Immunotoxicity Testing: Methods and Protocols (Methods in Molecular Biology, 598) 9781607614005, 9781607614012, 1607614006

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Methods

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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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195 207 221 233 241

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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immunity for hazard identification purposes in rats in a standard toxicology study. Toxicology 126:137–152 House RV, Luebke RW (2007) Immunotoxicology: thirty years and counting. In: Luebke R, House R, Kimber I (eds) Immunotoxicology and immunopharmacology, 3rd edn. CRC, Boca Raton, FL, pp 3–20 EPA, Biochemical Test Guidelines (1996) OPPTS 880.3550 immunotoxicity. US/EPA, Washington DC Kuper CF, Harleman JH, Richter-Reichelm HB, Vos JG (2000) Histopathologic approaches to detect changes indicative of immunotoxicity. Toxicol Pathol 2:454–466 Germolec DR, Nyska A, Kashon M, Kuper CF, Portier C, Kommineni C, Johnson KA, Luster MI (2004) The accuracy of extended histopathology to detect immunotoxic chemicals. Toxicol Sci 82:504–514 Haley P, Perry R, Ennulat D, Frame S, Johnson C, Lapointe JM, Nyska A, Snyder P, Walker D, Walter G (2005) STP Immunotoxicology Working Group. Best practice guideline for the routine patholgy evaluation of the immune system. Toxicol Pathol 33:404–408 Descotes J, Nicolas B, Pham E (1997) Sentinel screening for human immunotoxicity. In: Environment and immunity. Proceedings of a Workshop held in Brussels on 20–21 May 1996. Air Pollution Epidemiology Reports Series. S. R NRC (1992) Biologic markers in immunotoxicology. A report by the US National Research Council. National Academy Press, Washington DC WHO (1996) Principles and methods for assessing direct immunotoxicity associated with exposure to chemicals. A report of the International Programme on Chemical Safety (Environmental Health Criteria 180). World Health Organization, Geneva

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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

Risk of Autoimmune Disease: Challenges for Immunotoxicity Testing

43

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

Risk of Autoimmune Disease: Challenges for Immunotoxicity Testing

45

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

Risk of Autoimmune Disease: Challenges for Immunotoxicity Testing

47

(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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obese chickens: influence of age and sex as measured by ELISA. Dev Comp Immunol 9:107–118 41. Sorg H, Lorch B, Jaster R, Fitzner B, Ibrahim S, Holzhueter SA, Nizze H, Vollmar B (2009) Early rise in inflammation and microcircula­ tory disorder determine the development of autoimmune pancreatitis in the MRL/ Mp-mouse. Am J Physiol Gastrointest Liver Physiol 295:G1274–G1280 42. Gubbels Bupp MR, Jorgensen TN, Kotzin B (2008) Identification of candidate genes that influence sex hormone-dependent disease phenotypes in mouse lupus. Genes Immun 9:47–56 43. Hughes GC, Clark EA (2007) Regulation of dendritic cells by female sex steroids: relevance to immunity and autoimmunity. Autoimmunity 40:470–481 44. Stevens AM (2006) Microchimeric cells in sys­ temic lupus erythematosus: targets or inno­ cent bystanders? Lupus 15:820–826 45. Rak JM, Maestroni L, Balandraud N, Guis S, Boudinet H, Guzian MC, Yan Z, Azzouz D, Auger I, Roudier C, Martin M, Didelot R, Roudier J, Lamber NC (2008) Transfer of the shared epitope through microchimerism in women with rheumatoid arthritis. Arthritis Rheum 60:73–80 46. Selmi C, Invernizzi P, Gershwin ME (2006) The X chromosome and systemic sclerosis. Curr Opin Rheumatol 18:601–605 47. Smith-Bouvier DL, Divekar AA, Sasidhar M, Du S, Tiwari-Woodruff SK, King JK, Arnold AP, Singh RR, Voskuhl RR (2008) A role for sex chromosome complement in the female bias in autoimmune disease. J Exp Med 205:1099–1108 48. Invernuizzi P, Miozzo M, Oetelt-Prigione S, Meroni PL, Persani L, Selmi C, Battezzati PM, Zuin M, Lucchi S, Marasini B, Zeni S, Watnik M, Tabano S, Maitz S, Pasini S, Gershwin ME, Podda M (2007) X monosomy in female systemic lupus erythematosus. Ann N Y Acad Sci 1110:84–91 49. Nandula SR, Armarnath S, Molinolo A, Bandyopadhyay BC, Hall B, Goldsmith CM, Zheng C, Larsson J, Sreenath T, Chen W, Ambudkar IS, Karlsson S, Baum BJ, Kulkarni AB (2007) Female mice are more susceptible to developing inflammatory disorders due to impaired transforming growth factor beta sig­ naling in salivary glands. Arthritis Rheum 56:1798–1805 50. Jane-wit D, Altuntas CZ, Monti J, Johnson JM, Forsthuber TG, Tuohy VK (2008) Sexdefined T-cell responses to cardiac self deter­ mine differential outcomes of murine dilated cardiomyopathy. Am J Pathol 172:11–21

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

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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 ( 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

3. Blair HC, Zaidi M (2006) Osteoclastic differen­ tiation and function regulated by old and new pathways. Rev Endocr Metab Disord 7:23–32 4. Blyler G, Landreth KS, Lillis T et  al (1994) Selective myelotoxicity of propanil. Fundam Appl Toxicol 22:505–510 5. Mosser DM, Edwards JP (2008) Exploring the full spectrum of macrophage activation. Nat Rev Immunol 8:958–969

Evaluating Macrophages in Immunotoxicity Testing 6. Edwards JP, Zhang X, Frauwirth KA, Mosser DM (2006) Biochemical and functional char­ acterization of three activated macrophage populations. J Leukoc Biol 80:1298–1307 7. Raes G (2002) FIZZ1 and Ym as tools to dis­ criminate between differentially activated macrophages. Dev Immunol 9:151–159 8. Siracusa MC, Reece JJ, Urban JF Jr, Scott AL (2008) Dynamics of lung macrophage activa­ tion in response to helminth infection. J Leukoc Biol 84:1422–1433 9. MacKinnon AC, Farnworth SL, Hodkinson PS et al (2008) Regulation of alternative mac­ rophage activation by galectin-3. J Immunol 180:2650–2658 10. Gangadharan B, Hoeve MA, Allen JE et  al (2008) Murine gammaherpesvirus-induced fibrosis is associated with the development of alternatively activated macrophages. J Leukoc Biol 84:50–58 11. Anderson CF, Gerber JS, Mosser DM (2002) Modulating macrophage function with IgG immune complexes. J Endotoxin Res 8:477–481 12. Zhang X, Goncalves R, Mosser DM (2008) The isolation and characterization of murine macrophages. Curr Protoc Immunol Chapter 14:Unit 14.1.:14.1.1–14.1.14 13. Riedy MC, Stewart CC (2001) Characterization of human monocytes/macrophages. Curr Protoc Immunol Chapter 14:Unit 14.3.: 14.3.1–14.3.8 14. Wahl LM, Wahl SM, Smythies LE, Smith PD (2006) Isolation of human monocyte popula­ tions. Curr Protoc Immunol Chapter 7:Unit 7.6A.:7.6A.1–7.6A.10 15. Ustyugova IV, Frost LL, VanDyke K, Brundage KM, Schafer R, Barnett JB (2007) 3,4-Dichloropropionaniline suppresses nor­ mal macrophage function. Toxicol Sci 97:364–374 16. Ouadrhiri Y, Scorneaux B, Sibille Y, Tulkens PM (1999) Mechanism of the intracellular kill­ ing and modulation of antibiotic susceptibility of Listeria monocytogenes in THP-1 mac­ rophages activated by gamma interferon. Antimicrob Agents Chemother 43:1242–1251 17. Vieira P, O’Garra A (2007) Regula‘ten’ the gut. Nat Immunol 8:905–907 18. Schmid D, Munz C (2007) Innate and adap­ tive immunity through autophagy. Immunity 27:11–21 19. Swanson MS, Byrne BG, Dubuisson JF (2009) Kinetic analysis of autophagosome formation and turnover in primary mouse macrophages. Methods Enzymol 452:383–402 20. Geissmann F (2007) The origin of dendritic cells. Nat Immunol 8:558–560

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21. Don Porto Carero A, Hoet PH, Nemery B, Schoeters G (2002) Increased HLA-DR expression after exposure of human mono­ cytic cells to air particulates. Clin Exp Allergy 32:296–300 22. Furst SM, Luedke D, Gandolfi AJ (1997) Kupffer cells from halothane-exposed guinea pigs carry trifluoroacetylated protein adducts. Toxicology 120:119–132 23. Harding CV (2001) Choosing and preparing antigen-presenting cells. Curr Protoc Immunol Chapter 16:Unit 16.1.:16.1.1–16.1.14 24. Harding CV (2001) Presenting exogenous antigen to T cells. Curr Protoc Immunol Chapter 16:Unit 16.2.:16.2.1–16.2.15 25. Dahlgren C, Karlsson A (1999) Respiratory burst in human neutrophils. J Immunol Methods 232:3–14 26. Lewis TL, Brundage KM, Brundage RA, Barnett JB (2008) 3,4-Dichloropropionanilide (DCPA) inhibits T cell activation by altering the intracellular calcium concentration follow­ ing store depletion. Toxicol Sci 103:97–107 27. Uchida N, Weissman IL (1992) Searching for hematopoietic stem cells: evidence that Thy1.1lo Lin- Sca-1+ cells are the only stem cells in C57BL/Ka-Thy-1.1 bone marrow. J Exp Med 175:175–184 28. Metcalf D (1971) Antigen-induced prolifera­ tion of bone marrow precusors of granulocytes and macrophages. Immunology 20:727–738 29. Blyler G, Landreth KS, Barnett JB (1994) Gender-specific effects of prenatal chlordane exposure on myeloid cell development. Fundam Appl Toxicol 23:188–193 30. Smith CL (2001) Basic confocal microscopy. Curr Protoc Cell Biol Chapter 4:Unit 4.5.: 4.5.1–4.5.12 31. Jedeszko C, Sameni M, Olive MB, Moin K, Sloane BF (2008) Visualizing protease activ­ ity in living cells: from two dimensions to four dimensions. Curr Protoc Cell Biol Chapter 4:Unit 4.20.:4.20.1–4.20.15 32. Henjakovic M, Sewald K, Switalla S et  al (2008) Ex vivo testing of immune responses in precision-cut lung slices. Toxicol Appl Pharmacol 231:68–76 33. Neumann K, Eppler E, Filgueira L et al (2003) Listeria species escape from the phagosomes of interleukin-4-deactivated human mac­ rophages independent of listeriolysin. Immunol Cell Biol 81:431–439 34. Frost LW, Neeley YX, Schafer R, Gibson LF, Barnett JB (2001) Propanil inhibits tumor necrosis factor-alpha production by reducing nuclear levels of the transcription factor NF-kB in the macrophage cell line IC-21. Toxicol Appl Pharmacol 172:186–193

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35. Hornbeck P (2001) Enzyme-linked immuno­ sorbent assays. Curr Protoc Immunol Chapter 2:Unit 2.1.:Unit 36. Hornbeck P, Winston SE, Fuller SA (2001) Enzyme-linked immunosorbent assays (ELISA). Curr Protoc Mol Biol Chapter 11:Unit11.2.: Unit11 37. Lee EO, Lee JR, Kim KH et  al (2006) The methylene chloride fraction of Trichosanthis Fructus induces apoptosis in U937 cells through the mitochondrial pathway. Biol Pharm Bull 29:21–25 38. Pan MH, Liang YC, Lin-Shiau SY, Zhu NQ, Ho CT, Lin JK (2000) Induction of apoptosis by the oolong tea polyphenol theasinensin A through cytochrome c release and activation of caspase-9 and caspase-3 in human U937 cells. J Agric Food Chem 48:6337–6346 39. Nakadai A, Li Q, Kawada T (2006) Chlorpyrifos induces apoptosis in human monocyte cell line U937. Toxicology 224: 202–209 40. Brundage KM, Schafer R, Barnett JB (2003) Altered AP-1 (activating protein-1) activity

41.

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44. 45.

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

R.R. Dietert (ed.), Immunotoxicity Testing: Methods and Protocols, Methods in Molecular Biology, vol. 598 DOI 10.1007/978-1-60761-401-2_7, © Humana Press, a part of Springer Science + Business Media, LLC 2010

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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

6.

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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

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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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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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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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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