................................................. 367 Lymph node auricular lymph node................... 33, 224–227, 229, 235 mesenteric lymph node................ 70, 124, 138, 325, 333 popliteal lymph node...............................10, 28, 37, 333 submandibular lymph node...................................... 325 Lymphocryptovirus (LCV)............................................ 115 Lysis............................. 59, 90, 176, 182, 201, 242, 249, 251, 255, 267, 307–309, 312, 314, 320, 394, 396, 410

M Macrophage alveolar.............................................. 76, 78, 80, 83, 113 peritoneal...................................................78, 80, 81, 83 splenic..................................................78, 104, 105, 330 Major histocompatibility complex (MHC) MHC class I...................... 113, 144, 199, 235, 409, 410 MHC class II................................................63, 88, 235 Mantoux test.................................................................. 186 Marginal zone B cells...................... 100, 103–105, 329, 331 Mast cell................... 55, 56, 61, 64, 122, 123, 332, 412, 419 Medical devices........................................................ 17, 241 Megakaryocyte...............57, 58, 64, 335, 338, 353, 354, 404 Melanoma cell model......................................145–148, 150 Membrane blebbing....................................................... 242 Membrane potential........................ 244, 250–251, 256, 288 Memory................................................ 5, 66, 104, 187, 197, 234, 235, 350, 390, 396, 418 Mercury................................................................ 19, 41–44 Meta-analysis......................................................... 381, 390 Metastasis............................................................... 144–147 Methyl salicylate............................................................. 226 Microbial killing....................................................... 81, 377 Microglia.........................................................43, 64, 66, 78 Mitochondria...............................................7, 88, 242–244, 250–252, 256, 303, 377 Mitogen concanavalin A................................................. 286, 407 phytohemagglutinin.......................................... 286, 389 pokeweed.................................................................. 407 Mixed lymphocyte reaction.....................268, 279, 408–409 Moloney murine leukemia virus......................208, 246, 255 Monoclonal antibody...............................66, 103–105, 138, 215, 262, 292, 294, 310, 311, 368, 388 Monocyte.......................... 59, 62, 64, 65, 69, 76, 79, 80, 88, 91, 105, 123, 187, 286, 347, 405–407, 412, 418

Immunotoxicity Testing 429 Index

Mouse lymphoma cells...................................145, 147–148, 152–153, 208 Mouse mammary tumor virus........................................ 115 Mucosa...............................23, 121, 123, 198, 325, 333, 336 Multiparameter evaluation..............................198, 294, 311 Multiple sclerosis.........................................23, 41, 186, 367 Mycobacterium............................................................... 186 Myeloperoxidase....................................................... 55, 311

N Nanotechnology............................................................. 380 National Toxicology Program (NTP)...................................... 4, 5, 21, 186, 198 Natural infection.............................................................. 47 Natural killer cell.......................................... 43, 61, 99, 113, 144, 197, 207, 259, 342, 344, 376, 403, 410–411 Necrosis................................................ 27, 59, 88, 144, 196, 242, 327, 329, 330, 332, 334, 335, 337, 338 Nematode............................................................... 120–122 Neonatal..........................................................3, 22, 24, 366 Neoplastic disease.................................... 3, 97, 98, 119, 160 Neutrophil.............................. 55–57, 60, 69, 100, 102–105, 112, 120, 123, 187, 337, 358, 377, 403, 420 Nitric oxide..................................................78, 83, 406, 419 Non-human primate.............................. 110, 115, 286, 306, 307, 310–313, 319, 324, 330, 341–358, 365, 380 No observed effect level (NOEL).................................. 161 Norepinephrine.............................................................. 260 Nuclear antigens............................................................. 377 Nutritional status.................................................... 324, 326

O Oral exposure..................................................28, 29, 32, 36 Organization of Economic Cooperation and Development (OECD)..................6, 8, 222, 230 Organ weight...................................................324, 326, 403 Otitis media................................................................ 18, 23 Ovalbumin............................................................... 28, 367

P Paraphenylenediamine............................................ 226–228 Pathogens............................................. 43, 68, 97, 103, 104, 119, 120, 125, 145, 151, 160, 195, 260 Perforin............................................196, 207–209, 214–216 Peripheral blood leukocytes....................................210, 211, 213, 214, 376, 377 Pesticides chlordane.................................................................... 19 heptachlor................................................................... 19 hexaclorobenzene....................................................... 19 Peyer’s patches.........................................233, 325, 336, 403 Phagocytosis............................................... 7, 55, 62, 67, 68, 80–82, 109, 113, 234, 377, 389 Phorbol myristyl acetate (PMA).............284, 288, 293, 358

Phosgene........................................................................ 198 Phycoerythrin..................................................208, 279, 348 Picryl chloride.................................................234, 236, 237 Plaque forming cell (PFC).............. 173, 176, 183, 185, 186 Plasma cells.................................... 175, 176, 329, 332, 334, 335, 337, 354, 372, 374 Platelet activating factor............................................. 56, 66 Polychlorinated biphenyls (PCBs)...............18, 42, 197, 388 Polymerase chain reaction (PCR) reverse transcription – PCR........................71, 245–246, 254–256, 284, 289, 296–298, 407, 413 Preclinical drug development..........................303–320, 420 Pregnancy................................................................ 342, 367 Prenatal........................................................22, 24, 151, 368 Primers.................................... 246, 255, 256, 289, 296–298 Propidium iodide..............................................88, 113, 209, 243, 244, 247, 249, 351 Prostaglandin...................................................55, 56, 63, 83 Protein estimation.................................................. 252–254 Protein phosphorylation................................................... 89 Proteomics............................70, 71, 305, 414, 416, 420, 421 Pseudomonas...................................................100, 103, 104

R Radioimmunoassay................................................. 289, 373 RANTES................................................................... 64, 65 Reactive nitrogen species.....................................78, 83, 377 Reactive oxygen species............... 58, 78, 83, 84, 88, 89, 377 Receptor occupancy.................................309, 315–318, 320 Recombination homologous.............................................................. 379 nonhomologous........................................................ 379 Reovirus.......................................................................... 114 Reporter antigen..............................................10, 28, 31–33 Respiratory burst................................... 55, 83, 84, 109, 113 Rheumatoid factor.................................................. 300, 377 Risk assessment............................................ 4, 6, 11, 18, 45, 98, 110, 186, 198, 222, 224 Risk/benefit ratio............................................................ 366 Risk management........................................................... 222 RNA extraction.................................92, 245–246, 254–256 RNA quantitation.........................................71, 89–91, 284 Rodent mouse......................4, 6, 9, 102, 114, 163–165, 167–171 rat.......................................... 4, 6, 9, 102, 114, 160–171

S Scintillation counter................................223–227, 229, 356 Selectin E-selectin.................................................................... 61 L-selectin............................................................ 61, 187 P-selectin.................................................................... 61 Sensitization............................3, 7–9, 11, 47, 177, 178, 181, 187, 189–191, 221–231, 234–237, 402, 409, 412

Immunotoxicity Testing 430 Index

Sheep erythrocytes.......................................173–183, 188, 374 red blood cells.................6, 160, 175, 188, 390, 391, 412 Skin.................................................... 3, 7–9, 11, 32, 33, 37, 130, 149, 221–231, 234– 237, 250, 337, 373, 389–392, 394–396, 412 Spleen..........................................70, 78, 104, 105, 124, 138, 144, 175, 176, 178–180, 182, 183, 198, 199, 210, 233, 274, 275, 285, 324–326, 329–332, 354, 374, 389, 403, 404, 408–411, 415, 416 Splenocytes............................................. 182, 210, 211, 213, 216, 264, 274, 407–410, 412, 417 Stimulation index.......................................8, 222, 225–227, 236, 358, 392, 394 Streptavidin.............................................. 91, 129, 256, 263, 265, 267, 278, 288, 290, 292, 293, 308, 314, 315, 318, 344, 348 Streptococcus.............................................43, 100, 102–104 Strongyloides.................................................................. 121 Subcutaneous exposure......................................29, 151, 212 Synthetic immune system....................................... 377–378

T Target cells......................................... 65, 69, 113, 196, 197, 199–202, 207–209, 211, 212, 217, 300, 308, 316, 344, 349–351, 357, 376, 409–411, 419 T cell receptor diversity.....................................23, 235, 407 T-dependent antibody response (TDAR)...............5, 47, 99, 100, 109, 110, 114, 159–171, 173–183, 196, 198, 233, 350–352, 357, 374 Tetanus toxoid........................................................ 174, 350 Therapeutics............................................. 10, 18, 46, 47, 58, 100, 102–106, 115, 306, 314, 319, 358, 366, 368, 369, 380–382, 406 Thimerosal......................................................260, 367, 368 Thiols............................................................................. 260 Thymic atrophy.............................................................. 144 Thymidine......................................................222–226, 230, 235, 266, 276, 346, 356, 407, 408 Thymocyte..............................................407, 415, 416, 419 Thymus................................................. 6, 70, 103, 144, 233, 324–330, 389, 403, 411, 415, 416, 419 Tier testing..................................................................... 5, 7 T-independent antibody response (TIAR)............. 103, 104 T lymphocyte Th17 cell.................................................................. 197

T helper 1......................................................... 186, 197 T helper 2................................................................. 197 tregs...........................................................144, 187, 197 Tolerance................................43, 45, 47, 187, 366, 368, 377 Toll-like receptor........................................................ 10, 43 Toxicant.................................................... 18, 19, 39, 43–44, 47, 58, 62, 109–111, 113– 115, 143, 147, 151, 241, 251, 256, 260, 365, 368, 369, 406, 415 Toxicogenomics...................................................... 414–417 Transcriptional expression...............................197, 416–417 Transcriptional factor nuclear factor-kappa B......................................... 65, 76 Transforming growth factor beta (TGF- b).......................................... 63–65, 144 Transgenic mice...................................................... 260, 379 Tributyltin oxide................................ 19, 121, 415–417, 419 Trichinella................. 100, 101, 120, 121, 124, 129, 136–138 Trichloroethylene..................................................... 42, 188 Trinitrophenyl-ovalbumin................................................ 28 Tumor challenge..............................................101, 143–154 Tumor necrosis factor TNF-a......................................................112, 284, 290 TNF-b...................................................................... 411 TUNEL assay..................................................243–244, 256 Type 1 diabetes....................................................23, 41, 367 Tyrosine kinase....................................................... 102, 198

U Ultraviolet light.............................................................. 121 UV transilluminator....................................................... 250

V Vaccination.......................186, 352, 366–368, 372, 374, 375 Vaccine safety......................................................... 366, 368 Viral titer............................................... 5, 98, 111, 112, 114 Virus reactivation.............100, 101, 105, 106, 110, 114, 115 Vitamin A....................................................................... 186

W Western blot.....................82, 83, 89, 90, 245, 252–254, 311 World Health Organization (WHO)....................7, 46, 296 Wound healing......................................................64, 65, 78

X Xenogeneic............................................................. 185–192