Environmental Genomics

Environmental Genomics M E T H O D S I N M O L E C U L A R B I O L O G YTM John M. Walker, SERIES EDITOR 410. Env...

Author: C. Cristofre Martin

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Environmental Genomics

M E T H O D S

I N

M O L E C U L A R

B I O L O G YTM

John M. Walker, SERIES EDITOR 410. Environmental Genomics, edited by C. Cristofre Martin, 2007 409. Immunoinformatics: Predicting Immunogenicity In Silico, edited by Darren R. Flower, 2007 408. Gene Function Analysis, edited by Michael Ochs, 2007 407. Stem Cell Assays, edited by Mohan C. Vemuri, 2007 406. Plant Bioinformatics: Methods and Protocols, edited by David Edwards, 2007 405. Telomerase Inhibition: Strategies and Protocols, edited by Lucy Andrews and Trygve O. Tollefsbol, 2007 404. Topics in Biostatistics, edited by Walter T. Ambrosius, 2007 403. Patch-Clamp Methods and Protocols, edited by Peter Molnar and James J. Hickman, 2007 402. PCR Primer Design, edited by Anton Yuryev, 2007 401. Neuroinformatics, edited by Chiquito J. Crasto, 2007 400. Methods in Lipid Membranes, edited by Alex Dopico, 2007 399. Neuroprotection Methods and Protocols, edited by Tiziana Borsello, 2007 398. Lipid Rafts, edited by Thomas J. McIntosh, 2007 397. Hedgehog Signaling Protocols, edited by Jamila I. Horabin, 2007 396. Comparative Genomics, Volume 2, edited by Nicholas H. Bergman, 2007 395. Comparative Genomics, Volume 1, edited by Nicholas H. Bergman, 2007 394. Salmonella: Methods and Protocols, edited by Heide Schatten and Abe Eisenstark, 2007 393. Plant Secondary Metabolites, edited by Harinder P. S. Makkar, P. Siddhuraju, and Klaus Becker, 2007 392. Molecular Motors: Methods and Protocols, edited by Ann O. Sperry, 2007 391. MRSA Protocols, edited by Yinduo Ji, 2007 390. Protein Targeting Protocols, Second Edition, edited by Mark van der Giezen, 2007 389. Pichia Protocols, Second Edition, edited by James M. Cregg, 2007 388. Baculovirus and Insect Cell Expression Protocols, Second Edition, edited by David W. Murhammer, 2007 387. Serial Analysis of Gene Expression (SAGE): Digital Gene Expression Profiling, edited by Kare Lehmann Nielsen, 2007 386. Peptide Characterization and Application Protocols edited by Gregg B. Fields, 2007 385. Microchip-Based Assay Systems: Methods and Applications, edited by Pierre N. Floriano, 2007 384. Capillary Electrophoresis: Methods and Protocols, edited by Philippe Schmitt-Kopplin, 2007 383. Cancer Genomics and Proteomics: Methods and Protocols, edited by Paul B. Fisher, 2007 382. Microarrays, Second Edition: Volume 2, Applications and Data Analysis, edited by Jang B. Rampal, 2007

381. Microarrays, Second Edition: Volume 1, Synthesis Methods, edited by Jang B. Rampal, 2007 380. Immunological Tolerance: Methods and Protocols, edited by Paul J. Fairchild, 2007 379. Glycovirology Protocols edited by Richard J. Sugrue, 2007 378. Monoclonal Antibodies: Methods and Protocols, edited by Maher Albitar, 2007 377. Microarray Data Analysis: Methods and Applications, edited by Michael J. Korenberg, 2007 376. Linkage Disequilibrium and Association Mapping: Analysis and Application, edited by Andrew R. Collins, 2007 375. In Vitro Transcription and Translation Protocols: Second Edition, edited by Guido Grandi, 2007 374. Quantum Dots: edited by Marcel Bruchez and Charles Z. Hotz, 2007 373. Pyrosequencing® Protocols edited by Sharon Marsh, 2007 372. Mitochondria: Practical Protocols, edited by Dario Leister and Johannes Herrmann, 2007 371. Biological Aging: Methods and Protocols, edited by Trygve O. Tollefsbol, 2007 370. Adhesion Protein Protocols, Second Edition, edited by Amanda S. Coutts, 2007 369. Electron Microscopy: Methods and Protocols, Second Edition, edited by John Kuo, 2007 368. Cryopreservation and Freeze-Drying Protocols, Second Edition, edited by John G. Day and Glyn Stacey, 2007 367. Mass Spectrometry Data Analysis in Proteomics edited by Rune Matthiesen, 2007 366. Cardiac Gene Expression: Methods and Protocols, edited by Jun Zhang and Gregg Rokosh, 2007 365. Protein Phosphatase Protocols: edited by Greg Moorhead, 2007 364. Macromolecular Crystallography Protocols: Volume 2, Structure Determination, edited by Sylvie Doublié, 2007 363. Macromolecular Crystallography Protocols: Volume 1, Preparation and Crystallization of Macromolecules, edited by Sylvie Doublié, 2007 362. Circadian Rhythms: Methods and Protocols, edited by Ezio Rosato, 2007 361. Target Discovery and Validation Reviews and Protocols: Emerging Molecular Targets and Treatment Options, Volume 2, edited by Mouldy Sioud, 2007 360. Target Discovery and Validation Reviews and Protocols: Emerging Strategies for Targets and Biomarker Discovery, Volume 1, edited by Mouldy Sioud, 2007 359. Quantitative Proteomics by Mass Spectrometry edited by Salvatore Sechi, 2007 358. Metabolomics: Methods and Protocols, edited by Wolfram Weckwerth, 2007 357. Cardiovascular Proteomics: Methods and Protocols, edited by Fernando Vivanco, 2007

M E T H O D S I N M O L E C U L A R B I O L O G YT M

Environmental Genomics Edited by

C. Cristofre Martin Center for Advanced Research in Environmental Genomics (CAREG), Department of Biology, University of Ottawa, Ottawa, Ontario, Canada; and Department of Biochemistry, St. George’s University Medical School, St. George’s, Grenada, West Indies

Editor C. Cristofre Martin Center for Advanced Research in Environmental Genomics (CAREG) Department of Biology University of Ottawa, Ottawa, Ontario, Canada and Department of Biochemistry St. George’s University Medical School St. George’s, Grenada West Indies [email protected]

Series Editor John M. Walker University of Hertfordshire Hatfield, Herts United Kingdom

ISBN: 978-1-58829-777-8

e-ISBN: 978-1-59745-548-0

Library of Congress Control Number: 1588297772 ©2008 Humana Press, a part of Springer Science+Business Media, LLC 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, 999 Riverview Drive, Suite 208, Totowa, NJ 07512 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. Cover illustration: The cover background represents an image of Arabidopsis thaliana that in the case of this volume was utilized as a model system to study changes in genes expression associated with exposure of the plant to ozone. The overlay images depict the setup of a comet assay electrophoresis system that is employed to determine the relative amount of DNA breakage found in a single nucleated cell obtained from an environmental sample. Printed on acid-free paper 987654321 springer.com

Preface

Environmental genomics seeks to predict how an organism or organisms will respond, at the genetic level, to changes in their external environment. These genome responses are diverse and, as a result, environmental genomics must integrate molecular biology, physiology, toxicology, ecology, systems biology, epidemiology, and population genetics into an interdisciplinary research program. Environmental genomics is a generic term that applies to all studies examining the impact that environmental conditions have on gene transcription, protein levels, the stability of the genome itself, or the diversity of genomes in a population. Subsequent studies that follow these genomic surveys are typically rebranded to reflect more specific goals. For example, physiogenomics studies the dynamic changes in gene expression that can occur under different physiological or pathological conditions. Toxicogenomics investigates the effect of natural or human-made toxins on the genome, while metabolomics identifies alternations in metabolic byproducts. Ecological genomics assays the complement of genomes, called the biome, that are present in an environmental sample. Genomic polymorphisms in populations can also be assayed for susceptibility to adverse environmental conditions. In taking account the current fields that are critical for the full study of environmental genomics, we have divided this volume into three main parts: (1) gene expression profiling, (2) whole genome and chromosome mutation detection, and (3) methods to assay genome diversity and polymorphisms within a particular environment. Environmental genomic studies can be emulated in the laboratory using model systems (including humans) that are biologically well characterized and whose genome sequencing projects have been completed. The majority of environmental genomics research, however, involves the use of wild, nonmodel organisms whose genome information is limited or absent. The limitations imposed by studying a nonmodel organism make genomics particularly challenging. As a result, we have focused our attention on the genomic techniques that do not require whole genome sequence information. When considering possible technical strategies, to ask environmental genomic questions, it is apparent that there is not a discrete set of techniques that are utilized by these researchers. Rather, to date, most environmental genomics v

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studies have had to rely on relatively standard genomic and proteomic techniques that are not unique to this field. As a result, the contents of this volume may harbor some redundancy to the contents of certain other Methods in Molecular Biology volumes covering similar areas of genomics and proteomics. In addition, we are aware that many investigators entering the field of environmental genomics do not come from backgrounds in molecular biology and genomics. Instead, individuals are conducting much of this research with diverse backgrounds in environmental sciences, toxicology, and ecology. Consequently, we have tried to focus on protocols that are not overtly rigorous in technique and that can be accomplished in a reasonably standard molecular biology laboratory. Our goal is that this volume might ultimately serve as a manual for an environmental scientist who wishes to embrace genomics to answer environmental questions. Conversely, classical molecular biologists have begun to enter the foray. While these individuals have technological expertise they are often devoid of some of the special considerations that need to be made when conducting environmental studies. These experimental design and analysis considerations are particularly important as environmental studies often have important impacts on industrial activities, government policy, risk assessment models, and environmental health. We, therefore, have paid special attention, where possible, to include discussion on the importance of design, experimental controls, and interpretation of data. When conducted in a thoughtful manner, environmental genomics should provide information that is beneficial to our understanding of specific molecular targets of adverse or changing environmental conditions. Comparison of genomic data sets from model organism and those of wild, nonmodel organisms will allow us to understand better the value (if any) of extrapolating laboratory data to field determinations. Once ample data are collected, environmental genomics promises to facilitate the production of predictive models that may allow use to identify environmental threats prior to the appearance of an overt negative impact (e.g., toxin exposure). As a result, it is hoped that the sum of these studies will reduce the uncertainties associated with environmental risk assessment and provide a systematic framework for determining environmental impact and ensuring human health and the sustainability of natural populations. C. Cristofre Martin, B.Sc., M.Sc., PhD

Contents

Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Contributors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Part I: 1.

v ix

Gene Expression Proﬁling

High-Throughput Whole Mount In Situ Hybridization of Zebraﬁsh Embryos for Analysis of Tissue-Speciﬁc Gene Expression Changes After Environmental Perturbation Louise E. Coverdale, Lindsay E. Burton, and C. Cristofre Martin . . .

Fluorescent RNA Arbitrarily Primed Polymerase Chain Reaction Doug Crump, Suzanne Chiu, Vance L. Trudeau, and Sean W. Kennedy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3. Isolation of O3 -Response Genes from Arabidopsis thaliana Using cDNA Macroarray Masanori Tamaoki . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4. Use of cDNA Macroarrays and Gene Proﬁling for Detection of Effects of Environmental Toxicants Jason L. Blum, Melinda S. Prucha, Vishal J. Patel, and Nancy D. Denslow . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5. Constructing and Screening a cDNA Library Kevin Larade and Kenneth B. Storey . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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43 55

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Comparative Molecular Physiological Genomics Sean F. Eddy and Kenneth B. Storey . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 81 7. Proteomic Analysis of Neuroendocrine Peptidergic System Disruption Using the AtT20 Pituitary Cell Line as a Model Fumin Dong, Liming Ma, Michel Chrétien, and Majambu Mbikay . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 111 8.

Proteomics-Based Method for Risk Assessment of Peroxisome Proliferating Pollutants in the Marine Environment Susana Cristobal . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 123 9. Environmental Metabolomics Using 1 H-NMR Spectroscopy Mark R. Viant . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 137

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Part II:

Detection of Whole Genome Mutation

10.

Restriction Landmark Genome Scanning for the Detection of Mutations Jun-ichi Asakawa . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 153 11. Use of the Comet Assay in Environmental Toxicology Loren D. Knopper and James P. McNamee . . . . . . . . . . . . . . . . . . . . . . . . 171 12.

The Micronucleus Assay Determination of Chromosomal Level DNA Damage Michael Fenech. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 185 13. Fluorescence In Situ Hybridization for the Detection of Chromosome Aberrations and Aneuploidy Induced by Environmental Toxicants Francesca Pacchierotti and Antonella Sgura . . . . . . . . . . . . . . . . . . . . . . . 217 14. Laboratory Methods for the Detection of Chromosomal Structural Aberrations in Human and Mouse Sperm by Fluorescence In Situ Hybridization Francesco Marchetti, Debby Cabreros, and Andrew J. Wyrobek . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 241

Part III:

Determination of Species Diversity

15.

Assembling DNA Barcodes Jeremy R. deWaard, Natalia V. Ivanova, Mehrdad Hajibabaei, and Paul D. N. Hebert . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 275 16. Application of Suppressive Subtractive Hybridization to Uncover the Metagenomic Diversity of Environmental Samples Elizabeth A. Galbraith, Dionysios A. Antonopoulos, and Bryan A. White . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 295 17. 16S rRNA Targeted DGGE Fingerprinting of Microbial Communities Vesela A. Tzeneva, Hans G. H. J. Heilig, Wilma Akkermans van Vliet, Antoon D. L. Akkermans, Willem M. de Vos, and Hauke Smidt . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 335 18.

An Emulsion Polymerase Chain Reaction–Based Method for Molecular Haplotyping James G. Wetmur and Jia Chen . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 351

Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 363

Contributors

Antoon D. L. Akkermans • Laboratory of Microbiology, Wageningen University, Wageningen, The Netherlands Wilma Akkermans van Vliet • Laboratory of Microbiology, Wageningen University, Wageningen, The Netherlands Dionysios A. Antonopoulos • Michigan State University, Department of Microbiology & Molecular Genetics, East Lansing, MI Jun-ichi Asakawa • Department of Genetics, Radiation Effects Research Foundation, Hiroshima, Japan Jason L. Blum • Department of Pharmacology and Therapeutics, University of Florida, Gainesville, FL Lindsay E. Burton • Center for Advanced Research in Environmental Genomics (CAREG), Department of Biology, University of Ottawa, Ottawa, Ontario, Canada Debby Cabreros • School of Public Health, University of California at Berkeley, Berkeley, CA Jia Chen • Department of Community and Preventive Medicine, Mount Sinai School of Medicine, New York, NY Suzanne Chiu • National Wildlife Research Center, Canadian Wildlife Service, Carleton University, Ottawa, Ontario, Canada Michel Chrétien • Ottawa Health Research Institute, University of Ottawa, Faculty of Medicine, Ottawa, Ontario, Canada Louise E. Coverdale • Center for Advanced Research in Environmental Genomics (CAREG), Department of Biology, University of Ottawa, Ottawa, Ontario, Canada Susana Cristobal • Department of Biochemistry and Biophysics, Stockholm University, Stockholm, Sweden Doug Crump • National Wildlife Research Center, Canadian Wildlife Service, Carleton University, Ottawa, Ontario, Canada Nancy D. Denslow • Center for Environmental and Human Toxicology, Department of Physiological Sciences, University of Florida, Gainesville, FL Jeremy R. deWaard • Department of Integrative Biology, Biodiversity Institute of Ontario, University of Guelph, Guelph, Ontario, Canada ix

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Fumin Dong • Ottawa Health Research Institute, University of Ottawa, Faculty of Medicine, Ottawa, Ontario, Canada Sean F. Eddy • Women’s Health Interdisciplinary Research Center, Department of Biochemistry Boston University School of Medicine, Boston, MA Michael Fenech • CSIRO Human Nutrition, Adelaide BC, South Australia, Australia Elizabeth A. Galbraith • Agtech Products, Inc., Waukesha, WI Mehrdad Hajibabaei • Department of Integrative Biology, Biodiversity Institute of Ontario, University of Guelph, Guelph, Ontario, Canada Paul D.N. Hebert • Department of Integrative Biology, Biodiversity Institute of Ontario, University of Guelph, Guelph, Ontario, Canada Hans G. H. J. Heilig • Laboratory of Microbiology, Wageningen University, Wageningen, The Netherlands Natalia V. Ivanova • Department of Integrative Biology, Biodiversity Institute of Ontario, University of Guelph, Guelph, Ontario, Canada Sean W. Kennedy • National Wildlife Research Center, Canadian Wildlife Service, Carleton University, Ottawa, Ontario, Canada Loren D. Knopper • Jacques Whitford, Ottawa, Ontario, Canada Kevin Larade • Brigham and Women’s Hospital, Harvard Medical School, Hematology Division, Boston, MA Liming Ma • Ottawa Health Research Institute, University of Ottawa, Faculty of Medicine, Ottawa, Ontario, Canada Francesco Marchetti • Life Sciences Division, Lawrence Berkeley National Laboratory, Berkeley, CA C. Cristofre Martin • Center for Advanced Research in Environmental Genomics (CAREG), Department of Biology, University of Ottawa, Ottawa, Ontario, Canada; and Department of Biochemistry, St. George’s University Medical School, St. George’s, Grenada, West Indies Majambu Mbikay • Ottawa Health Research Institute, University of Ottawa. Faculty of Medicine, Ottawa, Ontario, Canada James P. McNamee • Health Canada, Consumer and Clinical Radiation Protection Bureau, Ottawa, Canada Francesca Pacchierotti • Section of Toxicology and Biomedical Sciences, ENEA CR Casaccia, Rome, Italy Vishal J. Patel • Bionomics Research and Technology Center, Environmental and Occupational Health Sciences Institute, Rutgers, The State University of New Jersey, Piscataway, NJ Melinda S. Prucha • Department of Pharmacology and Therapeutics, University of Florida, Gainesville, FL

Contributors

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Antonella Sgura • Department of Biology, Università Roma Tre, Rome, Italy Hauke Smidt • Laboratory of Microbiology, Wageningen University, Wageningen, The Netherlands Kenneth B. Storey • Institute of Biochemistry, Departments of Biology and Chemistry, Carleton University, Ottawa, Ontario, Canada Masanori Tamaoki • Biodiversity Conservation Research Project, National Institute for Environmental Studies, Tsukuba, Ibaraki, Japan Vance L. Trudeau • Centre for Advanced Research in Environmental Genomics (CAREG), University of Ottawa, Ottawa, Ontario, Canada Vesela A. Tzeneva • NIZO Food Research B. V., Ede, The Netherlands Mark R. Viant • School of Biosciences, University of Birmingham, Birmingham, UK Willem M. de Vos • Laboratory of Microbiology, Wageningen University, Wageningen, The Netherlands James G. Wetmer • Department of Microbiology, Mount Sinai School of Medicine, New York, NY Bryan A. White • University of Illinois at Urbana-Champaign, Departments of Animal Sciences & Pathobiology, Division of Nutritional Sciences, North American Consortium for Genomics of Fibrolytic Ruminal Bacteria, Urbana, IL Andrew J. Wyrobek • Life Sciences Division, Lawrence Berkeley National Laboratory, Berkeley, CA

I Gene Expression Proﬁling

1 High-Throughput Whole Mount In Situ Hybridization of Zebraﬁsh Embryos for Analysis of Tissue-Speciﬁc Gene Expression Changes After Environmental Perturbation Louise E. Coverdale, Lindsay E. Burton, and C. Cristofre Martin

Summary Whole mount in situ hybridization is a process that allows the visualization of gene expression (mRNA) within the cells of an intact organism. By comparing gene expression domains between organisms that have been subjected to different environmental conditions, an understanding of the cellular and tissue-specific effects of these environmental exposures can be identified. This technique is complementary to gene expression profiling techniques such as DNA microarrays which can usually provide information only on the differential levels of gene expression within an organism or tissue. In the case of whole mount in situ hybridization there is the added ability to detect differences in the distribution of cells, within a whole organism, expressing a particular gene. Subtle changes in the distribution of cells expressing a gene may not be reflected in the overall level of gene expression when RNA samples are retrieved from a whole organism and assayed. Exploitation of automation technology has made whole mount in situ hybridization a procedure that is amiable to high-throughput genomic studies. Combining automation with computer-aided image analysis makes this an efficient strategy for quantifying subtle changes in tissues and genes expression that can result from sublethal exposures to environmental toxins, for example. Key Words: Assay; hybridization; zebrafish.

automated;

gene

expression;

high-throughput;

From: Methods in Molecular Biology, vol. 410: Environmental Genomics Edited by: C. Cristofre Martin © Humana Press, Totowa, NJ

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1. Introduction Urbanization and human activities such as agriculture, forestry, and mining have introduced an ever increasing number of compounds into water supplies, groundwater, and natural aquatic environments at levels that have never previously been observed (1–3). As a result, governments, industry, and chemical producers are under increasing pressure to assay the potential effects of these chemicals in our environment. Traditional toxicological analysis provides parameters of acute toxicological effect (typically death); however, LC50 determinations are unable to identify the detrimental effects of low-level chronic exposure (4). Exposure of embryos to some compounds, even at low levels, can result in developmental perturbations that affect later life stages and the ability of organisms to mature and reproduce. Further, it is becoming increasingly apparent that obvious external abnormalities are not a requirement for decreased reproductive fitness in an organism. Thus, traditional types of analysis may not be able to identify all underlying problems that might occur as a result of these compounds being in our environment. The small- and large-scale gene expression profiling of embryos exposed to environmental toxins will allow us to understand common cellular mechanisms of toxicity as well facilitate the formulation of predictive models of toxicity for chemical families. Toxicological studies by our laboratory and others, using small-scale in situ hybridization analysis, have been very successful in identifying the effects that various compounds can have on vertebrate embryonic development (5–7). In situ hybridization is a process that facilitates the visualization of cell and tissue specific gene expression. This process labels cells, which contain mRNA that is complementary to an antisense gene probe, with a colored precipitate. Traditionally a very labor-intensive and time-consuming process, methodology efficiencies and recent advances in automation technology can be applied to this technique to assess the effects that a broad group of compounds may have on the embryonic development of invertebrate and vertebrate systems. Our ability to assay the expression patterns of a relatively large number of genes on a large sample size gives us the ability to identify compounds whose effects on embryonic development may not be overtly pathological but that can be identified only using statistical methods. Further, computeraided microscopy and image analysis software allow the automation of data collection such as morphometric measurements of gene expression domains. By using digital image analysis strategies such as pixel threshold techniques, measurements can be taken in an accurate, quantitative, and unbiased fashion. The large number of samples that can be analyzed fulfills the need for statistical significance and as a result facilitates the identification of subtle differences that might not be observed using qualitative methods.

In Situ Hybridization of Zebraﬁsh

5

The procedure outlined in this chapter has been optimized for use with zebrafish embryos; however, it can be utilized with other species with minimal modifications. 2. Materials All solutions should be made using RNase-free reagents and diethyl pyrocarbonate (DEPC)-treated distilled water. Instruments and tools that will make contact with embryos and solution reagents should be treated to be RNase free (see Note 1). 2.1. Equipment 1. Heat block for making in situ baskets. 2. In situ baskets: Cut off pointed tips of 1.5-mL microcentrifuge tubes or other cylindrical object and discard. Melt a piece of fine nylon mesh over one open end of remaining tube using the heat block at a low heat. Cut around the edges so no overhanging mesh remains. Be careful not to overheat, as this will melt the nylon mesh. 3. Six-well plates (Becton Dickinson, cat. no. 1146). 4. Flat tweezers to move in situ baskets between wells. 5. Glass pipets to move embryos from baskets and tubes.

2.2. Preparation of the Digoxigenin (DIG) Probes 1. 1 μg of purified linearized plasmid that contains the gene of interest (see Note 2). 2. 10× DIG label mix: 10 mM ATP, 10 mM CTP, 10 mM GTP, 6.5 mM UTP, 3.3 mM DIG-11-UTP (Roche). 3. Transcription buffer and bovine serum albumin (BSA)/dithiothreitol (DTT; supplied with enzyme). 4. Phenol–chloroform–isoamyl alcohol (PCI): ratio of 25:24:1 of phenol, chloroform, and isoamyl alcohol, respectively. Store in the dark at 4°C. 5. Chloroform–isoamyl alcohol (CIA): ratio of 24:1 of chloroform and isoamyl alcohol, respectively. Store at –20°C.

2.3. Preparation of Embryos 1. 4% PFA–PBS solution: 4% paraformaldehyde in 80 ml. of 1× phosphate-buffered saline (PBS) solution. Slowly add NaOH until paraformaldehyde is completely dissolved. Adjust pH to 7.2 with HCl. Add distilled water (dH2 O) to 100 mL. Store in 4°C for short term storage (1–2 weeks) and at –20°C for long-term storage. 2. 5× PBS: Dissolve 40 g of NaCl, 1 g of KCl, 14.4 g of Na2 HPO4 , and 2.4 g of KH2 PO4 . Add DEPC-treated water to 800 mL and adjust pH to 7.4 with HCl. Adjust the final volume to 1 L with DEPC-treated water. Dilute to make 1× PBS when appropriate.

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2.4. Whole Mount In Situ Hybridization 1. PBS Tween (PBST): 1× PBS and 0.1% Tween-20. 2. Proteinase K (50 μg/mL): dissolve 50 mg of proteinase K powder in water. Aliquot the proteinase K solution into tubes and store at –20°C. 3. Glycine solution: 2.7 mM glycine in 1× PBS. 4. 5× Preabsorbed anti-DIG antibody solution: anti-DIG antibody (Roche) 1:1000 ratio, 2% calf serum, 2 mg/mL BSA, and 20–50 fixed embryos in 1× PBST. Let solution sit for at least 1 h at room temperature (RT) or overnight at 4°C. Store at 4°C. Before use, dilute to 1× in PBST (see Note 3). 5. 20× SSC: 3 M NaCl and 0.3 M sodium citrate, pH to 7. Sterilize by autoclaving. 6. Hybridization mixture: 50% formamide, 5× SSC, 0.1% Tween-20, 50 μg/mL of heparin, 100 μg/mL of yeast tRNA, 9 mM citric acid. Store at –20°C. 7. Post-hybridization mixture (post-hyb mix): 50% formamide, 5× SSC, 0.1% Tween20, 50 μg/mL of heparin, 9 mM citric acid. Store at –20°C. 8. 100 ng probe (1–2 μL of resuspended probe) (Subheading 3.1.). 9. Blocking solution: 2% calf serum, 2 mg/mL of BSA, and add PBST to volume. 10. Staining buffer: 100 mM Tris-HCl, pH 9.5, 50 mM MgCl2 , 100 mM NaCl, 0.1% Tween-20, 0.4 mM levamisol. (See Note 4.) 11. NBT stock solution: 70 mg/mL of nitroblue tetrazolium (NBT) in dimethylformamide. Store at 4°C in the dark. (See Note 5.) 12. BCIP stock solution: 50 mg/mL of 5-bromo-4-chloro-3-indolyl (BCIP) in 70% dimethylformamide. Store at 4°C in the dark. (See Note 5.) 13. Embryo preservation solution: 1× PBS with 0.025% sodium azide.

3. Methods 3.1. Preparation of DIG-Labeled RNA Probe 1. Prepare the DNA template by digesting 10 μg of plasmid DNA (containing your target sequence) using the appropriate restriction endonuclease and according to the manufacturer’s instructions. The restriction enzyme should cut at the 3 end of the gene sequence to synthesize an antisense probe and cut at 5 end of the gene sequence to synthesize a sense control probe. (See Note 6.) 2. The template is purified further by using a commercially available column system or can be extracted with an equal volume of PCI and CIA. 3. The extracted template is them precipitated by the addition of 1/10 volume of 3 M sodium acetate and 2 volumes of 100% cold ethanol. 4. The solution is gently vortex-mixed followed by centrifugation at maximum speed for 30 min. 5. The DNA pellet is washed in cold 75% ethanol, briefly air dried, and resuspended in 10 μL of DEPC-treated water. 6. Establish the concentration of the resuspended (in DEPC-treated water) template using a spectrophotometer or by agarose gel electrophoresis with appropriate standards.

In Situ Hybridization of Zebraﬁsh

7

7. Synthesis of the probe requires the following reaction, which should not exceed a total volume of 20 μL. The probe synthesis reaction consists of 1 μg of linearized DNA template, 1× DIG label mix, 1× transcription buffer (supplied with enzyme), DTT or BSA (supplied with enzyme), 40 U RNasin (Promega), and 20–50 U of RNA polymerase. Incubate at the appropriate temperature and time (typically 1 h) for the particular RNA polymerase used. 8. Add another 20–50 U of RNA polymerase and incubate for an additional hour. 9. Add 40 U of DNase I enzyme and incubate at 37°C for 10 min. (See Note 7.) 10. Precipitate the probe by adding to a final concentration 0.02 M EDTA, 0.5 M LiCl, and 70% Ethanol at –80°C for 1–2 h or O/N. 11. Pellet the probe through centrifugation at maximum speed for 30 min at 4°C. 12. Wash the pellet with 99% ethanol, air dry, and resuspend in 50 μL of DEPC-treated water or less. Store the probe at –80°C. Test the quality of the probe by running 2 μL on an ethidium bromide stained agarose gel. (See Note 8.)

3.2. Preparation of Zebraﬁsh Embryos Unless indicated all embryo manipulations take place in small homemade baskets contained within a single well of a six-well plate (Fig. 1). A single well can hold six baskets. Baskets are moved between the wells containing different incubation solutions. Use a pair of flat tipped forceps to move baskets between wells.

Fig. 1. Schematic diagram depicting homemade baskets made by cutting the ends of either a BEEM capsule or 1.5-mL microcentrifuge tube and adhering a nylon mesh to one end. The mesh can be adhered using a glue adhesive or by gently melting the tube end. Embryos are contained within these baskets and solution incubations are conducted in a six-well plate. Baskets are moved between wells using forceps.

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1. Fix dechorionated embryos overnight in 4% PFA at 4°C. 2. Wash embryos in 1× PBS before dehydrating and storing in methanol at –20°C. 3. Embryos should be stored in methanol at least overnight before being used for in situ hybridization. Embryos can be stored in methanol at –20°C for years and still be used for in situ hybridization.

3.3. Whole Mount in situ Hybridization 1. Selected embryos are rehydrated with successive incubations of 5 min in 75% MeOH–25% PBS, 50% MeOH–50% PBS, 25% MeOH–75%PBS, and 4 incubations of 5 min in PBST. The embryos that are rehydrated will be used for the in situ hybridization process and for preparing the preabsorbed antibody solution. Prepare the preabsorbed antibody with the rehydrated embryos according to Subheading 2.3.4. 2. Treat experimental embryos 24 h and older with proteinase K (0.1 μg/mL) in PBST for a predetermined time. (See Note 9.) 3. Quickly wash embryos in PBST and incubate in 2 mg/mL of glycine in PBST for 5 min. 4. Fix embryos in 4% PFA–PBS for 20 min before washing again in PBST. 5. Using a pipet, embryos are transferred to 1.5-mL microcentrifuge tubes and prehybridized in hybridization buffer for at least 1 h at 65°C. 6. Hybridize overnight at 65°C in fresh hybridization solution with 1–2 μL of probe per 100 ul. 7. Using a pipet, the embryos are transferred to baskets for the remainder of processing. 8. Embryos are washed in successive incubations of 10 min at 65°C in: 75% post hyb-mix–25% 2× SSC 50% post hyb-mix–50% 2× SSC 25% post hyb-mix–75% 2× SSC 100% 2× SSC 9. The embryos are then washed twice for 30 min in 0.2× SSC at 60°C. The embryos are then washed in successive incubations of 5 min at room temperature in: 75% 0.2× SSC–25% PBST 50% 0.2× SSC–50% PBST 25% 0.2× SSC–50% PBST 100% PBST

3.4. Staining and Detection 1. The embryos are incubated in the blocking solution at room temperature for 1–4 h, and subsequently incubated in the preabsorbed anti-DIG antibody (1:5000) for 3–4 h at room temperature.

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2. To remove any excess antibody, the embryos are washed 6 times in PBST at room temperature for 15 min. (See Note 10.) 4. The embryos are then pre-incubated in the staining buffer for 5 min at room temperature. 5. The staining reaction occurs in staining buffer containing 0.40 mM NBT and 0.40 mM BCIP). The duration of the staining reaction is dependent on the probe and expected of gene expression. The staining reaction should be monitored every 15–30 min to determine the proper duration of the reaction. Embryos can be viewed while remaining in the baskets using a standard stereo dissecting microscope. The staining process is conducted in the dark and exposure to light should be minimized. The longer the embryos are in the stain mix the darker the staining gets, but if stained for too long high background staining can result and thus making it hard to differentiate between true staining and background. This is particularly true when the level of gene expression being assayed is low. (See Note 11.) 6. Following a satisfactory stain, the embryos are washed 2× in PBST at room temperature for 15 min. 7. Postfixed in 4% PFA for 2 h at RT or overnight at 4°C. 8. The embryos are then washed in PBS and can be stored in 0.025% sodium azide– PBS at 4°C for an indefinite amount of time.

3.5. Automated In Situ Hybridization The following automated protocol utilizes an Insitu Pro Automated In Situ Hybridization system by Intavis Bioanalytical Instruments AG (Koeln, Germany). For this procedure the embryos have been rehydrated and treated with proteinase K before being put into the machine (up to step 3.3.3) and requires staining to be done manually (starting at step 3.4.4). The program begins at room temperature (T0[OFF]) with the rinsing of the machine. The embryos are then incubated in 125 μL of hybridization buffer (C) for 20 min at room temperature (step 5) before the temperature is raised to 65ºC (T2[HIGH]) and incubated for 90 min in fresh hybridization solution. The probe is then added to the embryos and they are incubated at 65ºC for 14 h (step 8). The embryos are washed in successive 20-min intervals in 150 μL of solutions D, H, F, and E, which are varying concentrations of hybridization solution and SSC (steps 9–13). The temperature once again returns to room temperature (T0(OFF)) and the embryos are further washed for 20 min in 150 ul of solutions I, J, K, and A. The embryos are then incubated for 60 min in 120 μL of solution L (blocking solution) before being incubated for 4 h in 120 μL of solution M (DIG antibody). The embryos are then washed 7 times in 150 μL of solution A (PBST) for 20 min before the program waits for user input (step 30 waiting NTMT). The user can then remove the embryos from the machine and proceed directly to step 3.4.4. The machine then cleans itself before the end of the program. Using this protocol up to 500 embryos can be processed in less than 48 h.

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The holding tubes used with the Insitu Pro system and the solutions placed within each of these containers are indicated below (see Note 12): A: PBST B: 50% methanol C: hybridization buffer D: 75% hybridization buffer + 25% 2× SSC E: 0.2× SSC F: 2× SSC H: 25% hybridization buffer + 75% 2× SSC I: 75% 0.2× SSC + 25% PBST J: 50% 0.2× SSC + 50% PBST K: 25% 0.2× SSC + 75% PBST L: Blocking solution M: DIG antibody PROBE: DIG-labeled RNA probe

The program code is indicated below: EMBRYO PROGRAM: Name

Param

1 SetTempReg 2 Rinse 3 Aliquot 4 Wait 5 Incubate 6 SetTempReg 7 Incubate 8 Incubate 9 Incubate 10 Incubate 11 Incubate 12 Incubate 13 Incubate 14 SetTempReg 15 Wait 16 Incubate 17 Incubate 18 Incubate 19 Incubate 20 Incubate 21 Incubate 22 Incubate 23 Incubate

T0(OFF) 5000 / 5000 ul 100 C-SAMPLE 5 min 20 min 125 C-SAMPLE T2 (HIGH) 90 min 140 C-SAMPLE 14 h 150 Probe-SAMPLE 20 min 150 D-SAMPLE 20 min 150 H-SAMPLE 20 min 150 F-SAMPLE 20 min 150 E-SAMPLE 20 min 150 E-SAMPLE T0 (OFF) 20 min 20 min 150 I-SAMPLE 20 min 150 J-SAMPLE 20 min 150 K-SAMPLE 20 min 150 A-SAMPLE 20 min 150 A-SAMPLE 60 min 120 L-SAMPLE 4 h 120 M-SAMPLE 20 min 150 A-SAMPLE

In Situ Hybridization of Zebraﬁsh 24 Incubate 25 Incubate 26 Incubate 27 Incubate 28 Incubate 29 Incubate 30 WaitForKey 31 Rinse 32 SetTempReg

11

20 min 150 A-SAMPLE 20 min 150 A-SAMPLE 20 min 150 A-SAMPLE 20 min 150 A-SAMPLE 20 min 150 A-SAMPLE 20 min 150 A-SAMPLE Waiting NTMT 5000/5000 μL T0 (OFF)

3.6. Quantiﬁcation of In Situ Hybridization Signal Quantification of the in situ hybridization signal is accomplished by the use of digital photomicroscopy and relatively basic image analysis software. A digital image is taken of the processed embryos and saved in a format usable by image analysis software. To ensure that no bias is entered into the data collection, imaging, and analysis should be conducted in a blinded fashion. In addition, it is necessary before image capture to ensure that all the embryos are oriented in a similar way so that perspective does not alter any of the measurements taken from the images. To accomplish this, embryos can be stabilized for digital photograph by placing them in a Petri dish containing a layer of agarose. Using small forceps or pipet, small grooves or holes can be made in the agar and embryos can be placed in these depressions to ensure stable orientation. Finally, uniform lighting should be applied so color values of your in situ signal are as uniform across samples as possible. After the acquisition of the embryos image, the image is analyzed using any number of image analysis software packages. These may include Adobe Photoshop (www.adobe.com), NIH Image (http://rsb.info.nih.gov/nihimage), SimplePCI (http://www.cimaging.net/), or Metamorph (http://www. moleculardevices.com/pages/software/metamorph.html). All of these softwares possess the ability to measure various image parameters (area, length, intensity etc.) and in most cases export this data to a usable format such as a spreadsheet for further analysis. Pixel thresholding techniques can be used to automatically identify cells which possess the in situ hybridization signal. This is typically accomplished by instructing the imaging software to select pixels of a certain spectral color (usually blue in the case of a NBT/BCIP staining protocol). Once pixels are automatically selected, the software itself can make measurements based on the selected pixels. Figure 2 shows an example of an analysis conducted on zebrafish embryos exposed to cadmium chloride. To determine whether or not cadmium exposure might affect embryonic brain development, we conducted an in situ hybridization using a probe whose

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Fig. 2. Treatment of zebrafish embryos with cadmium chloride results in abnormal development of the brain. (A) Whole mount in situ hybridization of a 24-h postfertilization zebrafish embryo using a eng2 antisense probe. Eng2 mRNA is localized to the midbrain/hindbrain boundary (dark stain). (B) Pixel threshold techniques are utilized to define (shaded area) and measure the expression domain of eng2. (C) Histogram showing the area (in arbitrary pixel units) of the eng2 expression domain in cadmium chloride treated and untreated 24 hour post fertilization zebrafish embryos. (*p < 0.05). (Reprinted from Coverdale and Martin, 2004, with permission) (8).

expression is limited to a discrete domain in the midbrain/hindbrain boundary. By analyzing 100 control and 100 treated embryos, and measuring the area of the Eng2 expression domain, we were able to demonstrate that cadmium exposure resulted in a statistically significant expansion in this tissue. 4. Notes 1. Dry solid reagents should be taken from a new unopened bottle and used exclusively for RNase-free work. Most dry reagents can be purchased as certified RNase free. DEPC-treated water is made by adding 1 mL of diethyl pyrocarbonate (DEPC) per 1000 mL of distilled water. The solution is shaken vigorously to dissolve the DEPC and allowed to stand for 1 hr. The solution is then autoclaved and allowed to cool before use. Alternatively, DEPC-treated water can be purchased from commercial suppliers. Storage bottles and other instruments should be treated with commercially available RNase decontamination solutions such as RNA Zap (Ambion). 2. Plasmid used for in vitro transcription of the RNA probes will contain RNA polymerase promoter sites such as T7, T3, or SP6. Examples of these plasmids are pCR2 (Invitrogen) or Bluescript II (Stratagene).

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3. The embryos used to prepare the preabsorbed antibody should be of the same developmental stage as the embryos that are going to be subjects of the in situ hybridization procedure. In addition, embryos can be chopped or ground and added to solution. In this case, the preabsorbed antibody should be allowed to settle of a period of 24 h or briefly centrifuged at low speed to remove embryonic debris from solution. 4. Levamisol stock solutions should be made fresh. Levamisol is added to reduce the activity of endogenous alkaline phosphatase activity within embryonic tissues and can reduce background staining. It does, however, reduce the overall efficiency of the staining process. Therefore if background staining is not a problem in certain samples the levamisol can be omitted. 5. Exposure of these reagents to light should be minimized as much as possible. 6. Avoid enzymes that digest to leave 3 overhangs. This type of overhang can apparently reduce the efficiency of the in vitro transcription reaction. 7. The addition of DNase following the riboprobe synthesis has been found to reduce background in some cases. Certified RNase-free DNase should be used in this case. If background is not a problem then this step can be eliminated. 8. Often a smear of RNA fragments may be observed when checking the probes on an agarose gel. Often these probes are still usable; however, ideally a single riboprobe species observed on the gel is best. 9. Sources of proteinase K vary in their activity. It is necessary to empirically test each batch of protein K stock solution to obtain optimal incubation times. Because of their fragility, 24-h embryos should be left for a lesser time (1 min) than 48-h embryos (2–4 min) and adult tissues (5–10 min). If the embryos are left in the proteinase K for too long they will be more prone to disintegration. Inadequate treatment with proteinase K will result in significantly reduced staining signal. 10. The embryos can be stored in PBST overnight at 4°C if needed. 11. Embryos should be left with a purple colored stain. If after approx 3 h the embryos have not stained, repeat the staining procedure with fresh staining mix and BCIP– NBT. Embryos can be left overnight in stain mix at 4°C. 12. The volume of these solutions required needs to be calculated and is based on the number of samples running on the machine.

Acknowledgments This work was supported by a grant to C. C. Martin from the Natural Sciences and Engineering Research Council of Canada (NSERC). References 1. Kolpin, D. W., Furlong, E. T., Meyer, M. T., Thurman, E. M., Zaugg, S. D., Barber, L. B., and Buxton, H. T. (2002) Pharmaceuticals, hormones, and other organic wastewater contaiminants in U.S. streams, 1999–2000: a national reconnaissance. Environ. Sci. Technol. 36, 1202–1211.

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2. Mandal, R., Hassan, N. M., Murimboh, J., Chakrabarti, C. L., Back, M. H., Rahayu, U., and Lean, D. R. S. (2002) Chemical speciation and toxicity of nickel species in natural waters from the Sudbury area (Canada). Environ. Sci. Technol. 36, 1477–1484. 3. Squillace, P. J., Scott, J. C., Moran, M. J., Nolan, B. T., and Kolpin, D. W. (2002) VOCs, pesticides, nitrate, and their mixtures in groundwater used for drinking water in the United States. Environ. Sci. Technol. 36, 1923–1930. 4. Nagel, R. and Isberner, K. (1998) Testing of chemicals with fish – a critical evaluation of tests with special regard to zebrafish. In: Braunbeck, T., Hinton, D. E. and B. Streit (Eds.) Fish Ecotoxicology, Birkh¨auser Verlag Basel/Switzerland, 338–352. 5. Lele, Z., Hartson, S. D., Martin, C. C., Whitsell, L., Matts, R. L., and Krone, P. H. (1999) Disruption of zebrafish somite development by pharmacologic inhibition of Hsp90. Dev. Biol. 210, 56–70. 6. Martin, C. C., LaForest, L., Akimenko, M-A., and Ekker, M. (1999) A role for DNA methylation in gastrulation and somite patterning. Dev. Biol. 206, 189–205. 7. Ellies, D. L., Langille, R. M., Martin, C. C., Akimenko, M-A, and Ekker, M. (1997) Specific craniofacial cartilage dysmorphogenesis coincides with a loss of dlx gene expression in retinoic acid treated zebrafish embryos. Mech. Dev. 61, 23–36. 8. Coverdale, L. E. and Martin, C. C. (2004) Not just a fishing trip—environmental genomics using zebrafish. Curr. Genom. 5, 299–308.

2 Fluorescent RNA Arbitrarily Primed Polymerase Chain Reaction A New Differential Display Approach to Detect Contaminant-Induced Alterations of Gene Expression in Wildlife Species Doug Crump, Suzanne Chiu, Vance L. Trudeau, and Sean W. Kennedy

Summary Differential display polymerase chain reaction (PCR) can facilitate the identification of novel molecular end points related to contaminant exposure in a wide range of species. To date, various differential display methodologies have been described in detail. Herein, we describe a modification of the RNA arbitrarily primed PCR (RAP-PCR) method that involves the fluorescent labeling of cDNA transcripts via 5 rhodamine-labeled 18-mer arbitrary primers. These arbitrary primers typically bind to the coding regions of cDNA, which simplifies the downstream identification of contaminant-responsive genes. The technique has been aptly named fluorescent RNA arbitrarily primed PCR, FRAP-PCR, and has been successfully utilized with several avian species and RNA sources (e.g., cultured cells, tissue). This straightforward, safe, and cost-effective approach represents a useful alternative to the radiometric-based RAP-PCR method. Key Words: Differential display; fluorescence; gene expression; method; RAP-PCR; toxicogenomics.

1. Introduction The “omics” revolution has provided a unique opportunity to explore and develop techniques that enable novel endpoint discovery at the level of the transcriptome, proteome, and metabolome. More specifically, toxicogenomics has emerged as an essential link between the transcriptome and the From: Methods in Molecular Biology, vol. 410: Environmental Genomics Edited by: C. Cristofre Martin © Humana Press, Totowa, NJ

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impacts of environmental toxicants. Differential display polymerase chain reaction (DD-PCR) represents a modern molecular biological tool that was developed to facilitate the characterization of differentially expressed genes and has realized use in a wide range of studies since its discovery (1). RNA arbitrarily primed PCR (RAP-PCR) is a related approach in which primers of arbitrary sequence are annealed with RNA to generate reproducible profiles that detect polymorphisms in expressed transcripts (2,3). The technique is amenable for use with a wide range of species and does not require any previous knowledge of the species’ genome. This allows for novel gene discovery as it pertains to exposure of laboratory-reared and free-living organisms to environmental toxicants. To date RAP-PCR has been used in gene expression studies with species such as the goldfish (4), leopard frog (5), snapping turtle (6), Fundulus heteroclitus (7), and the cladoceran, Daphnia magna (8). Similar to the trend observed in the field of microarrays, DD-PCR technologies have incorporated the use of commercially available fluorescent dyes in order to minimize the reliance on radioisotopes for labeling. The initial attempts of fluorescent DD-PCR involved 3 -anchored primers labeled at the 5 end with rhodamine, fluorescein, or FITC (9–12). Diener et al. (8) developed a modified RAP-PCR method in which PCR products were labeled with a fluorescent (Cy5) adapter primer following PCR. To streamline the technique and remove the necessity of a secondary labeling step, we have developed a method called fluorescent RNA arbitrarily primed PCR (FRAP-PCR), which incorporates a fluorescently labeled arbitrary primer into the PCR. Throughout method development, we wanted to assess the versatility of FRAP-PCR with several different species and tissue types after exposure to various environmental contaminants. Novel gene targets have been identified in neuronal and hepatic avian cells as well as brain and liver tissue in species such as the herring gull, mallard duck and chicken following exposure to contaminants including polybrominated diphenyl ethers, the rodenticide Brodifacoum, and dioxin. This technique extends and improves the radiometric-based RAP-PCR and is an attractive approach for researchers trying to identify novel molecular mechanisms of action of various environmental contaminants in a wide range of species. 2. Materials All materials can be stored at room temperature (RT) unless otherwise stated. 2.1. RNA Isolation from Tissue or Cultured Cells 1. TRIzol Reagent (Invitrogen). This solution contains phenol and thiocyanate compounds and should be handled wearing gloves and a lab coat. Store at 4°C.

Fluorescent RNA Arbitrarily Primed PCR 2. 3. 4. 5. 6.

17

Chloroform. Isopropanol. Ethanol (EtOH). Diethyl pyrocarbonate (DEPC)-treated water. DNA-free kit (Ambion). Store at –20°C.

2.2. cDNA Synthesis 1. SuperScript™ II, RNase H-Reverse Transcriptase (200 U/μL; Invitrogen). Stable for more than 2 yr at –20°C. 2. 5× First-strand buffer: 250 mM Tris-HCl, pH 8.3, 375 mM KCl, 15 mM MgCl2 . Store at –20°C. 3. 0.1 mM dithiothreitol (DTT). Store at –20°C. 4. dNTP mix: 10 mM of each dNTP. Store at –20°C. 5. 18-mer arbitrary primers (25 μM): A3, 5 -AATCATGAGCTCTCCTGG-3 ; B3, 5 -CATACACGCGTATACTGG-3 ; C3, 5 -CCATGCGCATGCATGAGA-3 . Store at –20°C. 6. RNase OUT™ Recombinant Ribonuclease Inhibitor (40 U/μL; Invitrogen). Store at –20°C.

2.3. FRAP-PCR 1. Qiagen Taq DNA polymerase (5 U/μL; Qiagen). Store at –20°C in a constant temperature freezer. 2. Qiagen 10× PCR buffer: Tris-HCl, KCl, (NH4 )2 SO4 , 15 mM MgCl2 , pH 8.7. Store at –20°C. 3. 25 mM MgCl2 . Store at –20°C. 4. DEPC-treated water. 5. dNTP mix: 10 mM of each dNTP. Store at –20°C. 6. Arbitrary primers labeled at the 5 end with rhodamine (10 μM; A3, B3, or C3). Store at –20°C and keep away from direct light. 7. QIAquick PCR Purification Kit (Qiagen).

2.4. Gel Electrophoresis 1. Gel solution: 6% acrylamide (29:1 acrylamide–bis-acrylamide) containing 7 M urea and 1× Tris–borate–EDTA (TBE) buffer. 2. 1× Tris–borate–EDTA (TBE) buffer: 89 mM tris base, 89 mM boric acid, 2 mM disodium EDTA, pH 8.3. 3. Loading dye: 99% formamide, 1 mM EDTA, pH 8, 0.009% xylene cyanol FF, and 0.009% bromophenol blue. 4. N,N,N ,N -Tetramethylethylenediamine (TEMED). 5. 25% Ammonium persulfate. 6. FDD Vertical Electrophoresis system with 60-well shark-tooth comb and lowfluorescence glass plates (GenHunter).

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7. 8. 9. 10.

Sigmacote (Sigma-Aldrich). Store at 4°C. External DC voltage power supply that is rated for >1700 V. Fluorescence imager: Typhoon 9210 Variable Mode Imager (Amersham). Inkjet printer that has the capacity for 11-in. × 17-in. paper for printing large gel images in actual size. 11. Razor blades.

2.5. PCR Product Isolation, Reampliﬁcation, and Cloning 1. 2. 3. 4. 5. 6. 7.

8. 9. 10. 11. 12. 13.

Glycogen (20 μg/μL). Store at –20°C. 3 M sodium acetate. EtOH. DEPC-treated water. PCR reagents as in Subheading 2.3. (Exception – 10 μM unlabelled arbitrary primers). TOPO® TA cloning kit (Invitrogen). One Shot® Chemically Competent E. coli should be kept at –80°C and the pCR-TOPO® vector at –20°C. Add 125 μL of SOC medium (2% tryptone, 0.5% yeast extract, 10 mM NaCL, 2.5 mM Kcl, 10 mM MgCl, 10 mM MgSO4 , 20 mM glucose) and shake sample horizontally (200 rpm) at 37°C for 1h. Luria-Bertani (LB) media. Ampicillin (10 mg/mL). Store at 4°C. Agar. X-gal (20 mg/mL). Store at –20°C away from direct light. M13 primers: forward (5 -GTAAAACGACGGCCAG-3 ) and reverse (5 CAGGAAACAGCTATGAC-3 ). Store at –20°C. QIAprep Spin Miniprep Kit (Qiagen).

3. Methods 3.1. RNA Isolation from Tissue or Cultured Cells RNA can be isolated from a variety of sources including cultured cells (e.g., neurons or hepatocytes) or whole tissue (see Note 1). Purification of mRNA is not required for FRAP-PCR which is ideal because mRNA typically comprises only about 3–5% of the total cellular RNA pool (13). Thus, the methodology described in this chapter is amenable to situations where yields of total RNA are expected to be limited (e.g., minute quantities of cells/tissue). Total cellular RNAs are easily purified but must be treated with DNase before use in FRAP-PCR (see Note 2) and aliquoted into single-use tubes (see Note 3). Total RNA input should be standardized within experiments and should be approx 750 ng (see Note 4). 1. Lyse individual wells of primary cells (approx weight = 1.2 mg) in 100 μL of TRIzol Reagent or add 1 mL per 50–100 mg of tissue and dissociate using the

Fluorescent RNA Arbitrarily Primed PCR

2. 3. 4. 5.

6. 7. 8. 9.

10. 11.

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Mixer Mill MM 300 (Retsch) and tungsten carbide beads for 2 min at 20 Hz. (See Note 5.) Incubate the homogenized samples for 5 min at RT to permit the complete dissociation of nucleoprotein complexes. Add 0.2 mL of chloroform per 1 mL of TRIzol, shake tubes vigorously by hand for 15 s, and incubate them at RT for 2–3 min. Centrifuge at 12,000g for 15 min at 4°C. Transfer the aqueous phase (60% of initial volume of TRIzol used) to a fresh tube. Add 0.5 mL of isopropyl alcohol per 1 mL of TRIzol and mix by inversion several times. Incubate samples at RT for 10 min and centrifuge at 12,000 g for 20 min at 4°C. Remove the supernatant. Wash the RNA pellet once with 1 mL of 75% EtOH per 1 mL of TRIzol, mix by vortex-mixing and centrifuge at 7500g for 5 min at 4°C. Dry the RNA pellet at RT and dissolve it in ≤100 μL of DEPC-treated water. Determine the RNA concentration by spectrophotometry. To remove genomic DNA from RNA, add 5 μL of DNase I buffer and 1 μL of recombinant DNase I (2 U/μL) to 10 μg of RNA in a 50-μL reaction. Incubate at 37°C for 20–30 min. Add 10 μL of resuspended DNase inactivation reagent and incubate for 2 min at RT. Centrifuge at 10,000g for 1.5 min and transfer the RNA to a fresh tube. Determine the new, DNase-treated RNA concentration by spectrophotometry, dilute to 75 ng/μL, and store 10-μL aliquots at –80°C.

3.2. cDNA Synthesis 18-mer arbitrary primers are used rather than oligo-dT primers in this technique to ensure that the transcripts synthesised are not biased towards the 3 untranslated region. This allows internal RNA fragments to be sampled, including the open reading frame, which facilitates downstream identification of gene products. This is especially important in applications with species that have relatively uncharacterized genomes (e.g., herring gull, mallard duck). A control without the reverse transcriptase enzyme should be performed to verify the absence of contaminating genomic DNA in subsequent PCR steps (no RT control). In addition, a commercially available control RNA (e.g., DNA-free total RNA from transformed rat embryo fibroblasts; GenHunter) should be included as a control for reverse-transcription dependent amplification of mRNAs. 1. Combine 10 μL of 75 ng/μL total RNA, 1 μL of dNTP mix, and 1 μL of arbitrary primer (A3, B3, or C3; 25 μM). Incubate the 12-μL mixture at 65°C for 5 min in a thermocycler followed by a quick chill on ice and a brief centrifugation. 2. Add 4 μL of 5× first-strand buffer, 2 μL of DTT, 1 μL of RNase Out, and 1 μL of SuperScript™ II Reverse Transcriptase to the mixture. 3. Incubate at 25°C for 10 min followed by 50 min at 42°C. 4. Terminate the reaction by heating at 70°C for 15 min. 5. Store cDNA at –20°C in 5-μL aliquots for subsequent PCR (see Note 6).

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3.3. FRAP-PCR Rhodamine is a light-sensitive dye. Thus, for all procedures involving primer stocks, working solutions or PCR products containing labeled primers, work must be conducted under low-light conditions. 1. Each 25-μL reaction should contain: 5 μL of cDNA, 2.5 μL of 10× PCR buffer, 1.5 μL of MgCl2 , 0.5 μL of dNTP mix, 1.25 μL of appropriate rhodamine-labeled primer (A3, B3, or C3), 0.1 μL of Qiagen Taq DNA polymerase and 14.15 μL of DEPC-treated water. 2. PCR is performed in a thermocycler under the following conditions: Initial Denaturation

94°C, 5 min

Stage I (1 cycle) (See Note 7)

94°C, 36°C, 72°C, 94°C, 54°C, 72°C, 72°C,

Stage II (30 cycles)

Final primer extension 3. 4. 5. 6. 7. 8. 9. 10. 11.

1 min 5 min 5 min 1 min 2 min (See Note 8) 2 min 10 min

Store samples at 4°C in the dark. Use QIAquick Nucleotide Removal Kit for PCR cleanup. (See Note 9.) Add 125 μL of buffer PB to 25 μL of PCR sample and mix. Pipet the entire sample onto a QIAquick column placed in a 2-mL collection tube and centrifuge (17,900g) for 30–60 s. Discard flow-through and place the QIAquick column back in the same tube. Add 0.75 mL of buffer PE to QIAquick column and centrifuge for 30–60 s. Discard flow-through and place the QIAquick column back in the same tube. Centrifuge for an additional 1 min to remove residual buffer PE. Place QIAquick column in a clean 1.5-mL microcentrifuge tube. To elute DNA, add 30 μL of buffer EB to the center of the QIAquick membrane, let the column stand for 1 min, and then centrifuge.

3.4. Gel Electrophoresis 1. Clean the inside surfaces of the glass plates thoroughly and treat the inside of the notched plate with Sigmacote. Assemble the glass plate sandwich and tape the edges to prevent leakage of the gel solution prior to polymerization. 2. Prepare 6% denaturing polyacrylamide gel. For the vertical system (45 cm long, 25 cm wide), approx 50 mL of the polyacrylamide solution is used. To this add 100 μL of fresh 25% ammonium persulfate and 50 μL of TEMED and mix. 3. Cast the gel at a slight angle to the bench top, lower slowly to a horizontal position, insert the casting comb, and let it polymerize overnight. Cover the top of the gel sandwich with a wetted Kimwipe and Saran Wrap to avoid drying out.

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4. Following polymerization, pre-run the gel for 45 min at 1500 V. Approx 750 mL of 1× TBE running buffer is necessary to fill the upper and lower buffer reservoirs. 5. Mix 30 μL of each sample and appropriate controls (no RT and control RNA) with 15 μL of FDD loading dye and incubate at 80°C for 2 min before loading 5 μL onto the gel (see Note 10). Before loading, flush urea out of sample wells using a Pasteur pipet. Load samples in triplicate. 6. Run the gel at 1700 V for 6 h, remove the Sigmacote-treated plate, and scan the gel on a Typhoon Imager (at 100 μm) according to the manufacturer’s instructions for rhodamine (excitation filter 532 nm; emission filter 580 BP 30). Ensure that the no RT control wells are blank and that there is banding in the control RNA lanes. 7. Print the actual size image on an 11-in. × 17-in. page and place the bottom plate on top of the printout with the gel facing up. Cover the gel with Saran Wrap to make band excision easier and prevent drying out. 8. Excise the bands of interest using a clean razor blade and scan the gel again to ensure the appropriate band was obtained. Cut only those bands that display a presence/absence pattern (Fig. 1).

3.5. PCR Product Isolation, Reampliﬁcation, and Cloning 1. Soak the gel slice in 100 μL of water for 10 min at RT in a 1.5-mL tube. 2. Incubate, with cap tightly closed, for 15 min at >95°C and centrifuge for 2 min. 3. Transfer the supernatant to a fresh tube; add 10 μL of 3 M sodium acetate, 2.5 μL of glycogen (20 μg/μL), and 450 μL of 100% EtOH. Place the tubes at –80°C for at least 30 min. 4. Centrifuge the tubes for 10 min to pellet the DNA. Remove the supernatant, and rinse the pellet with 200 μL of ice-cold 85% EtOH. Centrifuge briefly and remove the residual EtOH. 5. Dissolve the pellet in 10 μL of DEPC water and use 4 μL for reamplification. Reamplification should be done using the same PCR reagents used in the FRAPPCR except unlabeled primers (10 μM) are used under the following thermocycle conditions: 30 cycles of 94°C for 30 s, 54°C for 1 min, 72°C for 1 min. 6. Run 15 μL of the reamplified PCR products on a 1% agarose gel stained with ethidium bromide to verify the size of insert. (See Note 11.) 7. PCR products are subcloned into pCR 2.1-TOPO® vector using the TOPO® TA Cloning Kit (Invitrogen) as follows: 2 μL of PCR product, 0.5 μL of pCR 2.1-TOPO® vector and 0.5 μL of salt solution. (See Note 12.) 8. Incubate at RT for 5 min and store at -20°C or proceed directly to transformation (steps 9–12). 9. Combine 2 μL of the ligation reaction with 25 μL (1/2 vial) of One Shot® Chemically Competent E. coli and mix gently (do not mix by pipetting). 10. Incubate on ice for 5–30 min. Heat shock at 42°C for 30 s and transfer to ice immediately.

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Fig. 1. FRAP-PCR product profiles comparing a control and treated sample. Total RNA from control and treated chicken embryonic neuronal cells was reverse transcribed with arbitrary primer A3 and amplified in the presence of 5 rhodaminelabeled primer A3. The arrow indicates a band that exemplifies the presence/absence pattern. In addition, several bands are constitutively expressed regardless of treatment.

11. Add 125 μL of SOC medium (2% tryptone, 0.5% yeast extract, 10 mM NaCl, 2.5 mM KCl, 10 mM MgCl2 , 10 mM MgSO4 , 20 mM glucose) and shake sample horizontally (200 rpm) at 37°C for 1 h. 12. Spread 50–75 μL from each transformation on a LB agar plate containing 50 μg/mL of ampicillin and 40 μg/mL of X-gal. Incubate at 37°C overnight. 13. Touch the edge of three white colonies with a pipet tip, resuspend the partial colonies individually in 50 μL of water, and use 1 μL in a PCR with M13 primers to directly analyze positive transformants.

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14. One positive transformant from each band of interest is cultured in LB/ampicillin media overnight at 37°C. 15. Isolate the plasmid DNA with QIAprep Spin Miniprep Kit from Qiagen and send the product for sequencing. 16. Confirmation of differential gene expression. (See Note 13.)

4. Notes 1. Primary cell culture experiments must be conducted under sterile conditions and immediately following the removal of culture medium from the cells, the plates must be placed directly on dry ice or transferred to a –80°C freezer until subsequent RNA isolation. Tissue harvesting in the laboratory should be conducted using sterile, RNase-free instruments that are cleaned with 3% hydrogen peroxide followed by nuclease-free water between each sample. Tissue samples must be frozen (on dry ice or in liquid nitrogen) as quickly as possible to avoid any RNA degradation. Alternatively, dissected tissue (<0.5 cm in one dimension) can be stored in RNAlater (0.1–0.2 g/mL; Ambion) at 4°C for a month or –20°C indefinitely. This is the ideal storage scenario for field collected samples as it eliminates the need for dry ice or a liquid nitrogen dewar at field sites. 2. Removal of trace amounts of chromosomal or other contaminating genomic DNA is crucial to the success of the FRAP-PCR technique. Genomic DNA can compete with the cDNA population during amplification and lead to false-positive “hits” in the RNA fingerprint. Following up on these potential targets represents a waste of reagents and time. We treat RNA samples with the DNA-Free kit (Ambion) which removes single and double-stranded genomic DNA while maintaining the integrity of the RNA template. In follow-up real-time reverse transcription PCR validations, we did not observe amplification in the no-reverse transcriptase controls. Another benefit to the DNA-Free kit is that it includes a novel DNase Removal Reagent that effectively removes DNase and divalent cations from the reaction mixture so that there is no chance they will interfere with downstream steps. The DNase/cation removal step is quick and does not require organic extraction, EDTA addition or heat inactivation (all of which might affect RNA integrity). 3. Preparing RNA aliquots in appropriate volumes and concentrations for subsequent cDNA synthesis avoids the necessity of repeated freeze–thaw cycles that can negatively affect the quality and quantity of the RNA. A decrease in signal intensity on gels has been observed following repeated freeze–thaw cycles of total RNA samples. 4. The optimal total RNA input for FRAP-PCR was determined by simultaneously comparing RNA profiles made from 200, 500, 750, and 1000 ng of total RNA. The most reproducible and clear banding patterns were observed above 750 ng. However, we have encountered situations where there is a large noise:signal ratio and decreasing the RNA input to 375 ng depletes the noise and improves band clarity.

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5. We have used both tungsten carbide and stainless steel beads for homogenization depending on the lysis buffer and tissue type. We found that TRIzol rusted the stainless steel beads, whereas the RNeasy Lysis buffer (Qiagen) did not. Gonads, heart, lung, and liver tissue was more effectively dissociated using the tungsten carbide beads whereas brain and thyroid were effectively dissociated using the stainless steel beads. 6. cDNA aliquots are prepared for the same reason as RNA aliquots; to avoid freeze/thaw degradation. A 20-μL cDNA reaction was used in four different FRAPPCR reactions and by the third run (i.e., 3 freeze–thaw cycles), gel profiles were negatively affected, especially the faint products. 7. Following first-strand synthesis, a low-stringency PCR step is performed (stage I) to incorporate the arbitrary primer into both ends of the PCR products. This secondstrand synthesis step ensures optimal amplification of cDNA products during the subsequent high-stringency PCR. 8. It is important to determine the optimal annealing temperature of the specific 18-mer primer used for FRAP-PCR. For A3 and B3, the ideal temperature is 54°C whereas for C3, it is 56°C. Before conducting experiments with 18-mer primers other than those described in this chapter, researchers should characterize their primers using primer analysis software (e.g., OligoAnalyzer; http://molbioltools.ca/molecular_biology_freeware.htm) to determine melting temperature, %GC content, and optimal PCR temperature. Empirical determination of the selected PCR conditions should follow. 9. During the initial development of the FRAP-PCR technique, rhodamine-labeled PCR products were not purified before electrophoresis. The result was large, nonspecific streaks across all wells (Fig. 2a). We hypothesized that the nonspecific binding was due to unincorporated nucleotides and excess, unbound rhodamine. The QIAquick PCR Purification Kit (Qiagen) recovers fragments between 100 bp and 10 kbp and removes those ≤40 bp. The quick and easy column-based kit was effective at removing the smears and improving resolution (Fig. 2b) and should be used in all FRAP-PCR experiments. Potential bands of interest could be missed as a result of the large nonspecific products in an unpurified sample. 10. Samples should not be loaded in the five outermost wells at either edge of the vertical electrophoresis system if possible. “Smiling” or “arcing” is sometimes encountered and this makes it difficult to compare the profiles to the center lanes, which tend to have bands that are horizontal, sharp, and well separated. It is also useful to leave one or two wells between treatment groups to reduce any diffusion of sample between treatments. 11. A rhodamine-labeled DNA ladder was not simultaneously run on the sequencing gel and thus, product size determination was performed during the reamplification PCR step. Products cut from the middle section of the sequencing gel ranged in size from 300–750 bp after 6 h of electrophoresis at 1700 V. In most cases, bands excised from the sequencing gel produce single products in the reamplification PCR that can be immediately subcloned into the TOPO vector. However, some bands that appear to represent single products reveal multiple bands in the agarose

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Fig. 2. Effect of purifying FRAP-PCR products with the QIAquick PCR Purification Kit before gel electrophoresis. Total RNA from chicken embryonic neuronal cells was reverse transcribed with arbitrary primer A3 and amplified in the presence of 5 rhodamine-labeled primer A3. One subset of 8 samples was loaded onto the gel immediately (a) while the other was purified using the QIAquick Kit (b). The results indicate the importance of including a PCR purification step prior to proceeding with the separation of the RNA fingerprint as many bands of interest may be missed.

gel screening step. This comigration or nonspecific amplification can be dealt with in two main ways: (1) excise the band from the agarose gel that corresponds to the right size on the sequencing gel, purify the product with the QIAquick Gel purification kit (Qiagen), and subclone the purified product or (2) increase the

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electrophoresis time on the sequencing gel to further resolve single bands that may in fact represent more than one PCR product. 12. Cloning and transformation can represent a significant proportion of the cost of FRAP-PCR depending on the number of potential “hits” that are explored. We have determined that cloning reactions can be conducted efficiently using half the recommended reagent volumes. That is, in the ligation reaction, only 0.5 μL of vector is used and the resulting ligation reaction is transformed into half a vial, or 25 μL, of competent cells. This helps save on costs and reagents and does not jeopardize the efficiency of TOPO TA cloning. 13. There are several postcloning methods for confirming differential gene expression. We use real-time reverse transcription PCR (MX4000 instrument and SYBR Green Master Mix; Stratagene) as this is one of the most sensitive and reliable methods for measuring relative mRNA expression. Real-time PCR primers are designed based on the sequences obtained using FRAP-PCR. 18S rRNA, -actin, or glyceraldehyde-3-phosphate dehydrogenase (GAPDH) serve as internal control genes to compare the relative abundance of the amplicon of interest to.

Acknowledgments This work was supported in part by a Pesticide Science Fund grant (Dr. Pierre Mineau, Environment Canada) and by funding from Environment Canada’s Strategic Technology Applications of Genomics for the Environment (STAGE). We acknowledge Stephanie Jones and Magdalena Jagla for critically reading the chapter. References 1. Liang, P. and Pardee, A.B. (1992) Differential display of eukaryotic messenger RNA by means of the polymerase chain reaction. Science 257, 967–971. 2. McClelland, M., Honeycutt, F., Vogt, T., and Welsh, J. (1997) Fingerprinting by Arbitrarily Primed PCR, in Methods in Molecular Biology, Vol. 85: Differential Display Methods and Protocols (Liang, P. and Pardee, A.B., eds.), Humana Press, Totowa, NJ, pp. 13–24. 3. Welsh, J., Chada, K., Dalal, S. S., Cheng, R., Ralph, D., and McClelland, M. (1992) Arbitrarily primed PCR fingerprinting of RNA. Nucleic Acids Res. 20, 4965–4970. 4. Blazquez, M., Bosma, P. T., Chang, J. P., Docherty, K., and Trudeau, V. L. (1998) Gamma-aminobutyric acid up-regulates the expression of a novel secretograninII messenger ribonucleic acid in the goldfish pituitary. Endocrinology 139, 4870–4880. 5. Crump, D., Lean, D., and Trudeau, V. L. (2002) Octylphenol and UV-B radiation alter larval development and hypothalamic gene expression in the leopard frog (Rana pipiens). Environ. Health Perspect. 110, 277–284.

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6. Trudeau, V. L., Chiu, S., Kennedy, S. W., and Brooks, R. J. (2002) Octylphenol (OP) alters the expression of members of the amyloid protein family in the hypothalamus of the snapping turtle, Chelydra serpentina serpentina. Environ. Health Perspect. 110, 269–275. 7. Picard, D. J. and Schulte, P. M. (2004) Variation in gene expression in response to stress in two populations of Fundulus heteroclitus. Comp. Biochem. Physiol. Part A 137, 205–216. 8. Diener, L. C., Schulte, P. M., Dixon, D. G., and Greenberg, B. M. (2004) Optimization of differential display polymerase chain reaction as a bioindicator for the cladoceran Daphnia magna. Environ. Toxicol. 19, 179–190. 9. Ranamukhaarachchi, D. G., Rajeevan, M. S., Vernon, S. D., and Unger, E. R. (2002) Modifying differential display polymerase chain reaction to detect relative changes in gene expression profiles. Anal. Biochem. 306, 343–346. 10. Cho, Y. J., Meade, J. D., Walden, J. C., Chen, X., Guo, Z., and Liang, P. (2001) Multicolor fluorescent differential display. Biotechniques 30, 562–571. 11. Ito, T. and Sakaki, Y. (1997) Fluorescent differential display, in Methods in Molecular Biology, Vol. 85: Differential Display Methods and Protocols (Liang, P. and Pardee, A.B., eds.), Humana Press, Totowa, NJ, pp. 37–44. 12. RNAspectra™ Kit for Fluorescent mRNA differential display: Protocol. GenHunter Corporation, Nashville, TN, USA. (www.genhunter.com) 13. Alberts, B., Bray, D., Lewis, J., Raff, M., Roberts, K., and Watson, J. D. (eds.) (1994) Molecular Biology of the Cell, 3rd edition. Garland, New York.

3 Isolation of O3 -Response Genes from Arabidopsis thaliana Using cDNA Macroarray Masanori Tamaoki

Summary Nylon membrane-based cDNA macroarrays are a widely available alternative to cDNA microarrays for the collection of large-scale gene expression data. cDNA macroarrays are used in many areas of molecular biology research for applications ranging from gene discovery to gene expression profiling. Although degree of location of DNA spot in cDNA macroarray is lower than that in cDNA microarray, it can be used to detect expression of a large number of genes because it uses radiolabeled cDNA as a probe. Thus, cDNA macroarray technology can be applied to obtain the gene expression profile in organs that show wide variety in mRNA expression, such as meristems in plant species and brain tissue. To carry out hybridization experiments with a cDNA macroarray, I describe here how to prepare macroarray filters on a small or large scale, as well as how to analyze macroarray experiments and determine the statistical significance of the gene expression data obtained. Key Words: Arabidopsis thaliana; cDNA macroarray; cDNA microarray; gene expression; ozone; statistical analysis; subset microarray.

1. Introduction Recently, measurements of gene expression levels have been carried out on a large scale by using both cDNA microarray (1) and cDNA macroarray (2) technology. cDNA expression arrays generally consist of a nylon membrane (cDNA macroarray) or glass slides (cDNA microarray) spotted with polymerase chain reaction (PCR) products produced from cDNA clones. The arrays are hybridized with radioactively or fluorescently labeled probes transcribed from mRNA. Both types of cDNA array have become very From: Methods in Molecular Biology, vol. 410: Environmental Genomics Edited by: C. Cristofre Martin © Humana Press, Totowa, NJ

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useful in the elucidation of mRNA expression patterns among different tissues, the determination of differential gene expression during growth or as the result of environmental conditions, and the identification of novel genes (see ref. 3 for review). Apart from the high number of genes screened, an advantage of cDNA microarrays is the sensitivity of detection because of the high probe concentration, which is a result of the small incubation volumes used. However, in a comparative study, cDNA macroarrays used in conjunction with 33 Plabeled probes showed higher sensitivity than cDNA microarrays because of the larger amount of DNA spotted on the membrane (4). Given the generally held opinion that 300,000 mRNAs are expressed per cell, an mRNA whose expression level is only 0.005% (i.e., 15 molecules per cell) theoretically can be detected using cDNA macroarrays (Table 1). Currently, the cDNA macroarray method is more accessible to researchers because, unlike microarray technology, it can be carried out using standard laboratory equipment. However, a disadvantage of cDNA macroarrays is that only one sample can be analyzed per membrane, resulting in variations in signal intensity because of the potential presence of different amounts of cDNA probe when comparing two or more samples (5). In contrast, cDNA microarrays on glass slides allow for direct comparison of two samples in the same array because each sample is labeled with a different fluorescent probe, such as Cy3 or Cy5. This practice allows accurate measurement of each signal. Therefore, a 2-fold up- or down-regulated gene can be identified using a single cDNA microarray (6). Contrastingly, comparative experiments using cDNA macroarray are easily affected by experimental variations. Consequently, statistical analysis of data from replicate experiments is needed to reduce the effects of such variations. In this chapter, I describe a protocol for cDNA macroarray analysis. This description includes (1) making a cDNA macroarray, (2) hybridization and Table 1 Comparison of cDNA macroarray and cDNA microarray cDNA microarray Support Degree of location Detection system Possibility of reuse Sensitivity Dynamic range Deviation

Slide glass < 500 genes/cm2 Radioisotope (33 P) Yes (see Note 9) 0.005% mRNA 104 -fold 20%

cDNA macroarray Nylon membrane 500–10,000 genes/cm2 Fluorescent dye (Cy3, Cy5) no 0.05% mRNA 102 -fold 100%

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data collection, and (3) the statistical analysis involved when evaluating a set of three independent experiments. By way of example, I present my macroarray analysis that compared the gene expression patterns of ozone (O3 )- and ambient air-treated Arabidopsis thaliana. 2. Materials 2.1. Preparation of Samples for cDNA Macroarray 1. Culture medium: 1:2 (v/v) Gamborg B5 Complete medium (GibcoBRL, Rockville, MD), 1:1000 (v/v) Hyponex (Hyponex 10:5:10; Hyponex Japan, Osaka, Japan), 1% sucrose; pH adjusted to 5.7.

2.2. Preparation of Probe for cDNA Macroarray (see Notes 1–4) 1. 4 M guanidium thiocyanate buffer: 4 M guanidium thiocyanate, 0.1 M Tris-HCl, pH 7.5, 10 mM EDTA, 0.5% Sarkosyl, 0.1% -mercaptoethanol. 2. Phenol–chloroform: 50% (v/v) liquid phenol, pH 8.0, 48% (v/v) chloroform, 2% (v/v) isoamyl alcohol. 3. 5.7 M CsCl cushion: 5.7 M CsCl, 0.1 M EDTA; pH adjusted to 7.5. 4. DEPC-treated water: 0.1% diethyl pyrocarbonate. 5. 10× Hybridization buffer: 1.2 M NaCl, 0.1 M Tris-HCl, pH 8.0, 50 mM EDTA, 10% sodium dodecyl sulfate (SDS). 6. Array BPB solution: 10 mM Tris-HCl, (pH 7.5, 0.25% bromophenol blue, 1 mM EDTA, pH 8.0, 60% (v/v) glycerol. 7. Alkaline solution: 0.5 N NaOH, 1.5 M NaCl. 8. Neutralizing solution: 1.5 M NaCl, 0.5 M Tris-HCl; pH adjusted to 7.4. 9. 2×SSC: 300 mM NaCl, 30 mM sodium citrate.

2.3. Hybridization of cDNA Macroarray Filters 1. 5× M-MLV RT buffer: 250 mM Tris-HCl, pH 8.3, 375 mM KCl, 15 mMMgCl2 . 2. dNTP mixture without dCTP: 20 mM dATP, 20 mM dTTP, 20 mM dGTP. 3. Hybridization solution (Church phosphate buffer): 0.5 MNa2 HPO4 , 1 mM EDTA, 1 μg/mL. poly dA [Roche Diagnostics, Indianapolis, IN], 7% SDS. Adjust pH to 7.2. 4. 0.2× SSC: 30 mM NaCl, 3 mM sodium citrate.

3. Methods 3.1. Preparation of Samples for cDNA Macroarray 1. Arabidopsis thaliana (L.) Heyhn accession Columbia (Col-0; ABRC, Columbus, OH) is used for analysis. 2. Plants for constructing cDNA library are grown in soil under a 16:8-h light:dark cycle at 22°C.

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3. Above ground organs, flower buds, and green siliques are harvested from 2- to 6-week-old plants. For liquid culture, sterile seeds are sown in culture medium and grown for 2 weeks under continuous light at 22°C with rotation. Seedlings and roots are collected from liquid-cultured plants. 4. For isolation of mRNA, plants are germinated on blocks of rock wool and grown in a growth chamber at 25°C at a relative humidity of 50–60% under a photosynthetic photon flux density (PPFD) of 100 μmol m−2 s−1 in 14:10-h light:dark cycles. 5. Plants are watered with a 5:10:5 liquid fertilizer (Hyponex, Hyponex Japan) diluted 2000-fold. Two-week-old plants are exposed to a single dose of 200 nL/L O3 for 12 h in an O3 chamber, as described previously (7). The O3 chamber is maintained at 25°C and a relative humidity of 70% under a PPFD of 100 μmol m−2 s−1 in continuous light. Plants remaining in ambient air serve as the controls. O3 is generated with an O3 generator (Sumitomo Seika Chemicals, Tokyo, Japan).

3.2. Probe Preparation for cDNA Macroarray (8) 3.2.1. Isolation of Poly(A)+ RNA 1. Total RNA is extracted from above ground organs, flower buds, roots, 2- to 6-wk-old plants, and liquid-cultured seedlings by the guanidium thiocyanate–CsCl ultracentrifugation method (9). 2. Four grams of frozen tissue are ground to powder in liquid nitrogen and treated with 25 mL of 4 M guanidium thiocyanate buffer. 3. Centrifuge the homogenate at 5,000g for 10 min at room temperature. 4. Transfer the supernatant to a fresh tube and add an equal volume of phenol– chloroform. Mix well. 5. Centrifuge the mixture at 5,000g for 5 min at room temperature. 6. Transfer the supernatant to a fresh tube, and repeat steps 4 and 5 four times. 7. Layer the supernatant onto a 15-mL 5.7 M CsCl cushion in a clear ultracentrifuge tube and centrifuge for 22 h at 20°C and 271,900g using a RP50VF rotor (Hitachi Koki, Tokyo, Japan). 8. Invert the tube to discard the supernatant, and allow any remaining fluid to drain into the paper towel. 9. Wash the pellet of RNA with 70% ethanol at room temperature. Invert the tube and drain off the ethanol. 10. Allow the pellet of RNA to dry, and dissolve it in DEPC-treated water. 11. Poly(A)+ RNA is purified from total RNA by using an mRNA purification kit according to the manufacturer’s instructions (Amersham Biosciences, Piscataway, NJ) (See Note 5).

3.2.2. Construction of cDNA Library 1. 1.5 to 7.5 μg poly(A)+ RNA is primed with oligo(dT)18 primer carrying an XhoI site, and the first-strand synthesis reaction is performed for 30 min at 52°C using SuperScript II (Invitrogen, Groningen, Netherlands) reverse transcriptase according to the manufacturer’s instructions. (See Note 6.)

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2. After second-strand synthesis, cDNA is blunt-ended with using DNA Blunting Kit (TaKaRa Bio, Ohtsu, Japan), and EcoRI adaptors are ligated to both ends. 3. The XhoI sites at the 3 ends of the cDNA are generated by digestion with XhoI. 4. Size selection of cDNAs before vector ligation is performed as follows. Synthesized cDNAs are separated by 1% agarose gel electrophoresis, and fractions ranging from 1 to 3 kb are recovered from the gel using a QIAquick Gel Extraction Kit (QIAGEN, Düsseldorf, Germany) as described by the manufacturer’s instructions. 5. The recovered fragments are cloned into the EcoRI–XhoI sites of pBluescript II SK− plasmid vector (Stratagene, La Jolla, CA) and transformed into Escherichia coli (E. coli) XL1-Blue MRF’ strain (Stratagene) by electroporation.

3.2.3. Normalization of cDNA Library 1. For normalization of the library, a single-stranded library is prepared from 5 μg of plasmid DNA. First, a single-stranded nick is introduced into the plasmid DNA using 20 U of gene II endonuclease of phage F1 (M13 gpII protein; Life Technologies, Gaithersburg, MD) for 30 min at 37°C (see Note 7). This enzyme makes a nick at the f1 replication origin in the plasmid DNA. Then, a singlestranded plasmid is generated with digestion of 250 U of exonuclease III (New England Biolabs) that has 3 to 5 exonuclease activity. The reaction is performed using attached buffer (TaKaRa Bio, 50 mM Tris-HCl, pH 8.0, 5 mM MgCl2 , 10 mM 2-mercaptoethanol), and the volume adjusted to 50 μL with water. Then incubate the reaction mixture for 1 h at 37°C. 2. The cDNA inserts are allowed to self-hybridize by using an excess (about 1 μg) of cDNA inserts generated by PCR amplification. For amplification of the inserts, about 5 ng of DNA template (the single-stranded library plasmid produced in Subheading 3.2.3.1.) was mixed with 1 μL of 100 μM T7 primer. 5 -TAATACGACTCACTATAGGG-3 and 1 μL of 100 μM SK primer 5 -GCTCTAGAACTAGTGGATC-3 , and then PCR was performed using Ex-Taq DNA polymerase (TaKaRa Bio) in a Perkin-Elmer 9600 Thermal Cycler (Applied Biosystems, Foster City, CA) according to the following program: 7 min ramp from room temperature to 94°C; 20 cycles of 1 min at 94°C, 2 min at 55°C, and 3 min at 72°C. A final extension of 7 min at 72°C follows. 3. The PCR product is ethanol-precipitated, dissolved in 1.5 μL of water, and then mixed with 5 μL of the single-stranded library cDNA (50 ng) dissolved with formamide, 0.5 μL(10 μg) of the 5 -blocking oligo nucleotide mixture. (5 -GCTCTAGAACTAGTGGATCCCCCGGGCTGCAGGAATTCG-3 and 5 -AATTCGGCACGAG-3 ), and 0.5 μL (10 μg) of the 3 -blocking oligo nucleotide mixture (5 -CTCGAGGGGGGGCCCGGTA-3 and 5 -GTA CCCAATTCGCCCT ATAGTGAGTCGTATTA-3 ). 4. The mixture is kept at 80°C for 3 min, 1 μL of 10× hybridization buffer and 1.5 μL of water are added, and hybridization is performed at 30°C for 24 h. 5. The remaining single-stranded circles are purified using hydroxyapatite chromatography (10) and converted to double strands using the Klenow fragment (TaKaRa

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3.2.4. Template Preparation and Sequencing 1. Plasmid DNA is prepared by the alkaline lysis method in a 96-well plate (11). 2. Clones are digested with ApaI and SmaI to release the inserts, whose sizes are measured by agarose gel electrophoresis of the digestion reactions. 3. Clones undergo nucleic acid sequencing with a BigDye Terminator Cycle Sequencing Ready Reaction Kit (Applied Biosystems) according to the manufacturer’s instructions and are electrophoresed using automated DNA sequencers (ABI PRISM 373 and 377XL; Applied Biosystems). 4. Random cDNA clones are chosen from the normalized cDNA libraries constructed (See Note 8). In the example experiment, 14,026 clones in total were sequenced from the 5 -end, and 39,207 clones were sequenced from the 3 -end. As a result, 39,207 3 -expressed sequence tag (EST) sequences were clustered into 12,028 independent groups (8) . 5. 12,028 independent EST clones were used for construction of a massive cDNA macroarray filter. Inserts of EST clones were amplified using 5 -GTAATACGACTCACTATAGGGC-3 and 5 -TCATTAGGCACCCCAGGC TTTACAC-3 as the primers. PCR was performed using a Perkin-Elmer 9600 Thermal Cycler (Applied Biosystems) in 80-μL reaction mixtures, each containing 1 μL of cultured E. coli cells as the PCR template, 1× Ex-Taq Buffer (TaKaRa Bio), 2.5 mMMgCl2 , 0.2 mM dNTPs, 0.2 μM each primer, and 0.5 U of Ex-Taq DNA polymerase (TaKaRa Bio). Amplification conditions are 35 cycles of denaturation at 94°C for 45 s, annealing for 45 s at 55°C, and elongation at 72°C for 2 min; the final extension is for 4 min at 72°C. 6. Five microliters of each PCR-amplified product is electrophoresed through a 1.5% agarose gel to check amplification of the EST fragment.

3.3. Preparation of Large-Scale cDNA Macroarray Filters 1. The remaining 75 μL of PCR product is mixed with 15 μL of array BPB solution and spotted onto an 8 × 12 cm nylon filter (Biodyne A, Pall, East Hills, NY) in duplicate by using a Biomek 2000 Laboratory Automation Workstation (Beckman Instruments Inc., Fullerton, CA). This pin blotter spots 2,880 EST clones on a filter. Therefore, for the example experiment, five independent filters were made to accommodate the 12,028 independent EST clones. An example cDNA macroarray filter is shown in Fig. 1A. 2. DNA (10 mg/μL) is spotted in duplicate as negative controls. 3. Spotted filters are denatured in a shallow tray of alkaline solution for 2 min, after which the filters are transferred into neutralizing solution for 2 min. 4. Rinse the filter in 2× SSC and place it, DNA side up, on paper towels to dry. 5. After the filters are dry, the spotted ESTs are fixed on the filters by UV crosslinking (1.2 J/cm2 ).

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Fig. 1. An example of hybridization pattern of cDNA macroarray analysis. (A) A cDNA macroarray filter before using hybridization. The filter is made with a Biomek 2000 Laboratory Automation Workstation (Beckman Instruments Inc.) and contains 2,880 EST clones. (B) An example of hybridization image of a cDNA macroarray filter. Large-scale cDNA macroarray filter containing Arabidopsis EST clones is hybridized with radiolabeled cDNA probes, which is made from total RNA of O3 -exposed Arabidopsis. The hybridization image is incorporated with high-resolution scanner (Storm; Amersham Biosciences).

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3.4. Hybridization of cDNA Macroarray Filters 3.4.1. Preparing Radiolabeled Target (RNA Labeling) 1. Total RNA or mRNA that is isolated from sample of interest is prepared as described earlier (see Subheading 3.2.1.). For the example experiment, total RNA was isolated from O3 -exposed and -unexposed Arabidopsis plants. 2. Ten micrograms of total RNA (or 0.2 μg of poly(A)+ RNA, if available) is mixed with 2 μg of oligo-dT12−18 (Invitrogen) and the volume adjusted to 14 μL with DEPC-treated water. 3. Incubate the RNA solution for 10 min at 70°C. Chill the sample on ice for 2 min. 4. Add 6 μL of 5× M-MLV RT buffer, 1.5 μL of dNTP mixture without dCTP, 1 μL of 0.1 M dithiothreitol, 6 μL of [-33 P] dCTP (spec act > 2,500 Ci/mmol; Amersham Biosciences), and 7.5 U SuperScript II (Invitrogen). 5. Mix well and incubate the reaction mixture at 37°C for 90 min. Chill the sample on ice for 5 min. 6. Apply the sample to a G-50 spin column (Probe Quant G-50 Micro Column; Amersham Bioscience). Centrifuge at 820g for 2 min. 7. Collect the effluent (about 30 μL) and denature it at 95°C for 5 min. 8. Keep the denatured probe on ice until it is used.

3.4.2. Hybridization 1. Place the cDNA macroarray filters into a hybridization bag (Atto, Tokyo, Japan). 2. Pour in 10 mL of hybridization solution into the hybridization bag and then seal it with a heat-sealer. 3. Submerge the filters in a water bath at 65°C and incubate for 1–2 h. 4. Remove the bag containing the filters from the water bath. Open the bag by cutting off one corner with scissors. 5. Add the denatured probe (see Subheading 3.3.1.) to the hybridization solution and then squeeze as much air as possible from the bag. 6. Reseal the bag with the heat-sealer. 7. Submerge the bag in a 65°C water bath for more than 16 h. 8. After hybridization, open the bag by cutting off a corner with scissors. 9. Transfer the filters to a flat-bottom plastic box containing 250 mL of 0.2× SSC, 0.1% SDS. 10. Wash the filters for 15 min at 65°C (see Note 10) with gentle agitation. 11. Replace the solution with fresh 250 mL of 0.2× SSC, 0.1% SDS, and transfer the box to a water bath set at 65°C with gentle agitation for 15 min. 12. Remove most of the liquid from the filters by placing them on paper towels. 13. Cover the filters with plastic wrap and expose them to a bioimaging plate (Fuji Film, Tokyo, Japan) for more than 12 h.

3.5. Data Collection and Statistical Analysis 1. Scan autoradiographs on a high-resolution scanner (Storm; Amersham Biosciences) with 50-μm resolution, and quantify the signal intensity with ArrayVision software

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2.

3.

4.

5.

6.

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(Amersham Biosciences) as described in the manufacturer’s instructions. An example scanned image is shown in Fig. 1B. Export the quantified signal intensity as a ‘.csv’ format file and import the file into a data analysis program (e.g., Microsoft Excel Version X, Microsoft, Redmond, WA) for analysis. Normalize interfilter differences in signal intensity by adopting global normalization (see Note 11) in the following way. Average the intensities of all signals in the filter, and then calculate the relative signal intensity as the ratio of the signal of interest to the average intensity of each filter. Thus the estimated values are designated as expression ratios. Before further analysis, average the expression ratios of the duplicate spots. To select stimuli-response genes (O3 -responsive genes in the example experiment), first eliminate from analysis all genes whose expression ratios are less than 10 times the background signal (the average expression ratio of DNA); this level corresponds to 0.02% expression in total RNA. In the example experiment, this process removed about two-thirds of the 12,028 ESTs from analysis. Select for further analysis genes whose expression was increased or decreased at least 3-fold after treatment stimuli (O3 exposure in the example experiment; see Fig. 2). Genes that respond to stimuli with reproducible results are identified by oneway analysis of variance at a significance level of 0.05. This procedure involves the use of the F statistic, which is used for estimation of population variance on the basis of the information in two or more random samples, to test the

Fig. 2. O3 -induced gene expression profiles in a cDNA macroarray. Examples of gene expression profiles induced by ambient air (left) and ozone (right) are shown. Part of the cDNA macroarray grid is magnified to show an example of an O3 -inducible gene (enclosed with circle).

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Fig. 3. Scatterplot of signal intensities of all expressed sequence tags (ESTs) in the Arabidopsis macroarray (left) and the subset macroarray (right). Normalized expression ratios for each clone in the macroarray are plotted against signals from O3 -unexposed (air; x-axis) and -exposed plants (y-axis). The diagonal lines are cutoff lines at 3-fold induction or repression of gene expression. The dots in the shaded area represent ESTs whose signal intensities are not more than 10 times that of the background signal intensity. The 157 clones up-regulated by O3 were used to make the subset macroarray. statistical significance of the differences among the obtained mRNA expression levels under each experimental condition. In the example experiment, I identified 205 nonredundant ESTs that were regulated by O3 . The expression of 157 of these was induced, and that of 48 suppressed, by O3 (Fig. 3) (12). 7. (Optional) If hierarchical clustering analysis (13) and K-mean clustering are necessary, the GeneSpring software package (version 5.0, Silicon Genetics, Redwood, CA) is recommended. 8. In the case of Arabidopsis thaliana, search for names and annotations of genes by using the Database for Arabidopsis Research and Tools (DART) program (http://tabacum.agr.nagoya-u.ac.jp/dart/). Functional classification of genes is performed by first classifying ESTs by referring to MATDB (http://mips. gsf.de/proj/thal/db/index.html). Second, unclassified ESTs are re-sorted according to the putative functions assigned through annotation.

3.6. Preparation of Subset cDNA Macroarray Filters If genes that respond to various stimuli have been isolated, preparation of a subset macroarray using the isolated genes is recommended for further analysis. Because, in general, only about 10% of genes respond to the stimuli

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of interest, subset macroarrays are more convenient than the large-scale macroarray for further experiments. The subset macroarray typically contains 100–1,000 genes, but I have made a subset macroarray that comprised only 12 genes (14). The potential applications of subset macroarrays are diverse. For example, subset macroarrays are suitable to confirm whether the isolated genes correctly respond to target stimuli, because investigation of gene expression by using Northern blotting or real-time PCR analysis is time-consuming and costly. Moreover, subset macroarrays are useful to compare the differences in stimulus-associated gene expression patterns among mutant strains or different tissues (11). 1. Obtain cDNA clones corresponding to genes of interest from a public repository or clone them in-house. In the example experiment, the 157 O3 -upregulated ESTs from Arabidopsis were obtained from the Kazusa DNA Research Institute (Kisarazu, Japan). 2. PCR-amplify the inserts of these EST clones by using the primers 5 -GTTTCCCAGTCACGAC-3 and 5 -CAGGAAACAGCTATGAC-3 (these primers are annealed to pBR derived plasmid vectors) and the PCR conditions described in 3.2.3. 3. Mix 75 μL of PCR products with 15 μL of array BPB solution and spot onto a 9 × 12-cm nylon filter (Biodyne A, Pall) in duplicate by using a Multi Pin Blotter 96 (Atto). An example of subset macroarray filter is shown in Fig. 3. 4. Spot DNA (10 mg/μL) in duplicate as a negative control. 5. Fix spotted cDNAs as described earlier (see Subheading 3.2.). 6. Hybridize the filter and quantify signal intensity as described earlier (see Subheading 3.3.). 7. Obtain the signal intensity of each spot by subtracting the mean signal intensity of the negative control ( DNA) from the mean signal intensity of the duplicate spots. 8. Normalize the signal intensities of each gene according to that of a gene whose expression level remains unchanged with stimuli treatment. In the example experiment, AtTub4 was used as a normalization marker gene because its expression level was confirmed previously to be unchanged between stress and nonstress conditions (15).

4. Notes 1. All reagents should be prepared using double-distilled water that has a resistance higher than 17.6 M-cm. 2. For preparation of RNAs, glassware must be made RNase-free by baking at 180°C for 8 h or more. All solutions should be made with 0.1% DEPC-treated water, with the exception of the buffer (see Note 3). The primary source of contamination with RNase is the hands of the researcher. Disposable gloves should therefore be worn during manipulations involving RNA.

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3. DEPC reacts rapidly with amines and cannot be used to make solutions containing buffers such as Tris. Reserve a fresh, unopened bottle of Tris crystals for preparation of RNase-free solution. 4. DEPC is suspected to be a carcinogen and should be handled with care. 5. The concentration of the RNA can be determined by measuring the OD260 of an aliquot of the final preparation. An RNA solution whose OD260 is 1 contains approx 45 μg/mL. 6. 5 -methyl dCTP instead of dCTP is used for synthesis of the first-strand cDNA to make the internal XhoI sites resistant to digestion. 7. The gene II endonuclease of phage F1 (M13 gpII protein; Life Technologies) is no longer commercially available. Instead of this enzyme, site-specific nickase, N.BstT9 (BIORON GmbH, Ludwigshafen, Germany), is available for this reaction. For making a nick in plasmid DNA, mix 5 μg of plasmid DNA, 20 U of N.BstT9 and N. Bst9-buffer [BIORON GmbH, 10 mM Tris-HCl, pH 8.5, 10 mM MgCl2 , 150 mM KCl, 1 mM dithiothreitol (DTT), 0.1 mg/mL of bovine serum albumin (BSA)], and the volume adjusted to 50 μL with water. Then incubate the reaction mixture for 30 min at 55°C. 8. The number of clones picked out is dependent on genome size and/or purpose for experiment. If you want to prepare a perfect set of cDNAs that express in Arabidopsis thaliana (whole genome size is about 1.25 × 108 bp), it should be necessary to pick up more than 3 × 106 colonies with using normalized cDNA library (average cDNA size is about 2 kb). In general, picking up clones more than 5-fold of genome size is needed to obtain a perfect set of cDNAs. 9. For comparing multiple sets of gene expression data, filters. used in macroarray analysis are often reused. An effective membrane-stripping protocol is boiling the membrane for 30 min in 1% SDS. This procedure limits reuse to five times because of physical damage to the membrane and reduction in the amount of cDNA spotted on the membrane. A gentler stripping protocol is described by Horngerg et al. (16). Because of the lower melting temperature of the shorter cDNA strands, after oxidative breakdown, stripping by their method can be carried out at relatively low temperatures, thereby reducing heat-induced damage. According to this paper, the same filter is able to use more than 10 times without a measurable reduction in their performance. 10. The hybridization efficiency of macroarray filters can be affected by the strictness of DNA probe, quality of RNA, and number of times the filter has been reused. With high-quality RNA and new filters, the hybridization conditions described in Subheading 3.3. work well. However, in the event of poor signal intensity or for alternative applications, I recommend that filter washing start at a temperature lower than 65°C. For example, when cDNA macroarray filters constructed from an Arabidopsis thaliana cDNA library are applied to the detection of gene expression profiles in other species, such as Indian mustard (Brassica napus) or wheat (unpublished results), I recommend starting the washing temperature at 42°C. If the signal intensities of all spots are strong, rewash the filters at a higher temperature.

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11. In Subheading 3.4., I suggest using the global normalization method for normalization of array signals. Recently, a more sophisticated method (lognormal distribution fitting method) for evaluation of data quality and normalization has been published (17).

References 1. Ramsay, G. (1998) DNA chips: state-of-the-art. Nat. Biotechnol. 16, 40–44. 2. Chalifour, L. E., Fahmy, R., Holder, E. L., Hutchinson, E. W., Osterland, C. K., Schipper, H. M., and Wang, E. (1994) A method for analysis of gene expression patterns. Anal. Biochem. 216, 299–304. 3. Stoughton, R. B. (2005) Applications of DNA microarrays in biology. Annu. Rev. Biochem. 74, 53–82. 4. Bertucci, F., Bernard, K., Loriod, B., Chang, Y. C., Granjeaud, S., Birnbaum, D., Nguyen, C., Peck, K., and Jordan, B. R. (1999) Sensitivity issues in DNA arraybased expression measurements and performance of nylon microarrays for small samples. Hum. Mol. Genet. 8, 1715–1722. 5. Schuchhardt, J., Beule, D., Malik, A., Wolski, E., Eickhoff, H., Lehrach, H., and Herzel. H. (2000) Normalization strategies for cDNA microarrays. Nucleic Acids Res. 28, E47. 6. Hughes, T. R., Marton, M. J., Jones, A. R., Roberts, C., Stoughton, R., Armour, C. D., Bennett, H. A., Coffey, E., Dai, H., He, Y. D., Kidd, M. J., Meyer, M. R., Slade, D., Lum, P. Y., Stepaniants, S. B., Shoemaker, D. D., Gachotte, D., Chakraburtty, K., Simon, J., Bard, M., and Friend, S. H. (2000) Functional discovery via a compendium of expression profiles. Cell 102,109–126. 7. Tamaoki, M., Matsuyama, T., Kanna, M., Nakajima, N., Kubo, A., Aono, M., and Saji, H. (2003) Differential O3 sensitivity among Arabidopsis accessions and its relevance to ethylene synthesis. Planta 216, 552–560. 8. Asamizu, E., Nakamura, Y., Sato, S., and Tabata, S. (2000) A large-scale analysis of cDNA in Arabidopsis thaliana: generation of 12028 non-redundant expressed sequence tags from normalized and size-selected cDNA libraries. DNA Res. 30, 175–180. 9. Sambrook, J., Fritsch, E. F., and Maniatis T. (1989) Molecular Cloning: A Laboratory Manual, 2nd ed. Cold Spring Harbor Laboratory Press. Cold Spring Harbor, pp F.4–F.5. 10. Ko, M. S. H. (1990) An equalized cDNA library by the reassociation of short double-strand cDNAs. Nucleic Acids Res. 18, 5705–5711. 11. Stowers, L., Herrnstadt, C., Grothe, A., Pease, E., Osterlund, M., Cable, P., Brolaski, M., and Gautsch. J. (1992) Rapid isolation of plasmid DNA. Am. Biotechnol. Lab. 10, 48. 12. Tamaoki, M., Nakajima,N., Kubo, A., Aono, M., Matsuyama, T., and Saji, H. (2003) Transcriptome analysis of O3 -exposed Arabidopsis reveals that multiple signal pathways act mutually antagonistically to induce gene expression. Plant Mol. Biol. 53, 443–456.

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13. Eisen, M. B., Spellman, P. T., Brown, P. O., and Botstein, D. (1998) Cluster analysis and display of genome-wide expression patterns. Proc. Natl. Acad. Sci. USA 95, 14863–14868. 14. Tamaoki, M., Matsuyama, T., Nakajima, N., Aono, M., Kubo, A., and Saji, H. (2004) A method for diagnosis of plant environmental stresses by gene expression profiling using a cDNA macroarray. Environ. Pollut. 131, 137–145. 15. Matsuyama, T., Tamaoki, M., Nakajima, N., Aono, M., Kubo, A., Moriya, S., Ichihara, T., Suzuki, O., and Saji H. (2002) cDNA microarray assessment for ozone-stressed Arabidopsis thaliana. Environ. Pollut. 117, 191–194. 16. Hormberg, J. J., de Haas, R. R., Dekker, H., and Lankela, J. (2002) Analysis of multiple gene expression array experiments after repetitive hybridizations on nylon membranes. BioTech. 33, 108–117. 17. Obayashi, T., Okegawa, T., Sasaki-Sekimoto, Y., Shimada, H., Masuda, T., Asamizu, E., Nakamura, Y., Shibata, D., Tabata, S., Takamiya, K., and Ohta, H. (2004) Distinctive features of plant organs characterized by global analysis of gene expression in Arabidopsis. DNA Res. 11, 11–25.

4 Use of cDNA Macroarrays and Gene Proﬁling for Detection of Effects of Environmental Toxicants Jason L. Blum, Melinda S. Prucha, Vishal J. Patel, and Nancy D. Denslow

Summary The method we describe in this chapter describes the synthesis and use of cDNA macroarrays for determining changes in gene expression due to environmental toxicants as well as the methods and materials that are required to do this work. While the details are for investigators working with nontraditional species for which commercial arrays are unavailable, anyone can design and use their own custom arrays using these protocols. We have intentionally left out details for statistical analysis for the arrays as the methods for doing this are still being developed and would need to be specific to the experiment being done. In all, gene macroarrays are a relatively easy way to generate large amounts of data in a short amount of time. Key Words: cDNA array; gene expression analysis; gene macroarray; nontraditional model.

1. Introduction Complementary DNA (cDNA) macroarrays are useful for detecting differences in gene expression caused by exposure to environmental toxicants. In this procedure, cDNA clones of interest are selected and immobilized on nylon membranes. RNA is isolated from a selected tissue, reverse transcribed to cDNA in the presence of radioactive nucleotides, and then incubated with the prepared nylon membranes to allow the labeled cDNA to bind to its complement on the array. Intensity of the signal is proportional to the amount of message in the sample RNA pool. From: Methods in Molecular Biology, vol. 410: Environmental Genomics Edited by: C. Cristofre Martin © Humana Press, Totowa, NJ

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For many investigators, cDNA arrays have become useful tools to identify potential new biomarkers. By using a few well-designed arrays, hypotheses can be generated and tested to gain greater insight into the direct or indirect global effects of toxicants, an approach that has been termed “Systems Toxicology” (1). Often, a toxicant may exert influence on biochemical pathways not previously considered. Through the use of gene arrays, one can get a better understanding for the mechanisms by which toxicants have genome-wide effects. In this way, small-scale theories can be moved into a larger-scale view to allow for a better understanding of the mechanisms by which toxicants work. Gene arrays offer the investigator the ability to screen for many different responses simultaneously from a single RNA sample. What previously would have taken larger amounts of RNA and longer time periods can be done in a fraction of the time using this technology. cDNA arrays, like other techniques, have their own advantages and disadvantages. The greatest advantage is the ability to simultaneously screen a large number of genes for changes in expression. Another advantage is that the starting sample size can be much smaller (just two micrograms of total sample RNA per array) compared to the 10–30 μg required per lane for Northern blotting (2). An important disadvantage to this technology, however, is the inability to determine different transcript sizes or to locate transcriptional start sites, which is more easily accomplished using Northern blotting or RPA (RNase protection assay). RPA uses solution hybridization, which has the advantage of being quantitative and fast, but requires synthesis of a labeled probe that can sometimes be troublesome to synthesize. In today’s world of modern molecular biology, we have seen the advent of numerous genome projects allowing for gene arrays to be made based on putative genes. By doing this, companies have created microarrays that can theoretically contain all gene transcripts. Toxicological research examining gene expression within traditional models (like human cell lines, rodents, and zebrafish) has become easier and more efficient than for those using nontraditional species, such as most fish species (3–5) and invertebrates (6). For those who use nontraditional species, species-specific arrays need to be created using cDNA clones already on hand. These are the people for whom this chapter is written. People who are studying species that are nontraditional cannot simply order clones off the shelf, like those that are available for human and mouse. For species where gene clones are not commercially available, libraries of genes must be generated in house. One method gaining in popularity is suppressive–subtractive hybridization from Clontech. This procedure allows for “subtraction” of one pool of cDNA from another (these can be two treatments or a treatment from control). A second, older method is differential

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display (GenHunter Corporation, Nashville, TN is one supplier) that allows for direct selection of differentially expressed genes from a gel for cloning and identification by sequencing. While still a useful method, it has fallen out of regular use in the past few years. 2. Materials 2.1. Ampliﬁcation and Preparation of the Clones 1. 2. 3. 4. 5. 6. 7. 8. 9. 10.

11. 12. 13. 14. 15. 16. 17.

Glycerol stocks of E. coli containing plasmids with cDNA inserts of interest. Taq polymerase and buffer (New England Biolabs cat. no. M0267). 10 mM dNTP mix (any commercial source). Primers to amplify the insert in your cloning vector diluted to 10 μM (M13, SP6, T4 or T7 are common ones). 70% Ethanol in nuclease-free water. Tris-EDTA buffer: 10 mM Tris, 1 mM EDTA, pH 8.0. 96-Well culture plates (any commercial source, round-bottom plates are useful for overnight cultures). Breathe easy strips for 96-well plates (USA Scientific cat. no. 9123-6100). LB medium: 10 g of tryptone, 5 g of yeast extract, 10 g of NaCl/L, pH 7.0. Millipore filtration system for purifying the polymerase chain reaction (PCR) products (MultiScreen Filtration System Vacuum Manifold cat. no. MAVM0960R and Montage PCR96 Cleanup plates cat. no. MANU03010). 96-Well PCR plates that fit your thermocycler (any commercially available plates for PCR will serve the purpose). Multichannel pipet, 20-μL and 200-μL sizes will be very helpful. Flat bottom 96-well or 384-well plates (any commercial source). Agarose gel electrophoresis set up capable of running the PCR products. DNA molecular weight ladder that has size range from 300 base pairs to 2000 base pairs (from any commercial source). UV spectrophotometer, preferably a plate reader model. 96-Well plates capable of being read in a UV spectrophotometer if desired (any commercially available, check your machine for suggestions).

2.2. Printing the Arrays 1. Precut Pall Biodyne B neutral nylon membranes (Nunc, cat. no. 250385). 2. Spot Report Array Validation System (Stratagene, cat. no.252005-7) which contains several controls [poly(dA), human Cot-1, Arabodopsis cDNAs, empty cloning vector, etc.] which should be spotted on the array. 3. UV-transilluminator (we use a UV Stratalinker 1800, Stratagene). 4. Centrifuge with a rotor for holding 96-well plates (any model will serve the purpose). 5. Robot (i.e., a Biomek 2000, Beckman Coulter using 100-nL pins). 6. Distilled water.

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7. 8. 9. 10.

10% Bleach. 70% Ethanol. Bromophenol blue. 20× SSC: 3 M NaCl, 0.3 M sodium citrate, pH 7.6, in nuclease-free sterile water. Autoclave to sterilize and store at room temperature. 0.01 mM bromophenol blue is added after sterilization.

2.3. Labeling, Hybridizing, and Washing the Arrays 1. M-MuLV reverse transcriptase enzyme and buffer [New England Biolabs, cat. no. M0253S (200 U/ml)]. 2. Random hexamer primers (New England Biolabs, cat. no. S1254S) add 80 μL of nuclease-free sterile water to 1 A260 unit. 3. 10× dNTP mix: can be purchased commercially or can be made by combining 20 μL each of 100 mM dCTP, dGTP, dTTP, and 10 μL 100 mM dATP. Bring the volume to 400 μL by adding nuclease-free sterile water (gives a final concentration of 4 mM dCTP, dGTP, dTTP, and 2 mM dATP). 4. [–33 P]dATP: from any supplier (Perkin Elmer cat. no. NEG612H for 250 μCi). 5. Spike RNA mix: commercially available from Stratagene as a kit with control spikes for printing the arrays or individual spikes can be purchased (Spike 2 RCA and Spike 3 rbcL, cat. no. 252202 and 252203, respectively). 6. 20× SSC: 3 M NaCl, 0.3 M sodium citrate, pH 7.6, in nuclease-free sterile water. Autoclave to sterilize and store at room temperature. 7. 20% SDS: 200 g of sodium dodecyl sulfate in 1 L of sterile water. 8. Hybridization buffer: 0.375 M NaCl, 0.0375 M sodium citrate, 7% SDS and 25% fresh formamide in a final volume of 500 mL made up with sterile nuclease-free water. Add 100 mg of yeast tRNA. Store at 4°C. 9. Wash buffer 1: 2× SSC–0.5% SDS (100 mL of 20× SSC, 25 mL of 20% SDS, and 875 mL of water). 10. Wash buffer 2: 0.5× SSC–0.5% SDS (25 mL of 20× SSC, 25 mL of 20% SDS, and 950 mL of water). 11. 10 mM EDTA, pH 8.0. 12. Kit to remove excess nucleotides from the labeling reaction (like Qiagen QIAquick Nucleotide Removal Kit, cat. no. 28304). 13. Dry bath incubators (2). 14. Liquid scintillation counter and buffer. 15. Hybridization bottles (Fisher Scientific). 16. Hybridization oven (we use a MaxiOven from Labnet).

2.4. Scanning and Quantiﬁcation of the Arrays 1. 2. 3. 4.

Transparencies: from any office supply store. Phosphor screen: Molecular Devices. Phosphor imager: We use Molecular Devices Typhoon Scanner. ImageQuant software: Molecular Devices.

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3. Methods 3.1. Amplifying the cDNA Clones (Blum et al., 2004) 1. Dispense 98 μL of LB broth containing appropriate antibiotics into each well of a 96-well plate. 2. Add 2 μL of each glycerol stock containing the plasmid of interest to the LB (one clone per well). Also, include an empty vector as another control for your array (should have very low hybridization when hybridized with your sample). 3. Cover the plate with a breathe easy strip and replace lid on the plate. 4. Allow the cultures to grow overnight at 37°C with shaking (about 125 rpm). 5. The next morning, make up a PCR master mix for 90 μL per reaction (Table 1) (make enough for 100 reactions if using a full plate). See Fig. 1A for a schematic for this method. 6. Dispense 90 μL of PCR master mix to each reaction well, and add 10 μL of overnight bacterial culture to each well. 7. Cover the PCR plate with an appropriate cover and cycle as described in Table 1. 8. After cycling, analyze the PCR products on a 1% agarose gel to ensure that clones were amplified with 5 μL of each product along with a DNA ladder. Re-run the PCR for any clones that did not properly amplify.

Fig. 1. Schematic of array printing process and cDNA labeling reaction. (A) General flowchart for the major steps involved in printing a nylon cDNA gene array. Starting from glycerol stocks containing the genes of interest, one can print an array. (B) The major steps for starting from total RNA to having labeled single-stranded cDNA.

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Table 1 PCR Master Mix and Cycling Conditions for Array Plates

Component

Volume per Reaction (L)

Nuclease-free water 10× Polymerase buffer MgCl2 dNTPs (10 mM) Forward primer (10 M)

59 10 8 8 2

Reverse primer (10 μM) Taq polymerase Total

2 1 90

Segment (Step) 1 2 3 4 5: Go to step 2, repeat 34X 6 7

Temperature 95ºC, 10 min 95ºC, 1 min 57°C,a 1 min 72ºC, 2 min

72ºC, 7 min 4ºC, hold

a The annealing temperature is dependent on the primer set for the cloning vector you use. The temperature listed is for the M13 primer set.

9. After you know that the clones have all amplified, load them onto the Millipore filtration unit and allow the fluid to flow through the membrane. Wash once with 70% ethanol. 10. Turn off the vacuum and resuspend the product by adding 50 μL of TE buffer, and incubate at room temperature for several minutes, and then pipet (a multichannel pipet is useful for this) into another 96-well plate. 11. Measure the concentration using a UV-spectrophotometer by diluting 5 μL of purified DNA in 95 μL of water. 12. If necessary, reamplify any clones that did not provide enough material and these can be pooled with the first round if desired. You will need at least 160 ng/L for printing.

3.2. Printing the Arrays 1. Before starting this step, you should plan the layout of genes on your array. We always spot genes in duplicate (see Note 1), keeping in mind that you need to include controls on your array (see Notes 2 and 3). 2. Dilute purified DNA (3000 ng in 18.75 μL = 160 ng/L) and place into the desired well for printing. 3. Add 1.9 μL of 3 M NaOH to each well. 4. Cover the plate and incubate at 65°C for 15 min (we use the hybridization oven for this). 5. Immediately chill on ice for 2–3 min. 6. Centrifuge the plate briefly. 7. Remove the plate cover and add 9.35 μL of 20× SSC that contains bromophenol blue to each of the wells, pipet up and down (volume should now be ∼30 μL).

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8. Centrifuge the plate again briefly at 100g for 5 sec. (It is very important to do this.) 9. Set the plate, membranes, bleach, water, and ethanol onto the robot and begin printing the arrays. You will need to program your robot accordingly (see Note 4). 10. In between printing sets (when all of the arrays on the platform have been printed) it is necessary to clean the pins. Have the robot submerge the pins for 10 seconds each in bleach, then water, then ethanol, and then use the fan to dry the pins and then print again. While the pins are being cleaned, a new set of membranes can be made ready. 11. Crosslink the DNA to the membrane using a UV-transilluminator at the setting used for Southern blots (100 mJ). 12. Using 100-nL pins with the robot it is possible to print over 100 membranes with high fidelity.

3.3. Labeling Samples (see Fig. 1B for diagram of this procedure) 1. Remove radiation from freezer and keep behind a protective Plexiglas shield. Turn on the 37°C and 100°C water baths. 2. Turn on the hybridization oven and set it to 64°C and place the hybridization buffer in the oven to preheat. 3. In a microcentrifuge tube add 2 μL of random primers, 0.6 μL of spike RNA mix, 2 μg of sample RNA and add DEPC water to bring the volume up to 13 μL. 4. Place the tube at 64°C for 5 min, and then let it cool down at room temperature for 5–10 min to allow the primers to anneal. 5. After the tubes have cooled, add 2 μL of 10× RT buffer, 2 μL of 10× dNTP mix, 1 μL of M-MuLV enzyme, and 2 μL of [-33 P]dATP. (Total volume in the tube = 20 μL). 6. Incubate at 37°C for 1.5–2 h. 7. Centrifuge the tubes down before opening. 8. Purify the labeled samples using a nucleotide removal kit. 9. Check counts of the sample on a liquid scintillation counter (we count 2 μL of the labeled material).

3.4. Hybridization and Washing Arrays 1. Start prehybridizing the membranes. Place the membranes in the hybridization bottles, add 5–6 mL of hybridization buffer (see Note 5), and return the bottles back into in the hybridization oven and rotate (12–14 rpm) at 64°C for 1.5–2 h (Fig. 2A). 2. Calculate the required volume of radiolabeled cDNA to be used for hybridization. Each membrane needs to be hybridized in the same concentration of radioactivity (1 million cpm/mL). Formula 1 Calculation for diluting labeled cDNA Probe = 1000000 cpm/mL∗ V /cpm/μL

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Fig. 2. Schematic for hybridization and washing of the arrays. (A) The major steps for the hybridization procedure. (B) Procedure for how to wash the arrays the day after hybridization, and laying the membranes out on a phosphor screen. Note the asymmetrical layout of the membranes to facilitate identification after scanning.

where Probe = the number of milliliters of label required in L and V = volume of hybridization buffer in mL. Then add 10 mM EDTA. The amount of EDTA required is 20 times the volume of Probe calculated above. Formula 2 Calculation for the addition of EDTA Probe∗ 20 = μL 10mM EDTA to be added to the label

3. Add the volume of probe calculated from formula 1 to volume of 10 mM EDTA computed in formula 2. Combine these in either a 1.5-mL centrifuge tube that you can put a locking cap on or into a screw cap tube to prevent the tube top from opening during the denaturation step. 4. Denature probe at 100°C for 5 min and then place it on ice for 2 min. 5. Centrifuge the tubes at 2,700 × g for 1 min and add the contents directly into the buffer of the appropriate hybridization bottle, already containing the membrane. 6. Shake the bottle slightly and place it back into the hybridization oven at 64°C. Hybridize overnight, about 14 h at 12–15 rpm.

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7. Place wash buffers 1 and 2 in the oven at 64°C (overnight). There is usually enough room on the sides of the hybridization oven to accommodate the wash buffers. 8. The following morning you may begin washing the membranes. Instructions for each wash (see Fig. 2B): a. First, pour off the hybridization buffer into radioactive waste as required by your institution, then add a couple of inches of wash buffer 1 to each hybridization bottle. Make sure that the buffer covers at least half of the membrane. Return to the hybridization oven for 30 min rotating at 12–14 rpm. b. Repeat the wash with buffer 1 three more times and then wash 4 times using wash buffer 2 (also 30 min each). We collect all washes as radioactive waste to be safe, but follow the guidelines at your site. c. You should complete a total of eight wash steps. 9. Remove the membranes from the bottles, dry them, and then place them in the phosphor imager cassette (see Note 6). (Note: By placing the transparency sheet on the grid portion of the cassette and placing the membranes in alignment to the grid, it makes the membranes easier to analyze; see Note 7) Develop the image of the membrane on the phosphor screen for 48 h and scan it with a typhoon scanner.

3.5. Quantiﬁcation After exposure to the phosphor screen, the membranes can be removed from the cassette and the screen can be developed. For spot quantification, we have used the ImageQuant 5.1 software package (Molecular Devices). First, the image is adjusted for contrast (this is only for the eye and does not affect the numbers in later analysis) and then a pair of boxes is drawn around the largest or most intense pair of spots (single gene). This pair of boxes is then copied to the other genes so that the area is maintained for all of the analytes. 1. After the boxes are drawn and all of the data from all of the membranes are collected, compute the mean of each gene pair and subtract the mean of all of the blank spots that were designed into the array. 2. Next, the genes on each membrane need to be normalized in such a way that they can be compared to other membranes of the set. This may include scaling each membrane to a particular mean or median, usually of the highest membrane. 3. Next log transform the data set to make the data appear more normally distributed. A statistician or bioinformatics expert may be helpful for this. 4. Use the appropriate statistical model and method for making your comparisons that you have determined prior to running the array experiment. 5. After determining which genes are most interesting, they should be confirmed by other methods, such as real-time or quantitative PCR.

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3.6. Statistical Analysis and Data Mining The most important thing to consider before doing gene array experiments is to decide how to analyze the final data. This includes the number of individuals per treatment, whether to pool individuals, what comparisons are going to be made, and the statistical tests to be used to make these comparisons. Since the methods for analyzing array experiments are still evolving, it would be best to consult with a statistician or bioinformatics specialist for assistance before starting these experiments, rather than afterwards. There are several ways to depict the data as well and usually this depends on the type of experiment that was done.

4. Notes 1. We always spot the genes in duplicate in order to safeguard against mis-spotting that may occur with the robot. 2. When designing the layout of your array, it is important to keep in mind that you need to include controls. We use an empty vector amplicon, several spots with no DNA (only 20× SSC with bromophenol blue), as well as messages for ribosomal proteins, in addition to the spikes and other cDNAs that come in the array spot reporter kit from Stratagene. 3. Since printing the arrays can be costly, expense may be controlled if a targeted array is designed such that two or more arrays are printed on a single membrane. 4. A robot is not absolutely necessary as hand printing stamps are commercially available, but to print in this fashion would be difficult for large quantities of arrays and they would be less consistent. It may be possible also to use a vacuum driven dot blotter, but this has not been tried and would likely be very tedious. 5. We generally use 5 mL of hybridization buffer, but the volume used depends on the size of the hybridization bottles. The larger the bottle the more buffer you need, as long as half of the membrane remains in contact with the buffer. 6. If you do not have a phosphor imager available to you, you can also use regular X-ray film. However, the dynamic range of film is several logs less than that of the phosphor screen so resolving differences may become more difficult. If film is used, photodensitometry can be used in place of ImageQuant or other phosphor imaging software. 7. Placing the membranes in an asymmetrical pattern in the phosphor screen will also enable easier identification of which individual membrane is which when you develop the phosphor image.

References 1. Waters, M. D. and Fostel, J. M. (2004) Toxicogenomics and systems toxicology: aims and prospects. Nat. Rev. Gen. 5, 936–948.

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2. Sambrook, J., Fritsch, E. F., and Maniatis, T. (1989). Molecular Cloning: A Laboratory Manual, 2nd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York, pp. 7.43–7.45. 3. Blum, J. L., Knoebl, I., Larkin, P., Kroll, K. J., and Denslow, N. D. (2004) Use of suppressive subtractive hybridization and cDNA arrays to discover patterns of altered gene expression in the liver of dihydrotestosterone and 11-ketotestosterone exposed adult male largemouth bass (Micropterus salmoides). Mar. Environ. Res. 58, 565–569. 4. Knoebl, I., Blum, J.L., Hemmer, M.J., and Denslow, N.D. (2006) Using gene arrays to determine temporal gene induction in sheepshead minnows exposed to 17-estradiol. J. Exp. Zoolog A Comp Exp Biol. 305, 707–719. 5. Larkin, P., Sabo-Attwood, T., Kelso, J., and Denslow, N.D. (2002) Gene expression analysis of largemouth bass exposed to estradiol, nonylphenol, and p,p -DDE. Comp. Biochem. Phys. Part B 133, 543–557. 6. Brouwer, M., Larkin, P., Brown-Peterson, N., King, C., Manning, S., and Denslow, N. (2004) Effects of hypoxia on gene and protein expression in the blue crab, Callinectes sapidus. Mar. Environ. Res. 58, 787–792.

5 Constructing and Screening a cDNA Library Methods for Identiﬁcation and Characterization of Novel Genes Expressed Under Conditions of Environmental Stress Kevin Larade and Kenneth B. Storey

Summary Many organisms provide excellent models for studying disease states or for exploring the molecular adaptations that allow cells and organisms to cope with or survive different stresses. The construction of a cDNA library and subsequent screening for genes of interest allows researchers to select for genes that are likely to play key roles in the regulation or response to the condition or stress of interest, those that may not be expressed (or exist) in other systems. Determination of the open reading frame(s) of novel genes, and extensive analysis of the proteins they encode, can open up new avenues of research and promote intelligent design of downstream projects. Key Words: Bioinformatics; cDNA library; differential screen; expression analysis; functional genomics; gene characterization; novel genes; stress induction; up-regulated.

1. Introduction A majority of coding genes that make up an animal’s genome are under selective pressure, such that there is little room for evolving function due to the importance of the wild type gene. However, in some cases, genes that are dormant (in selected regions of heterochromatin) or genes that may have arisen as a result of whole or partial gene duplication (with switching of selected exons and/or introns), may confer a selective advantage to organisms living in a certain environment or exposed to certain conditions. Extra copies of From: Methods in Molecular Biology, vol. 410: Environmental Genomics Edited by: C. Cristofre Martin © Humana Press, Totowa, NJ

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a gene or “spliced copies” may become activated (or mutated) to code for new proteins, in effect supplying “novel” genes to the existing suite of genes present in an organism. Although a number of genome projects are complete and numerous others are ongoing, a wide variety of fascinating organisms exist for which only selected genes have been identified (1,2). By examining the response and regulation of gene expression in systems that not only survive, but, in many cases thrive under adverse conditions, insight can be gained into the prevention and potential treatment of these types of systemic stresses. The end goal is to determine the function of such genes. Advances in the field of functional genomics is helping to address this goal and advances in bioinformatics, combined with the availability of large data sets for wellannotated genes, has provided a basis for the generation of software and analysis programs that allow researchers to examine nucleotide and protein sequences using established “rules” and algorithms. Much information can be gleaned from the nucleotide sequence itself, as well as the primary amino acid sequence of the translated gene, such that informed hypotheses can be constructed and tested prior to initiating larger downstream projects. Such preliminary analysis sets the stage for further experiments, such as the cloning and expression of open reading frames, examination of the upstream promoter, construction of mutated constructs that affect gene expression, and possibly even the creation of a knock-out animal to examine the in vivo effects of the loss of select genes. The first step in this process is the identification of differentially expressed genes for the condition of interest. This chapter outlines the basic construction of a cDNA library from a select population of mRNA, including methods for mRNA/cDNA preparation and cloning, subsequent screening for differentially expressed genes, and downstream confirmation via Northern blotting. Methods for analysis of select sequences are also outlined using the previously unreported novel gene, sarp-2, which is included as an example. Sarp-2 is one of several novel clones (3,4) isolated from a cDNA library constructed with mRNA prepared from hepatopancreas of the marine snail, Littorina littorea; the gene is induced by anoxic exposure. Identification and characterization of this novel gene was performed as outlined in this chapter and the results are reported in the accompanying figures. 2. Materials All chemicals used are of molecular biology grade or their equivalent and of the highest purity. All plastic and glassware, including bottles and pipet tips, are autoclaved and gloves must be worn at all times during operations involving nucleic acid manipulation.

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2.1. Total RNA Isolation 1. RNase-free water. Add 1 mL of diethyl pyrocarbonate (DEPC) (Sigma-Aldrich) to 1 L of water (0.1% v/v), stir overnight (>12 h), autoclave. This destroys any RNases present and this water will be used to make up solutions in this section and to dissolve RNA samples. 2. Trizol reagent (Invitrogen). 3. Chloroform (Fisher Scientific). 4. Isopropyl alcohol (Fisher Scientific). 5. 70% Ethanol. Add 30 mL of DEPC-treated water to 70 mL of ethyl alcohol (200 proof; Pharmco). 6. Oligo(dT) cellulose (New England BioLabs). Dry oligo(dT) cellulose is combined with 0.1 M NaOH to create a slurry and poured into a sterile column or 1-mL syringe plugged with sterile cotton or glass wool. The column is equilibrated with loading buffer prior to adding sample. 7. Loading buffer: 1 M NaCl, 2 mM phosphate buffer, pH 7.2. 8. Middle wash buffer: loading buffer + 0.3 M NaCl. 9. Elution buffer: 10 mM Tris-HCl pH 7.2–7.4, 1 mM EDTA. 10. 3 M sodium acetate, pH 5.2. 11. Ethanol (200 proof; Pharmco).

2.2. cDNA Library Synthesis 1. Cloning vector for cDNA library synthesis. Many vectors exist, often designed for a specific application. Researchers should examine the features of those available and decide on a vector appropriate for their application. Some common vectors used in cDNA library synthesis include: Uni-ZAP XR (Stratagene), plTriplEx2 (Clontech), pSPORT1 (Invitrogen), lSCREEN-1 (Novagen/Merck). The Uni-ZAP XR cloning vector will be used for illustrative purposes in this chapter. 2. 1 μg of column purified mRNA. 3. Oligonucleotide linker-primer containing a poly (dT) region. (ex. 5 -NNNNN NNNCTCGAGdT(15)-3 ). 4. RNasin (20 U/μL; Promega). 5. AMV reverse transcriptase (10 U/μL) with 10× reverse transcriptase buffer (Promega). 6. Nucleotides (dATP, dGTP, dTTP; 10 mM each dNTP) 7. 5-Methyl cytosine analog (*dCTP; 5 mM). 8. Ribonuclease (RNase) H (5 U/μL; New England BioLabs). 9. DNA polymerase I (E. coli, 10 U/μL) and 10× DNA polymerase I buffer (New England BioLabs). 10. Eco RI adapters. An EcoRI adaptor can be purchased commercially or constructed using two primers of different lengths, which can be hybridized to form a duplex with a 5 phosphate on the blunt end. Include an overhang of 5 -AATTCNNNN-3 on one primer to create an EcoRI restriction site. The opposite end of the duplex (the blunt end of the adaptor) can be ligated to the ends of any cDNA containing

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

19.

20.

21.

Larade and Storey a 5 phosphate group. The adapters should be diluted in sterile ddH2 O to a final concentration of 5 μg/μL. 50× TAE buffer: 2 M Tris-acetate (Tris + acetic acid) pH 8.5, 100 mM EDTA. 1% TAE agarose gel: 1× TAE buffer, 1% agarose (1 g/100 mL), ethidium bromide (1 μg/mL). 100 mM EDTA. T4 DNA ligase (400 U/μL) and 10× T4 DNA ligase buffer (New England BioLabs). T4 polynucleotide kinase (10 U/μL) and 10× T4 polynucleotide kinase buffer (New England BioLabs). TSM buffer: 50 mM Tris-HCl, pH 7.5, 100 mM NaCl, 8 mM MgSO4 , 0.01% (w/v) gelatin. XL1-Blue (Stratagene). LB-ampicillin broth: 10 g of tryptone, 5 g of yeast extract, 5 g of NaCl, pH 7.5. Dilute up to 1 L with ddH2 O and autoclave. Broth is supplemented with 0.2% (w/v) maltose and ampicillin (100 μg/mL final concentration) after broth has cooled to room temperature. NZY agar: 5 g of NaCl, 2 g of MgSO4 7H2 O, 5 g of yeast extract, 10 g of peptone, 15 g of agar, pH 7.5. Dilute to 1 L with ddH2 O and autoclave. Pour plates when agar has cooled to approx. 50°C and store at 4°C until used. The agar can also be stored at 4°C and remelted in a microwave just before use. NZY top agar: 5 g of NaCl, 2 g of MgSO4 7H2 O, 5 g of yeast extract, 1 g of peptone/L, 0.7% w/v agarose, pH 7.5. Dilute up to 1 L with ddH2 O and autoclave. The top agar is stored at 4°C and remelted in a microwave just before use. The agar must be cooled to <45°C before addition of bacteria. Pfu polymerase (2.5 U/μL) (Stratagene).

2.3. Ampliﬁcation of the cDNA Library 1. NZY plates (24 × 24 cm), LB-ampicillin broth, XL1-Blue cells, NZY agar, NZY top agar (see Subheading 2.2., items 19 and 20). 2. TSM buffer (see Subheading 2.2., item 16). 3. Chloroform (Fisher Scientific). 4. Dimethyl sulfoxide (DMSO; Sigma-Aldrich).

2.4. Differential Screening of the cDNA Library 1. NZY plates (24 × 24 cm; see Subheading 2.2., items 19 and 20). 2. Hybond-N 0.45-μm nylon membranes (GE Healthcare). 3. 20× SSC: 3 M NaCl, 0.3 M Na-citrate (for 1 L: 175 g of NaCl, 88 g of sodium citrate); dilute as necessary with ddH2 O. 4. Denaturation buffer: 1.5 M NaCl, 0.5 M NaOH. 5. Neutralization buffer: 1.5 M NaCl, 0.5 M Tris-HCl, pH 8.0. 6. Rinse buffer: 0.2 M Tris-HCl, pH 8.0, 2× SSC. 7. Whatman filter paper or chromatography paper (Fisher Scientific). 8. dNTPs: dATP/dTTP/dGTP (10 mM each dNTP).

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9. Anchored Oligo(dT) primer: (5 -dT(15)(A/G/C)-3 ). Can be purchased commercially (Invitrogen; New England BioLabs). 10. RNasin (20 U/μL; Promega). 11. Dithiothreitol (DTT) (Sigma-Aldrich). Make up a 0.1 M stock with sterile ddH2 O. 12. AMV reverse transcriptase (10 U/μL) with 10× reverse transcriptase buffer (Promega). 13. [-32 P]dCTP (3000 Ci/mol; GE Healthcare). 14. RNase A (60 mg/mL in ddH2 O; Sigma-Aldrich). 15. Quick Spin columns (TE; Sephadex G-25, Fine) for radiolabeled DNA purification (Roche). 16. 1× T10 E1 buffer: 10 mM Tris-HCl, pH 8.0, 1 mM EDTA. 17. Hybridization buffer. Modified Church’s Buffer: 0.25 M Na2 HPO4 , 0.25 M NaH2 PO4 (pH 7.5), 1 mM EDTA, 7% sodium dodecyl sulfate (SDS; w/v); or buffer can be purchased commercially: Ultrahybe (Ambion). 18. Membrane washing buffer: 0.1× SSC, 0.1% SDS (w/v). 19. Membrane rinsing buffer: 0.1× SSC. 20. X-ray film (Kodak). 21. TSM buffer (see Subheading 2.2, item 16). 22. Chloroform (Fisher Scientific). 23. XL1-Blue culture. 24. NZY top agar (see Subheading 2.2, items 19 and 20).

2.5. In Vivo Excision of Clones 1. 2. 3. 4. 5. 6. 7.

XL1-Blue cells. 10 mM MgSO4 . ExAssist helper phage. Uni-ZAP XR phage stock. LB-ampicillin broth (see Subheading 2.2, item 18). SOLR cells. LB-ampicillin agar plates: Agar (15 g) is added to LB broth (1 L) and autoclaved. Cool to <50°C, add ampicillin (100 μg/mL), and pour into 10-cm plates at desired thickness.

2.6. Miniprep Plasmid Isolation 1. 2. 3. 4.

LB-ampicillin broth (see Subheading 2.2, item 18). Prelysis buffer: 50 mM glucose, 25 mM Tris-HCl pH 8.0, 10 mM EDTA. RNase A (see Subheading 2.4, item 14). Alkaline lysis buffer: 0.2 N NaOH, 1% SDS. This buffer must be made fresh just prior to use. 5. Neutralization buffer: 5 M potassium acetate, 30% acetic acid, 50 mM glucose, 25 mM Tris-HCl, pH 8, 10 mM EDTA. 6. Isopropanol (Fisher Scientific). 7. 70% Ethanol (see Subheading 2.1, item 5).

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2.7. Insert Isolation 1. Restriction enzymes: EcoRI and XhoI (New England Biolabs); these may be adjusted depending on the adaptors ligated. 2. 10× restriction enzyme buffer. Buffer will depend on restriction enzymes being used and is supplied with the enzyme. 3. DNA loading dye: 0.25% w/v bromophenol blue, 0.25% w/v xylene cyanol FF, 50% v/v glycerol. 4. 1% TAE agarose gel and 50× TAE buffer (see Subheading 2.2, item 11 and 12). 5. DNA ladder (Invitrogen). A ladder should be chosen based on the size of the clones expected (ranging from 100 bp up to several kilobases). 6. Filtered pipette tips (1 mL). 7. Sterile cotton or glass wool.

2.8. Synthesis of Labeled Probe 1. dNTPs without dCTP (dATP/dTTP/dGTP; 5 mM each). 2. Random primer (100 mM d(N)6 ; New England BioLabs). 3. 10× random primer buffer: 0.5 M Tris-HCl, pH 7.6, 20 mM DTT, 50 mM MgCl2 , 0.4 M KCl. 4. Klenow fragment of DNA polymerase I (5 U/μL; New England BioLabs). 5. [-32 P]dCTP (3000 Ci/mol; GE Healthcare). 6. Quick Spin columns (TE; Sephadex G-25, Fine) for radiolabeled DNA purification (Roche).

2.9. Northern Blotting 1. 10× MOPS buffer: 200 mM 3-(N-Morpholino)propanesulfonic acid (MOPS), 50 mM sodium acetate, 10 mM EDTA, pH 7. 2. 1.25% Agarose formaldehyde denaturing gel. Melt 3.75 g of agarose in 217 mL of ddH2 O containing ethidium bromide (1 μg/mL) in a sterile flask. Place solution in an incubator set to 60°C. Into a separate sterile flask, add 30 mL of MOPS 10× buffer and 53 mL of formaldehyde 37% (v/v) and place this solution at 60°C. Once both solutions have equilibrated to 60°C, combine the contents of both flasks together, gently swirl without introducing bubbles, and pour into large gel tray to desired thickness. 3. RNA sample buffer: 1× MOPS buffer, 2.2 M formaldehyde, 50% (v/v) formamide. 4. RNA loading buffer 6× stock: 1× MOPS buffer, 50% (v/v) formamide, 40% (v/v) glycerol. Add a few flakes of bromophenol blue and xylene cyanol as tracking dyes. 5. Hybond 0.45-μm nylon membrane (GE Healthcare). 6. 20× SSC (see Subheading 2.4, item 3). Dilute as necessary with ddH2 O. 7. Whatman filter paper or chromatography paper (Fisher Scientific). 8. Paper towels. 9. Flat piece of Plexiglas or sequencing gel plate. 10. Solid weight.

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11. Glass or plastic pipet (5- or 10-mL) for smoothing the gel/membrane/filter paper and removing air bubbles from transfer set-up. 12. Plastic wrap. 13. Northern blot fixing solution: 0.05 N NaOH.

2.10. Probing Northern Blots 1. Hybridization solution: Ultrahybe Hybridization buffer (Ambion) or modified Church’s buffer (see Subheading 2.4. item 17). 2. Radiolabeled probe (prepared in Subheading 2.8). 3. Northern blot washing solution: 0.1× SSC, 0.1% SDS in sterile ddH2 O. 4. X-ray film or phosphoimager screen. 5. Northern blot stripping solution: boil a solution of ddH2 O and 0.5% SDS and immerse blots to be stripped. Repeat as necessary until radioactive label (cpm) detected using a handheld Geiger counter is at or below background levels.

3. Methods Commercial products have been designed for specific types of experiments (high vs. low copy number genes, etc.) that examine and identify differentially expressed genes. Although the technology changes rapidly, cDNA library construction using mRNA isolated from a tissue/organ/organism of interest remains the standard method for identifying new and novel genes and proteins. Synthesis of such a library can be carried out using a wide variety of available products; a common standard method for library construction, screening and positive clone analysis is outlined in this chapter. These instructions assume the use of the Uni-Zap-cDNA synthesis kit (Stratagene; see ref. 5), but can be adjusted for the use of other vectors. Some common phagemid cloning vectors available include, but are not limited to: plTriplEx2 (Clontech), pSPORT1 (Invitrogen), and lSCREEN-1 (Novagen). Following construction of the library, differential screening will identify selected clones that are induced (or repressed) in the condition being tested. In the example provided here, we follow the steps used to find and characterize the novel gene sarp-2, which is induced in the hepatopancreas of Littorina littorea in response to anoxia exposure. This up-regulation is confirmed with Northern blotting. Following sequencing of the sarp-2 clone, sequence analysis can be performed using the applications outlined in Subheading 3.11. 3.1. Total RNA Isolation 1. A number of kits are available today for total and mRNA isolation (Ambion, Roche, Qiagen, Invitrogen, Novagen, etc.). These kits often provide high quality, pure RNA, but are relatively expensive. A traditional method for RNA isolation is outlined in this section.

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2. Homogenize tissue samples in Trizol reagent (1 mL per 100 mg of tissue) and add chloroform (0.2 mL/mL of reagent). Invert samples repeatedly (15 s), incubate at room temperature (2–3 min), and centrifuge at 12,000g at 4°C for 15 min. 3. Remove the aqueous layer and transfer to a sterile Eppendorf tube. Add an equal volume of isopropanol to precipitate the RNA. Incubate samples at room temperature for 10 min. 4. Pellet the RNA by centrifuging the samples at maximum speed in a microcentrifuge for 10 min at 4°C. Aspirate the supernatant and wash the pellet with 70% ethanol (250 μL). Recentrifuge the RNA at maximum speed for 5 min. Aspirate the ethanol and air-dry the pellet for 10 min on the lab bench. Dissolve the RNA in an appropriate amount of DEPC-treated water (assuming you get 10 μg for every 10 mg of tissue) and store at 4°C (if it will be processed within a week) or at –20°C (long-term storage). 5. Determine RNA concentration and purity spectrophotometrically by measuring absorbance at 260 and 280 nm. For RNA of acceptable quality, the A260 /A280 ratio should fall between 1.6 and 2.0. (See Note 1.) 6. Purify the poly(A)+ mRNA from total RNA samples using oligo(dT) cellulose. chromatography. Heat total RNA to 65°C, mix with loading buffer (1:1 v/v), and load onto an oligo(dT) cellulose column. 7. Wash the column with loading buffer (1 bed volume) and collect the eluate in a sterile Eppendorf tube. Load the recovered eluate onto the column, collect it as it elutes, then reload onto the column (repeat twice). 8. Wash the column with loading buffer (1 bed volume), followed by middle wash buffer (1 bed volume), and finally with elution buffer (1.5 bed volume). This last step elutes the poly(A)+ mRNA, to which 0.1 volumes of 3 M sodium acetate, pH 5.2, and 2.5 volumes of absolute ethanol are added. The poly(A)+ mRNA is placed at –20°C and left to precipitate overnight. The protocol can be stopped here. 9. The following day, centrifuge the tubes at 13,000g for 45 min at 4°C. Discard the supernatant and wash the pellet with 70% ethanol (250 μL) and centrifuge at the same speed for 10 min. The resulting pellet consists of mRNA, which is air-dried and resuspended in DEPC-ddH2 O (see Note 2). If the mRNA is not going to be used immediately, store at –80°C until further use.

3.2. cDNA Library Synthesis 1. Combine mRNA (5 μg; <30 μL) with the oligonucleotide linker-primer (3 μg) and RNasin (5 μL) in an RNase-free Eppendorf tube. Synthesize a complementary first strand of cDNA, using the mRNA as template, with the addition of dNTPs (3 μL), 5-methyl-dCTP (3 μL), 10× reverse transcriptase buffer (5 μL), and adjusting the total volume up to 45 μL. Mix the contents gently and allow primer and template to anneal at room temperature for 10 min. 2. Add reverse transcriptase to the reaction tube (5 μL; 50 U) to bring the final volume up to 50 μL. Gently mix the contents of the tube, briefly centrifuge to collect contents at the bottom of the tube, and then place at 42°C for 1 h. (See Note 3.)

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3. Second-strand cDNA synthesis (complementary to the first strand) is carried out in the same tube. The following components are added to the first strand reaction: dNTPs (6 μL), 10× DNA polymerase I buffer (20 μL), [-32 P]dCTP (1 μL), RNase H (1 μL; 5 U), DNA polymerase I (10 μL; 100 U). Adjust the final reaction volume to 200 μL with sterile ddH2 O and incubate for 2 h at 16°C. 4. Place the newly double-stranded cDNA reaction on ice and blunt-end the termini by adding 22.5 μL of dNTPs (A/G/C/T) and Pfu DNA polymerase (2.5 μL) into the tube and incubate at 72°C for 30 min. 5. Add an equal volume of phenol–chloroform (1:1 v:v, pH 7.4) and vortex-mix the tube briefly. Centrifuge the sample at maximum speed in a desktop centrifuge for 10 min at room temperature. Transfer the aqueous layer to a fresh Eppendorf tube and add an equal volume of chloroform. Vortex-mix the tube briefly and centrifuge as above for 5 min. Transfer the aqueous layer to a fresh tube. 6. Add an equal volume of isopropanol to the sample, followed by addition of sodium acetate (to a final concentration of 0.1 M). Place the tube at –20°C overnight to allow the cDNA to precipitate. The protocol can be stopped at this step. 7. The following day, centrifuge the precipitated cDNA for 1 h at maximum speed in a microcentrifuge at 4°C. Transfer the supernatant into a new tube and store until cDNA precipitation has been confirmed. Using a Geiger counter, confirm radiolabel incorporation into the precipitated cDNA pellet. Check the supernatant to confirm that most of the radioactivity is in the pellet. 8. Wash the pellet by adding 70% ethanol (250 μL), followed by gentle agitation. Centrifuge the tube at maximum speed in a microcentrifuge for 10 min. Carefully remove and discard the ethanol, taking care not to disturb the pellet, and air dry for 10 min at room temperature. The location of the pellet should be marked on the outside of the tube with a labelling marker because it will “disappear” once it is dried. 9. Resuspend the cDNA in sterile ddH2 O (10 μL). Gently flick the sample and briefly centrifuge to collect contents at the bottom of the tube. The tube is kept on the bench for 10 min at room temperature (or at 4°C for 30 min) to allow the cDNA to fully dissolve. Transfer the cDNA to a new tube and check the old tube with a Geiger counter to confirm cDNA was completely resuspended. (See Note 4.) 10. Ligate EcoRI adapters (1.5 μL) to the blunt-ended termini by adding T4 DNA ligase (5 U) and 10× T4 DNA ligase buffer (1.5 μL). Final reaction volume is 15 μL (make up the final volume with sterile ddH2 O if required). Incubate the reaction for 16 h at 10°C. 11. Heat inactivate the ligase by placing the tube at 75°C for 15 min, followed by brief centrifugation to collect the contents at the bottom of the tube. Place the sample on ice and add the following components: T4 polynucleotide kinase (10 U), 10× T4 polynucleotide kinase buffer (0.5 μL). Adjust the final reaction volume in the tube to 20 μL. Incubate the reaction at 37°C for 30 min, followed by heat inactivation of the kinase at 75°C for 15 min. Centrifuge the tube briefly to collect the contents at the bottom and place tube on ice.

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12. Digest the linker primer’s XhoI site by adding 4 μL of the appropriate 10× restriction enzyme buffer, 10 μL of sterile ddH2 O, and 6 μL of XhoI enzyme to the cDNA (to a final volume of 40 μL). Mix the reaction by flicking, centrifuge briefly to collect contents at the bottom of the tube, and place at 37°C for 2 h. Add an equal volume of isopropanol and place tube at –20°C overnight to precipitate the digest. (See Note 5.) 13. Centrifuge the precipitated cDNA at maximum speed in a microcentrifuge for 1 h at 4°C. Aspirate and discard the supernatant, taking care not to disturb the pellet. 14. Wash the pellet with 70% ethanol (250 μL) and centrifuge for 10 min at maximum speed at 4°C. Carefully aspirate the supernatant and air-dry the pellet on the benchtop for 10 min (see Note 6). Resuspend the pellet in the appropriate volume of ddH2 O (in this case we will use 50 μL) and store at 4°C. 15. Prepare 10 mL of 1% TAE agarose gel, pour into a 10-cm plate and allow the gel to polymerize. Create a grid on the bottom of the plate with 10 squares. 16. Using a 100 mM EDTA solution, make serial dilutions (from 10 ng/μL up to 250 ng/μL) of a DNA standard with a known concentration (such as the DNA ladder in Subheading 2.7.5.). 17. Label a square for each of the serial dilutions (0, 5, 10, 25, 50, 75, 100, 150, 200, 250 ng/μL). 18. Carefully pipet 1 μL of each dilution onto the surface of the plate. 19. Below each dilution, spot 1 μL of the cDNA sample and allow all the liquid to absorb into the plate for approx. 20 min. 20. View the plate using ultraviolet (UV) light and estimate the concentration of the cDNA sample by comparing with the standards. 21. Combine an aliquot of the resuspended cDNA (∼100 ng) with the vector arms (1 μg), 10X DNA ligase buffer (1 μL), T4 DNA ligase (2.5 U), and sterile ddH2 O up to 10 μL. Incubate the reaction for 16 h (overnight) at 10°C. 22. The ligation is packaged into phage using a commercially available packaging kit, compatible with the phagemid vector being used, according to the manufacturer’s instructions (in this example, GigapackIII XL from Stratagene would be used). Experimental DNA is combined with the packaging extract, mixed and incubated for a set amount of time, chloroform separated, and then the phage is ready to be titered.

3.3. Titration and Ampliﬁcation of the cDNA Library 1. From a stock vial, streak XL1-Blue cells onto an LB-ampicillin agar plate and incubate overnight at 37°C. The following day, prepare 25 mL of LB-ampicillin broth culture by inoculating a single colony into a flask and incubating on a shaker at 37°C until the OD600 = 1. 2. Centrifuge the XL1-Blue culture at 1000g for 10 min at 4°C. Resuspend the cell pellet in sterile 10 mM MgSO4 and adjust the volume until the OD600 is 0.5. 3. Prepare aliquots (100 μL each) of serial library dilutions (1, 1:10, 1:100, 1:1000, 1:10,000) in TSM buffer, mix each with an equal volume of freshly cultured XL1-Blue cells, and incubate at 37°C for 20 min.

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4. Mix the contents of each tube with NZY top agar (∼3 mL for each dilution; cooled to <50°C) and immediately spread on 10-cm plates. Allow plates to cool (until top agar has solidified), then invert and incubate at 37°C until plaques are observed to form. Count the number of plaques on each plate and determine library titer using the following formula: pfu/mL = (number of plaques × dilution factor)/volume of extract plated (see Note 7). 5. To amplify the cDNA library, prepare 10 NZY plates (24 × 24 cm) and an overnight LB-ampicillin broth culture of XL1-Blue cells (as described in steps 1 and 2 of this subheading). 6. Pipet XL-1 Blue cells (100 μL) into 10 sterile 1.5-mL tubes, add an aliquot of packaged phage mixture (containing ∼50,000 phage particles) to each tube, and incubate for 15 min at 37°C. 7. Combine the contents of each tube with 7 mL of molten NZY top agar and spread evenly onto each of the NZY plates. 8. Incubate the plates at 37°C overnight to allow plaque formation on the bacterial lawn. Following plaque formation, spread TSM buffer (10 mL) over the surface of each plate, followed by incubation at 4°C overnight with gentle shaking to release the phage from the solid media and into solution. 9. Pool the TSM media containing bacteriophage from the 10 plates and add chloroform to 5% v/v. Incubate this mixture at room temperature for 15 min, centrifuge at 500g for 10 min, then transfer the supernatant to a new sterile container. 10. Add DMSO to the supernatant (7% v/v final) and aliquot the amplified library into sterile 1.5-mL tubes and store at –80°C. The titer of each amplified library should be determined as described above.

3.4. Differential Screening of the cDNA Library 1. Prepare NZY plates (10 plates; 24 × 24 cm) as described in Subheadings 3.3.1–3.3.4. Adjust the amount of phage library used to infect the bacteria using the calculated titer, so each plate contains approx. 20,000–50,000 pfu. 2. Plaques will develop over the course of approx. 12 h and should be viewed regularly until they are round in shape and measure approx. 1 mm across. 3. Place a Hybond-N nylon membrane onto each plate containing the newly developed plaques for one min (adsorption time). Orientation marks should be made on the membrane and on the plate so that the correct plaques can be chosen when going back to select positive clones. 4. Soak each membrane in denaturation buffer for 1 min, neutralization buffer for 5 min, and rinse buffer for 0.5 min. 5. Allow membranes to air dry on filter paper and UV cross-link the phage DNA to membranes using an ultraviolet (UV) crosslinker. 6. Repeat steps 3–5 using a second membrane on each plate, with the adsorption time lengthened to 2 min. 7. Bake the plaque lifts in a gel dryer for 1 h at 80°C between two fresh pieces of filter paper.

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8. Screening requires the synthesis of two sets of probe. One set is synthesized using control mRNA as a template and a second set using mRNA from the stress or condition being examined. 9. Synthesize probe using a protocol similar to that used for first strand cDNA synthesis (Subheading 3.2.; materials listed in Subheading 2.2.). Aliquot poly(A)+ mRNA template (about 1 μg) into an autoclaved, DEPC-treated, 1.5-mL microfuge tube and heat at 65°C for 5 min. 10. Cold-snap the mRNA on ice, with subsequent addition of 10× reverse transcriptase buffer (5 μL), dNTPs (3 μL, without dCTP), oligo(dT) primer (3 μg), RNasin (5 μL) into each tube. Adjust the total volume up to 40 μL. Flick the tubes gently to mix, centrifuge briefly to collect contents at the bottom of the tube, and allow the primers to anneal to the RNA at room temperature for 10 min. 11. Add 5 μL of reverse transcriptase and 5 μL of [-32 P]dCTP (3000 Ci/mol) to each tube and incubate at 42°C for 1 h. Transfer the tubes to a water bath preset to 16°C. 12. To degrade the RNA, add RNase A (5 μL) and incubate at 37°C for 30 min. This will leave a radiolabeled first strand of cDNA, which is loaded onto a quick spin column and centrifuged at 400g for 3 min. The eluant (labeled cDNA) is used to probe membranes. 13. Plaque lift membranes are pre-hybridized in hybridization solution (enough to cover the membrane) for 30 min in a rotating hybridization tube at 55°C. 14. Boil the radiolabeled probe for 5 min (see Note 8) and place immediately on ice. Add probe directly to the existing hybridization solution (see Note 9) to a concentration of 1 × 106 cpm/mL. To allow for direction comparison, add identical levels of radioactivity (cpm) to paired blots (control vs. experimental). Hybridize blots overnight (∼16 h) with rotation at 45°C. 15. After hybridization, wash the plaque-lift membranes in 0.2× SSC, 0.1% w/v SDS at 55°C for 10 min. Repeat this wash step 3 additional times to reduce background radioactivity. If background levels remain high (as detected using a handheld Geiger counter), washes can be repeated as desired. 16. Place washed membranes radioactive side down on plastic wrap while still damp and seal tightly so that liquid does not leak out. Wrap the membranes a second time using a plastic wrap and smooth the surface using a paper towel (to remove wrinkles or air bubbles). Expose the blots to X-ray film (or a phosphoimager screen) for an appropriate length of time. Exposure time depends on the cpm of the specific probe bound to each membrane and should be adjusted accordingly. 17. Develop the X-ray film (or phosphoimager screen) and analyze the resulting pattern of spots on the autoradiograms. An example of a primary screen autoradiogram is shown in Fig. 1A. Each spot represents a plaque where probe has hybridized to cDNA. 18. Using the orientation markers, trace differentially expressed clones back to their corresponding phage plaques on the NZY plates and excise. A sterile Pasteur pipet is used to excise each section of agar, which is then placed into a 1.5-mL Eppendorf tube containing 500 μL of TSM buffer. If it is not possible to isolate a single clone (especially common during the primary screen) putatively positive “regions” can be excised.

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19. Add chloroform to each tube (to 5% w/v), vortex-mix for 30 s, and leave at room temperature for 30 min, allowing the phage to desorb from the agar into the buffer. 20. Centrifuge samples for 5 min at 3000g at 4°C and transfer the supernatants (containing isolated phage) to fresh Eppendorf tubes and store at 4°C. 21. Perform a spot titer for each putative clone or “region” to determine the approx. pfu/mL (as described in Subheading 3.3.). 22. Re-plate each putative positive clone or “region” on a separate circular (10 cm diameter) NZY plate at 50–100 pfu/plate. 23. Construct plaque lifts for this secondary screen using small, circular nylon membranes and follow the same steps outlined above for a primary screen (starting at step 3). Repeat differential screening in an identical manner (starting at step 8.). Select positive clones representing genes of interest (differentially expressed) by carefully examining the resulting autoradiograms. An example of a secondary screen autoradiogram is shown in Fig. 1B.

3.5. In vivo Excision of Clones 1. The procedure followed in this section depends upon the cDNA library vector used. Components must be purchased commercially (generally in the form of a kit from the same manufacturer that created the vector). A brief overview of the procedure used with the Uni-Zap cDNA libraries (ExAssist Helper Phage with SOLR cells; Stratagene, see ref. 6) is provided. 2. Prepare a stock of XL1-Blue cells as outlined in Subheading 3.3.1. Prepare a stock of SOLR cells using the same protocol as that for XL1-Blue cells. Cells are grown at 37°C in LB-ampicillin broth overnight (∼16 h) and are centrifuged at 3000g for 5 min. 3. Discard the supernatant and resuspend each bacterial cell pellet in 10 mM MgSO4 to an OD600 = 1.0. Store the SOLR cell pellet at 4°C until step 8. 4. Aliquot 200 μL of each phage stock into a separate Eppendorf tube, add an equal volume of XL1-Blue Cells, and 1 μL of helper phage. 5. Incubate the tubes at 37°C for 15 min. Prepare an equal number of 15-mL tubes containing 3–5 mL of LB-ampicillin broth. Following the incubation, transfer the contents of each Eppendorf tube into labeled 15-mL tubes. Incubate these tubes at 37°C with gentle shaking for approx. 3 h. 6. Centrifuge the tubes for 15 min at 400g to pellet cells and debris and transfer the supernatant, containing the intermediate filamentous phage, to a fresh tube and incubate at 70°C for 15 min. 7. Centrifuge the new tubes for 15 min at 3000g to pellet any remaining cells, and then transfer the supernatant containing the filamentous phage stock to fresh 15-mL tubes and store at 4°C. 8. Add filamentous phage stock (100 μL) representing each selected phagemid into labeled tubes. Resuspend SOLR cells with brief mixing and aliquot 200 μL into each tube and incubate at 37°C for 15 min.

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Fig. 1. (Continued)

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9. Pipette an aliquot of cells (100–200 μL) from each tube onto an LB-ampicillin plate and incubate at 37°C overnight. 10. Streak a single colony from each plate onto a fresh LB-ampicillin plate to create a master plate of SOLR cells containing the pBluescript plasmid of interest. Incubate these plates overnight at 37°C. 11. Inoculate a single colony from a master plate into 3 mL of LB-ampicillin broth. When the OD600 = 1.0, a glycerol stock is created by combining an aliquot of the log-phase bacterial culture with sterile glycerol (15% v/v final concentration). Each tube is mixed and kept at –4°C for 30 min, then stored at –80°C.

3.6. Miniprep Plasmid Isolation 1. A variety of kits are commercially available that provide a quick and clean plasmid isolation. Such kits are available from a number of companies including, but not limited to Nucleospin (BD Biosciences), Qiagen, and Invitrogen. A standard “non-kit” protocol is outlined here. 2. Select a single colony from a master plate and inoculate 3 mL of LB-ampicillin broth in a 15-mL tube. Incubate overnight (∼16 h) at 37°C with shaking. 3. Pipet 1.5 mL of the overnight culture into an Eppendorf tube and centrifuge at 3000g at 4°C for 10 min. Aspirate and discard the supernatant. 4. Resuspend the pellet in 400 μL of pre-lysis buffer, transfer to a sterile Eppendorf tube, add RNase A (5 μL) and vortex-mix. 5. Add an equal volume of freshly diluted alkaline lysis buffer and keep tube at room temperature for 5 min. 6. Add an equal volume of neutralization buffer and vortex-mix the tube until the white precipitate (protein, genomic DNA, and cell debris) is broken up. Keep on ice for 5 min and then centrifuge in a microcentrifuge at maximum speed for 10 min at 4°C.

Fig. 1. Differential screening of a cDNA library. (A) Primary differential screen of duplicate plaque lift membranes using [32 P]dCTP labelled cDNA synthesized from mRNA isolated from the hepatopancreas of the marine snail, Littorina littorea. Normoxic: mRNA isolated from animals kept at 4°C under normal aerated conditions. Anoxic: mRNA isolated from animals kept under a nitrogen gas atmosphere for 1 h, 12 h, and 24 h (equal amounts of mRNA from each time-point combined). (B) Secondary differential screen of duplicate plaque lift membranes using the same probe as in (A) to confirm putative anoxia-responsive clones. (C) In vivo excision of plasmids from phage stock, with subsequent plasmid purification from bacteria, allowed insert isolation from putative anoxia-induced clones. Lane 1 shows a particular clone of interest, encoding snail anoxia-responsive protein 2 (sarp-2); various inserts of other nonspecific genes are in the subsequent lanes. Positive clones should be confirmed as differentially expressed with Northern blots and then further characterized as outlined in Fig. 2.

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7. Transfer the supernatant to a fresh 1.5-mL Eppendorf tube and add an equivalent volume of isopropanol. Invert tube repeatedly to mix and incubate for 10 min at room temperature. 8. Centrifuge the tube at maximum speed in a microcentrifuge for 10 min at room temperature. Aspirate and discard the supernatant and wash the pellet (DNA plasmid) with 70% ethanol. Recentrifuge as described previously for 5 min, Aspirate ethanol and allow the pellet to air-dry for 10 min at room temperature. 9. Resuspend the plasmid DNA in an appropriate volume of sterile ddH2 O (generally start with 20 μL) and quantify using spectrophotometry (1 OD = 50 μg/mL of double- stranded DNA). Plasmid is stored at 4°C (short term) or –20°C (long term) until further use.

3.7. Insert Isolation 1. Digest the plasmid DNA with appropriate restriction enzymes (in this case, EcoRI and XhoI). 2. Combine an aliquot of plasmid DNA (5 μL) with 0.5 μL of each enzyme, 1 μL of 10× reaction buffer, and adjust the final volume up to 10 μL with sterile ddH2 O. Incubate digest at 37°C for at least 1 h. 3. Stop the restriction digest by heating at 94°C for 5 min and then adding 0.1 volumes of DNA loading dye. 4. Load samples onto a 1% TAE agarose gel and separate the restriction fragments by running the gel at 90 V in 1× TAE buffer. An appropriately sized DNA ladder should be used as a molecular weight standard, allowing the size of the insert to be estimated. An example of digested inserts separated using agarose gel electrophoresis is shown in Fig. 1C. 5. Carefully slice each insert band out of the gel using a sterile razor blade and place each into a labeled Eppendorf tube. To isolate the DNA from the gel agarose, a number of options are available. Commercial kits using filtered columns or glass milk are available, however a quick and easy protocol is outlined here. 6. Place the gel slices at –80°C until frozen (∼15 min). The collar of a 1-mL filtered pipet tip is removed using a razor blade and the bottom half of the tip is fixed inside it. This apparatus is then placed firmly into the top of a labeled Eppendorf tube. Each frozen gel slice is transferred quickly into the opening of the tip on top of the filter and the tube is placed into a microcentrifuge set to spin at 10,000g for 3 min. Remove the tubes from the centrifuge and discard the filter apparatus. Insert DNA is present in the eluant and can be stored at 4°C (short term) or –20°C (long term).

3.8. Synthesis of Labeled Probe 1. Transfer an aliquot of the insert DNA (12 μg) into a sterile 1.5-mL Eppendorf tube and adjust the final volume of 8 μL. Denature the DNA by placing at 94°C for 3 min, centrifuge briefly to collect contents at the bottom of the tube and place immediately on ice. Add the following to the tube on ice: dNTPs (ATP, TTP, GTP; final concentration = 1 mM each), 1 μL random primer (100 mM d(N)6 ;

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NEB), 2 μL of 10× random primer buffer, 2.5 U of Klenow fragment of DNA polymerase I (NEB), 2.5 μL of [-32 P]dCTP (3000 Ci/mol; Amersham). 2. Gently mix the tube and place in a heating block at 37°C for 45 min. Load the labeled reaction onto a quick spin column and centrifuge at 400g for 3 min. The eluted radiolabeled probe is used for Northern blotting.

3.9. Northern Blotting 1. Prepare a 1.25% agarose formaldehyde denaturing gel and submerse it in enough 1× MOPS buffer to cover the wells. Pre-run the gel for 15 min (as RNA samples are being prepared). 2. Aliquot the appropriate volume of total RNA, containing between 10 and 20 μg, into labeled tubes on ice. Add an equal volume of RNA sample buffer into each tube. 3. Incubate samples at 55°C for 10 min and place immediately on ice. Add the appropriate amount of RNA loading buffer (final concentration = 1×) to each sample. 4. Mix the RNA samples gently and briefly centrifuge tubes to collect the entire sample at the bottom of each Eppendorf tube. 5. Load the entire contents of each tube into the wells of the gel and record sample order for reference. 6. Perform gel electrophoresis at 100 V until the loading dye front reaches the end of the gel. 7. Place the gel on plastic wrap and visualize the RNA using a UV light. A photograph (digital or Polaroid) should be used to maintain a photographic record. The ribosomal RNA (rRNA) bands are used as an indicator of degradation and as a rough guide to ensure equal loading of the wells. Fig. 2A (lower panel) provides an example of ethidium bromide stained ribosomal RNA visualized with UV light. 8. The separated RNA will be transferred from the gel onto a 0.2-μm nylon membrane by the neutral transfer method. Soak the gel in 10× SSC for 10–15 min to remove excess formaldehyde and ethidium bromide. 9. While the gel is soaking, set up the transfer apparatus. Fold a large square of plastic wrap over onto itself (2 ft × 3 ft) and place on the benchtop. 10. Center a square piece of Plexiglas or glass sequencing plate (larger than the gel being transferred) on the plastic wrap. The wrap should extend 3–4 in. beyond the edges of the plate. 11. Cut two pieces of filter paper (used as a wick) such that they are approx. 1–2 inches longer than the plate on all sides. 12. Prewet the plate with 10× SSC and place each filter paper sequentially over the plate. Remove wrinkles or air bubbles by smoothing each layer with a sterile pipet. 13. Orient the gel face down on the second piece of filter paper, directly in the center. 14. Cut a membrane to the exact size of the gel. Briefly rinse the membrane in sterile water, then rinse in 10× SSC, and place it over the entire gel. Use a pipet to smooth the membrane over the gel, removing any wrinkles or air bubbles. 15. Cut two pieces of filter paper (the same size as the gel and membrane) and place each sequentially on top of the membrane. Prewet the filter paper in 10× SSC and roll smooth using the pipet.

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Fig. 2. Characterization of the novel gene from L. littorea, sarp-2. (A) Following insert isolation from a clone designated sarp-2 (as described in Fig. 1), expression of the sarp-2 transcript was monitored over a time course of anoxia exposure by Northern blotting. The expression of sarp-2 was normalized by examining the expression of a control gene (-tubulin) on the same blot, as well as by examining the intensities of the ribosomal RNA band on the gel before transfer. (B) Sequencing of the insert sarp-2 showed a 550 nucleotide sequence containing the full open reading frame for a 131 amino acid protein. Multiple common protein domains and motifs were identified in the sarp-2 mRNA and deduced SARP-2 protein sequences using the applications outlined in Subheading 3.11. and selected results are illustrated here.

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16. Place a piece of plastic wrap (the same size as that on the benchtop) over the entire transfer apparatus. Using a sharp razor blade, cut a “window” around the gel/filter paper. Remove this square of plastic wrap and place a stack of paper towels (5 in. high) on top of the filter paper. Add 10× SSC (10 mL) into each end of the transfer apparatus (between the inner and outer layers) and fold over the plastic wrap to seal. Place a weight (250–500 g) on top of the paper towels and allow the transfer to occur overnight (see Note 10). 17. Following transfer, cross-link the RNA to the membrane following the same protocol described in Subheading 3.4.5., or the membrane can be soaked for 5 min in 0.05 N NaOH and then rinsed in sterile ddH2 O.

3.10. Probing Northern blots 1. Probing Northern blots is similar to probing plaque lift membranes. Prehybridize Northern blots at 55°C using hybridization solution (enough to cover the surface of the blot) in a hybridization oven for a minimum of 30 min. 2. Heat radiolabeled cDNA probe at 94°C for 5 min, cold-snap on ice, then add directly to the hybridization solution. 3. Incubate Northern blots with probe at 45°C for at least 16 h (overnight). Following hybridization, wash blots with Northern blot washing solution at 55°C for 10 min. Repeat a total of 4 times. 4. If radioactive background appears high when scanned with a handheld Geiger counter, wash steps can be repeated as desired. 5. Wrap Northern blots in plastic wrap, expose to X-ray film or a phosphoimager screen for the appropriate length of time, and develop. 6. Perform densitometric analysis on the resulting autoradiogram using appropriate imaging software (Example, Imagequant (GE Healthcare) or QuantityOne (BioRad)). An example of a Northern blot is shown in Fig. 2A (upper panel). 7. As needed, probe can be removed from a blot by boiling a solution of ddH2 O and 0.5% SDS and pouring it on top of the blots in a wash container. Repeat until radioactive counts reach (or are below) background levels. Rinse stripped blots in sterile water (to remove residual SDS) and re-probe as desired. Reprobing for a constitutively expressed or “housekeeping” gene is used to normalize the relative levels of your differentially expressed gene against a gene whose expression does not change under the experimental condition being examined. The Northern blot in Fig. 2A was stripped and reprobed with -tubulin to control for loading, as illustrated in the Fig. 2A (middle panel). 8. Additional experimental protocols can and should be performed to further characterize a novel gene. The promoter of the gene can be examined to determine if stress or condition responsive elements are present which may play a role in gene expression. Transcriptional analysis can be examined with nuclear runoff assays to determine if a gene is transcriptionally regulated or if the mRNA is stabilized. The gene can be expressed as a protein and basic biochemistry can be carried out to determine a possible role or function in vitro. The protein can also be used as an antigen for producing an antibody that can be used for Western

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Larade and Storey blotting and subcellular localization experiments. A knockdown (using siRNA) or knockout (using genetic manipulation) of a gene can be performed to determine the effects and/or phenotype resulting from loss of the novel gene. These methods and techniques are outside the scope of this chapter and they represent downstream experiments from the “starting point” outlined here. Refined protocols can be found in other volumes of the Methods in Molecular Biology series, as well as in classic laboratory manuals such as Molecular Cloning: A Laboratory Manual (7). Figure 3 provides a basic view of experiments used to examine the novel gene, kvn, discovered and characterized by the authors.

3.11. Clone Analysis Differentially expressed clones should be bidirectionally sequenced using standard methods (either in the laboratory or by a DNA/Genomics facility). Subsequent sequence analysis is performed using commercially available computer software and a wide array of analysis tools available online. An index of some excellent bioinformatics programs and applications available to the scientific public for analysis of cDNA and putative amino acid sequences is listed in this subheading. As a first step, DNA sequences should be copied and pasted into NCBI’s BLAST server for comparison to existing and annotated sequences. Genes sharing sequence identity or similarity to existing expressed sequence tags (ESTs) will be identified with an E-value. The E-value represents the probability of each alignment occurring due to chance, where a value <e−5 signifies that an alignment is not due to error. Novel or unknown genes will often come back with no determinable match, although some conserved domains and/or motifs may be identified and provide a short sequence match. 3.11.1. National Center for Biotechnology Information (NCBI) http://www.ncbi.nlm.nih.gov/ 1. Basic Local Alignment Search Tool (BLAST): http://www.ncbi.nlm.nih.gov/ BLAST/ BLAST first lists four letter “words” of residues in one sequence; then the words are expanded to segments (profile based). The programs used for nucleotides are blastx (compares a nucleotide query sequence translated in all reading frames against a protein sequence database) and tblastx (compares the six-frame translations of a nucleotide query sequence against the six-frame translations of a nucleotide sequence database), whereas blastp (compares an amino acid query sequence against a protein sequence database) is used for protein analysis. 2. ORF (open reading frame) finder: http://www.ncbi.nlm.nih.gov/gorf/gorf.html ORF finder allows identification of potential coding regions in all 6 reading frames of a nucleotide sequence. Selected ORFs can then be transferred directly into

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Fig. 3. Flowchart outlining the characterization of a novel gene. Starting with genomic DNA, the transcription of a gene can be measured using a nuclear runoff assay (top left panel). The gene examined here (kvn) is actively transcribed in nuclei isolated from the hepatopancreas of L. littorea kept under anoxic conditions for 48 h. This gene has been characterized previously (4) and contains all of the features common to eukaryotic genes that are identifiable in mRNA.

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3.11.2. Gene Translation and Characterization of a Protein Sequence Determining the function of novel genes is a challenge currently facing many of the large genome projects. An mRNA sequence contains much information, but it is traditionally the functioning protein that performs the important task. In order to anticipate the function of a particular gene, its sequence is usually translated into a predicted protein, which is then analyzed for structural homology and conserved regions (known as domains or motifs) that have been identified in proteins having known functions. Many times a cDNA clone will represent a known homologue. The genes/proteins represented by these clones can be assimilated into an existing scheme, since they have often been characterized in other systems, organs or tissues (as well as during various stresses or conditions) and perform a known role or function. However, if the clone of interest does not have any known homologues, it must be further analyzed and characterized using the tools available. Some tools for analyzing a novel gene/protein include the following: PredictProtein: http://www.embl-heidelberg.de/predictprotein/predictprotein.html The translated protein sequence of interest is queried against characterized sequences using the PredictProtein server, which retrieves similar sequences in the database and predicts aspects of protein structure. Protein Sequence Analysis (PSA): http://bmerc-www.bu.edu/psa/request.htm PSA predicts protein secondary and tertiary structure based on amino acid sequence. SOSUI: http://sosui.proteome.bio.tuat.ac.jp/sosui_submit.html This tool predicts secondary structure of membrane proteins from an input protein sequence by examining the physicochemical properties (such as hydrophobicity and charges) of amino acid sequences.

Fig. 3. (Continued) Expression of kvn was monitored over a time course of anoxia exposure as shown by Northern blotting (middle panel). Northern blots were normalized by examining the expression of a control gene (-tubulin) on the same blot, as well as the intensity of ribosomal RNA on the gel before transfer. Expression of the KVN protein was measured over a time course of anoxia exposure as shown by Western blots (bottom left panel). Western blots were standardized by re-probing the blot with an antibody for a control protein (-actin), as well as comparing the protein levels in each sample lane after the membrane is stained with a reversible Ponceau S stain.

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PROSITE: http://www.expasy.ch/prosite/ PROSITE is a database of conserved protein motifs. The query output provides matches for biologically significant patterns and profiles based on the primary sequence of the protein. PSORT: http://psort.nibb.ac.jp/ After a gene is translated into a protein, the amino acid sequence often contains a region that represents a “transport” or “localization” signal. This signal determines the cellular localization of the protein and may help in determining its function. PSORT compares an input sequence to known consensus sequences to determine if a sorting signal is present. SignalP: http://www.cbs.dtu.dk/services/SignalP/ The SignalP server predicts the presence and location of signal peptide cleavage sites in input amino acid sequences. Submitting DNA Sequences to NCBI GenBank http://www.ncbi.nlm.nih.gov/Genbank/submit.html Prior to publication of research that has retrieved and characterized DNA sequences, the sequence must be annotated and submitted to a sequence database. The submission may include such information as coding regions, start and stop codons, or any information provided by the applications listed here. Using an internet browser, the new sequence is submitted to Genbank through Bank It (web based tool) or Sequin (stand alone submission tool).

4. Notes 1. Total RNA sample quality can also be assessed via denaturing agarose gels (see Subheading 3.9.). Both the integrity of the ribosomal RNA (rRNA) and the distribution of mRNA in each lane are indicators of degradation. The 28S rRNA band should be approximately twice as intense as the 18S rRNA (although with many invertebrates, only a single rRNA band is observed due to a cryptic nick in the 28S rRNA, resulting in a single rRNA band migrating at the approximate size of the 18S rRNA). There should be an even distribution of mRNA throughout the lane (as indicated by a light pink ethidium bromide smear when viewed using UV light). 2. The final reaction volume in the cDNA library synthesis step (3.2.1) will be 50 μL, so the RNA must be resuspended in the appropriate volume of ddH2 O. If the RNA is not concentrated enough, the sample can be concentrated using a SpeedVac centrifuge (preferred method) or reprecipitated using isopropanol as before (some loss will occur). 3. Remove 5 μL of the first-strand synthesis reaction and aliquot it into a new tube. Add 0.25 μL of [-32 P]dCTP and incubate with the first-strand synthesis reactions.

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4.

5. 6.

7.

Larade and Storey Place the radioactive first-strand synthesis control reaction at –20°C until ready to resolve by electrophoresis on an alkaline agarose gel. (See Note 4.) Remove an aliquot of the second strand cDNA synthesis reaction (1 μL) and analyze it with the aliquot saved from the first strand reaction (see Note 3) using an alkaline agarose gel. These radiolabeled “control” reactions allow determination of the size and quality of synthesized cDNA. Gel electrophoresis is performed similar to the procedure outlined in Subheading 3.7 or 3.9. Melt agarose (3.75 g) in ddH2 O (270 mL) containing ethidium bromide (1 μg/mL) and cool to 60°C. Add 0.1 volumes of 10× alkaline agarose electrophoresis buffer (500 mM NaOH, 10 mM EDTA, pH 8.0), swirl without introducing bubbles, pour gel, and allow it to polymerize. Submerse the gel in enough 1× alkaline agarose electrophoresis buffer to cover the wells and prerun for 5 min. To each of the samples (first- and second-strand reactions, 1 μL), add 10 μL of 1× alkaline agarose electrophoresis buffer and 2 μL of alkaline loading buffer (300 mM NaOH, 6 mM EDTA, 18% Ficoll 400, 0.15% bromocresol green, 0.25% xylene cyanol FF). If possible, electrophorese gel in a cold room (or at low voltage) to prevent overheating. Following electrophoresis, fix the gel in 7% trichloroacetic acid (w/v), dry in a gel dryer, and perform autoradiography. To enhance precipitation, add NaCl to a final concentration of 25 mM. Depending on how mRNA was originally prepared, the quality may be suitable to create a library without size fractionation. If transcripts above or below a certain size are desired, cDNA fractionation can be performed. One method for this is briefly outlined here. A Sepharose drip column (∼1 mL) can be used to separate cDNA by size; longer cDNA sequences will elute off the column first followed by progressively shorter sequences. Columns can be made in the lab using Sepharose having the desired molecular weight cutoff (often S500), or can be purchased from a company such as Invitrogen). Equilibrate column with loading buffer (10 mM Tris-HCl, pH 7.5, 0.1 mM EDTA, 25 mM NaCl) by loading 1 mL onto the column and allowing it to drain completely. Repeat 3–5 times. Load sample cDNA onto the column (in a volume of ∼100 μL of loading buffer) and allow it to enter the Sepharose. Aliquot 100 μL of loading buffer onto the column, allow it to enter the Sepharose and collect the eluant into the first tube. Continue adding 100-μL aliquots of loading buffer on top of the column (allow each to enter the Sepharose prior to adding subsequent aliquots). Collect the eluant in approx. 50 μL fractions. A few microliters of a sample dye containing glycerol (50%) and bromophenol blue (50%) can be added after the sample is loaded to track the progress of the cDNA elution. Each collected fraction can be analyzed by loading 5 μL on a 5–6% standard nondenaturing acrylamide gel, followed by autoradiography (wet or dry gel), to determine cDNA size in each. Combine the fractions containing the desired lengths of cDNA into a single tube and then process as described in Subheading 3.2.5–3.2.8. Resuspend the dried pellet in the appropriate volume of ddH2 O and proceed to Subheading 3.2.15. The Uni-ZAP vector contains -galactosidase within the multiple cloning site, allowing a “blue-white” color assay to be performed to determine the ratio of

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recombinants (white plaques) to nonrecombinants (blue plaques) in the cDNA library. Transfer an aliquot of the final packaged reaction (1 μL) into two Eppendorf tubes. Label the first tube 1 (undiluted). Add 9 μL of ddH2 O to the second tube, then aliquot a single microliter of this 1:10 dilution into a fresh Eppendorf tube labeled 2 (1:10 dilution). Add an aliquot of XL1-Blue cells (100 μL; prepared as described in Subheading 3.3.1) into each of the labelled tubes and incubate at 37°C for 15 min. Combine IPTG (15 μL of a 0.5 M stock) and X-gal (50 μL of a 250 mg/mL stock) with 3 mL of molten NZY top agar. Cool the agar to <50°C, add the infected XL1-Blue cells, spread the mixture on top of a standard NZY plate, and allow top agar to solidify. Incubate the plate at 37°C overnight (∼16 h) until colored plaques develop. Examine both plates (one of which should contain ∼250–500 plaques) and count blue vs. white plaques to determine the background of the library. 8. Sheared salmon sperm DNA (Sigma-Aldrich) can also be included as a nonspecific, unlabelled DNA source to reduce background binding to membranes. Add sheared salmon sperm DNA (0.2 mg DNA/mL hybridization solution) to the Eppendorf tube containing probe and follow the protocol from Subheading 3.4.14. This note is also applicable to Subheading 3.10. 9. When adding probe to the hybridization tube, take care not to “spray” the membrane. If concentrated probe contacts the membrane, the result can be a “hot spot” which may obscure important regions of the blot. Probe should be dispensed directly into hybridization solution (using longer pipet tips) or added to an aliquot of hybridization solution (∼1 mL), which is carefully transferred to the tube drop by drop. 10. A layer of plastic wrap can be placed around the stack to prevent a potential “leak” (and resulting short circuit) in the transfer setup between the paper towel and the wicks. This extra layer ensures that buffer must flow through the gel and membrane to reach the paper towel, improving Northern blot transfer.

Acknowledgments We express special thanks to Jan Storey for critical reading of the manuscript, along with offering helpful advice and providing excellent editing.

References 1. Larade, K. and Storey, K. B. (2002) A profile of the metabolic responses to anoxia in marine invertebrates, in Cell and Molecular Responses to Stress, Vol. 3: Sensing, Signaling and Cell Adaptation (Storey, K. B. and Storey, J. M., eds.), Elsevier Press, Amsterdam, pp. 27–46. 2. Storey, K. B. and Storey, J. M. (2004) Physiology, biochemistry and molecular biology of vertebrate freeze tolerance: the wood frog, in Life in the Frozen State (Benson, E., Fuller, B., and Lane, N., eds.) CRC Press, Boca Raton, pp. 243–274.

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3. Larade K. and Storey K. B. (2004) Anoxia-induced transcriptional up-regulation of sarp-19: cloning and characterization of a novel EF-hand containing gene expressed in hepatopancreas of Littorina littorea. Biochem. Cell. Biol. 82, 285–293. 4. Larade K. and Storey K. B. (2002) Characterization of a novel gene up-regulated during anoxia exposure in the marine snail, Littorina littorea. Gene 283, 145–154. 5. Stratagene Instruction Manual Revision #073013b. 2003. For: cDNA Synthesis Kit, ZAP-cDNA Synthesis Kit, and ZAP-cDNA Gigapack III Gold Cloning Kit. 6. Stratagene Instruction Manual Revision no. 073002f. 2003. For: ExAssist Interference-Resistant Helper Phage with SOLR Strain. 7. Sambrook, J., Russell, D.W. (2001) Molecular Cloning: A Laboratory Manual, 3rd lab edition. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY.

6 Comparative Molecular Physiological Genomics Heterologous Probing of cDNA Arrays Sean F. Eddy and Kenneth B. Storey

Summary The use of DNA microarrays has gained wider acceptance as a standard tool for molecular biology studies over the past decade. In particular, biomedical studies embraced this technology as soon as arrays were produced for the common laboratory species. Slower to develop, however, has been the use of microarray screening with non-standard animal models, even though these species present fascinating physiological phenomena for study. The very high cost and huge amount of work involved in developing and producing a DNA array or microarray for a new species is prohibitive for most researchers working in comparative biology. The alternative is to explore the use of heterologous array hybridization, screening for stress-induced gene expression in one species using an array developed for another species. This chapter provides a comprehensive review of the current literature on heterologous DNA array hybridization and explores the factors that must be taken into account when performing heterologous microarray analysis on nonstandard species. Changes in methodology (e.g. hybridization conditions, stringency of washing) to optimize the percent cross reaction, the potential for false positives and false negatives to occur, and techniques for downstream analysis and confirmation of array data are all discussed. Examples of cross-hybridization using human microarrays are discussed using phylogenetically diverse species ranging from ground squirrels to frogs to snails. As with any new technology, the willingness to grasp cross-species analysis has been slow but the future looks bright for heterologous DNA hybridization and microarray analysis now that the initial hurdles have been overcome. Key Words: Comparative genomics; cross-species DNA array hybridization; mammalian hibernation; semiquantitative PCR.

From: Methods in Molecular Biology, vol. 410: Environmental Genomics Edited by: C. Cristofre Martin © Humana Press, Totowa, NJ

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1. Introduction The vast majority of research in biochemistry and molecular biology is centered on a very low number of model species; human, mouse and rat are the main mammalian models whereas fruit flies (Drosophila melanogaster) and nematodes (Caenorhabditis elegans) are primary invertebrate models. However, the genetic information programmed within numerous other species offers researchers unique insights into the molecular mechanisms that underlie organismal responses to numerous stresses, conditions and diseases. For example, study of appropriate animal model systems is the only way to determine how cells and organisms have evolved to endure environmental stresses including, but not limited to, oxygen deprivation, extremes of hot or cold temperature, freezing, and high or low salinity. Regulating these biological processes requires a concerted effort put forth by multiple signaling pathways eventually targeting specific genes to activate or repress expression in order to maintain sustained cellular function during stress. The development of DNA array screening technology has given researchers an immensely powerful new tool with which to study cellular responses to stress. Array screening can take a snapshot of the total gene expression patterns within a cell at any given time point providing insights into the responses of individual genes as well as an integrated view of the responses of functional units (e.g., metabolic pathways, signal transduction cascades, etc.). Since the advent in the 1970s of the Southern blot that used DNA bound to nitrocellulose or nylon membranes, researchers have searched for ways to immobilize DNA on smaller and smaller scales and platforms. cDNA arrays and microarrays are the latest nucleic acid immobilization tool for molecular biologists. Since their introduction their physical size has decreased dramatically at the same time as the information they possess has greatly increased. The field has grown immensely since the introduction of DNA arrays in the mid-1990s (1–5), with researchers applying DNA array screening to nearly all areas of biology and medicine. Their initial use was in studying the mRNA expression profiles, or transcriptomes, of cells, tissues or organisms to gain insight into the changes in gene expression between two or more metabolic states, however, recent advances made using microarrays include identifying and genotyping numerous pathogens within clinical samples, also called metagenomics (6–8). There are two primary methods for producing DNA arrays. These are largely the same today as they were a decade ago except that the cost to produce arrays has dropped significantly and the arrays themselves have become higher density and more complete, often covering entire genomes. The two methods are: 1. Photolithography; the synthesis of specific oligonucleotides on a support medium such as glass. Developed by Affymetrix, this method invokes light-directed oligonucleotide synthesis directly onto a glass slide or support (3).

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2. Spotting cDNAs onto a glass or nylon support. The method, originally developed by Patrick Brown’s lab at Stanford University, involves amplification and purification of cDNAs that are then spotted onto the support (1). Advances in robotics now allow production of high density cDNA arrays and microarrays.

Comparative molecular biology and physiology, particularly areas dealing with non-traditional model organisms, has only begun to grasp the power of DNA array technology, largely due to a lack of experimental platforms with which to proceed. Prevailing thought in the field has been that a homologous cDNA platform must be used in order to generate useful data. However, this is not the case, especially considering that large mixed populations of bacterial and viral pathogens can now be genotyped and identified on a single array (6–8). If this is the case, then heterologous cDNA array screening between closely related organisms should not pose a problem. The production of a DNA array for each and every new species is expensive and cost-prohibitive for the amount of information that is produced so studies have been slow to expand beyond the traditional model species. Recently, however, arrays have become available for more and more model species, representing widely differing groups of organisms and, thereby, broadening the range of studies for which homologous array screening can be used. For example, Affymetrix currently has platforms for a number of nonmammalian animals (the frog Xenopus laevis, the fruit fly D. melanogaster, the nematode C. elegans, zebrafish Danio rerio), plants (Arabidopsis thaliana, barley, grapes, maize, soybeans, tomato), and prokaryotes (Bacillus subtilis, Escherichia coli, Pseudomonas aeruginosa, Staphylococcus aureus). Thus, there is a large existing platform of pre-fabricated arrays that many comparative researchers can exploit for cross-species heterologous analysis. The diversity of experimental animal models in use by researchers around the world is huge. Currently, it is simply not possible to construct and screen homologous DNA arrays for each and every species of interest at a cost that is reasonable for most researchers. Instead, approaches aimed at exploiting existing microarray technology (using microarrays fabricated for model organisms or other closely related organisms) need to be embraced. Heterologous probing, screening cDNA from one organism using an array produced from another species, has significant potential as a gene discovery tool. Although cross-species hybridization will never be 100%, with appropriate optimization of conditions, heterologous hybridization can allow the analysis of expression responses by thousands of genes. Thus, heterologous DNA hybridization is not only feasible and valid on scientific principles, but provides comparative biologists with a means to study the diversity of gene regulation without the prohibitive costs of having to create a microarray platform for each and every species under scientific study. To understand

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the range of opportunities provided by this approach, we must first look at similarities at the genetic level between nonmodel organisms and the model organisms for which array platforms have already been fabricated and then decide whether or not such an undertaking is feasible. The success of cross-species array hybridization depends largely on the degree of identity between genes from nontraditional model species and homologous genes in model organisms. Detailed analysis of orthologous human and rodent gene sequences has shown that, on average, protein coding regions share 85% nucleic acid identity, 5 untranslated regions are approx 70% identical, and 3 untranslated regions are approx 71% identical (9,10). When this analysis was extended to compare mammalian (human and mouse) to nematode (C. elegans) genes, 44% of human/rodent orthologs were found to have nematode counterparts that had a mean approx. 50% identity at the nucleotide level (11). This strong identity between orthologous gene sequences across the animal kingdom give us a good basis for predicting that heterologous cDNA array probing will succeed. Our own studies have cloned and sequenced a variety of specific genes from hibernating small mammals including ground squirrels (Spermophilus tridecemlineatus.) and bats (Myotis lucifugus) and the results have shown that the degree of identity between gene orthologs of humans and hibernators is virtually the same as that noted above for humans and rodents. For example, analysis of the open reading frame sequences of the heart isoform of fatty acid binding protein (h-fabp) from S. tridecemlineatus, M. lucifugus, human, mouse and rat showed 91% nucleotide identity between the five species (12–14). Notably, fatty acid binding proteins are of key importance to hibernation as they facilitate the intracellular transport of fatty acids from the plasma membrane to the mitochondria, lipids being the primary fuel used during hibernation. Similar results have been obtained for the full or partial cDNA sequences of several other genes cloned from hibernators including atpase6/8, coxII, hif1, mlc2(v), nd2, pag, pgc-1 and ppar- (12,14–19). Other researchers have also shown a high degree of nucleotide identity among genes from hibernating mammals compared with their human, mouse or rat counterparts (20,21). Gene cloning from other nonmodel species also supports the suggestion that the sequences of many genes are highly conserved across a broad range of vertebrate and invertebrate species. We have recently sequenced a number of genes from the wood frog, Rana sylvatica, a species that survives the freezing of its body fluids over the winter months. The genes for fibrinogen and fibrinogen are up-regulated in the liver of freezing frogs and sequencing of the partial cDNA for wood frog fibrinogen showed that it was 67 % identical with human fibrinogen over the same region (22). The wood frog aat gene encoding the ATP/ADP translocase showed 70% nucleic acid

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identity compared with the human gene (23) and other cloned genes from wood frogs showed remarkably high identity with their mammalian counterparts including the inorganic phosphate carrier, pic, and the acidic ribosomal protein, P0 (24,25). Mitochondrial genes cloned from the turtle, Chrysemys picta marginata, also displayed high homology to the corresponding mammalian genes (26). The genes for ribosomal protein L26 and ferritin, cloned from the marine snail, Littorina littorea (27–29) provide even more evidence that genetic similarity between non-model and model species can be utilized for heterologous cDNA array screening. Lastly, a muscle-LIM gene cloned from the cold hardy gall insect, Epiblema scudderiana, shows gene homology of 60% over the entire mRNA and 75% identity within the open reading frame (ORF) compared to that of the appropriate model species, in this case D. melanogaster (30). Thus, our data on gene homologies strongly suggest that comparative DNA array hybridization will work. Actual array screening results from our lab and others documents this, as discussed below. Given the demonstrated high degree of nucleotide identity in comparisons of cloned hibernator genes with other mammals, the capacity to utilize crossspecies array hybridization as a means of gene discovery in hibernation has never been better. The same applies to many other heterologous pairings. Thus, cross-species array hybridization has become a rational approach to studying nontraditional model animals as a means of deciphering the complex modifications in the transcriptome under different stresses. In addition to our studies, many experiments have been performed using other species as well, harnessing the high-throughput capabilities of cDNA arrays. To this end, we have carried out gene expression studies on animal species using heterologous probing with either Clontech ATLAS™ rat cDNA arrays and human 19K cDNA arrays (University Health Network, Toronto). These studies have been highly effective for identifying: a) previously unrecognized target genes that participate in environmental stress tolerance, and b) tracing the cellular signaling pathways that are active in stress response. Our first studies used ATLAS™ nylon arrays containing rat cDNAs to assess changes in gene expression during mammalian hibernation in the ground squirrel, S. tridecemlineatus. Cross-hybridization was very high between the two rodent species and the results showed, among others, a clear up-regulation of a-fabp (the adipose isoform of fatty acid binding protein) in brown adipose tissue (14) and a striking 50% suppression of genes encoding numerous ribosomal proteins, including S12, L21, and L36a, in skeletal muscle during hibernation (31). Subsequent work from our lab has shown that mammalian (bat or ground squirrel) hybridizations to cDNA arrays can give up to 85–90% hybridization (31), lower vertebrates (wood frogs) on human arrays will give 60–80% hybridization (32) and the marine snail Littorina littorea will hybridize to 18% of genes on 19K human

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arrays (33) using low stringency hybridizations. Such cDNA array studies with the heart of wood frogs have identified a number of genes that are putatively up-regulated in the organ during freezing. Interestingly, three genes designated as up-regulated on the cDNA microarray experiments were previously identified as freeze-responsive genes by other means, specifically the ATP/ADP translocase, glucose transporters and glucose-6-phosphate dehydrogenase (22,34,35). Other labs have also made good use of mammalian cross species hybridizations proving that this method of comparative analysis is sound (14,31,36–44). Indeed, the study by Rinaudo and Gerin (40) showed that heterologous screening of a 4000 human gene microarray with woodchuck liver cDNA resulted in no false positives and only 29 false negatives (0.7%); a false positive was defined as a gene that shows putative up- or down-regulation on the array but no differential expression on downstream analysis whereas a false negative was a gene that does not cross react on the array. The false negatives appeared to occur because of high genetic distance between humans and woodchucks for these particular genes. The most promising aspect of this work was the complete lack of false positives, suggesting that rigorous heterologous microarray analysis yields real useable data. Analyzing hybridization patterns of woodchuck liver samples in comparison to human liver samples on nylon filter arrays, it was found that human liver samples hybridized to 20–60% of array probes (depending on temperature and salt concentrations during washing), whereas woodchuck liver samples hybridized to nearly the same degree, showing 18–53% hybridization to array probes under identical conditions (40). Another study used human cDNA arrays containing 4400 genes to study UV-induced melanoma in the opossum, Monodelphis domestica. This heterologous probing worked very well and the results showed that 79 genes were up-regulated by UV-treatment whereas 28 were down-regulated (43). It was noted that the majority of published gene sequences for this evolutionarily distant marsupial mammal showed 70–80% identity with the corresponding human genes. A study on a porcine model of vascular remodeling analyzed gene expression on a human ATLAS cDNA platform (45). The resulting analysis and downstream characterization found differential expression of col1a1 and col3a1. More recently, studies on pig mRNA expression have been performed on human nylon DNA arrays reproducibly detecting the expression of 4324 porcine genes (36). Investigators have also begun to use high-density microarrays to investigate the possibility of using heterologously probed cDNAs produced from porcine samples. Initial results using human Affymetrix high density oligonucleotide arrays are promising (41) as are those using porcine samples on human UniGEM microarrays (46).

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Studies analyzing gene probes from cattle, pig, and dog on human and mouse Affymetrix high-density oligonucleotide arrays showed that successful hybridization can be performed with a high degree of statistical significance (47). By slightly lowering the sensitivity of array hybridizations and washings, mainly through altering the salt concentration in washing buffers, hybridization patterns were detected for 2972 transcripts from cattle heart and liver on human U133A GeneChips covering 18,000 human transcripts, with a correlation coefficient of 0.792 across all 2972 genes when compared to expression profiles of human heart and liver (47). In another study, the response of bovine macrophages to Escherichia coli OH157:H7 LPS treatment was investigated on human UniGEM microarrays developed by Incyte (44) as a model for the effects of pathogen invasion. These researchers found that nearly 80% of genes located on the array produced a sufficient and detectable hybridization signal for analysis, 5644 hybridizations out of 7075 total targets. Of genes randomly selected for downstream analysis, 90% gave results in agreement with the microarray data and homology between cow and human genes ranged from 76% to 96%, with a mean of 86%, once again suggesting that the high degree of homology between humans and other mammals provides a strong basis for the use of cDNA arrays in comparative studies. What cross-species DNA array analysis has told us is that different species display a remarkable degree of conservation at the gene level that is evidenced by a high degree of cross-hybridization on DNA arrays. Thus, from studies to date with cross-hybridization between two mammalian species, we can expect up to 85–90% cross-reactivity of genes found on any set of microarrays (31) or by limiting probe hybridizations via increased stringency during washing (40), we can limit and focus our attention on downstream targets that are likely to be significantly up- or down-regulated genes eliminating the possibility of pursuing false positives. Indeed, the use of whole genome or near whole genome microarrays in a cross-hybridization analysis can actually allow a broader coverage of the transcriptome than if researchers were to create arrays from their own cDNA libraries at either high or low density. Since the focus of many researchers is to use the arrays to identify genes that would be good subjects for downstream analysis, researchers are better served by using existing microarray platforms to do their initial screening and direct their major research efforts into downstream analysis of gene regulation and protein function. A variety of companies now produce commercial microarrays and some of them produce arrays for multiple species; Table 1 lists a few of these major microarray producers that make microarrays at a reasonable cost. By no means is this list intended to be complete as new companies and institutes produce arrays at a rapid pace. As described above and in the discussions that

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Table 1 Companies Producing DNA Arrays. Companies and institutes offering a wide variety of DNA array platforms that can be exploited for comparative purposes Company Affymetrix Amersham Clontech Ontario Cancer Institute Superarray Bioscience

High Density Yes Yes Yes and No Yes No

Probe Cy3, Cy3, Cy3, Cy3, 32 P

Cy5 Cy5 Cy5 and Cy5

32

P

follow, we show that a number of different cDNA arrays have been used in comparative research with a high degree of success. Before considering the use of DNA arrays for comparative molecular biology, researchers must first identify the likelihood that the cDNA produced from their model species will cross-react with a particular array and generate sufficient hybridization signals to produce meaningful data. Successful cross-species microarray analysis does not just apply to mammalian genes and models, but broadly across the animal kingdom. For example, researchers using rainbow trout, Oncorhynchus mykiss, as a model for zinc exposure in the environment, assessed gene expression by probing a gill array containing 18,432 cDNA clones made from the puffer fish, Fugu rubripes. Genes involved in energy production, protein synthesis and the inflammatory response were up-regulated in gills after 6 days of exposure to zinc (37). 2. Materials All chemicals used are of molecular biology grade or their equivalent and of the highest purity. All plastic and glassware, including bottles and pipette tips, are autoclaved and gloves must be worn at all times during operations involving nucleic acid manipulation. cDNA ATLAS arrays are purchased from Clontech. Human 19K cDNA arrays are purchased from the Ontario Cancer Institute. 2.1. Total RNA Isolation 1. Diethyl pyrocarbonate (DEPC) (Sigma-Aldrich, St. Louis, MO) is added to water at a concentration of 0.1% (v/v), stirred overnight (>12 h), autoclaved. Tips, tubes and other plastic or glassware may be purchased as certified RNase-free or treated by stirring overnight in DEPC-treated water to destroy any RNases present. DEPCtreated water and RNase-free plastic or glassware are used to make up all solutions in this section and to dissolve the final RNA samples.

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Trizol reagent (Invitrogen, Carlsbad, CA). Chloroform (Fisher Scientific, Fairlawn, NJ). Isopropanol (Fisher Scientific). 70% Ethanol. Add 30 mL of DEPC treated water to 70 mL of 100% ethanol (Pharmco, Brookfield, CT).

2.2. Denaturing RNA Gel Electrophoresis 1. Stock 10× MOPS buffer: 200 mM 3-(N-morpholino) propanesulfonic acid (MOPS), 50 mM sodium acetate, 10 mM EDTA, pH 7. 2. 1% (w/v) agarose formaldehyde denaturing gel: Melt 3 g of agarose in 217 mL of ddH2 O containing ethidium bromide (1 μg/mL) in a sterile flask. Place solution in an incubator set to 55°C. Into a separate sterile flask, add 30 mL of MOPS 10× buffer and 53 mL of formaldehyde 37% (v/v) and place this solution at 55°C. Once both solutions have equilibrated to 55°C, combine the contents of both flasks together in a fumehood and gently swirl without introducing bubbles, and pour into large gel tray to desired thickness. 3. RNA sample buffer: 1× MOPS buffer, 2.2 M formaldehyde, 50% (v/v) formamide. 4. RNA Loading buffer 6× stock: 1× MOPS buffer, 50% (v/v) formamide, 40% (v/v) glycerol. Add a few flakes of bromophenol blue and xylene cyanol as tracking dyes.

2.3. mRNA Isolation 1. Oligotex poly(A)+ mRNA isolation kits (Qiagen). Note: the three buffers listed below come with the kit. 2. Oligotex binding buffer (OBB): 20 mM Tris-HCl, pH 7.5, 1 M NaCl, 2 mM EDTA, 0.2% w.v sodium dodecyl sulfate (SDS; Sigma-Aldrich). 3. Oligotex wash buffer (OWB): 10 mM Tris-HCl, pH 7.5, 150 mM NaCl, 1mM EDTA 4. Oligotex elution buffer (OEB): 5 mM Tris-HCl, pH 7.5.

2.4. cDNA Probe Synthesis 1. 1 μg of mRNA sample. 2. Polymerase chain reaction (PCR) Thermalcycler (e.g., Bio-Rad iCycler [Bio-Rad], PTC-100 [MJ Research]). 3. Oligo-5 -dT20 N-3 (Bio S&T, Montreal, QC). 4. Random primer (100 mM d(N)6 ; New England Biolabs). 5. CDS Primer mix (Clontech). 6. [-32 P]dATP (3000 Ci/mol; GE Healthcare). 7. dNTP mix 1 (dCTP/dTTP/dGTP; 2.5 mM each). 8. 20 mM dNTP mix 2 (6.67 mM each of dATP, dGTP, dTTP). 9. 2 mM dCTP. 10. Cy3-dCTP, Cy5-dCTP (GE Healthcare).

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

Dithiothreitol (DTT) (Sigma-Aldrich). Make up a 0.1 M stock with sterile ddH2 O. Superscript II RNase H reverse transcriptase (200 U/μL) (Invitrogen). RNasin (20 U/μL; Promega). 0.5 M EDTA (Sigma-Aldrich). 10 M NaOH (Sigma-Aldrich). 5 M acetic acid (Sigma-Aldrich). Isopropanol (Fisher Scientific). 70% Ethanol. Prepare as described in Subheading 5.1. TE buffer : 10 mM Tris-base, pH 8.0, 1 mM EDTA.

2.5. DNA Array Hybridization and Washing 1. 2. 3. 4. 5. 6.

Church’s buffer: 0.25 MNa2 HPO4 , 0.25 MNaH2 PO4 , pH 7.5, 7% SDS w/v. 20× SSC: 3.0 M NaCl, 0.3 M sodium citrate (Sigma-Aldrich). 20% SDS w/v. Yeast tRNA (10 mg/mL) (Invitrogen). Calf thymus DNA (10 mg/mL) (Sigma). DIG Easy Hybe Solution (Roche).

2.6. DNA Array Analysis 1. X-ray film or phoshorimaging plate (for ATLASTM cDNA arrays). 2. Microarray reader (for Human 19K cDNA arrays). A number of companies (AlphaInnotech, Affymetrix, VersArray Chip Reader) have array readers available for purchase but some companies or services (and many core facilities at research institutions) will scan and read arrays for a fee, which is considerably more cost effective that purchasing your own array reader. 3. Downloadable analysis software (e.g., Scanalyze; http://rana.lbl.gov/).

2.7. Conﬁrmation of Results: Semiquantitative Reverse Transcriptase-Polymerase Chain Reaction (RT-PCR) 1. 2. 3. 4.

DNAman software (Lynnon Biosoft). Primer Designer software (Scientific and Educational Software). Bio-Rad iCycler (Bio-Rad) or other PCR thermalcycler with gradient capabilities. 50× TAE buffer: 242 g of Tris-base, pH 8.5, 57.1 mL of glacial acetic acid, 37.2 g of EDTA, 1 L of ddH2 O. 5. 1% TAE agarose gel: 1× TAE buffer, 1% agarose (w/v), ethidium bromide (1 μg/mL) in 100 mL of water. 6. DNA loading dye: 0.25% w/v bromophenol blue, 0.25% w/v xylene cyanol FF, 50% v/v glycerol. 7. DNA ladder (Invitrogen). A ladder should be chosen based on the size of the expected PCR products (ranging from 100 bp up to several kilobases).

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3. Methods Before beginning any microarray experiment, it is essential to have proper controls and time points so that the data obtained is biologically meaningful. For considerations on choosing proper controls see Notes 1 and 2. 3.1. Total RNA Isolation 1. Poly(A)+ mRNA isolation kits can be used to harvest poly(A)+ mRNA from frozen tissue samples; however, our experience with one-step isolations is that the yield is generally low when using kits. Instead, we recommend using the traditional method of total RNA isolation, followed by checking to ensure that the RNA is of good quality, and then proceeding with mRNA isolation and array hybridization. 2. Homogenize control and experimental samples in Trizol reagent (1 ml per 100 mg of frozen tissue) and then add chloroform immediately (0.2 mL/mL of Trizol). Invert samples rapidly and repeatedly (15 s) and incubate at room temperature (5 min). Centrifuge at maximum speed for 15 min at 4°C. 3. Remove the aqueous (top) layer and transfer to a sterile RNase-free Eppendorf tube. Add an equal volume of isopropanol to precipitate the RNA. Incubate at room temperature for 10 min. 4. Pellet the RNA by centrifuging the samples at maximum speed in a microcentrifuge for 10 min at 4°C. Carefully, aspirate the supernatant and wash the pellet with 70% ethanol (250 μL). Recentrifuge the RNA at maximum speed for 5 min. Aspirate the ethanol and air-dry the pellet. Be careful not to overdry the pellet, as it will be difficult to resuspend. Dissolve the RNA in DEPC-treated water in a ratio of approx 10 μL of water per 10 μg of pellet and store at –20°C (if it will be processed within a week) or at –80°C (long-term storage). 5. Determine RNA concentration and purity spectrophotometrically by measuring absorbance at 260 and 280 nm. The A260 :A280 ratio should fall between 1.6 and 2.0 for RNA of good purity. Proceed to analyze the quality via RNA gel electrophoresis. Calculate how much total RNA is required for 10–20 μg using the standard of 40 μg of RNA gives an A260 reading of 1.00.

3.2. Denaturing RNA Gel Electrophoresis 1. Prepare a 1% agarose formaldehyde denaturing gel and submerse it in enough 1× MOPS buffer to cover the wells. Prerun the gel for 15 min (while RNA samples are being prepared). 2. Aliquot appropriate volumes of total RNA, containing between 10 and 20 μg, into labeled tubes on ice and dilute up to 15 μL with DEPC treated water. Add 15 μL of RNA sample buffer into each tube and 6 μL of 6× RNA loading buffer. 3. Incubate samples at 55°C for 10 min and place immediately on ice. Add the appropriate volume of RNA loading buffer to each tube to give a final 1× concentration of loading buffer in each sample. 4. Mix the RNA samples gently and briefly centrifuge tubes to collect the entire sample at the bottom of each Eppendorf tube.

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5. Load the entire contents of each tube into the wells of the gel and record sample order for reference. 6. Perform gel electrophoresis at 100 V until the loading dye front reaches the end of the gel. Place the gel on plastic wrap and visualize the RNA using a UV light. The 28S and 18S ribosomal RNA (rRNA) bands are used as an indicator of RNA quality and should be found in a ratio of approx 2:1. This step ensures that you have good quality total RNA before isolating mRNA for DNA array hybridization. Although total RNA can be used for probe synthesis, isolating mRNA is recommended. If RNA is not of good quality the ratio of the 28S rRNA and 18S rRNA bands will be much less than 2:1 or there may be smearing of the sample. (see Note 3 for considerations on maximal amount of RNA to be used for microarray analysis.)

3.3. mRNA Isolation Using Oligotex Mini Kit (Qiagen) 1. Heat Oligotex suspension to 37°C in a water bath, mix by vortex-mixing, and then place at room temperature. 2. Set a heating block to 70°C and heat the OEB. 3. Begin with 0.25–0.5 mg of good quality total RNA although this may need to be modified subsequently for selected tissues or animals if the mRNA yield is not sufficient. 4. Pipet total RNA into an RNase-free 1.5-mL Eppendorf tube and adjust the volume to 500 μL with RNase-free water. 5. Add 500 μL of OBB and 30 μL of Oligotex suspension. 6. Incubate the samples for 3 min at 70°C in a heating block to disrupt secondary structure. 7. Remove samples from the heating block and place at room temperature for 10 min to allow the oligo dT30 on the Oligotex particles to hybridize to poly(A)+ tails of mRNA. 8. Pellet the Oligotex–mRNA complex by centrifugation at maximum speed in a microcentrifuge for 2 min and then carefully remove the supernatant. 9. Resuspend the Oligotex–mRNA pellet in 400 μL of OWB by pipetting up and down, pipet the suspension onto a spin column (provided with the kit) and place the column in an RNase-free 1.5-mL Eppendorf tube. Centrifuge for 1 min at maximum speed. 10. Transfer the spin column to a fresh RNase-free 1.5-mL Eppendorf tube and apply 400 μL of OWB to the column, centrifuge at maximum speed, and discard the flow through. 11. Transfer the spin column to a new RNase-free 1.5-mL Eppendorf tube and apply 20 μL of hot (70°C) OEB onto the column, pipet up and down 3–4 times to resuspend the Oligotex–mRNA resin, and centrifuge for 1 min at maximum speed and save the eluant which contains purified mRNA. 12. Repeat step 11 for maximal mRNA yield and pool the eluants from the two centrifugations. (see Note 4 as further mRNA treatment may be inadvisable at this point.)

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3.4. cDNA Probe Synthesis 1. Preheat thermal cycler to 70°C and add 1 μg (at least 0.5 μg/μL) of each mRNA sample (control and experimental) into separate 0.5 mL (or 0.2 mL depending on block size of thermal cycler) PCR tubes. To each tube, add 100 ng of Oligo-5 dT20 N-3 and 100 ng of random primers (200 ng total) to ensure labeling of the full mRNA pool. 2. Clonetch ATLAS™ kits recommend using 1 μL of the CDS primer mix included with their kits, which includes sequence specific primers for the genes on their arrays. Although the CDS primer mix works relatively well with mRNA samples from hibernating mammal species, we have found that by replacing the CDS primer mix with 100 ng of Oligo-5 -dT20 N-3 and 100 ng random primers (200 ng total), we get similar results for highly expressed genes and a better representation of all genes in samples. This is because the CDS primer mix contains species-specific primers that may fail to label genes that have sequence differences within the region covered by the CDS primer sequence. For screening nonmammalian species that are more distant on the phylogenetic tree, the use of Oligo-5 -dT20 N-3 and random primers is absolutely required. 2. Dilute the reaction to 3 μL total by adding the necessary amount of DEPC-water and incubate for 2 min on the thermal cycler before reducing the temperature to 50°C for 2 min to allow for sufficient hybridization of the primers to mRNA. 3. To make the ATLAS™ master mix for Clontech ATLAS™ arrays, add to a 0.5-mL Eppendorf tube per reaction: 2 μL of 5× reaction buffer (included with Superscript II, Invitrogen), 1 μL of dNTP mix 1 (2.5 mM each of dTTP, dCTP, and dGTP; Invitrogen), 0.5 μL of 100 mM DTT, and 3.5 μL of [-32 P]dATP (3000 Ci/mol; GE Healthcare). Alternatively, other radiolabeled nucleotides can be used provided that they are not included in the dNTP mix (i.e., [-32 P]dGTP can be included with a dNTP mix consisting of dATP, dTTP, and dCTP). To create a master mix for synthesizing fluorescently labeled cDNAs (see Note 5 for other considerations when using fluorescently labeled dNTPs), add per reaction: 8 μL of 5× reaction buffer, 3 μL of 20 mM dNTP mix 2 (6.67 mM each of dATP, dGTP, dTTP), 1 μL of 2 mM dCTP, 1 μL of 1 mM Cy3 or Cy5 dCTP (use one to label the control sample and the other to label experimental sample), 4 μL of 0.1 M DTT and 20 μL of water to bring the reaction up to 37 μL for each reaction. Keep master mixes on ice. 4. Finish preparing the master mix by adding Superscript II reverse transcriptase (RT) to the master mix tubes. ATLAS™ array kits include an MMLV-RT enzyme that we have found to contain little very little activity from time to time. We recommend replacing the 1 μL of MMLV-RT with 1 μL of Superscript II RT per reaction is thus. The addition of 1 μL of Rnasin (Promega) per reaction is also advisable to prevent RNA degradation. Mix by pipetting up and down several times. 5. After the 2 min of incubation of mRNA at 50°C, add 8 μL of the ATLAS™ master mix for synthesis of radiolabeled cDNAs or 37 μL of fluorescent master mix for synthesis of fluorescent cDNAs to each reaction and incubate at 42°C. For radiolabeled probe, the reaction should be incubated for at least 25 min. For

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fluorescently labeled probe, reaction times are significantly longer (at least 2–3 h) due to the poor incorporation of the Cyanine dyes. The reactions are stopped by adding 1 μL of 0.5 M EDTA. Once this is done, the stopped reactions can be stored overnight at –20°C. 6. Probe cleanup varies depending on whether fluorescent probes or radiolabeled probes are made. Radiolabeled probes are cleaned up using the spin columns included with the Clontech ATLAS™ kits to remove unincorporated nucleotides. Each individually prepared radiolabeled cDNA probe is then hybridized to its own individual array. Fluorescent probes are purified first by RNA hydrolysis: add 2 μL of 10 M NaOH and incubate at 65°C for 20 min, then neutralize the reaction with 4 μL of 5 M acetic acid. The labeled cDNA is then precipitated using 100 μL of isopropanol on ice for 30 min followed by centrifugation and washing with 70% ethanol. The labeled Cy3 and Cy5 labeled cDNA samples are then dried, each diluted in 5 μL of water or TE (10 mM Tris-base, pH 8.0, 1 mM EDTA) buffer, and then the two samples are combined into a cDNA pool before hybridization to an array.

3.5. DNA Array Hybridization and Washing 3.5.1. Hybridization and washing of Clontech ATLASTM Arrays (See Note 6) 1. The suggested hybridization temperature for the arrays is 68°C for homologous hybridization. However, for heterologous hybridization, we find that the temperature should be lower. For mammalian hibernator samples, hybridizing at 68°C can generate a signal but better hybridization occurred at 55°C and we ultimately found that overnight hybridization at 44°C in Church’s buffer generated the best hybridization result. For heterologous hybridization with nonmammalian species, reducing the temperature to 40°C gives the best hybridization signal with very little background. 2. After hybridization, the washing steps need to be modified and monitored to ensure that no cross-hybridization signal is lost with heterologous systems. Washes are started at 5× SSC (diluted from the 20× stock), 1% SDS, followed by washes at 2× SSC, 1% SDS, then 1× SSC, 0.5% SDS, and finally 0.5 × SSC, 0.5% SDS. After each wash step, ATLAS™ arrays should be checked for hybridization signal using a Geiger counter. When a wash results in the signal dropping to 500–1000 cpm, washing should be stopped and ATLAS™ arrays exposed to X-ray film or phosphorimager plates. After developing the film or reading the phosphorimager plate, the two films or two images (control vs. experimental) can be overlaid to identify differentially expressed targets first by visual inspection. To obtain a more quantitative result, each image generated from a phosphorimager or from scanned ×-ray film should be converted into .tiff files to be compatible with image analysis software.

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3. ATLAS™ arrays can be reused at least three times. Arrays are stripped by boiling in 10% SDS for 10 min and washing with 2× SSC to remove all the SDS. Then the arrays are wrapped in cellophane and stored at –20°C until reuse.

3.5.2. Hybridization and Washing of Human 19 K microarrays (adapted from the Ontario Cancer Institute protocol, www.microarray.ca/, See also Note 6) 1. Prepare the hybridization solution by taking 100 μL of DIG Easy Hyb solution per hybridization and adding 5 μL of yeast tRNA (10 mg/mL) (Invitrogen) and 5 μL of calf thymus DNA (10 mg/mL) (Sigma) to reduce nonspecific binding. The mixture is heated to 65°C for 2 min and cooled to room temperature. 2. Add 80 μL of the prepared hybridization solution to a pooled pair of Cy3 and Cy5 dye-labeled cDNA samples and again heat to 65°C for 2 min followed by cooling the mixture to room temperature. 3. When using the 19K human microarrays, the genes are arrayed over two slides, so care must be taken when adding the prepared probe to the slides. One slide will lie on top of the other with the array sides facing inwards. Carefully apply the probe mixture in hybridization solution slowly and evenly along one of the edges ensuring that there are no bubbles present. 4. Place the extra hybridization solution inside a hybridization chamber (a sealable slide chamber placed horizontally in a 37°C incubator) to ensure that humidity levels are maintained so the hybridization reaction does not dry out. Place the slides inside the chamber and incubate overnight at 37°C. No modifications in hybridization temperature are needed. 5. After hybridization, wash slides in 2× SSC to remove hybridization buffer and perform subsequent washes by placing the microarray slides in a slide rack and washing in pre-warmed (50°C) 2× SSC, 0.1% SDS for 10 minutes followed by a wash in prewarmed (50°C) 1× SSC, 0.1% SDS. Finally, dip slides in 1× SSC, followed by a brief wash in isopropanol and centrifuge at 500 × g to remove any unbound fluorescent cDNAs. The microarrays can then be scanned at two wavelengths to quantify the different fluors. Two image files are generated and each is analyzed for fluorescent intensities. In cases where evolutionary distance is a concern for generating a good hybridization signal, it is advisable to wash much less stringently. For example, in studies utilizing cDNAs that are only 60–80% identical, it is advisable to lower the washing temperature to 45°C and only perform the 2× SSC wash. In our experience and in the experience of others (40), the salt concentration of the wash buffer has the greatest effect on removing probe from the arrays.

3.6. Array Analysis Analyzing cDNA arrays has become easier with time. Our analysis has primarily been done using the Scanalyze program developed by Michael Eisen, which is available free of charge to academic researchers (http://rana.lbl.gov/),

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coupled with visual inspection of target spots on the arrays themselves. Scanalyze allows users to input two array images, usually one image generated from scanning Cy3 hybridized targets and one image generated from Cy5 targets. Further information can be found at http://rana.lbl.gov/manuals/ScanAlyzeDoc.pdf. Other DNA array analysis programs abound; for further information see Note 7. 1. Open tiff files corresponding to Cy3 and Cy5 scanned images of 19K cDNA arrays in Channel 1 and Channel 2, respectively. 2. Once the images are loaded, click on the redraw button and adjust the gain and normalization of each image such that they have the same brightness and intensity upon visual inspection. 3. Grid the images such that each of the 19K cDNA “spots” is outlined by a Scanalyze generated circle. For each new batch of arrays, a new grid must be created. Click on the New Grid button on the Grid control form and select between 1 and 32 grids. 4. Enter the number of columns and rows per grid, column and row spacing and column and row height. 5. Because array printing is sometimes not entirely perfect, the grid may or may not fit the array exactly. In this case, the directional buttons in Scanalyze can be used to adjust the array grid up, down, left or right as well as stretching the grid in the same direction. When the array grid is close to being perfectly overlaid on each image, Scanalyze can perform fine tuning of the grid by pressing the refine button. If selected spots are misaligned, they can be further manipulated by selecting the “spot” option and using the directional buttons to align spots individually. 6. Once the grid has been made and fits the array, hit the save data button. Scanalyze will calculate the output information for each spot on the array and provide it in a tab delimited format that can be opened in Microsoft Excel. 7. By far the quickest and easiest type of analysis is to determine the hybridization ratio of intensities generated Channel 1: Channel 2 (e.g., control vs. hibernating). This will give a general indication of the ratio of gene levels between one state and another. Because the data is in Microsoft Excel, it can then be sorted based on highest to lowest (or lowest to highest) ratios by clicking on the Data tab and then selecting Sort. The genes corresponding spots on the arrays that show the greatest up- or down-regulation are then identified and downstream analysis is performed (see Note 8).

3.7. Conﬁrmation of Results: Semiquantitative PCR ( See Note 9) 1. For each gene of interest, obtain sequences for the homologous gene from other animals by downloading gene sequences from National Center for Biotechnology Information (www.ncb.nlm.nih.gov). 2. Select nucleotide from the drop down menu and enter the gene interest either by name or abbreviation. 3. Once one gene sequence is obtained, it is often easier to do a BLAST search (www.ncbi.nlm.nih.gov/BLAST/) to obtain the sequence of the gene from other species. Download the sequence of the gene from multiple sources. For example,

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5.

6.

7.

8.

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for studies of ground squirrel genes, other rodent and/or mammalian sequences (e.g., mouse, rat, human) might be chosen. For example: with hibernator genes, we typically use Homo sapiens, Mus musculus and Rattus norvegicus as a starting point for determining homologous regions. For animals of greater evolutionary distance, it is helpful to have sequences that are more diverse and/or from species that are more closely related phylogenetically to the species of interest. For example, if analyzing a turtle gene, a choice of frog, chicken and rat sequences might be more appropriate for initial analysis. Three alternate sequences is often sufficient, but sometimes with a well-studied gene, more species can be added to the analysis. Open each sequence (H. sapiens, M. musculus, and R. norvegicus) in DNAman. Select Edit, Select All from the drop down menu and enter each sequence (Sequence, load channel) into its own channel. Once sequences are loaded, select Sequence, Multiple Alignment, Add From Channel and enter in all the sequences. Select the full alignment button and hit OK. A comprehensive gene alignment will be displayed with regions of homology identified by dark shading. Once regions of high homology are found, Primer Designer software (Scientific and Educational Software) is used to generate putative primer sequences. Primer sequences showing no mismatched bases within the last 10 bases at the 3 end can be used to generate primers. In some cases where sequence variability is high, a degenerate PCR primer is designed instead and used to obtain a cDNA sequence from the organism of interest. With this newly obtained sequence, species-specific primers can then be designed for use in expression analysis. The species-specific primer can also be used in the technique of rapid amplification of cDNA ends (RACE-PCR) in order to obtain the full sequence of the open reading frame of the gene of interest (12). The protocol for creating first strand cDNA is identical to that used in DNA array probe synthesis section (Subheading 3.4.; see also Note 10) with the exception that no labeled nucleotides are added. The master mix contains 2 μL of 5× reaction buffer (Invitrogen), 0.5 μl 100 mM DTT, and 1 μl 10 mM dNTPs (2.5 mM each dATP, dTTP, dCTP, dGTP). To optimize PCR conditions for a new sequence from a new animal, we routinely set up a temperature gradient on a gradient enabled thermal cycler (Bio-Rad iCycler) in a range of 50–70°C. The heating blocks are usually set up to run a gradient across eight samples so the temperature increments are 2.5°C. Set up a master mix on ice for each 50 μl reaction containing 5 μL of 10× reaction buffer, 2.5 μL of 50 mM MgCl2 , 1 μl 10 mM dNTPs, 1 μl 0.5 μM PCR primers, 1 μL of template, 0.25 μL of Taq DNA polymerase (Invitrogen) and 39.25 μL of water. The PCR protocol generally used is one initial denaturation step at 95°C for 2 min, and then 35 cycles of 95°C for 45 s, annealing (50°C to 70°C for 45 s), and extension at 72°C. The extension time at 72°C depends on the size of the product being amplified. Taq polymerase is a highly active so the general rule of 1 min per kilobase of DNA being amplified is more than sufficient but the time can be scaled back as required for shorter amplifications. After the 35 cycles are performed, a final 72°C extension step is carried out for 10 min and then the reaction is set on hold at 4°C or placed at 4°C in a refrigerator.

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9. Run all of the reactions on a 1% TAE agarose gel, stain with ethidium bromide and visualize under UV light and take a picture for a photographic record. Identify the PCR annealing temperature (and/or extension time) that gave the highest amount of amplification and use these conditions for subsequent work. 10. Sequence the PCR product to confirm that it is indeed the gene of interest. 11. Once the annealing conditions are worked out and the PCR product has been confirmed as being the gene of interest, the control and experimental samples can be compared. Prepare serial dilutions of the first-strand cDNA for both control and experimental samples (i.e., 10−1 , 10−2 , 10−3 , 10−4 ) for the gene of interest. Prepare an identical set of serial dilutions to be used to amplify a control gene. Commercial primers for genes such as -tubulin, -actin, or glyceraldehyde-3phosphate dehydrogenase (more commonly referred to as GAPHD on commercial websites) may work for species with high sequence identity but for animals further separated through evolution, designing your own primers for control genes from highly identical regions is advisable. 12. PCR is performed on each dilution in the control and experimental samples. When complete, the products are electrophoresed on a 1% TAE agarose gel, which is stained with ethidium bromide and visualized under UV. Band intensities in the different lanes are measured using imaging software (Imagequant). Bands from one or more of the lower dilutions will likely be saturating in intensity so choose a band that is subsaturating but gives sufficient signal as the one to be quantified. This procedure ensures that the band chosen for quantification has an intensity that falls within the linear range of the imaging software used. Band intensity of the gene of interest in each lane can be normalized against the intensity of the corresponding control gene band to normalize for any unequal loading.

3.8. Outlooks: Comparability of Microarray Data, Comparative Genomics, and Hibernation One of the early concerns with DNA array analysis was the lack of useful public domains to house the wealth of information that was produced (48–50). The initial hope was to create unique public databases that would allow researchers free access to microarray data in order facilitate more rapid discovery in areas that may be seemingly unrelated. For example, a researcher studying a particular gene, would be able to look at various microarray profiles and determine where and when the gene is up- or down-regulated and formulate a hypothesis about it’s regulation in relation to other genes. The introduction at the NIH of the gene expression omnibus (GEO) (http:// www.ncbi.nlm.nih.gov/geo/) has allowed researchers to do just this (51,52). Other microarray databases also exist. The Stanford University Microarray Database (http://genome-www5.stanford.edu/) lists published data, references and the organisms from which the data were obtained and ArrayExpress (http://www.ebi.ac.uk/arrayexpress/), a part of the European Bioinformatics

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Institute (53–56), performs similarly to the NIH GEO database containing data from over 12,000 hybridizations covering at least 35 species. Currently, GEO remains the largest and fully open database that allows scientists free access to data acquired by high-throughput technology, including those relating to mRNA expression, genomic DNA analysis, serial analysis of gene expression (SAGE), mass spectroscopy, and proteomics. While these databases are useful, particularly for researchers working on model species, they have only just begun to be utilized by researchers undertaking comparative studies. It is clear from that cross-species microarray analysis allows comparative researchers an opportunity to further research output immensely. Two areas that cross-species array screening does not address are (1) the case of genes displaying low homology between the array and the target organism, and (2) the occurrence of novel genes that are specific to a particular organism and, therefore, not represented on commercial arrays. However, arrays are being produced for more and more species all the time so, to some extent, both issues may dissipate with time as arrays become available for species that are phylogenetically closer to the species of interest. In the hibernation field, for example, the laboratory of Matt Andrews (57) has recently produced a DNA array with over 4000 cDNAs derived from a S. tridecemlineatus cDNA library and used this array to analyze the heart transcriptome in hibernation. With 4000 genes arrayed, a high percentage of the genome remains unrepresented and, hence, at the present time, heterologous probing still gives a broader result; e.g., we achieved 85–90% hybridization by ground squirrel cDNA with commercial 19,000 gene human arrays. However, the species-specific array offers the potential opportunity to find novel genes that occur only in hibernators (i.e., not found in the human genome) and, hence, the species-specific array could make a unique contribution toward the complete genetic analysis of the hibernation phenotype. Because data from array screening must be followed up with rigorous downstream analysis, regardless of which array platform is used, the advantage for comparative biologists clearly currently lies with the use of cross-species cDNA array hybridizations. This is particularly true in the cases where the species of interest displays remarkable gene identity to a model species. Because the genomes of a number of non-traditional model species are currently being sequenced around the world, including S. tridecemlineatus, which is slated for full genome sequencing by the Human Genome Research project (http://www.genome.gov/), the field of cross-species microarray analysis should bloom in the coming years. Annotation and analysis of genes and gene structures from many species will shed more light into precise gene identity and homology, further confirming the usefulness of microarrays for future cross-species work by comparative biologists.

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4. Notes 1. A critical part of any array study is establishing guidelines and using proper controls. As in all scientific endeavors, the choice of the appropriate control situation is the key to being able to properly interpret the gene expression changes that occur in the experimental situation. This seems especially important in array screening studies because these are analyzing mRNA pools and mRNA typically has a very short half-life in cells so for these studies it is best to have control and experimental samples that are as closely matched in time and in preexperimental state as possible. For example, a current controversy in the hibernation field illustrates this. We want to know how torpor is regulated and what genes need to be up-regulated to help an animal enter torpor and/or stabilize metabolism for long term survival the torpid state. Hence, we choose control and experimental animals that are as closely matched as possible: in this case, controls are euthermic animals at 37°C body temperature that have not entered torpor in the 5°C cold room versus animals in torpor in the same cold room with body temperature near ambient. This shows us the pool of gene expression changes that define the difference between active and torpid states. By contrast, some other groups advocate comparisons of summer active animals with winter torpid ones (58). This could show seasonal differences in the pools of mRNA in organs but is inappropriate for investigating the control of torpor because there are too many other differences between summer and winter animals including environmental conditions (e.g., photoperiod, thermoperiod), physiological states (e.g., actively feeding or not; active above ground vs. sleeping in burrows), and reproductive states that make it impossible to “dissect out” the gene expression changes that are torporspecific. Hence, summer active animals are, at best, an extremely poor biological control and, at worst, a time point that is erroneous and detrimental to hibernation research as a whole. Indeed, the experimental protocol that we use (euthermic vs. torpid winter animals) for our gene screening has shown that a wide variety of genes are specifically up-regulated when animals enter torpor; these appear to perform essential biological functions in the torpid state. We also find extensive organ-specific activation of stress-induced signal transduction pathways in torpid versus euthermic animals including different classes of mitogen-activated protein kinases (17,59) which shows that organs maintain substantial metabolic activity during torpor. This actually goes against some previous “conventional wisdom” in hibernation research, which had the notion that most biological processes were turned down or off during torpor. 2. It is worth noting that our stress marker screening using Kinexus Kinetworks™ phospho-protein screens (Table 2) also revealed that selected proteins show an altered phosphorylation state during torpor in liver of S. tridecemlineatus further supporting the idea that hibernators do maintain metabolic activity during torpor. The results included an unchanged phosphorylation status of p38MAPK (Thr180 /Tyr182 ), elevated phosphorylation of JUN (Ser73 ), decreased phosphorylation of AKT at Ser473 but not Thr308 . This data are in agreement with previous studies in S. richardsonii that showed that during torpor p38MAPK activity was

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Table 2 Phospho-protein screen for S. tridecemlineatus liver Protein Name N-methyl-d-aspartate glutamate receptor subunit 1 (112) Adducin (121) Adducin (80) Oncogene SRC (49) Signal transducer and activator of transcription 5 (94) p38 MAP kinase (38) Protein kinase C (S657) Oncogene SRC (49) Protein kinase C / (T368) MAP kinase kinase 6 (MEK6) (36) MAP kinase kinase 3 (MEK3) (33) S6 kinase p70 (80) Protein kinase C Signal transducer and activator of transcription 3 (83) Oncogene JUN (40) Oncogene Raf 1 (69) Oncogene Raf 1 (63) Protein kinase C Protein kinase B- (Akt1) (T308) Protein kinase B- (Akt1) (S473) Glycogen synthase kinase 3a (45) dsRNA dependent protein kinase (68) Glycogen synthase kinase 3 (45) Glycogen synthase kinase 3 (40)

Abbreviation

Epitope

Euthermic Torpid

NR1

S896

133

—

Adducin a Adducin g SRC STAT5

S724 S662 Y529 Y694

1000 485 347 616

894 404 834 —

p38 MAPK PKCa SRC PKCa/b MEK6

T180/Y182 S657 Y418 T638/641 S207/T211

205 1658 714 1128 188

136 3710 1413 1356 240

MEK3

S207/T211

295

321

p70 S6K PKCe STAT3

T389 S719 S727

688 618 1255

905 921 2315

JUN RAF1 RAF1 PKCd PKBa PKBa GSK3a

S73 S259 S259 T505 T308 S473 S21

— 656 986 407 341 1193 172

426 411 943 947 417 547 54

PKR

T451

243

653

GSK3a

Y279

437

286

GSK3b

Y216

—

317

Extracts were prepared from liver of euthermic and hibernating ground squirrels and assessed using the Kinexus Kinetworks™ screens for phosphoprotein status to identify activation or suppression of signaling proteins during hibernation. The epitope to which the antibody is derived is given and relative expression levels of each protein in the euthermic and torpid states are given in arbitrary units.

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unaltered, while JNK activity, the kinase that phosphorylates JUN, was highly elevated (59) and studies in S. tridecemlineatus that showed that Ser473 but not Thr308 was decreased during hibernation and torpor (60). Hence, data from array screening agrees well with data gathered by more traditional assays. 3. cDNA probes for microarray analysis are labeled using either fluorescence or radioactivity. Fluorescence probes are used exclusively for high-density DNA microarrays whereas 32 P-labeled probes are generally used for macroarrays, such as Clontech ATLAS™ arrays. In situations of heterologous probing, our experience has shown that outcomes can be highly successful when several conditions are met: a. There is a high percent identity of genes between the two species (the test species vs. the one used to make the array), b. Optimal amounts of starting material are available, and c. Slightly less stringent hybridization conditions are applied than would be used in homologous probing. From tissue samples, we first purify RNA using the Trizol (Invitrogen) method of RNA extraction followed by mRNA [or poly(A)+ ] purification using Qiagen Oligotex purification kits according to manufacturer’s protocols. Our work has also found that while hybridizations can be performed with limited starting material (0.25–0.5 μg mRNA or 1–2 μg of total RNA at concentrations of at least 0.5 μg/μL), significantly higher hybridization intensity is obtained using purified mRNA isolated using Qiagen Oligotex–mRNA purification kits according to the manufacturer’s protocols. The maximum mRNA suggested by each array protocol is usually at least 1 μg of mRNA. cDNA probes for microarray analysis are then prepared from the mRNA using either fluorescence or radioactivity. Fluorescence probes are used exclusively for high density DNA microarrays whereas 32 P-labeled probes are generally used for macroarrays, such as Clontech ATLAS™ arrays. In order to compare two samples (e.g., control vs. experimental) with 32 P-labeled probes, one set of radiolabeled cDNA is hybridized to one array and another probe set is hybridized to a duplicate array. 4. Once the mRNA is prepared, it is suggested by most protocols that DNase treatment be performed prior to array hybridization to rid samples of any contaminating genomic DNA. While it is advisable to rid the sample of any contaminations, we found that eliminating this step had essentially no effect on array hybridization. In fact, when we eliminate this step, we rarely see binding above background levels (areas of the arrays spotted with buffer and no DNA) to genomic DNA array spots found on arrays. Thus, in cases where the mRNA sample is extremely limited, we feel that the possibility of RNases being introduced through increased handling outweigh the risk of some a small fraction of genomic DNA contaminating the reaction. 5. For fluorescence labeling, cyanine dyes (Cy3 and Cy5 for short), which are extremely light sensitive, are the preferred choice due to their relative ease of hybridization to microarrays (e.g., these dyes show little steric hindrance compared

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to other fluorescent tags). In this case, one sample is Cy3 labeled and the other is Cy5 labeled and then the two probes are hybridized on a single chip. Further, for microarray studies involving fluorescently labeled cDNA, we generally perform two sets of hybridizations, in the first set labeling the control sample with Cy3 dye and the experimental sample with Cy5 dye and in the second set reversing this to eliminate any bias in the labeling procedure. Although the absolute fluorescence generated from each individual hybridization may vary, similar expression ratios are typically found on reciprocally probed arrays. 6. For our studies using Clontech ATLAS™ arrays, we first used the suggested conditions given in the protocols included with the array kits to prepare radiolabeled cDNA. For our hybridization to microarrays, we began by following guidelines set out by the Microarray Center of the Ontario Cancer Institute. Subsequently, we found that altering some of the protocols increased our ability to detect transcripts on the arrays during heterologous probing. In general, preparation of cDNAs incorporating 32 P and fluorescent tags are the same with the major exception being that fluorescent probes should be made in the dark or reduced light to minimize excitation and degradation of the fluorescent signal. 7. While our analysis using Scanalyze has proven sufficient for identifying target genes for further studies, other analysis programs may offer more to other researchers. Other freeware applications available to researchers include the HTML based programs known as “Bullfrog” for Affymetrix arrays and “Spot” for custom and other types of cDNA arrays (61). Also available is the Gene Expression Open Source System (GEOSS) formerly known as Gene X Va (62), which allows users to input hybridization ratios and prepares data for clustering by easily converting hybridization ratios into color intensities for visualization and producing Venn diagrams (if multiple experimental samples are used). Table 3 lists companies and other researchers that offer analysis software. Open source platforms are generally made freely available to academic and nonprofit researchers, usually with registration and referencing of the program. Other programs available are licensed to researchers at a price but do not necessarily improve upon the large number of freeware applications available to date. 8. A 1.5-fold change in gene expression seems to be sufficient to document a statistically significant difference between control and experimental situations in most cases but may still miss some important clues. Another layer of analysis can be used in which groups of related genes are assessed for overall changes in pathway response to stress. For example, a study by Mootha et al. (53) found that when analyzing 22,000 genes from skeletal muscle of age-matched human males falling into three categories (normal glucose tolerance, impaired glucose tolerance, diabetic), no genes were differentially regulated according to prior standards of statistical analysis or more simply, on a gene by gene basis, there was little detectable expression difference between genes in the three groups. However, by enriching the genes into sets for a pathway-based analysis, they found that genes involved in oxidative phosphorylation were co-coordinately down-regulated in diabetics and this was traced to the action of the transcription factor PGC-1that

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Table 3 Microarray analysis programs Array Program ArrayPro Bullfrog and Spot F-Scan GenePix GEOSS ImaGene MatArray

P-scan Quantarray Scanalyze

Developer Media Cybernetic Zapala et al. Munson et al., NIH Axon Instruments Lee et al. BioDiscovery Wang et al., National Research Center for Juvenile Diabetes Munson, et al., NIH Packard Bioscience Michael Eisen

Open Source No Yes Yes No Yes No Yes

Yes No Yes

Reference (61) (64) (52) (65)

(64)

showed a approx 20% decrease in expression in diabetics (63). The development the Gene Set Enrichment Analysis (GSEA) approach for clustering genes into ordered groups according to known signaling pathways aids in the identification of biochemical functions that are associated with a particular stress. Of course, this analysis requires prior knowledge of signaling pathways and their downstream effects on gene expression, which may or may not be available for comparative systems. 9. While the preferred method of downstream microarray analysis is generally stated to be quantitative RT-PCR (Q-PCR), in reality, we feel that Q-PCR is unnecessary for downstream analysis of data derived from microarray screening in comparative studies. This is because the key outcome being sought is the relative change in gene expression between control and experimental conditions (i.e., is a gene upor down-regulated), not the quantitative amount of each mRNA transcript type in each sample. Thus, semiquantitative RT-PCR is more than sufficient. Further, the term “quantitative” applied to PCR is not only incorrect, but also misleading. For a result to be truly quantitative, controls must be performed at every step of the analysis, including controlling for % yields during RNA isolation through to spiking the isolated RNA with a control mRNA of known quality and quantity so that downstream analysis can be performed on the control to give a precise standard for all downstream applications. For confirmation of gene expression changes highlighted by heterologous screening, the two methods most often used are Northern blotting (including dot or slot blotting) or semiquantitative RT-PCR. In cases where a differentially regulated target gene has already been cloned and is available to researchers, Northern blotting or dot blotting would be the preferred method of downstream analysis. In most cases, however, comparative

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biologists would choose semiquantitative RT-PCR which is highly effective for two reasons: (a) PCR is conducive to high-throughput analysis of gene expression, and (b) the technique also generates material for a partial sequence analysis which, after translation, allows researchers to assess changes to amino acid sequence and putative structure/function differences of the protein in the species of interest as compared with its homologues in Genbank. 10. When using RT-PCR for validation, it is important to amplify as much of the total transcript population as possible. Oligo-5 -dT20 N-3 priming works for many small mRNAs (<2 kB) but larger mRNAs and mRNAs containing a lot of secondary structure require additional priming. Thus, it is wise to also include random primers for first strand cDNA synthesis.

Acknowledgments Appreciation and thanks are extended to J. Storey for critical reading and suggestions for the manuscript. This work was supported in part to funding to S.F.E. (Ontario Graduate Scholarship) and K.B.S. (NSERC). K.B.S. holds a Canada Research Chair in Molecular Physiology. References 1. Schena, M., Shalon, D., Davis, R. W., and Brown, P. O. (1995) Quantitative monitoring of gene expression patterns with a complementary DNA microarray. Science 270, 467–470. 2. Schena, M., Shalon, D., Heller, R., Chai, A., Brown, P. O., and Davis, R. W. (1996) Parallel human genome analysis: microarray-based expression monitoring of 1000 genes. Proc. Natl. Acad. Sci. USA 93, 10614–10619. 3. Pease, A. C., Solas, D., Sullivan, E. J., Cronin, M. T., Holmes, C. P., and Fodor, S. P. (1994) Light-generated oligonucleotide arrays for rapid DNA sequence analysis. Proc. Natl. Acad. Sci. USA 91, 5022–5026. 4. DeRisi, J., Penland, L., Brown, P. O., Bittner, M. L., Meltzer, P. S., Ray, M., Chen, Y., Su, Y. A., and Trent, J. M. (1996) Use of a cDNA microarray to analyse gene expression patterns in human cancer. Nat. Genet. 14, 457–460. 5. Shalon, D., Smith, S. J., and Brown, P. O. (1996) A DNA microarray system for analyzing complex DNA samples using two-color fluorescent probe hybridization. Genome Res. 6, 639–645. 6. Urisman, A., Fischer, K. F., Chiu, C. Y., Kistler, A. L., Beck, S., Wang, D., and DeRisi, J. L. (2005) E-Predict: a computational strategy for species identification based on observed DNA microarray hybridization patterns. Genome Biol. 6, R78. 7. Wang, D., Coscoy, L., Zylberberg, M., Avila, P. C., Boushey, H. A., Ganem, D., and DeRisi, J. L. (2002) Microarray-based detection and genotyping of viral pathogens. Proc. Natl. Acad. Sci. USA 99, 15687–15692. 8. Wang, D., Urisman, A., Liu, Y. T., Springer, M., Ksiazek, T. G., Erdman, D. D., Mardis, E. R., Hickenbotham, M., Magrini, V., Eldred, J., Latreille, J. P.,

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7 Proteomic Analysis of Neuroendocrine Peptidergic System Disruption Using the AtT20 Pituitary Cell Line as a Model Fumin Dong, Liming Ma, Michel Chrétien, and Majambu Mbikay

Summary Environmental pollutants may affect the activities of many cellular enzymes. The effect on the proteome of enzymatic inhibitors can be determined using two-dimensional (2D) gel electrophoresis. In neuroendocrine cells, proprotein convertases 1 and 2 (PC1 and PC2) mediate the proteolytic activation of many precursors to peptide hormones and neuropeptides. Enzymatic activities of these calcium-dependent proteinases are readily regulated by chelating agents and by heavy metals ions found in the environment. Such an inhibition could result in a potentially pathological disruption of the peptidergic system. We are interesting in finding out to what extent specific inhibition of these enzymes could affect the proteome of a neuroendocrine cell. To address this question, we used the mouse pituitary AtT20 cell line as a model. We compared the proteomic pattern of control cells to that of cells overexpressing proSAAS, a PC1-specific inhibitor. The comparison was conducted using two-dimensional (2D) gel electrophoresis, mass spectrometric identification of differing proteins and immunoblotting to confirm their identity. The 2D analysis revealed a number of alterations in the proteome of proSAAS-overexpressing cells. Mass spectrometric analysis of tryptic peptides identified two proteins found in more abundance in these cells as proSAAS and Ephrin type A receptor 2. Key Words: AtT20 cells; immunoblotting; mass spectrometry; PC1; proprotein convertase; proSAAS; proteomics; 2D gel.

From: Methods in Molecular Biology, vol. 410: Environmental Genomics Edited by: C. Cristofre Martin © Humana Press, Totowa, NJ

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1. Introduction Whole animal systems can be useful for understanding the phenotypic effect of exposure to toxicants. The use of whole animal systems for expression profiling, genomic or proteomic, can be complicated due to the fact that they are composed of many different cell types that may be differentially affected by environmental conditions. As a result, the use of cell culture systems is often required for initial studies. The AtT-20 cell line was derived from a mouse pituitary adenoma (1). It expresses pro-opiomelanocortin (POMC), the precursor of several bioactive hormones, including the stress hormone adrenocorticotropin (ACTH), the satiety hormone -melanocyte-stimulating hormone (-MSH) and the analgesic hormone -endorphin (2). These hormones are generated by multiple internal cleavages of POMC by PC1 and PC2 (3,4), two members of the family of calcium-dependent subtilases named proprotein convertases (PCs) (5). PC1 and PC2 are widely expressed in neuronal and endocrine tissues (6–9). Their intracellular activities are modulated by the levels of co-resident specific inhibitors, proSAAS for PC1 and 7B2 for PC2 (10–13). Together, these enzymes undoubtedly influence the neuroendocrine peptidergic system that regulates many important biological functions including growth, energy homeostasis, appetite and satiety, nociception and analgesia and, possibly, cognition and behavior. Genetic deficiency for PC1 and PC2 has been shown to affect some of these functions in mice and humans (14–16). PC1 and PC2 enzymatic activities are sensitive to potential environmental pollutants, most particularly to calcium chelating agents such as EDTA and to heavy metal ions such as Hg, Cu and Zn (17,18). It is not known how their inhibition by these common pollutants would alter the proteome of neuroendocrine cells. Detection of such alterations could be a measure of the potential disruptive effect of a PC-inhibitory compound on the neuroendocrine peptidergic system. The AtT20 cell line has been extensively used to study the biosynthesis of PC1 and PC2 and their roles in POMC processing into multiple bioactive peptides (3,4). It represents therefore a valid model to evaluate the effects of pollutants on the peptidergic neuroendocrine cells. As a first step towards determining the usefulness of proteomics as an assessment method for neuroendocrine peptidergic disruption, we examined changes induced in the proteome of AtT20 cells by overexpression of proSAAS, the PC1-specific inhibitor [AtT20(proSAAS) cells] (11). Two-dimensional (2D) gel profiling of AtT20 and AtT20(proSAAS) cells revealed many differences in the relative protein abundance. Mass spectrometric analysis identified two proteins found in greater abundance in AtT20(proSAAS) cells as proSAAS and Ephrin type A receptor 2 (EphA2). EphA2 identity was confirmed by 2D immunoblotting. This receptor

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tyrosine kinase is known to play a role in neural outgrowth (19) and in cancer (20). 2. Materials 2.1. Cell Culture and Lysis 1. Mouse pituitary AtT20 and AtT20(proSAAS) cell lines. AtT20(proSAAS) were originally obtained by transfection into AtT20 cells of a proSAAS expression vector containing a neomycin resistance gene, followed by selection for cells expressing this gene by their resistance to the cytotoxic drug G418 (11). 2. Dulbecco’s modified Eagle’s medium (DMEM) (Gibco/BLR, Grand Island, NY) supplemented with 10% fetal bovine serum (FBS) (Wisent Inc., Quebec, Canada) and 30 μg/ml gentamicin sulfate (Gibco/BLR), with or without 500 μg/mL of G418 (Gibco/BLR). 3. Phosphate-buffered saline (PBS): 1.35 M NaCl, 28 mM KCl, 80 mM Na2 HPO4 ·7H2 O, 15 mM KH2 PO4 , pH 7.2. 4. Versine: PBS containing 0.1 mM EDTA. ´ 5. Buffer B: 10 mM N-2-hydroxyethylpiperazine-N-2-ethanesulfonic acid (HEPES)KOH at pH 7.4, 10 mM KCl, 1.5 mM MgCl2 , 0.5 mM sodium EGTA, 1 mM dithiothreitol (DTT). 6. Protease Inhibitor Cocktail (PIC) tablets (Roche Diagnostics GmbH, Mannheim, Germany). One tablet is dissolved in 10 mL of buffer B. 7. Corning cell lifter (Sigma-Aldrich, St. Louise, MO). 8. Syringe (3-mL) and needle (26-gauge). 9. Protein Assay Kit (Bio-Rad, Hercules, CA).

2.2. Two-Dimensional (2D) Gel Electrophoresis 1. 2. 3. 4. 5. 6. 7. 8.

9. 10. 11.

IPGphor strip holder, 18 cm (Amersham Biosciences, Piscataway, NJ). IPGphor isoelectric focusing system (Amersham Biosciences). Hoeffer DALT Electrophoresis System (Amersham Biosciences). Immobilized pH gradient (IPG) strip of pH 4–7L, 18 cm (Amersham Biosciences, Uppsala, Sweden). Store at –20 ºC. Rehydration buffer (RB): 7 M urea, 2 M thiourea, 4% CHAPS, 1% DTT. Bio-Lyte 3/10 Ampholyte (Bio-Rad). Store at 4 ºC. Mineral oil (Bio-Rad). IPG strip equilibration buffer 1 (EB1): 50 mM Tris-HCl, pH 8.8, 6 M urea, 30% (v/v) glycerol, 2% sodium dodecyl sulfate (SDS), 1% DTT, 1 × 10−4 % bromophenol blue. IPG strip equilibration buffer 2 (EB2): 50 mM Tris-HCl, pH 8.8, 6 M urea, 30% (v/v) glycerol, 2% SDS, 4% iodoacetamide, 1 × 10−4 % bromophenol blue. Acrylamide–bis-acrylamide solution (30.8%T) (neurotoxin, work with gloves!): 30% Acrylamide, 0.8% bis-acrylamide. Store at 4 ºC. 10% SDS.

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12. 13. 14. 15.

10% Ammonium persulfate. N, N, NN -Tetramethyl-ethylenediamine (TEMED, Bio-Rad). SDS electrophoresis buffer: 25 mM Tris, 192 mM glycine, 0.1% SDS. Low melting agarose (LMT) (Gibco/BLR). Dissolve 1% LMT in SDS electrophoresis buffer by heating in a microwave oven. Keep at 50ºC before use. 16. Silver staining solutions: (1) fixative: 30% ethanol, 5% acetic acid; (2) sensitizer: 0.02% sodium thiosulfate pentahydrate; (3) silver reagent: 0.2% silver nitrate; (4) Developer: 4% potassium carbonate, 0.025% formaldehyde (37%); (5) stop reagent: 4% Tris base, 2% acetic acid. 17. Umax Powerlook 1100 scanner (Umax Technologies, Dallas, TX).

2.3. Matrix-Assisted Laser Desorption-Ionization Time-of-Flight Mass Spectrometry (MALDI-TOF MS) 1. Trypsin (cat. no. V5111, Promega, Madison, WI). 2. ZipTip C18 pipet tips (Millipore, Bedford, MA). 3. Matrix solution: 10 mg/mL of -cyano-4-hydroxycinnamic acid in 50% acetonitrile containing 0.1% trifluoroacetic acid. 4. Voyager DETm -PRO matrix-assisted laser desorption ionization time-of-flight (MALDI-TOF) mass spectrometer (PerSeptive Biosystems, Framingham, MA). 5. MALDI plate (PerSeptive Biosystems, Framingham, MA).

2.4. Immunoblotting 1. 2. 3. 4. 5. 6.

Nitrocellulose membrane (Bio-Rad). Towbin buffer: 25 mM Tris, 192 mM glycine, 0.1% SDS, 20% (v/v) methanol. 1× Tris-buffered saline (1× TBS): 10 mM Tris-HCl, pH 8.0, 150 mM NaCl. TBS-T: 1×TBS, 0.1% Tween-20. 5% Skim milk solution: TBS-T, 5% skim milk. ECL™ anti-rabbit IgG, horseradish peroxidase linked F(ab )2 fragment (from donkey) (Amersham Biosciences). 7. Western Lightning Chemiluminescence reagent Plus (PerkinElmer, Boston, MA).

3. Methods 3.1. Sample Preparation 1. Plate AtT20 and AtT20(proSAAS) cells in 150-mm dishes (three dishes per cell type), in DMEM medium. Let them grow to confluence at 37°C in a 5% CO2 – 95% air humidified atmosphere. The culture medium for AtT20(proSAAS) was supplemented with G418 at 500 μg/mL to maintain selection. 2. Wash cell monolayers 3 times with 10 mL of PBS each time. 3. Cover cells with Versine; scrape them off the plate with a cell lifter and transfer the suspensions into centrifuge tubes. 4. Sediment cells by centrifugation at 1300g at 4°C for 5 min. Remove and discard the supernatants.

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5. Centrifuge again as described above to remove the remaining Versine. 6. Suspend cells into 0.5 mL of PIC-containing buffer B and disrupt them by 30 pull/push through a 26-gauge needle attached to a syringe. 7. Centrifuge the resulting lysates at 1000g at 4°C for 7 min. Transfer supernatants into fresh tubes and discard pellets. 8. Centrifuge supernatants at 105 g at 4°C for 30 min; transfer the supernatants (cytosolic fraction) into fresh tubes and keep the pellet (microsomal fraction). 9. Add 3 volumes of acetone to each cytosolic fraction and place at -20°C for 1 h to allow protein precipitation. Centrifuge at 15,800g at 4°C for 15 min. Discard the supernatants and keep the pellet. 10. Dissolve each pellet of cytosolic or microsomal proteins in 500 μL of RB by sonication (see Note 1). 11. Determine the protein concentration by the Bradford method using the Bio-Rad Protein Assay kit.

3.2. 2D Electrophoresis 3.2.1. Isoelectric Focusing (IEF) 1. Dissolve 0.8 mg of cytosolic proteins or 1 mg of microsomal proteins in 340 mL of RH buffer containing 0.5% Bio-Lyte 3/10 Ampholyte and 0.5% (v/v) of 0.1% bromophenol blue. 2. Pipet each mixture into an IPG strip holder (see Note 2). 3. Remove the protective cover from an 18-cm long IPG strip of pH 4–7L. Place it with gel side down in the IPG strip holder. Make sure that the acidic end of the IPG strip faces toward the pointed end of strip holder (see Note 3). 4. Dropwise add 800 μL of mineral oil to both ends of the strip holder. Tilt the strip holder to let the mineral oil cover the entire IPG strip. Close the lid. 5. Rehydrate the IPG strip on the IPGphor System at 25°C for 12 h. 6. Perform IEF at 25°C at 50 μA per strip with the following settings: 500 V for 1 h, 1000 V for 1 h, 8000 V for 8 h and, finally, hold at 500 V until electrophoresis in the second dimension.

3.2.2. Sodium Dodecyl Sulfate-Polyacrylamide Gel Electrophoresis (SDS-PAGE) 1. Prepare 1-mm thick, 19 × 23 cm, 12%T polyacrylamide gels in the Hoeffer DALT Gel Caster. Vertical slab SDS gels are cast in a Multiple Gel Caster. For assembly of the gel cassette, refer to manufacturer user manual (Hoefer DALT System User Manual, Amersham Biosciences). To cast 25 slab gels, mix the following solutions: 600 mL of Acrylamide/bis-acrylamide solution (30.8%T), 375 mL of 1.5 M Tris-Cl, pH 8.8, 15 mL of 10% SDS, and 15 mL of 10% ammonium persulfate. Add water (see Note 4) to 1500 mL. Before gel casting, add 215 μL of TEMED. 2. Equilibrate each IPG strip in 10 mL of EB-1 solution with shaking for 10 min, and then in 10 mL of EB-2 for another 10 min (see Note 5).

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3. Place the strip on top of the vertical SDS gel and seal it with 1% LMT agarose (see Note 6). 4. Perform electrophoresis overnight, under 90 V constant voltage in the Hoeffer DALT Electrophoresis Tank (Amersham Biosciences) cooled with circulating water bath at 12°C. 5. Unload the gel cassette. Make sure that the gel sticks to one of the plates when opening it. Place the gel into a glass tray containing the fixative for silver staining.

3.3. Silver Staining (19) 3.3.1. Silver Staining (See Note 7) The following steps are carried out with shaking. 1. 2. 3. 4. 5. 6. 7. 8. 9. 10.

Soak the gel in the fixative for 30 min. Change the fixative and shake for another 30 min. Wash the gel in water 5 times for 5 min each. Incubate the gel in the sensitizer for 1 min. Wash the gel in water 2 times for 1 min each. Incubate the gel in the silver reagent for 30 min. Wash the gel in water 2 times for 1 min each. Incubate the gel in the developer until protein spots appear. Stop the development with the stop reagent for 30 min. Wash the gel in water 3 times for 5 min each.

3.3.2. Capturing of 2D Image Silver-stained gels are scanned on Umax Powerlook 1100 scanner (Fig. 1) and the digitized images are saved in a computer. 3.4. Silver Gel Destaining (20) 1. 2. 3. 4. 5.

Excise the gel spots of interest. Mix 30 mM potassium ferricyanide and 100 mM sodium thiosulfate in a 1:1 ratio. Cover the gel pieces with 100 μL of the above solution. Shake at room temperature until the silver stain is removed. Wash the gel pieces with several changes of 500 μL of water each to remove the yellow reagent. 6. Incubate the gel pieces in 200 μL of 200 mM ammonium bicarbonate with shaking for 20 min. 7. Wash the gel pieces in 500 μL of water twice for 1 min. 8. Centrifuge and remove the water.

3.5. Trypsin Digestion (21,22) 1. Incubate the gel pieces in 200 μL of CH3 CN for 10 min. 2. Centrifuge and remove CH3 CN.

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Fig. 1. Representative illustration of 2D profiles of silver-stained proteins from the cytosolic fractions of AtT20 and AtT20(proSAAS) cells. Numerous proteins spots were more stained in one profile that in the other. The most prominent are indicated with arrows. Note the circled clusters of high molecular weight molecular proteins found in more abundance in AtT20(proSAAS) cells. Their abundance may be due, directly or indirectly, to reduced activity of PC1 caused by proSAAS overexpression. 3. SpeedVac the gel pieces for 15 min. 4. Incubate the gel pieces in 100 μL of 100 mM NH4 HCO3 , 10 mM DTT at 56°C for 45 min. 5. Centrifuge and remove the DTT solution. 6. Add 100 μL of 100 mM NH4 HCO3 , 55 mM iodoacetamide and incubate at room temperature in the dark for 30 min. 7. Centrifuge and remove iodoacetamide solution. 8. Wash the gel pieces in 100 μL of 100 mM NH4 HCO3 with shaking for 10 min. 9. Centrifuge and remove NH4 HCO3 solution. 10. Dehydrate the gel pieces in 100 μL of CH3 CN at room temperature for 5 min. 11. Centrifuge and remove CH3 CN. 12. Wash the gel pieces in 100 μL of 100 mM NH4 HCO3 with shaking for 5 min. 13. Centrifuge and remove NH4 HCO3 solution. 14. Dehydrate the gel pieces in 100 μL of CH3 CN at room temperature for 5 min. 15. Centrifuge and remove CH3 CN. 16. SpeedVac the gel pieces for 15 min. 17. Swell the gel pieces in 20 μL of 50 mM NH4 HCO3 containing 500 ng of trypsin on ice for 30 min. 18. Remove remaining trypsin solution from the tube and add 50 mM NH4 HCO3 without trypsin to cover gel pieces. 19. Digest proteins in gel at 37°C overnight. 20. Centrifuge and transfer the supernatant into a new tube.

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21. Add 50 μL of 50% CH3 CN, 0.1% trifluoroacetic acid to the gel pieces and sonicate in a cold water bath for 3 min. 22. Centrifuge and transfer the supernatant to the primary supernatant. 23. Repeat the steps 21 and 22 twice more. 24. SpeedVac the supernatant until the volume is smaller than 10 μL. 25. Add 0.1% trifluoroacetic acid to 10 μL.

3.6. MALDI-TOF MS of Tryptic Peptides 1. Before MALDI-TOF analysis, purify tryptic digest peptides with ZipTip C18 pipet tips following the manufacturer’s instructions. 2. Elute the purified peptides onto the MALDI plate with 2 μL of matrix solution. 3. Perform mass spectrometry in a delayed extraction and reflectron mode with the following instrument setting: 20 kV acceleration voltage, 150 ns delay time, 70% grid voltage, 0.05% guide wire voltage, 128 laser shots, laser intensity of 2000, and a mass gate from 800 to 2800. Trypsin autodigestion products of 842.51 m/z and 1045.56 m/z are used for internal mass calibration (Fig. 2). 4. The masses of monoisotopic peaks are chosen to identify the proteins by searching Swiss-Prot and TrEMBL databases using the search engine PeptIdent (http://www.expasy.ch/tools/peptident.html) with a mass match within 50 ppm to the corresponding theoretical mass.

3.7. 2D Immunoblotting 1. Repeat the 2D gels as described under Subheading 3.2. 2. Transfer proteins spots from the unstained gels onto nitrocellulose membranes in a Hoefer SE 600 tank containing the Towbin buffer, according to the manufacturer’s user manual. 3. Block the membranes with 5% skim milk solution. 4. Incubate the membranes in 5% skim milk solution containing anti-EphA2 primary antibody at room temperature for 1 h. 5. Wash the membranes in TBS-T 4 times for 5 min each. 6. Incubate the membranes in 5% skim milk solution containing HRP-conjugated donkey IgG against rabbit IgG for 1 h. 7. Wash the membrane in TBS-T 4 times for 5 min. 8. Reveal immunoreactive spots by Western Lightning Chemiluminescence reagent Plus as described in the manufacturer’s protocol (Fig. 3).

4. Notes 1. Complete solubilization, desegregation, denaturation and reduction of proteins in the sample are critical for the success of protein separation by 2D gel. Therefore, protein pellet is dissolved into RB by sonication for at least 3 min. Since urea can hydrolyze to isocyanate at elevated temperatures, resulting in protein modification

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Fig. 2. MALDI-TOF mass spectra of peptides from proSAAS and Ephrin type receptor 2 (EphA2). Proteins of cytosolic fractions of AtT-20 and AtT-20 (proSAAS) cells were resolved on 2D gel. Proteins were detected by silver staining. The protein spot indicated with an arrow was excised, digested by trypsin and analyzed by MALDI-TOF mass spectrometry. Peptides from tryptic autodigestion are marked with stars. The masses of monoisotopic peaks were searched against mouse entries in the protein databases Swiss-Prot and TrEMBL, using PepIdent search engine. The proteins were identified as proSAAS (A) and EphA2 (B) based on peptide masses and protein pI as well as protein molecular weight matched to a particular hit in the database. EphA2 is a receptor tyrosine kinase implicated in morphogenesis (23) and oncogenesis (24).

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Fig. 3. 2D immunoblotting for EphA2-related proteins. Proteins from AtT20 and AtT20(proSAAS) cells were fractionated by 2D gel. They were transferred onto nitrocellulose and probed with an antibody against the C-terminal end of mouse EphA2 (C-20, Santa Cruz Biotechnology). Immunoreactive proteins, staining with differential intensities between the two samples are indicated with white arrows. Mouse EphA2 is a 952-residue polypeptide, with a theoretical molecular mass of approx 120 kDa. Lower mass immunoreactive proteins may represent products of EphA2 processing/degradation; those with the same mass and different pIs may represent differentially phosphorylated forms of the same proteins.

2. 3. 4. 5. 6. 7.

by carbamylation, the sonication is carried out in a cuphorn sonicator cooled with an ice water bath. IPG strip holder is made up of ceramic. Handle with care. To avoid contamination, work with gloves. Do not trap bubbles under the IPG strip. Water used here and for preparing solutions must have a resistivity of more than 18 M-cm. Glass tubes, 20-cm long, with a cap are ideal for the equilibration. Avoid trapping bubbles between IPG strip and SDS gel. Silver staining is one of the most sensitive visualization methods. For high-quality results, use high-purity reagents and avoid contamination by wearing gloves.

Acknowledgments The authors thank Dr. Nabil G. Seidah for the gift of AtT20(proSAAS) cells and Dr. Ajoy Basak for his critical review of this manuscript. The work was supported by grants from the Canadian Institute of Health Research. References 1. Eipper, B. A. and Mains, R. E. (1975) High molecular weight forms of adrenocorticotropic hormone in the mouse pituitary and in a mouse pituitary tumor cell line. Biochemistry 14, 3836–3844.

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2. Bertagna, X. (1994) Proopiomelanocortin-derived peptides. Endocrinol. Metab. Clin. North Am. 23, 467–485. 3. Benjannet, S., Rondeau, N., Day, R., Chrétien, M., and Seidah, N. G. (1991) PC1 and PC2 are proprotein convertases capable of cleaving proopiomelanocortin at distinct pairs of basic residues. Proc. Natl. Acad. Sci. USA 88, 3564–3568. 4. Zhou, A., Bloomquist, B. T., and Mains, R. E. (1993) The prohormone convertases PC1 and PC2 mediate distinct endoproteolytic cleavages in a strict temporal order during proopiomelanocortin biosynthetic processing. J. Biol. Chem. 268, 1763–1769. 5. Seidah, N. G. and Chretien, M. (1999) Proprotein and prohormone convertases: a family of subtilases generating diverse bioactive polypeptides. Brain Res. 848, 45–62. 6. Marcinkiewicz, M., Day, R., Seidah, N. G., and Chrétien, M. (1993) Ontogeny of the prohormone convertases PC1 and PC2 in the mouse hypophysis and their colocalization with corticotropin and alpha-melanotropin. Proc. Natl. Acad. Sci. U SA 90, 4922–4926. 7. Schäfer, M. K., Day, R., Cullinan, W. E., Chrétien, M., Seidah, N. G., and Watson, S. J. (1993) Gene expression of prohormone and proprotein convertases in the rat CNS: a comparative in situ hybridization analysis. J. Neurosci. 13, 1258–1279. 8. Seidah, N. G., Chrétien, M., and Day, R. (1994) The family of subtilisin/kexin like pro-protein and pro-hormone convertases: divergent or shared functions. Biochimie 76, 197–209. 9. Zheng, M., Streck, R. D., Scott, R. E., Seidah, N. G., and Pintar, J. E. (1994) The developmental expression in rat of proteases furin, PC1, PC2, and carboxypeptidase E: implications for early maturation of proteolytic processing capacity. J. Neurosci. 14, 4656–4673. 10. Muller, L. and Lindberg, I. (1999) The cell biology of the prohormone convertases PC1 and PC2. Prog. Nucleic Acids Res. Mol. Biol. 63, 69–108. 11. Fricker, L. D., McKinzie, A. A., Sun, J., Curran, E., Qian, Y., Yan, L., et al. (2000) Identification and characterization of proSAAS, a granin-like neuroendocrine peptide precursor that inhibits prohormone processing. J. Neurosci. 20, 639–648. 12. Qian, Y., Devi, L. A., Mzhavia, N., Munzer, S., Seidah, N. G., and Fricker, L. D. (2000) The C-terminal region of proSAAS is a potent inhibitor of prohormone convertase 1. J. Biol. Chem. 275: 23596–23601. 13. Mbikay, M., Seidah, N. G., and Chretien, M. (2001) Neuroendocrine secretory protein 7B2: structure, expression and functions. Biochem. J. 357, 329–342. 14. Furuta, M., Yano, H., Zhou, A., Rouillé, Y., Holst, J. J., Carroll, R., et al. (1997) Defective prohormone processing and altered pancreatic islet morphology in mice lacking active SPC2. Proc. Natl. Acad. Sci. USA 94, 6646–6651. 15. Zhu, X., Rouille, Y., Lamango, N. S., Steiner, D. F., and Lindberg, I. (1996) Internal cleavage of the inhibitory 7B2 carboxyl-terminal peptide by PC2: a potential mechanism for its inactivation. Proc. Natl. Acad. Sci. USA 93, 4919–4924.

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16. Jackson, R. S., Creemers, J. W., Farooqi, I. S., Raffin_Sanson, M. L., Varro, A., Dockray, G. J., et al. (2003) Small-intestinal dysfunction accompanies the complex endocrinopathy of human proprotein convertase 1 deficiency. J. Clin. Invest. 112, 1550–1560. 17. Jean, F., Basak, A., Rondeau, N., Benjannet, S., Hendy, G. N., Seidah, N. G., et al. (1993) Enzymic characterization of murine and human prohormone convertase1 (mPC1 and hPC1) expressed in mammalian GH4C1 cells. Biochem. J. 292, 891–900. 18. Zhou, Y. and Lindberg, I. (1993) Purification and characterization of the prohormone convertase PC1(PC3). J. Biol. Chem. 268, 5615–5623. 19. Rabilloud, T., Kieffer, S., Procaccio, V., Louwagie, M., Courchesne, P. L., Patterson, S. D., et al. (1998) Two-dimensional electrophoresis of human placental mitochondria and protein identification by mass spectrometry: toward a human mitochondrial proteome. Electrophoresis 19, 1006–1014. 20. Gharahdaghi, F., Weinberg, C. R., Meagher, D. A., Imai, B. S., and Mische, S. M. (1999) Mass spectrometric identification of proteins from silver-stained polyacrylamide gel: a method for the removal of silver ions to enhance sensitivity. Electrophoresis 20, 601–605. 21. Shevchenko, A., Wilm, M., Vorm, O., and Mann, M. (1996) Mass spectrometric sequencing of proteins silver-stained polyacrylamide gels. Anal. Chem. 68, 850–858. 22. Beranova-Giorgianni, S. and Desiderio, D. M. (2000) Mass spectrometry of the human pituitary proteome: identification of selected proteins. Rapid Commun. Mass Spectro.m 14, 161–167. 23. Klein, R. (2004) Eph/ephrin signaling in morphogenesis, neural development and plasticity. Curr. Opin. Cell Biol. 16, 580–589. 24. Walker-Daniels, J., Hess, A. R., Hendrix, M. J., and Kinch, M. S. (2003) Differential regulation of EphA2 in normal and malignant cells. Am. J. Pathol. 162, 1037–1042.

8 Proteomics-Based Method for Risk Assessment of Peroxisome Proliferating Pollutants in the Marine Environment Susana Cristobal

Summary Pollution in aquatic environment is of increasing concern for its impact on both human and natural populations. Applying proteomics to monitor marine pollution is a new approach to evaluate the effects of environmental pollutants on the biota. Aquatic organisms living in coastal and estuarine areas are particularly prone to exposures to a variety of pollutants, some of which can act as peroxisome proliferators. However, peroxisomal responses in particular and biomarker responses in general can be influenced by several biotic and abiotic factors. Utilizing proteomics-based techniques that permit the evaluation of hundreds to thousands of proteins in a single experiment can circumvent those drawbacks. Applying this method, the peroxisomal proteome from digestive glands of mussels Mytilus sp. can be analyzed by two-dimensional electrophoresis (2-DE) and the 2-DE maps from control samples and samples obtained in a polluted area can be compared. The up- and down-regulated proteins compose the protein expression signature (PES) associated with exposure to peroxisome proliferating pollutants. This method generates highly reproducible patterns that can be applied to laboratory or field experiments. Key Words: Biomarker; marine pollution assessment; peroxisome; peroxisome proliferation; protein expression signature; 2-DE.

1. Introduction Biological monitoring involves the evaluation of the physiological status of sentinel organisms, such as mussels, living in the monitored environment. This is done by determining the values of selected biological parameters that are known to vary in response to the toxic effects of pollutants (1). Among the From: Methods in Molecular Biology, vol. 410: Environmental Genomics Edited by: C. Cristofre Martin © Humana Press, Totowa, NJ

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chemical compounds affecting the marine environment, it has been verified that some can induce morphological and functional changes in peroxisomes and also peroxisome proliferation (2,3). Peroxisome proliferators cause increases in the amount and number of peroxisomes as well as the induction of certain peroxisomal enzymes, especially those of the fatty acid -oxidation. It has been shown that induction of the acyl-CoA oxidase (AOX) activity occurs after exposure to peroxisome proliferating pollutants, both in laboratory and field experiments ( 2,4–6). However, one of the major obstacles encountered using any single parameter biomarker is how to distinguish polluted-related changes from “natural” variations (7). The environmental monitoring field is starting to transform from the traditional panel of single-parameter biomarkers into new methods that could sum this multivariable information. Proteomics-based techniques have been introduced in the field in an attempt to solve these problems. The first proteomic studies reported in this field focused on fish species (8,9). However, attention is now moving to the identification of a set of mussel proteins commonly expressed after exposure to toxic contaminants (10). In several laboratory experiments, the utility of protein expression signatures (PES) has been reported. This methodology has been explored in mussels exposed to copper, Aroclor, and salinity stress (11,12); in rainbow trout treated with diazinon, nonylphenol, propetamphos, and exposed to sewage treatment plant effluents (13); and in clams exposed to model pollutants (14). More recently, the use of surface-enhanced laser desorption/ionization-time of flight (SELDI-TOF) has been introduced in field and laboratory experiments to establish PES in mussels exposed to pollutants (15,16). Peroxisomal proteomics is a recently developed method that provides a complex PES and can be applied to assess marine pollution from peroxisome proliferating pollutants both in laboratory and field experiments (17,18). One additional advantage of proteomic-based methods over conventional biomarkers is that previous knowledge of pollutant mechanisms of action are not required (19). The method presented here has provided the larger number of identified proteins from PES of mussels (17,18). The identification of these proteins by mass spectrometry opens up the possibility to improve our understanding of toxicological pathways. The major procedures involved in this protocol are isolation of a peroxisomeenriched fraction by density gradient centrifugation, precipitation of peroxisomal proteins, and solubilization and separation of the peroxisomal proteins by 2-DE. The subproteomic 2-DE maps obtained can be analyzed and quantified via different software packages available in the market that permit the gel visualization, detection, and quantification of the spot proteins, gels matching, data analysis, and data integration. The proteins that change expression after exposure to a polluted environment comprised the PES.

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2. Materials 2.1. Animals Mussels, Mytilus galloprovincialis Lmk, 35–45 mm in length or Mytilus edulis, 5 cm in shell length, were collected at the same time of year at low tide in different sampling sites. One of the sampling sites corresponded to a well documented clean area, and animals from that site were used as controls. 2.2. Isolation of Peroxisome-Enriched Fraction All buffers in the fractionation process were prepared fresh and maintained in ice during the manipulations. 1. Solubilization buffer (HM): 250 M sucrose, 5 mM 3-(N-morpholino) propanesulfonic acid (MOPS), 1 mM EDTA Na2 , 0.1% ethanol, 0.2 mM phenylnethylsulfonyl fluoride (PMSF), 2 μM leupeptin, 2 μM pepstatin, 1 mM -aminocaprioic acid, 0.2 mM dithiothreitol (DTT), pH 7.2. (See Note 1.) 2. Iodixanol gradient buffer: 250 M sucrose, 30 mM MOPS, 6 mM EDTA Na2, 0.6% ethanol, pH 7.2. 3. Iodixanol solutions at 28% and 50% concentrations for the gradient are prepared with the commercial solution Optiprep (60% iodixanol) diluted with iodixanol gradient buffer.

2.3. Precipitation of Proteins 1. 20% (v/v) trichloroacetic acid (TCA) in cold acetone and 0.07% (v/v) -mercaptoethanol. Store in the –20°C freezer. 2. Cold acetone 100%. Store in the –20°C freezer.

2.4. Solubilization, Rehydration 1. Solubilization buffer: 7 M urea, 2 M thiourea, 2% (w/v) 3-((3-cholamidopropyl) dimethylammonio)-1-propanesulfonic acid (CHAPS), 0.5% Triton X-100, 1% -mercaptoethanol, 1% (v/v) Pharmalyte (3–10), 1% (w/v) dithiothreitol (DTT) (20). (See Note 2.) 2. Rehydration buffer: 8 M urea, 2% CHAPS (w/v), 15 mM DTT, 1% -mercaptoethanol, 0.2% (v/v) Pharmalyte (3–10). 3. Iodoacetamide (IAA) stock solution at 0.95 M.

2.5. Isoelectric Focusing (IEF) and Equilibration 1. Equilibration buffer (EQ): 6 M urea, 50 mM Tris, 30% glycerol (v/v), 2% (w/v) sodium dodecyl sulfate (SDS), 0.002% (w/v), Coomassie brilliant blue (CBB), pH 8.8. 2. EQ containing 1% DTT. 3. EQ containing 4% (w/v) IAA.

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2.6. Sodium Dodecyl Sulfate-Polyacrylamide Gel Electrophoresis (SDS-PAGE) These instructions assume the use of ready-made gels 12.5% Tris-HCl, type Criterion. If SDS-PAGE gels are prepared in the laboratory, 1 cm thickness is recommended. 1. 0.5% Agarose dissolved in running buffer. 2. Running buffer (5×): 125 mM Tris, 960 mM glycine, 0.5% (w/v) SDS, pH 8.3.

2.7. Colloidal Coomassie Staining 1. Fixing solution: methanol–acetic acid–water (45:1:54). 2. Staining solution: 17% ammonium sulfate, 345 methanol, 3% (v/v) O-phosphoric acid, 0.1% (w/v) Coomassie G250.

3. Methods Proteins that change in expression after exposure to a polluted environment comprise the PES obtained with this method. In field experiments, the selection of a well documented unpolluted area as a control station is essential to obtain reliable results. Ideally, the animals collected from control and polluted stations should be exposed to equivalent biotic (age, selected by shell size) and abiotic conditions (salinity, pH, temperature). 3.1. Isolation of Peroxisome-Enriched Fraction Homogenization of minced tissue and subcellular fractionation by differential and density gradient centrifugation in iodixanol is performed according to an established method (21) with a few modifications outlined below. The main subcellular fractions are termed according to the nomenclature used by Völkl and Fahimi (22). Thus, the total homogenate is termed A, the heavy mitochondrial fraction B, the light mitochondrial or enriched peroxisomal fraction D, the cytosolic fraction E, and the microsomal fraction F (Fig. 1). 1. The digestive glands from 50 mussels (M. galloprovincialis), approx 5 g, are dissected for each isolation procedure. Experiments from all sampling areas are performed in parallel. 2. Cut the tissue into small pieces into a Potter tissue grinder held in an ice bath containing 3 mL/g (wet tissue weight) of ice-cold buffer HM. 3. Homogenize the tissue at low speed with three down-and-up strokes using a loosefitting pestle and pour the homogenate into a tube to start the fractionation. 4. To remove debris, unbroken cells, and concomitantly most of the nuclei, centrifuge at 70g for 10 min in a refrigerated low-speed centrifuge.

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Fig. 1. Diagrammatic representation of the procedure used to isolate peroxisomeenriched fractions from mussels digestive glands.

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5. Remove the supernatant and resuspend the pellet in 2 mL/g of ice-cold HM and rehomogenize and centrifuge again under the same conditions. 6. Pour off the second the second supernatant and combine it with the first one, composing together the postnuclear fraction A. 7. Centrifuge the postnuclear fraction A at 1950g for 10 min in a refrigerated highspeed centrifuge. 8. Decant the supernatant and centrifuge it again at 1950g for 10 min. The final pellet contains the majority of mitochondria fraction B and the supernatant contains the postmitochondrial fraction C. 9. Subject the latter supernatant (postmitochondrial fraction C) to 39,000g for 45 min, remove the supernatant that contains cytosol and microsomes, fractions E and F; and resuspend the pellet in approx 2 mL of ice-cold HM using a glass rod. This pellet comprises the enriched heavy peroxisomes and light mitochondria fraction, termed D. 10. One milliliter of fraction D was layered carefully on the top of 6 mL of 28% (v/v) iodixanol, 5 mM MOPS, 0.1% ethanol, 1 mM tetrasodium EDTA solution (pH 7.3, density 1.16 g/mL) and 1 mL of 50% (v/v) iodixanol, 5 mM MOPS, 0.1% ethanol, 1 mM tetrasodium EDTA solution (density 1.27 g/mL) and centrifuged at 40,000 rpm (131,000gavg ) for 2 h in a Beckman L7-55 centrifuge using a TFT50.2 Ti rotor. The peroxisome-enriched fraction is obtained from the interface between 28% and 50% of iodixanol. 11. To evaluate the enrichment in peroxisomes of the fraction obtained, biochemical techniques can be applied. It is essential to essential to obtain a peroxisomeenriched fraction of the same quality from the different sampling areas. Otherwise, the possible inclusion of higher amount of contaminant proteins in any sample would interfere in the analysis. The activities of the following marker enzymes were measured across the fractionation procedure: catalase (CAT) for peroxisomes, succinate dehydrogenase (SD) for mitochondria and acidic phosphatase (AP) for lysosomes. (23). Protein concentration can be determined according to Bradford (24) or equivalent methods. In addition, protein gel blot analysis with different commercial polyclonal antisera, according to standard procedures using chemoluminescence for detection, can be performed (Fig. 2).

3.2. Precipitation of Proteins 1. To precipitate the sample, add 1 volume of 20% (v/v) TCA in cold acetone and 0.07% (v/v) -mercaptoethanol to 1 volume of sample. This solution must be prepared freshly. The final TCA concentration in the assay will be 10%. 2. Precipitate proteins for at least 45 min at –20°C. Vortex-mix every 15 min. (See Note 3.) 3. Centrifuge at 12,000g for 10 min in a refrigerated centrifuge. 4. Discharge the supernatant, and make a pulse centrifugation at approx 12,000g to remove the rest of the TCA that otherwise could alter the pH.

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Fig. 2. The enrichment of specific organelle proteins was followed by loading the same amount of protein (20 μg each) of total homogenate (lane A), mitochondrial pellet (lane B), postmitochondrial fraction (lane C), light mitochondrial fraction (lane D), cytosolic and microsomal fraction (E + F), peroxisome-enriched fraction (lane P) onto a 12.5% T polyacrylamide gel and stained by Coomassie blue. Note the distinct polypeptide pattern characterizing the corresponding subcellular compartments. CAT, catalase. In the immunoblot of CAT, it was found intensively stained in the fraction P.

5. Wash the pellet with 1 mL of acetone and 0.07% (v/v) -mercaptoethanol (freshly prepared) and vortex-mix. Break the protein pellet carefully and afterwards centrifuge at 12,000g for 10 min in the cold room. Repeat this step if the supernatant obtained is yellow. 6. Discharge the supernatant, and make a pulse of 12,000g to remove the acetone. This additional step reduces the drying time. 7. Dry open tubes at room temperature for 20–30 min. It is convenient to cover the tubes with aluminum paper or similar to avoid contamination by dust and others.

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3.3. Solubilization and Rehydration 1. Add solubilization buffer to the precipitated samples. Add approximately half of the loading volume of buffer required for isoelectric focusing (IEF). Depending on the length of the IPG strip, those are the recommended loading volume (7 cm ∼125 μL, 11 cm ∼185 μL, 18 cm ∼250 μL) + 1 μL of 0.01% (w/v) CBB. Vortex-mix at room temperature for 15 min. 2. Add 30 mM IAA from concentrated solution at 0.95 M and vortex-mix at room temperature for 15 min. This solution should be freshly prepared. 3. Add rehydration buffer (∼1/2 final loading volume) and vortex-mix at room temperature for 30 min. 4. Centrifuge at room temperature at 5000g for 10 min. 5. Measure the protein concentration. We recommend to use only 1–2 μL of sample by applying any micro-method such as Bradford (24) or Smith (25). 6. If the sample will not be loaded immediately, keep it at 4o C.

3.4. IEF and Equilibration These instructions assumed the use of a Protean IEF cell (Bio-Rad). 1. Load the supernatant into the IEF tray and place the commercial Immobiline dry (IPG) strip on top, ensuring that there are no air bubbles under the strip. 2. Methods for IEF: For 11-cm IPG strip: passive rehydration 12–15 h, rapid voltage slope. Step 1: 250 V for 15 min Step 2: 8000 V for 2 h 30 min Step 3: 35,000 V · h at 8000 V (or until 45,000 V · h at 8000 V if necessary) After 1 of rehydration add mineral oil to cover the IPG strip. 3. After IEF, the IPG strip can be stored at –20°C, or equilibrated immediately. 4. Place the IPG strip in the equilibration buffer containing 1% DTT for 15 min at room temperature with shaking. 5. Place the IPG strip in a new equilibration buffer containing 4% (w/v) IAA for 15 min at room temperature with shaking. Prepare fresh IAA for the experiment.

3.5. SDS-PAGE These instructions are based on using Criterion SDS-PAGE cell or Dodeca cell and Criterion gels (Bio-Rad). (See Notes 4 and 5.) 1. Place the IPG strip on the SDS-PAGE gel, ensuring there are no air bubbles at the interface. 2. Add 0.5% agarose dissolved in the running buffer, to the top of the gel to seal the strip. 3. The process is run in the cold room, using magnetic stir bars to maintain the same temperature in the running buffer. The temperature of the running buffer should be room temperature or lower.

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Fig. 3. Fractions enriched in peroxisomes were obtained and proteins from these fractions were run by 2-DE. In this example two representative 2–DE maps from a control area and from a polluted area are shown. The gels were scanned and analyzed by an image analysis program called Image Master Platinum. This program detects the spots from the gel and determines their optical density values, which are in accordance with the protein concentration. These values are changed later to the spot-volume values. Finally, there is a normalization process that determines the volume percentage of each spot from the map, from the total volume of spots. The vol % values are the ones used in the next steps for the comparison of the different 2-DE maps obtained from the different stations. In this figure, vol % for two representative spots is depicted. The spot marked with a circle on the gel left side corresponded to a protein downregulated 2.6-fold and the example from the gel right side corresponded to a protein up-regulated 3-fold after exposure to the polluted environment. The three-dimensional reports represents the real “visual” difference because the scale is identical for all the spots.

4. For 11-cm SDS-PAGE gels run at 120 V until the CBB has reached the bottom of the gel. For a 18-cm gel, constant current will be used; for the first 30 min, at 16 mA, then change to 24 mA until the CBB mark reaches the bottom.

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Fig. 4. The general schema of a protein expression signature obtained in M. edulis is summarized in an artificial 2-DE map. The up-regulated spots were marked in grey and the down-regulated spots in black.

3.6. Colloidal Coomassie Staining 1. 2. 3. 4.

Fixing the gel for 30 min in fixing solution. Stain for 12–18 h in colloidal Coomassie solution. Destain in methanol for 2–3 min. Destain in water for 12 h or until the background is clear. It is good to change water regularly, so the background removal increases. Store in the cold room until the gel is scanned and analyzed or spots are picked for mass spectrometry analysis (Figs. 3 and 4).

3.7. Image Acquisition and Analysis 1. The gel images can be obtained using Image Scanner (Amersham Biosciences). 2. The data are analyzed using Image Master 2D Platinum 6.0 from Amersham Biosciences or other software available in the market. 3. Image analysis included spot detection, spot quantification and normalization, background subtraction, and spot matching, followed by statistical analysis. 4. The amount of protein per spot was expressed as the volume of the spot that is defined as sum of the intensities of all the pixels that make up the spot. 5. To correct the variability due to CBB staining and to reflect the quantitative variations between spots, the spot volumes are normalized as a percentage of the total

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volume of all spots in the gel. Thereby % values for all protein-spots generated are evaluated for significant differences between groups. The 2-DE maps from polluted stations are analyzed by matching to the reference 2-DE map (uncontaminated station). To focus on the spots that are more drastically altered (up- or downregulated), statistical two-sample t-test are utilized, and so only 95% or higher significances between control and polluted stations are taken into consideration.

Notes 1. Leupeptin, pepstatin, -aminocaproic acid, and DTT stocks solution are dissolved in dH2 O. PMSF is a hazardous material. Prepare a stock solution in isopropanol at 100 m M concentration. 2. To prepare the solubilization and hydration buffers, it is recommended to dissolve the urea in a small amount of MilliQ water at 37°C and add thiourea afterwards. The urea–thiourea solution is washed with Amberlite ion exchanger beads 10 mg/mL solution. Vortex-mix, allow to stand for 10 min, and remove the supernatant with the small tip. 3. In the precipitation of proteins by TCA, different yields will be obtained depending on the protein concentration. The best results are obtained if the concentration of the protein sample is approx 1 mg/mL. 4. Running SDS-PAGE. To ensure that there are no air bubbles at the interface between IPG strip and the gel, an alternative method is to pour the agarose solution in the well and introduce slowly the IPG strip on top of it. 5. Running SDS-PAGE. If the running buffer is very cold, the process will slow very much, and if very warm, the gel running pattern gel could appear discontinuous.

Acknowledgments This work was partially supported by grants from the Swedish Research Council and Carl Trygger foundation, Magnus Bergvalls foundation. I thank Prof. Miren Cajaraville and her research group for the collaboration. References 1. Shugart, L. R., Mccarthy, J. F., and Halbrook, R. S. (1992) Biological markers of environmental and ecological contamination: an overview. Risk Anal 12, 353–360. 2. Cajaraville, M. P., Cancio, I., Ibabe, A., and Orbea, A. (2003) Peroxisome proliferation as a biomarker in environmental pollution assessment. Microsc. Res. Tech. 61, 191–202. 3. Fahimi, H. D., and Cajaraville, M. P. (1995) Induction of peroxisomal proliferation by some environmental pollutants and chemicals in animal tissues. Cell Biology in Environmental Toxicology, Vol. (Cajaraville, M. P., ed.), University of Basque Country Press Service, Bilbo, Spain, pp. 221–225.

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4. Cancio, I., Orbea, A., Volkl, A., Fahimi, H. D., and Cajaraville, M. P. (1998) Induction of peroxisomal oxidases in mussels: comparison of effects of lubricant oil and benzo(a)pyrene with two typical peroxisome proliferators on peroxisome structure and function in Mytilus galloprovincialis. Toxicol. Appl. Pharmacol. 149, 64–72. 5. Porte, C., Biosca, X., Sole, M., and Albaiges, J. (2001) The integrated use of chemical analysis, cytochrome P450 and stress proteins in mussels to assess pollution along the Galician coast (NW Spain). Environ. Pollut. 112, 261–268. 6. Cajaraville, M. P., and Pal, S. G. (1995) Morphofunctional study of the haemocytes of the bivalve mollusc Mytilus galloprovincialis with emphasis on the endolysosomal compartment. Cell Struct. Funct. 20, 355–367. 7. Cajaraville, M. P., Bebianno, M. J., Blasco, J., Porte, C., Sarasquete, C., and Viarengo, A. (2000) The use of biomarkers to assess the impact of pollution in coastal environments of the Iberian Peninsula: a practical approach. Sci. Total. Environ. 247, 295–311. 8. Martínez, I., Solberg, C., Lauritzen, K., and Ofstad, R. (1992) Two-dimensional electrophoretic analysis of cod (Gadus morhua, L.) whole muscle proteins, watersoluble fraction and surimi. Effect of the addition of CaCl2 and MgCl2 during the washing procedure. Appl. Theor. Electrophor 2, 201–206. 9. Piñeiro, C., Vazquez, J., Marina, A. I., Barros-Velazquez, J., and Gallardo, J. M. (2001) Characterization and partial sequencing of species-specific sarcoplasmic polypeptides from commercial hake species by mass spectrometry following twodimensional electrophoresis. Electrophoresis 22, 1545–1552. 10. Aardema, M. J., and Macgregor, J. T. (2002) Toxicology and genetic toxicology in the new era of toxicogenomics”: impact of “-omics” technologies. Mutat. Res. 499, 13–25. 11. Shepard, J. L., and Bradley, B. P. (2000) Protein expression signatures and lysosomal stability in Mytilus edulis exposed to graded copper concentrations. Mar. Environ. Res. 50, 457–463. 12. Shepard, J. L., Olsson, B., Tedengren, M., and Bradley, B. P. (2000) Protein expression signatures identified in Mytilus edulis exposed to PCBs, copper and salinity stress. Mar. Environ. Res. 50, 337–340. 13. Bradley, B. P., Shrader, E. A., Kimmel, D. G., and Meiller, J. C. (2002) Protein expression signatures: an application of proteomics. Mar. Environ. Res. 54, 373–377. 14. Rodriguez-Ortega, M. J., Grosvik, B. E., Rodriguez-Ariza, A., Goksoyr, A., and Lopez-Barea, J. (2003) Changes in protein expression profiles in bivalve molluscs (Chamaelea gallina) exposed to four model environmental pollutants. Proteomics 3, 1535–1543. 15. Knigge, T., Monsinjon, T., and Andersen, O. K. (2004) Surface-enhanced laser desorption/ionization-time of flight-mass spectrometry approach to biomarker discovery in blue mussels (Mytilus edulis) exposed to polyaromatic hydrocarbons and heavy metals under field conditions. Proteomics 4, 2722–2727. 16. Manduzio, H., Cosette, P., Gricourt, L., Jouenne, T., Lenz, C., Andersen, O. K., Leboulenger, F., and Rocher, B. (2005) Proteome modifications of blue

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mussel (Mytilus edulis L.) gills as an effect of water pollution. Proteomics 5, 4958–4963. Mi, J., Orbea, A., Syme, N., Ahmed, M., Cajaraville, M. P., and Cristobal, S. (2005) Peroxisomal proteomics, a new tool for risk assessment of peroxisome proliferating pollutants in the marine environment. Proteomics 5, 3954–3965. Apraiz, I., Mi, J., and Cristobal, S. (2006) Identification of proteomics signatures of exposure to marine pollutants in mussels (Mytilus edulis). Mol. Cell Proteomics 5, 1274–1285. Kennedy, S. (2002) The role of proteomics in toxicology: identification of biomarkers of toxicity by protein expression analysis. Biomarkers 7, 269–290. Gorg, A., Obermaier, C., Boguth, G., Harder, A., Scheibe, B., Wildgruber, R., and Weiss, W. (2000) The current state of two-dimensional electrophoresis with immobilized pH gradients. Electrophoresis 21, 1037–1053. Ghosh, M. K., and Hajra, A. K. (1986) A rapid method for the isolation of peroxisomes from rat liver. Anal. Biochem. 159, 169–174. Volkl, A., and Fahimi, H. D. (1985) Isolation and characterization of peroxisomes from the liver of normal untreated rats. Eur. J. Biochem. 149, 257–265. Graham, J. M., and Higgens, J. A. (1993) Methods in Molecular Biology, Vol 19, (Graham, J. M. H., J. A., ed.), Humana Press, Totowa, NJ, pp. 1–28.. Bradford, M. M. (1976) A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal. Biochem. 72, 248–254. Smith, P. K., Krohn, R. I., Hermanson, G. T., Mallia, A. K., Gartner, F. H., Provenzano, M. D., Fujimoto, E. K., Goeke, N. M., Olson, B. J., and Klenk, D. C. (1985) Measurement of protein using bicinchoninic acid. Anal. Biochem. 150, 76–85.

9 Environmental Metabolomics Using 1 H-NMR Spectroscopy Mark R. Viant

Summary Environmental metabolomics is a subdiscipline of metabolomics and focuses on the study of metabolic changes in organisms in response to environmental challenges. This approach is ideal for studying multiple species within an ecosystem because it is not dependent on knowledge of an organism’s genome. Unbiased measurements of an organism’s metabolic composition can in principle be used to identify novel biomarker profiles and modes of action of stressors. This chapter presents protocols for the extraction of metabolites from biological samples, the measurement of metabolites using 1 H nuclear magnetic resonance (NMR) spectroscopy, and finally the analysis of the metabolic data using multivariate statistical methods. First, the preparation of biofluids (e.g., blood and urine) for NMR analysis is described together with a methanol–chloroform protocol for extracting metabolites from tissue samples. Next the NMR methods are presented, comprising a standard one-dimensional (1-D) 1 H-NMR method and a two-dimensional (2-D) 1 H-1 H J-resolved NMR experiment. The advantages and limitations of each method are discussed. Finally, two methods for analyzing the multivariate metabolic NMR data are presented. These include a traditional fingerprinting approach that comprises of a spectral preprocessing step followed by multivariate statistical analysis. Although reliable and proven, this method often produces results in terms of unidentified metabolites that are of limited value to the biologist. The second and newer method is based on metabolic profiling in which NMR spectra are deconvoluted into a list of metabolites and their concentrations. Although more biologically insightful, this latter approach can prove labor intensive. Key Words: Fingerprinting; J-resolved; metabolomics; methanol–chloroform; NMR; p-JRES; profiling; ProMetab; tissue extract.

From: Methods in Molecular Biology, vol. 410: Environmental Genomics Edited by: C. Cristofre Martin © Humana Press, Totowa, NJ

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1. Introduction Metabolomics is the study of low molecular weight, endogenous metabolites in biological systems including cells, tissues or biofluids (1–3). These metabolites are the products of metabolism and include carbohydrates, lipids, amino acids, and many other compounds. The collection of all metabolites within a cell is called the metabolome. Environmental metabolomics is a subdiscipline of metabolomics and focuses on the study of metabolic changes in organisms in response to environmental challenges (4). This approach is ideal for studying the impact of stressors such as pollution, not least because no species-specific DNA sequence information is required. The application of metabolomics to ecotoxicology also shares many of the advantages that are exploited by transcriptomics and proteomics including unbiased measurement of extensive molecular profiles that can facilitate biomarker discovery and identify modes of action. A further critical advantage, which is specific to metabolomics, is the link between metabolism and organism physiology, i.e., the metabolic phenotype can provide a direct reflection of the physiology of the organism. For example, ATP and glycogen levels inform directly on energetic status, while glutathione and ascorbate are indicators of oxidative status. Some potential applications for environmental metabolomics include characterizing the effects of stressors such as diseases and/or pollutants on both terrestrial (5–8) and aquatic organisms (4–10). In addition, metabolomics will likely have a major impact in the environmental risk assessment of pharmaceuticals, pesticides, and other household and industrial chemicals (11–13). There are three main components to a metabolomics experiment comprising sample collection and preparation, the measurement of metabolites, and analysis of the metabolic data. The methods in this chapter are based around this structure. Metabolites are present in biofluids, such as blood and urine, as well as in tissues. The choice of sample is dependent on the biological question being asked, but is often dictated by the design of the experiment (e.g., the size of the organism). Biofluids are in general easier to collect than tissue samples, and facilitate time-course studies of changes in metabolism within individual organisms. The metabolic information obtained from a biofluid is effectively an integration of the metabolic changes occurring within each of the animal’s organs and can be strongly influenced by the environment. It is in principle an ideal sample for obtaining information on the overall metabolic condition of an organism. This advantage, however, can also be a disadvantage in that biofluid metabolite concentrations can vary so dramatically between individuals that statistically significant differences between stressed and unstressed organisms are hard to achieve with realistic numbers of samples. This is less of a problem with blood (vs. urine) as it is under greater homeostatic regulation. Tissues are in general under even greater homeostatic regulation than biofluids and can

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provide highly consistent metabolic measurements between individuals, with the disadvantage that they are less sensitive to environmental change. Methods to prepare biofluids and tissue extracts for metabolomics are described in Subheading 3.1. Although the protocols for tissues and biofluids differ, there are a number of considerations that are important to both. In particular, as soon as samples are collected they need to be frozen and stored at –80ºC to prevent metabolic decay. Because high-resolution NMR spectroscopy requires samples to be in a liquid state, biofluids require minimal sample preparation. Tissue samples, however, require the extraction of the metabolites into an appropriate solution. The two most widely used analytical techniques in metabolomics are 1 H-NMR spectroscopy and mass spectrometry (1,2,14). In Subheading 3.2 we focus on two NMR methods that have been used extensively in environmental metabolomics. The benefits of an NMR approach include: (1) observation of all high abundance metabolites that contain nonexchangeable hydrogen atoms; (2) potentially quantitative metabolite measurements with a high degree of reproducibility; (3) relatively high throughput and automated analyses; and (4) robust and established NMR technology with minimal instrument downtime. The greatest disadvantage of NMR spectroscopy is its relatively poor sensitivity, which limits the observation to an estimated hundred or so metabolites per sample. This likely accounts for less than 10% of an organism’s metabolome. Mass spectrometry has significantly higher sensitivity and hence can complement NMR measurements by facilitating the analysis of less abundant metabolites. To date, however, extremely few mass spectrometry based environmental metabolomic studies have been reported, which reflects the nascent state of this field. In summary, NMR-based metabolomics provides a rapid, unbiased, top-down (or screening) approach that can provide insight into a broad array of metabolism. Two strategies have evolved for the analysis of NMR metabolic data. The traditional method, termed fingerprinting, is based on the analysis of an intact NMR spectrum that can be considered a “fingerprint” of unassigned peaks arising from low molecular weight metabolites. This requires an initial preprocessing of the NMR fingerprint to convert it to an appropriate format for multivariate statistical analysis. Multivariate methods such as principal components analysis (PCA) or partial least squares regression (PLS) can be conducted to identify similarities and differences between the NMR fingerprints. A plethora of other algorithms exist for multivariate analyses (15–17), but most have the common goals of constructing classification models and/or biomarker discovery. The second and newer strategy for analyzing NMR data, termed profiling, uses computational methods to deconvolute each NMR spectrum into a list of metabolites and their concentrations. After deconvolution the

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same PCA and PLS multivariate methods of analysis can be applied directly to these data. The advantage of this approach is that the data is considerably more meaningful to a biochemist as it consists of metabolite identities instead of unassigned peaks. In Subheading 3.3. we provide an introduction to both approaches. 2. Materials 2.1. Tissue Collection and Storage 1. Liquid nitrogen and a benchtop dewar.

2.2. Bioﬂuid Collection and Storage 1. Liquid nitrogen and a benchtop dewar. 2. Heparinized blood collection tube (e.g., BD Vacutainer Heparin Tubes).

2.3. Combined Extraction of Polar and Lipophilic Metabolites from Tissue Using Methanol– Chloroform 1. Straight-sided glass vials with aluminum lined black urea screw caps, including both “large” (height × diameter of 46 mm × 12.5 mm) and “small” (36 mm × 11 mm) vials (Fisher Scientific). 2. Methanol (HPLC grade; Fisher Scientific): place on ice during procedure. 3. Chloroform (pesticide analysis grade; Fisher Scientific): place on ice during the procedure.

2.4. Preparation of Bioﬂuids and Polar Tissue Extracts for NMR Spectroscopy 1. NMR buffer: sodium phosphate buffer (composed of NaH2 PO4 and Na2 HPO4 salts; Fisher Scientific) at pH 7.0, made up in D2 O (99.9% purity; Goss Scientific Instruments, Great Baddon, UK), containing 1 mM sodium 3-(trimethylsilyl)proprionate2,2,3,3-d4 (TMSP; 98% purity; Goss Scientific Instruments) as an internal chemical shift standard. Store at room temperature in a desiccator. 2. Norell 5-mm NMR tubes and caps.

2.5. Preparation of Lipophilic Tissue Extracts for NMR Spectroscopy 1. Deuterated NMR solvent: 2:1 mixture of chloroform-d (CDCl3 ; 99.8% purity; Goss Scientific Instruments) and methanol-d4 (CD3 OD; 99.8% purity; Goss Scientific Instruments), containing 0.5 mM tetramethylsilane (TMS; 99.9% purity; Goss Scientific Instruments) as an internal chemical shift standard. Store at room temperature in a desiccator. 2. Norell 5-mm NMR tubes and caps.

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2.6. High-Resolution 1 H NMR Spectroscopy 1. 500-MHz or 600-MHz NMR spectrometers are typically used for metabolomics. 2. A conventional NMR probe is acceptable, although the elevated sensitivity provided by an NMR cryo- or cold-probe is a definite advantage.

3. Methods 3.1. Sample Collection and Preparation The preparation of biological samples for metabolomics involves three principal steps. The first is to rapidly collect and then freeze the sample so as to quench metabolism and preserve the metabolite concentrations (see Subheadings 3.1.1. and 3.1.2.). Samples should then be stored at –80ºC to prevent any metabolic decay (18). The second step is required only for tissue samples and involves mechanically disrupting the tissue in the presence of a solvent to extract the low molecular weight metabolites and to remove the proteins (see Subheading 3.1.3.). Although several extraction methods could be used, here we recommend a methanol–chloroform procedure that separates the polar and lipophilic metabolites into two fractions (19). The final step is to optimize the solution for high-resolution NMR spectroscopy. For biofluids and polar tissue extracts this entails buffering the sample pH (to minimize variation in the chemical shifts of the NMR resonances), adding D2 O (to provide a frequency lock for the NMR spectrometer), and adding an NMR chemical shift standard (see Subheading 3.1.4.). For lipophilic tissue extracts, this requires addition of deuterated solvents for frequency lock and addition of a chemical shift standard (see Subheading 3.1.5.). 3.1.1. Tissue Collection and Storage 1. Rapidly dissect the tissue (ideally 100 mg wet mass, although the methods presented here have successfully been applied to 20 mg of tissue) and then immediately freeze in liquid nitrogen to quench metabolism. Transfer the sample to a labeled cryovial and return to liquid nitrogen or place in dry ice. (See Note 1.) 2. Samples should always be maintained at –80ºC (or colder) either in liquid nitrogen, dry ice or a freezer. Long-term storage in a –80ºC freezer is typically most convenient. (See Note 2.)

3.1.2. Bioﬂuid Collection and Storage 1. Collect the biofluid (ideally 250–500 μL) using an appropriate method, for example with a syringe and needle. 2. For blood, transfer each sample into a heparinized blood collection tube. (See Note 3.) Centrifuge the tube to remove the cells and then transfer the plasma sample to a cryovial. Freeze the vials in liquid nitrogen or dry ice.

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3. For urine, transfer each sample directly into a cryovial and then freeze in liquid nitrogen or dry ice. 4. Samples should be stored at –80ºC (or colder).

3.1.3. Combined Extraction of Polar and Lipophilic Metabolites from Tissue Using Methanol–Chloroform 1. Label four glass vials (three small and one large) for each tissue sample. 2. Remove the tissue samples from the freezer and place on dry ice. Weigh each sample rapidly (ideally 100 mg), taking care not to let them thaw. 3. Add 4 mL/g (wet weight) ice-cold methanol and 0.85 mL/g ice-cold de-ionized water to a large glass vial, add the first tissue sample, and then homogenize for 5–10 s using a Polytron homogenizer. 4. Add 2 mL/g of ice-cold chloroform to the homogenized sample, vortex-mix for 30 s, and then place on ice. Repeat this process for all the samples. 5. Shake all the samples, on ice, using an orbital shaker for 10 min. All solutions will be monophasic. (See Note 4.) 6. Centrifuge the samples at 1800g, for 5 min, at 4°C. Transfer each supernatant to a small glass vial. 7. Add 2 mL/g of chloroform and 2 mL/g of deionized water to each sample (see Notes 4 and 5) and vortex-mix for 30 s. 8. Centrifuge the samples at 1800g, for 10 min, at 4°C. The solutions will separate into an upper methanol–water phase (with polar metabolites) and a lower chloroform phase (with lipophilic compounds) separated by a thin layer of cellular debris. 9. Using two Hamilton syringes with metal needles, transfer the upper and then lower layers of each sample into separate small glass vials. (See Note 6.) 10. Remove the solvents from all samples using a speed vacuum concentrator and then store at –80 °C until required.

3.1.4. Preparation of Bioﬂuids and Polar Tissue Extracts for NMR Spectroscopy 1. For the dried polar tissue extracts (from the methanol phase in Subheading 3.1.3.), resuspend the samples in 550 μL of NMR buffer, and vortex-mix for 10 s (see Note 7). In this particular case, the NMR phosphate buffer concentration should be 100 mM. 2. For the biofluids, mix 300 μL of each sample with 300 μL of the NMR buffer, and vortex-mix for 10 s. In this particular case, the original NMR phosphate buffer concentration should be 200 mM such that after dilution the concentration is 100 mM. 3. Centrifuge at 12,000-g, for 5 min, at room temperature. 4. Transfer 520 μL into each labeled NMR tube.

3.1.5. Preparation of Lipophilic Tissue Extracts for NMR Spectroscopy 1. Resuspend the lipophilic tissue extracts (from the dried chloroform phase in Subheading 3.1.3.) in 550 μL of deuterated NMR solvent, and vortex-mix for 10 s.

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2. Centrifuge at 1000g, for 5 min, at room temperature. 3. Transfer 520 μL into each labeled NMR tube.

3.2. High-resolution 1 H-NMR Spectroscopy NMR spectrometers are specialized analytical instruments that require significant operator training before use. The methods described here assume a basic working knowledge of NMR spectroscopy. While a number of companies market NMR systems, including Bruker BioSpin, JEOL and Varian, the methods described below have been optimized using Bruker spectrometers (for which the author has most experience). These methods, however, are directly applicable to other NMR systems. Subheadings 3.2.1. and 3.2.2. describe specific one-dimensional (1-D) and two-dimensional (2-D) NMR experiments used in environmental metabolomics, including the advantages and limitations of each approach. The general optimization strategy for a generic sample before data collection is described in Subheading 3.2. (See Note 8.) 1. Load the 5-mm NMR tube into the spectrometer. 2. Set the sample temperature to 300 K and allow the solution a couple of minutes to thermally equilibrate within the spectrometer. (See Note 8.) 3. Tune and match the NMR probe. 4. Lock the spectrometer frequency to the deuterium resonance arising from the NMR solvent (either to D2 O when analyzing biofluids or polar tissue extracts, or to CDCl3 for nonpolar tissue extracts). 5. Shim the sample preferably using automated methods. (See Note 9.) 6. Determine the pulse duration for a 360º tip angle of a peak in the NMR spectrum, from which 60º and 90º tip angles can easily be calculated. 7. Determine the frequency of the water resonance and set the center of the spectrum to this frequency.

3.2.1. Standard 1-D 1 H-NMR Experiment The 1-D 1 H-NMR experiment is a standard pulse sequence used in metabolomics (4). The primary advantages include a rapid and relatively sensitive acquisition of a metabolic fingerprint in typically 3–10 min per sample. The spectrum produced will often, however, be composed of highly overlapping peaks and the baseline may contain several broad resonances from high molecular weight compounds (Fig. 1). A number of modifications to this method may be necessary, dependent on the sample, as described below. 1. Acquisition parameters: pulse sequence comprising [relaxation delay–60º–acquire], where the pulse power is set to achieve a 60º tip angle; 7 kHz spectral width; 2.5-s relaxation delay; typically 40–160 transients are collected into 32K data points; water suppression dependent upon the nature of the sample (see Note 10); optional T2 -spectral editing, again dependent on the type of sample (see Note 11).

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Fig. 1. Representative 1-D 1 H-NMR spectrum of the polar metabolites extracted from red abalone (Haliotis rufescens) foot muscle. The several hundred NMR peaks are estimated to arise from approx 100 low molecular weight endogenous metabolites.

2. Processing parameters: zero-fill to 64K data points; apply exponential line broadening of 0.5 Hz; apply Fourier transformation; manually phase spectrum (zero- and first-order corrections); manually correct the baseline using a polynomial function; calibrate the spectrum by setting TMSP or TMS peak to 0.0 ppm. 3. Record and process the spectrum using the parameters above.

3.2.2. 2-D 1 H-1 H J-Resolved NMR Experiment The 2-D 1 H-1 H J-resolved (JRES) NMR sequence can be used to generate a considerably less congested metabolic fingerprint, which is effectively a “proton broad-band decoupled” 1-D 1 H-NMR spectrum (termed p-JRES) (20). This is achieved by projecting the JRES spectrum, which itself is composed of chemical shift (F2) and spin–spin coupling (F1) axes, along the F1 axis. The reduced spectral congestion in the 1-D p-JRES spectrum increases the likelihood that a specific metabolite will appear as a well-resolved and identifiable peak, thereby maximizing the extraction of metabolic information from each spectrum. In addition, the p-JRES spectrum has a flat baseline (as a result of T2 -editing) and provides spin–spin coupling data that can aid metabolite identification. This approach is therefore the preferred method for NMR metabolomics (20). However, it requires longer acquisition times of typically 10–20 min.

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1. Acquisition parameters: pulse sequence comprising (relaxation delay–90º–[t1 /2]– 180º–[t1 /2]–acquire], where t1 is an incremented time delay; 7 kHz spectral width in F2 (chemical shift axis), and 50 Hz in F1 (spin–spin coupling constant axis); 3.5-s relaxation delay; typically 8 transients per increment for 32 increments in total are collected into 8K data points; water suppression dependent on the nature of the sample. (See Note 10.) 2. Processing parameters: zero-fill to 128 data points in F1 and to 16K points in F2; multiply both F1 and F2 dimensions by unshifted sine-bell window functions; apply double complex Fourier transformation; tilt the spectrum by 45º; symmetrize; calibrate the spectrum by setting TMSP or TMS peak to 0.0 ppm. 3. Record and process the spectrum using the parameters above. 4. Calculate the 1-D skyline projection (p-JRES) of the 2-D spectrum.

3.3. Analysis of NMR Metabolomics Data Two related strategies exist for the analysis of NMR metabolic data. These can be differentiated into a traditional method called fingerprinting (Subheading 3.3.1.) and a newer approach called profiling (Subheading 3.3.2.). 3.3.1. Fingerprinting Based Analysis of NMR Metabolic Data Fingerprinting is based on the multivariate analysis of whole NMR spectra, where each spectrum can be considered a “fingerprint” of unassigned peaks arising from low molecular weight metabolites in the biological sample. 1. Spectral preprocessing: this is the first step in fingerprinting and is used to convert 1-D NMR spectra (or 1-D p-JRES projections of 2-D NMR spectra) into a format for multivariate analysis. An increasing number of commercial and free software packages now exist for spectral preprocessing. We use software developed in our own laboratory using MATLAB (The MathWorks, Natick, MA) called ProMetab—Processing Metabolite profiles derived from NMR spectra (20). This software is freely available to the scientific community. ProMetab converts raw Bruker NMR spectra into a format for multivariate analysis by segmenting the spectra into chemical shift bins of a user-defined width. Following removal of unwanted spectral features, such as the residual water resonance, specific groups of bins can be compressed into single segments to minimize the effects of pHinduced shifting of the NMR peaks. Various normalization strategies are available, as is the generalized log transformation (21). 2. Multivariate statistical analysis: analysis of the pre-processed data can be performed using a commercial software package. The usual objective of such an analysis is to identify similarities and/or differences between multiple NMR fingerprints, which are typically represented using a scores plot (Fig. 2a). If meaningful differences between the spectra are discovered, for example, between a control and a diseased group of samples, an associated loadings plot (see Fig. 2b)

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Fig. 2. (a) Scores plot from a principal components analysis of multiple 1-D 1 HNMR spectra of red abalone foot muscle. Three groups of animals were included in the study comprising healthy abalone (), diseased animals (), and abalone showing stunted growth (•). The ellipses represent the mean ± SD (along PC1 and PC2) of each of the three groups. The tight grouping of the healthy animals confirms that their metabolic fingerprints are similar. The diseased abalone, however, have an altered metabolic fingerprint, and separate from the healthy animals along the PC1 axis. (b) The PC1 loadings plot from the same principal components analysis which indicates the metabolic differences between the healthy and diseased abalone (in terms of unidentified NMR peaks). Peaks with positive loadings arise from metabolites that are at elevated concentration in the diseased abalone, and vice versa.

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can be used to determine which peaks in the NMR spectra are producing this difference (i.e., biomarker discovery). A convenient strategy for multivariate analysis is to do this within MATLAB using, for example, PLS_Toolbox (Eigenvector Research, Manson, WA). This software package contains many chemometric routines including principal components analysis (PCA) and partial least squares (PLS) regression. Multivariate analyses are beyond the scope of the current chapter, but interested readers are advised to consult refs. 16,17. 3. Having determined which peaks in the NMR spectra appear to differentiate the groups of samples, the next task is to identify which metabolites give rise to these peaks. Metabolites are typically identified via comparison of peak positions to tabulated chemical shifts (e.g., 22) and then confirmed by 2-D NMR experiments such as 1 H–1 H correlation spectroscopy (COSY) and 1 H–13 C heteronuclear single quantum coherence (HSQC). Although relatively labor intensive (in fact this is currently a major rate-limiting step in NMR metabolomics), until the peaks are identified the data lack biochemical meaning and is therefore of little value to the biologist.

3.3.2. Proﬁling Based Analysis of NMR Metabolic Data 1. Profiling is a computational more challenging approach for analyzing NMR data, and is based on the deconvolution of the peaks in each NMR spectrum into a list of metabolites and their concentrations. Because the chemical shift of a peak in an NMR spectrum is subject to fluctuate based on the sample pH, temperature and other “matrix effects,” it is currently impossible to accurately identify and quantify the metabolites in a 1-D NMR spectrum using fully automated methods. Some amount of human intervention is required to guide the deconvolution process. One of the leading tools for the profiling based analysis of 1-D NMR metabolic data is the Chenomx NMR Suite software (Chenomx, Edmonton, Canada), which currently uses a library of 240 metabolite spectra to identify and quantify metabolites in biological fluids. 2. After spectral deconvolution the same multivariate statistical methods of analysis (e.g., PCA and PLS) used in Subheading 3.3.1. can be applied directly to these data. The advantage of this profiling approach is that the results of the multivariate analyses are considerably more meaningful to the biologist as it consists of metabolite identities instead of unassigned peaks. This is clearly more useful information and can be used for biomarker discovery, characterizing effects on specific metabolic pathways, and for instrument-independent data archiving.

4. Notes 1. When dissecting many samples it is fastest to place the dissected tissue directly in a cryovial and then freeze that vial immediately in liquid nitrogen. 2. Samples can be stored for many months. For example, plasma samples that were snap-frozen and then maintained at –80ºC showed no significant metabolic changes after storage for up to 9 months (18).

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3. Do not use EDTA, as the anticoagulant as it will generate interfering peaks in the NMR spectrum. 4. Bligh and Dyer (23) determined that the volumes of solutions should have the following ratios: monophasic solution for metabolite extraction, 2:1:0.8 methanol– chloroform–water, and the biphasic solution for separating the fractions, 2:2:1.8 methanol–chloroform–water. The total volume of water includes both the water already present in the tissue plus the added de-ionized water. 5. To partition better the polar (phospho)lipids into the chloroform layer the polarity of the methanol phase can be increased by replacing the water with 0.8% KCl aqueous solution. 6. All sample handling and storage must be conducted in glass because chloroform can leach compounds from plastic tubes and pipet tips, which can contaminate the NMR spectra. 7. For the extracts derived from the methanol–chloroform extraction only, after vortex-mixing transfer the sample from the glass vial to an Eppendorf tube. 8. Ideally, within a particular metabolomics study, all the samples should be consistent in terms of the extraction method, the NMR buffer, and the volume of solution in the NMR tube. In this case, minimal optimization of each sample in the NMR spectrometer is required. Specifically, only steps 1, 2, 4, and 5 are required, and the quality of optimization can be checked using the criteria in Note 9. 9. The quality of shimming, as measured using the full-width at half-maximum (FWHM) of the TMSP or TMS peak, should be less than 2.0 Hz (before any line broadening induced by apodization). 10. For samples that comprise 99–100% deuterated solvents (e.g., from the tissue extractions), basic water presaturation can be used to suppress the residual water resonance. For biofluid samples that contain a large percentage of H2 O, however, more powerful water suppression techniques must be used such as excitation sculpting. 11. If the standard 1-D 1 H-NMR spectrum exhibits broad resonances (that arise from high molecular weight macromolecules and motionally constrained compounds) then a modified pulse sequence can be used, facilitating observation of just the free, low molecular weight metabolites. This is often required for the analysis of plasma samples. The required pulse sequence is called the 1 H Carr–Purcell–Meiboom–Gill (CPMG) spin-echo NMR sequence.

Acknowledgments I am indebted to the Natural Environment Research Council, UK, for an Advanced Fellowship in metabolomics (NER/J/S/2002/00618). The methods presented here have been optimized in conjunction with several postdoctoral researchers and graduate students whom I thank, in particular Drs. Ching-Yu Lin and Huifeng Wu, and Mr. Adam Hines and Andrew Southam.

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References 1. Kell, D. B. (2004) Metabolomics and systems biology: making sense of the soup. Curr. Opin. Microbiol. 7, 296–307. 2. Griffin, J. L. (2003) Metabonomics: NMR spectroscopy and pattern recognition analysis of body fluids and tissues for characterisation of xenobiotic toxicity and disease diagnosis. Curr. Opin. Chem. Biol. 7, 648–654. 3. Nicholson, J. K., Connelly, J., Lindon, J. C., and Holmes, E. (2002) Metabonomics: a platform for studying drug toxicity and gene function. Nat. Rev. Drug Discov. 1, 153–161. 4. Viant, M. R., Rosenblum, E. S., and Tjeerdema, R. S. (2003) NMR-based metabolomics: a powerful approach for characterizing the effects of environmental stressors on organism health. Environ. Sci. Technol. 37, 4982–4989. 5. Griffin, J. L., Walker, L. A., Garrod, S., Holmes, E., Shore, R. F., and Nicholson, J. K. (2000) NMR spectroscopy based metabonomic studies on the comparative biochemistry of the kidney and urine of the bank vole (Clethrionomys glareolus), wood mouse (Apodemus sylvaticus), white toothed shrew (Crocidura suaveolens) and the laboratory rat. Comp. Biochem. Physiol. B 127, 357–367. 6. Bundy, J. G., Osborn, D., Weeks, J. M., Lindon, J. C., and Nicholson, J. K. (2001) An NMR-based metabonomic approach to the investigation of coelomic fluid biochemistry in earthworms under toxic stress. FEBS Lett. 500, 31–35. 7. Bundy, J. G., Lenz, E. M., Bailey, N. J., Gavaghan, C. L., Svendsen, C., Spurgeon, D., Hankard, P. K., Osborn, D., Weeks, J. M., and Trauger, S. A. (2002) Metabonomic assessment of toxicity of 4-fluoroaniline, 3,5-difluoroaniline and 2-fluoro-4-methylaniline to the earthworm Eisenia veneta (Rosa): identification of new endogenous biomarkers. Environ. Toxicol. Chem. 21, 1966–1972. 8. Bundy, J. G., Spurgeon, D. J., Svendsen, C., Hankard, P. K., Weeks, J. M., Osborn, D., Lindon, J. C., and Nicholson, J. K. (2004) Environmental metabonomics: applying combination biomarker analysis in earthworms at a metal contaminated site. Ecotoxicology 13, 797–806. 9. Rosenblum, E. S., Viant, M. R., Braid, B. M., Moore, J. D., Friedman, C. S., and Tjeerdema, R. S. (2005) Characterizing the metabolic actions of natural stresses in the California red abalone, Haliotis rufescens using 1 H NMR metabolomics. Metabolomics 1, 199–209. 10. Stentiford, G. D., Viant, M. R., Ward, D. G., Johnson, P. J., Martin, A., Wei, W., Cooper, H. J., Lyons, B. P., and Feist, S. W. (2005) Liver tumours in wild flatfish: a histopathological, proteomic and metabolomic study. OMICS J. Integrat. Biol. 9, 281–299. 11. Viant, M. R., Bundy, J. G., Pincetich, C. A., de Ropp, J. S., and Tjeerdema, R. S. (2005) NMR-derived developmental metabolic trajectories: an approach for visualizing the toxic actions of trichloroethylene during embryogenesis. Metabolomics 1, 149–158. 12. Viant, M. R., Pincetich, C. A., Hinton, D. E., and Tjeerdema, R. S. (2006) Toxic Actions of Dinoseb in Medaka (Oryzias latipes) Embryos as Determined

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by in vivo 31 P NMR, HPLC-UV and 1 H NMR Metabolomics. Aquat. Toxicol. 76, 329–342. Viant, M. R., Pincetich, C. A., and Tjeerdema, R. S. (2006) Metabolic effects of dinoseb, diazinon and esfenvalerate in eyed eggs and alevins of Chinook salmon (Oncorhynchus tshawytscha) determined by 1 H NMR metabolomics. Aquat. Toxicol. 77, 359–371. Dunn, W. B. and Ellis, D. I. (2005) Metabolomics: current analytical platforms and methodologies. Trends Anal. Chem. 24, 285–294. Lindon, J. C., Holmes, E., and Nicholson, J. K. (2001) Pattern recognition methods and applications in biomedical magnetic resonance. Prog. Nucl. Magnet. Reson. Spec. 39, 1–40. Eriksson, L., Johansson, E., Kettaneh-Wold, N., and Wold, S. (2001) Multiand Megavariate Data Analysis—Principles and Applications. Umetrics, Umea, Sweden. Wise, B. M., Gallagher, N. B., Bro, R., Shaver, J. M., Windig, W., and Koch, R. S. (2004) PLS_Toolbox Version 3.5 Manual. Eigenvector Research, Manson, US. Deprez, S., Sweatman, B. C., Connor, S. C., Haselden, J. N., and Waterfield, C. J. (2002) Optimisation of collection, storage and preparation of rat plasma for 1 H NMR spectroscopic analysis in toxicology studies to determine inherent variation in biochemical profiles. J. Pharm. Biomed. Anal. 30, 1297–1310. Weckwerth, W., Wenzel, K., and Fiehn, O. (2004) Process for the integrated extraction identification, and quantification of metabolites, proteins and RNA to reveal their co-regulation in biochemical networks. Proteomics 4, 78–83. Viant, M. R. (2003) Improved methods for the acquisition and interpretation of NMR metabolomic data. Biochem. Biophys. Res. Commun. 310, 943–948. Purohit, P. V., Rocke, D. M., Viant, M. R., and Woodruff, D. L. (2004) Discrimination models using variance stabilizing transformation of metabolomic NMR data. OMICS J. Integrat. Biol. 8, 118–130. Fan, W. M. T. (1996) Metabolite profiling by one- and two-dimensional NMR analysis of complex mixtures. Prog. Nucl. Magnet. Res. 28, 161–219. Bligh, E. G., and Dyer, W. J. (1959) A rapid method of total lipid extraction and purification. Can. J. Biochem. Physiol. 37, 911–917.

II Detection of Whole Genome Mutation

10 Restriction Landmark Genome Scanning for the Detection of Mutations Jun-ichi Asakawa

Summary There is a continuing need for more efficient methods to examine mutations in humans and other species resulting from exposures to environmental toxins and radiation. Environmental genomic studies, which often utilize nonmodel system species and as a result, there is a particular need for a method that does not rely on the availability of genome sequence information. Restriction landmark genome scanning (RLGS) is a twodimensional electrophoresis (2-DE) of end-labeled DNA fragments. A vertical giant gel 2-DE system has been developed and applied to the RLGS. On a single RLGS pattern of mouse or human DNA, approx 2000 DNA fragments (spots) varying in size from 1.0 to 5.0 kb in the first dimension and 0.2 to 3.0 kb in the second dimension are visualized. In principle, this system will detect genomic alterations of two types: (1) that due to gain or loss of a cut site for the three restriction fragment enzymes employed in the study and (2) that due to insertion/deletion/ rearrangement (I/D/R) events. After optimization of the sample preparation and electrophoresis conditions, the gel quality reached a level such that the electrophoresis patterns derived from a single DNA sample gave distribution patterns of spots able to be superimposed. This technology can visualize up to 3000 DNA fragments per gel without using any probes, and thus should be highly efficient in monitoring for mutations resulting in I/D/R events in DNA fragments distributed throughout the genome. This method relies on direct labeling of DNA fragments rather than hybridization and therefore precise information on genome sequences is not required. As a result this method is applicable to any species. Key Words: Deletion; genome scanning; insertion; mutation detection; rearrangement; restriction landmark genome scanning; two-dimensional electrophoresis.

From: Methods in Molecular Biology, vol. 410: Environmental Genomics Edited by: C. Cristofre Martin © Humana Press, Totowa, NJ

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1. Introduction The advent of restriction landmark genome scanning (RLGS) techniques permitting the two-dimensional separation of DNA fragments from a genomic digest is an important advance that has allowed researchers to study the occurrence of spontaneous and induced mutation in both somatic and germinal tissues. A unique gel holder for the first-dimension agarose disc gel was made from a Teflon tube and a glass tube. The development of the disc gel system resulted in high-resolution separation of DNA fragments with enhanced reproducibility. A new device for the in-gel restriction enzyme digestion of the fractionated DNA fragments was also made from a Teflon tube. The development of this vertical disc gel system has made the RLGS a practical method. The overview of the revised RLGS method is schematically represented in Fig. 1. In a revised protocol, the DNA samples were double digested with two restriction enzymes: EcoRV (GAT↓ ATC) and NotI (GC↓ GGCCGC). The NotI sites of the digests were specifically end-labeled with 32 P by fill-in reaction with a DNA polymerase and separated by high-resolution 2-DE. The DNA samples were first separated on agarose disc gels by size. After the first electrophoresis, the DNA fragments were again digested in situ with the third restriction enzyme Hinf I (G↓ ANTC) and cut into smaller fragments that were separated perpendicularly on polyacrylamide slab gels. Following the second electrophoresis the sheet gels were dried and the NotI fragments were visualized by autoradiography. The autoradiographic image of labeled DNA fragments represents the RLGS profile and is reproducible enough that profile comparisons can be made between two different DNA samples. This technique can be used to determine mutation events within a genome after exposure to an environmental contaminant. In addition, subsequent isolation and sequencing of the differential RLGS fragments found between profiles can be used to identify the specific locus that these mutations have occurred. Since earlier description of the methods (1–7) we employ, several modifications have introduced. Current protocol and the equipment will be described in this chapter. 2. Materials 2.1. Blocking: Repair of Adventitious Damage (RAD) 1. High molecular weight genomic DNA: dissolve at 330 μg/mL in TE buffer (10 mM Tris, 1 mM EDTA, pH 8.0). Store at 4ºC. (See Notes 1 and 2.) 2. DNA polymerase I (4 U/μL) (TOYOBO, Osaka). Store at –20°C. 3. High buffer (10× HB): 0.5 M Tris-HCl, pH 7.4, 100 mMMgCl2 , 1 M NaCl, 10 mM dithiothreitol (DTT). Store at –20ºC. 4. Deoxynucleotide -thiotriphosphates (dCTP []S, dGTP []S) (Amersham ). Store at –20°C.

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Prepare Genomic DNA NotI

NotI

NotI

EcoRV EcoRV

EcoRV

EcoRV

EcoRV

Digest with NotI/EcoRV

EcoRV site (blunt end)

NotI site

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GAT CTA

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Second-dimension electrophoresis: polyacrylamide gel

Fig. 1. A schematic flow chart of the refined RLGS method.

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5. Dideoxynucleotide triphosphates (ddATP, ddTTP) (Amersham). Store at –20°C. 6. RAD stock: Mix 130 μL of 10× HB, 13 μL of 1 M DTT, 30 μL of 1 mM ddATP, 30 μL of 1 mM ddTTP, 24 μL of 10 μM dCTP[]S, and 24 μL of 10 μM dGTP[]S. Store in aliquots at –20°C. 7. Wide-bore tips: Cut and remove 5 mm from the top of a 0.2-mL pipet tip.

2.2. Restriction Enzyme Digestion and Isotope Labeling 1. NotI (20 U/μL) (TOYOBO). Store at –20°C. (See Note 3.) 2. EcoRV (20 U/μL) (TOYOBO). Store at –20°C. (See Note 3.) 3. NotI additive: 0.5 M NaCl, 20 mMMgCl2 , 0.065% bovine serum albumin (BSA), 0.065% Triton X-100, 4 mM DTT. Mix 100 μL of 5 M NaCl, 20 μL of 1 MMgCl2 , 65 μL of 1% BSA, 6.5 μL of 10% Triton X-100, 4 μL of 1 M DTT, and 805 μL of H2 O. Store in aliquots at –20°C. 4. [-32 P]dCTP (3000 Ci/mmol; Amersham). 5. [-32 P]dGTP (3000 Ci/mmol; Amersham). 6. Seaquenase Ver. 2.0 (12 U/μL; USB, Cleveland). Store at –20°C. 7. Stop solution: 50 mM EDTA, pH 8, containing 50% sucrose and 0.5% each of bromophenol blue (BPB) and xylene cyanol (XC). Store at room temperature.

2.3. Two-Dimensional Electrophoresis 2.3.1. First Dimension (1st D) Several small equipment items necessary for making gels or in situ digestion are shown in Fig. 2. 1. 1st D buffer for 1-5 kb fragments (10× Boyer buffer): 1 M Tris, 0.4 M sodium acetate, 0.36 M NaCl, 40 mM EDTA, pH 8.15. Dissolve 242 g of Tris, 109 g of

Fig. 2. Small equipment items used for the agarose disc gel preparation and in situ digestion.

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3.

4.

5.

6. 7. 8. 9. 10. 11. 12.

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sodium acetate trihydrate, 42 g of NaCl, and 23.4 g of Na2 EDTA · dihydrate in 1.8 L of H2 O and adjust the pH to 8.15 with acetic acid (the pH of the 1× buffer is 8.05). Filter through a 0.45-μm nitrocellulose filter and store at room temperature. 1st D buffer for 5- to 12-kb fragments (10× marathon TBE buffer): 1.35 M Tris, 0.45 M boric acid, 0.1 mM EDTA, pH 8.8. Dissolve 327 g of Tris, 55.6 g of boric acid, and 0.74 g of Na2 EDTA · dihydrate in 1.8 L of H2 O. Filter through a 0.45-μm nitrocellulose filter and store at room temperature. Stacking agarose gel (0.6%): Add 2.5 g of sucrose, 0.3 g of SeaKem Gold agarose (FMC), and 5 mL of corresponding 10× 1st D buffer (Boyer or marathon TBE) to 40 mL of H2 O. Boil the mixture in a microwave oven. After the agarose was completely melted, add warm water to obtain the initial weight. When the temperature of the solution drops to 65ºC, remove undissolved particles via vacuum filtration through a 0.45-μm nitrocellulose filter. Divide 5 mL aliquot each in a Falcon 15-ml centrifuge tube and store at 4°C. Running agarose gel for 1- to 5-kb fragments (0.9%): Mix 10 g of sucrose, 0.9 g of SeaKem Gold agarose (FMC), 0.9 g of SeaKem GTG agarose (FMC), and 20 mL of 10× Boyer buffer and adjust the volume to 200 mL with H2 O. Boil the mixture and melt the agarose in a microwave oven. Adjust the volume and filtrate the solution as described above. Divide 30 mL aliquot each in a Falcon 50-mL centrifuge tube and store at 4ºC. (See Note 4.) Running agarose gel for 5- to 12-kb fragments (0.7%): Mix 10 g of sucrose, 0.4 g of SeaKem Gold agarose (FMC), 1.0 g of SeaKem GTG agarose (FMC), and 20 mL of 10× marathon TBE buffer and adjust the volume to 200 mL with H2 O. Boil the mixture and melt the agarose in a microwave oven. Adjust the volume and filter the solution as described above. Divide 30 mL aliquot each in a Falcon 50-mL centrifuge tube and store at 4ºC. (See Note 4.) Support stands with double-buret clamps. Disposable plastic syringes (1-mL and 6-mL). Needles (10-cm long 19-gauge, 90º cut). Three-way stopcocks: Fit the plastic syringes. Silicon-tubes [inner diameter (i.d.) of 3 mm and 5 mm]. Cutoff tips: Cut and remove 5 mm from the top and 10 mm from the bottom of 0.2-mL pipet tips to fit the plastic syringes. Size marker: mix 3 μL of 1-kb ladder (500 ng/μL; NEB), 30 μL of stop solution, and 67 μL of H2 O. Store at 4°C.

2.3.2. In Situ Restriction Enzyme Digestion 1. BPB solution (0.01%). Store at room temperature. 2. Reaction tubes: 36-cm long Teflon tubes with i.d. of 2.7 mm connected to 3 cm of Silicon-tubes (i.d. 3 mm). Attach ID labels with consecutive numbers to the tubes. 3. HinfI buffer (10×): 200 mM Tris-HCl, pH 8.3, 1 M NaCl, 100 mM MgCl2 , 10 mM DTT. Store at –20°C. 4. HinfI (20 U/μL) (TOYOB). Store at –20°C.

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2.3.3. Second Dimension (2nd D) 1. Repel-Silane ES (Amersham: dimethyldichlorosilane solution for glass plates coating). 2. Adhesive tape: Scotch 3M or compatible. 3. TBE buffer (5×): 400 mM Tris, 445 mM boric acid, 12.5 mM EDTA. Dissolve 108 g of Tris, 55 g of boric acid, 9.3 g of EDTANa2 ·2 H2 O in H2 O. Adjust the volume to 2 L. Filter through a 0.45-μm nitrocellulose filter and store at room temperature. 4. Glycerol solution (50%): Mix equal volumes of glycerol (BRL, ultrapure) and H2 O. Store at room temperature. 5. Acrylamide stock solution (30%): Dissolve 290 g of acrylamide (99.9% pure) and 10 g of N, N-methylene bis-acrylamide in H2 O. Adjust the volume to 1000 mL and filter thorough a 0.45-μm nitrocellulose filter. Protect from light and store at 4°C. (See Note 5.) 6. Ammonium persulfate: Prepare fresh 10% solution. 7. N,N,N ,N - Tetramethylethylenediamine (TEMED). 8. Water saturated 2-butanol: Mix 300 mL each of 2-butanol and H2 O, take out upper part and store in a spray bottle at room temperature. 9. Blue connecting agarose solution: Melt 0.5 g of SeaKem Gold agarose in 100 mL of 1× TBE buffer containing each of 0.05% BPB and XC dye. Divide into three Falcon 50-mL centrifuge tubes. Store at 4°C. 10. Sampling platform: a plastic plate 3 mm thick, 33 cm × 10 cm.

2.4. Autoradiography 1. Gel drier with vacuum pump: Bio-Rad model 583 or equivalent. 2. Filter paper: 35 cm × 43 cm, 0.7 mm of thick. (Chromatography paper No. 526, Advantech, Tokyo). 3. Saran Wrap (50 cm of width). 4. X-ray films (35 × 43 cm): Fuji RX-U, Kodak BioMax MS or equivalent. 5. Intensifying screens (35 × 43 cm): Fujifilm GRENEX HR-12, Kodak BioMax MS or equivalent. 6. Exposure cassette for 35 × 43 cm film. 7. X-ray film developer.

3. Methods 3.1. Repair of Adventitious Damage (“Blocking”) 1. Melt the RAD stock and prepare a master solution for blocking; mix 0.3 μL of DNA polymerase I and 1.8 μL of RAD stock per DNA sample. Mix via a gentle pipetting and keep on ice. (Do not vortex!) 2. Add 2.1 μL of the RAD master solution to 7 μL of DNA (∼2 μg). Mix via repetitive gentle pipetting with wide-bore tips. Incubate at 16°C for 30 min followed by 65°C for 30 min. (See Note 1.)

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3.2. Restriction Enzyme Digestion and Isotope Labeling 1. Add 2.1 μL of NotI additive, 1 μL each of NotI and EcoRV. Mix thoroughly to achieve complete digestion by gently pipetting up and down (10 times) via a wide-bore tip. Incubate at 37°C for 3 h. 2. Radiolabel the NotI-derived 5 protruding ends of the sample DNA by a fill in reaction with a DNA polymerase (Sequenase Ver. 2.0). Prepare label master stock: mix 1.6 μL each of [-32 P]dCTP and [-32 P]dGTP, 0.2 μL of 1 M DTT, and 0.3 μL of Sequenase per reaction. Add 3.7 μL of the label master stock to the DNA digest and mix thoroughly by pipetting. Incubate at 37°C for 30 min. 3. Add 5 μL of stop solution and allow the mixture to stand on ice. (See Note 6.)

3.3. Two-Dimensional Electrophoresis The development of the current first-dimensional electrophoresis procedures has involved considerable experimentation. The Biochemical Genetics Program at the Radiation Effects Research Foundation has in the past made extensive use of a disc gel electrophoresis system in the study of protein variants, and this system has been adapted for the DNA gels. After considerable experimentation, I have obtained the best results with agarose disc gels cast in Teflon tubes, with an inner diameter of 2.4 mm. I found that the inside of these tubes are uneven, which stabilizes the position of the agarose gel during long durations of electrophoresis. The gel consisted of a 1.5-cm 0.6% stacking agarose gel, and a 59-cm 0.9% running agarose gel. The vertical disc gel electrophoresis system is shown in Fig. 3. By using a single system, up to 10 DNA samples can be analyzed simultaneously. The development of the vertical disc gel system resulted in high-resolution separation of large DNA fragments. This system dramatically reduced the space for electrophoresis as well as the amount of expensive enzymes (e.g., HinfI; the original system required 10,000 U of enzyme but now we need 500 U per reaction) used for in situ digestion and thus made the RLGS method more practical and less costly. The first dimension agarose disc gel electrophoresis system and the latest version of the second dimension multiple giant gel vertical electrophoresis apparatus (for 10 or 8 gels) are shown in Figs. 3 and 4, respectively. Both systems are available from Ohtsuka-Rikagaku (Hiroshima, Japan) on request. 3.3.1. First Dimension 3.3.1.1. Preparation of Agarose Disc Gels

The gel preparation of the first electrophoresis gel is schematically represented in Fig. 5. Start the gel preparation before isotope labeling. Prepare two types of gels simultaneously if two patterns (1–5 kb and 5–12 kb) are to be studied. Preparation of softer gels for 5- to 12-kb long fragments is optional.

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Fig. 3. Electrophoresis apparatus used for the first dimension agarose disc gel electrophoresis.

1. Melt the running agarose and stacking agarose in a microwave oven and place the container in a heater block at 70°C. 2. Connect a 6-mL plastic syringe fitted with a three-way stopcock and the gel holder with 3 cm of silicon tubing (5 mm i.d.). 3. Dip the bottom of the gel holder to a 1-cm depth and slowly suck up the running agarose solution to reach the first line, 57 cm from the bottom. (See Note 7.) 4. Hold it approx 30 s and stand it on the double buret clump on a support stand. Solidify for about 10 min. 5. Turn the stopcock 90º to the right to open the left inlet, and carefully remove the mounted syringe with the stopcock from the gel holder. 6. Overlay the running gel solution to the second line (59 cm) with a warmed 1-mL syringe fitted with a 90º-cut long needle. Tap the upper part of the gel holder several times with a fingernail to make a flat surface. Solidify for about 3 min. 7. Overlay the stacking gel solution to the third line (60.5 cm). Tap the gel gently with a fingernail to make the top of the gel flat. Solidify for about 10 min.

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Fig. 4. Electrophoresis apparatus used for the second dimension polyacrylamide gel electrophoresis.

Fig. 5. A schematic representation of the preparation of the agarose disc gel in a Teflon tube.

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8. Fill the tube with 1× 1st D buffer. 9. Fit each gel holder into the anodal tank through a silicon plug with a 5-mm diameter hall. 10. Add 350 mL of 1× 1st D buffer to the bottom tank. 11. Place the top tank on the bottom tank and fill the tank with 300 mL of 1× 1st D buffer. (See Note 8.)

3.3.1.2. Electrophoresis 1. Load 10 μL of the labeled sample, containing about 1 μg of DNA, onto each gel. (See Note 9.) 2. Load 10 μL of size marker (150 ng) onto a gel. 3. When all of the samples are loaded, put the cover on the top tank and connect the electrodes to the power supply. 4. Run the gel at constant voltage. For short DNA fragments (1–5 kb), run at 130 V for 19 h followed by at 140 V for about 24 h (∼5800 V · h) until the center of the marker dye, BPB, migrates 50 cm. For long DNA fragments (5–11 kb), run at 70 V for a week (about 11,200 V · h) until the center of the marker dye, XC, migrates 50 cm. (See Note 10.)

3.3.1.3. In Situ Restriction Enzyme Digestion

The NotI/EcoRV fragments separated on the agarose disc gels need to be further digested into smaller fragments with frequently cutting endonucleases like HinfI before the second separation as shown in Fig. 1. Figure 6 schematically represents how to recover the necessary gel portions and Fig. 7 represents the steps of the in situ digestion. The enzyme reaction in a Teflon tube dramatically reduces the amount of expensive restriction enzyme needed and makes the process much easier as well as more reproducible. 1. Turn off the power supply and disconnect the electrodes from the apparatus. Take the top cover off and dispose the cathodal tank buffer with an aspirator. 2. Take out each gel holder from the cathodal tank. Rinse the bottom of the gel with deionized water to remove the traces of radioisotope (save the waste, it is radioactive!). 3. Expel the gel containing the marker DNA into a 50-mL Falcon tube. Add 30 mL of ethidium bromide solution (0.5 μg/mL) and stain the gel for 20 min. Examine the gel via ultraviolet light. Photograph with a transparent ruler to record the distance of each band of the marker DNA. (See Note 11.) 4. Assuming that 1-kb and 5-kb marker DNA have migrated to about 8 cm and 38 cm from the bottom, respectively, a 32-cm gel portion of 7–39 cm from the bottom (this portion contains BPB and XC at each end) will be used for in gel digestion. (In a similar way, for the longer fragments, a gel portion of 7–39 cm from the bottom containing 5- to 12-kb fragments is used.) Draw two lines as marks with dry ink on each gel holder at 7 cm and 39 cm from the top of the gel.

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Fig. 6. A schematic representation of expelling the agarose disc gel.

Fig. 7. A schematic representation of gel handling for in situ restriction enzyme digestion.

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5. Draw BPB solution into a 1-mL plastic syringe fitted with a cutoff tip, and slowly expel the gel by forcing through the blue BPB solution. Carefully let the blue solution to the first 7-cm line. Cut off the expelled 7-cm gel at a 45º angle of with a scalpel. Expel the gel portion of 39 cm into a 50-mL Falcon tube containing 20 mL of 1× HinfI buffer. When the BPB solution reaches the second 39-cm line, cut the gel at a 90º angle of. 6. Equilibrate the gel to 1× HinfI buffer at room temperature. Rock the tube occasionally and change the buffer two times at each 10 min. Incubate for 30 min in total. 7. Pour the gel noodle with buffer into a small tray placed at an angle about 10º. 8. Fit a cutoff tip to a 6-mL syringe and connect the tip to a 33-cm long 2.7-mm i.d. Teflon tube with a 3-cm long 3-mm i.d. silicon tube. Attach the Teflon tube to the edge of the noodle gel and slowly draw up the gel into the tube. Draw the buffer into the syringe and remove the buffer from the gel as much as possible. 9. Add 600 μL of 1× HinfI buffer containing 400 U of HinfI and 0.01% BSA to a 2-mL round-bottom microtube. Draw up the solution and fill the tube to cover the gel surface with the HinfI mixture. 10. Remove the syringe and loop the Teflon tube by connecting one end to the other with the 3-cm silicon tube. Put the looped tube into a nylon bag and incubate at 37ºC for 4 h. (See Note 12.)

3.3.2. Second Dimension 3.3.2.1. Gel Casting

Prepare gels at least 1 day before starting the second electrophoresis. By the system shown in Fig. 4, 10 gels (33 cm wide, 46 cm long and 0.8 mm thick) are made simultaneously. The acrylamide solution is toxic; wear plastic gloves during the gel preparation. 1. Clean each glass plate meticulously and treat the surface of the beveled side with Repel Silane. 2. Set up the apparatus and seal the bottom holes with adhesive tape (Scotch 3M). (See Note 13.) 3. Mix 348 g of 30% acrylamide stock, 504 g of 5× TBE, 23.3 g of 50% glycerol, 15 g of 10% APS, and 1110 g of H2 O in a flask to make 5.15% acrylamide solution in 1.25× TBE. Divide into two 1-L vacuum flasks. We find that measuring these reagents by weight is more accurate than measuring by volume using graduated cylinders. This accuracy is important in ensuring the reproducibility of the electrophoresis steps. 4. Deaerate the solution by applying vacuum, gently at the beginning. Swirl the flask and continue deaeration until no more air bubbles are generated. 5. Add 210 μL of TEMED to each 1-L acrylamide solution and mix thoroughly. (See Note 14.)

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6. Immediately fill the apparatus from the bottom entrance through a silicon tube. Clamp the entrance when the solution has reached to 1 cm from the top of glass plates. 7. Cover the surface by a gentle spraying of water-saturated 2-butanol. (See Note 15.) 8. After about 1 h of polymerization, replace the surface solution to 1× TBE. 9. Cover the TBE solution of the gel with Saran Wrap and store at room temperature until use. The gels are stable for about 5 days at room temperature. Avoid allowing the top of the gel to dry.

3.3.2.2. Electrophoresis 1. Melt the blue connection agarose and place the container in a heater block at 70°C. 2. After 4 h of digestion with HinfI, expel each gel into a 50-mL Falcon tube containing 20 mL of 1× TBE buffer. 3. After 10 min of equilibration, discard the buffer and pour the noodle gel onto a sampling platform laid on the top of the gels. Straighten the noodle with a ruler and slide to the edge of the platform. 4. Overlay the top of the gel with the connection agarose to reach to the beveled line of the glass plate. 5. Slip the noodle down along with the glass plate using a spatula and flow of melted blue connection agarose. 6. Cover the noodle enough connection agarose and solidify for 10 min. Remove the platform. 7. Add 2.5 L each of 1× TBE buffer to both reservoirs. Remove the 3M tape from the holes and contact the gel portion with the buffer. 8. Connect the electrodes to a power supply and run the gel at 150 V for about 40 h until the marker dye XC migrates to 35 cm. (See Note 16.)

3.3.3. Autoradiography 1. 2. 3. 4. 5.

6.

7. 8. 9. 10.

At the end of electrophoresis, turn off the power and disconnect the electrodes. Dispose of electrophoresis buffer. Lay the apparatus and remove the plastic cover. Slowly pry apart the top glass plate from the gel using a metal spatula. Place a piece of filter paper (35 cm × 43 cm, 2 cm wider than gel) on the gel. Apply gentle pressure so that the gel firmly attaches to the filter paper. Cut off the 3-cm portion of the gel with a scalpel. Hold the edge of paper, and quickly flip the gel over and lay it on another piece of filter paper on a gel drier (place the gel on the top). Cover the gel surface with Saran Wrap. Try to avoid creases and bubbles. Dry the gel under vacuum on a gel drier set at 80°C. Put an ID number of the gel on the edge of the filter paper. Establish an autoradiography by exposing the gel to X-ray film and intensifying screen in a metal cassette at –80ºC for about a week. Develop the autoradiogram.

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3.4. Image Analysis A digitized image of a BALB/c mouse RLGS pattern is shown in Fig. 8. In this method, end labeled DNA fragments are visualized by autoradiography. The spot intensity represents the copy number of the spot DNA. Except for sexchromosome-derived fragments and multiple-copy fragments such as ribosomal DNA, the intensities of most spots in an RLGS pattern represent two copies of autosomal DNA fragments. Genomic mutation detectable by this method consists of alterations in the length of the DNA fragments visualized, either by insertions or deletions within these fragments, or base changes at the recognition sites of the restriction enzymes used. Mutations can be detected by searching for new spots in patterns of progeny not present in either parent, or by detecting half-intensity (one-copy) spots in the progeny that are two-copy in the parents. A mutated fragment will shift position on the gel if the distance from the NotI site to the closest HinfI or EcoRV site is altered. The expected positions of mutant fragments are schematically represented in Fig. 9. In some cases, the altered fragment may not be found on this gel (although it might appear on a different gel) or may produce no spot if the NotI site is involved in a deletion or base substitution. In any event, the intensity of the normal spot should decrease by 50% in theory. To analyze the complex 2-DE patterns, computer programs have been utilized. The computer-based image analysis

Fig. 8. A digital image of RLGS pattern of BALB/c mouse. Fragment sizes shown for each dimension are based on sequence data of cloned spots.

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H

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Fig. 9. Restriction map diagrams showing possible explanations for the various spatial configurations for the allelic pairs of spots. Fragments labeled C indicate the common allele with V1, V2, and V3 representing variants. Restriction sites are labeled: N = NotI, H = HinfI, and E = EcoRV. Hatched rectangles labeled represent deletions.

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Fig. 10. Close-ups of regions showing examples of spots involved in mutations. Arrows point to spots where mutations were detected. Mutated spots are labeled with μ. In the leftmost panel, the heterozygous spot involved in a mutation disappeared.

detect the presence or absence or any changes in intensity of each spot; thus, we can detect deletion, amplification, insertion, and rearrangement throughout the genome efficiently. Examples of mutations detected in our mouse study are shown in Fig. 10. 3.5. Other Restriction Enzymes for RLGS The combination of restriction enzymes—NotI, EcoRV, and HinfI—gave a mostly evenly distributed spot pattern for human and mouse (rat also) DNA. The most reliable landmark enzyme currently available for human and mouse is NotI in our experience (we examined more than 10,000 gels). As for the second enzyme (in the standard method; EcoRV), PstI (C↓ TGCAG) and PvuII (CAG↓ CTG) are both usable. As for the third enzyme for in situ digestion, MboI (↓ GATC), BamHI (G↓ GATCC), PstI, and PvuII are usable (although MboI is somewhat expensive). These enzymes are usable in any combination without problems. RLGS can be applicable to other species and it is easier if the genome size of the study species is smaller than human or mouse. We have examined the following species: (1) Medaka (a fish; the genome size is 1 × 109 base pairs [bp]): NotI, EcoRV, and HinfI combination , (2) Sacchromyces (yeast; 3 × 107 bp): HindIII (A↓ AGCTT; used as landmark enzyme, labeled with only [32 P ]dATP) and HinfI, also HindIII and PstI, and (3) E. coli (4 × 106 bp): EcoRI (landmark enzyme, labeled with only [32 P]dATP) and HinfI. For the preparation of RLGS for small-genome species, many other enzymes can

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be used as the landmark enzymes, but for in situ digestion in agarose only limited enzymes, the five aforementioned enzymes are practical.

4. Notes 1. Intact high molecular weight DNA is very viscous and it is often difficult to measure its volume with a pipet. In such a case, digest the genomic DNA with EcoRV. Digest approx 5–10 μg of DNA with 5 U of enzyme in 100 μL 1× high buffer for 1 h at 37°C. Ethanol precipitate by adding 10 μL of 5 M NaCl and 250 μL of ethanol. Recover the partial digest by centrifugation and dissolve in 100 μL of TE. Estimate the concentration by measuring the OD260 . Ethanol precipitate the same again and dissolve in TE at a concentration of 300 μg/μL. Use this partially digested DNA instead of the high molecular weight DNA. 2. Unless stated otherwise, all buffers and solutions are prepared with Milli-Q H2 O that has a resistance value of 18 M. Deionized and distilled H2 O is also usable. 3. Rstriction enzymes (NotI, EcoRV, and HinfI) manufactured by NEB are also usable. All enzymes should be stored in a −20ºC freezer without defroster. 4. Measure the weight of the agarose solution before and after boiling and adjust the weight by adding H2 O after boiling. 5. Acrylamide and bis-acrylamide are often contaminated with metal ions. Stock solutions can be purified by stirring overnight with 1 g of a mixture of ionic exchange resins (Amberlite IRN-150L, Amersham) per liter of solution. Fresh solutions should be prepared every few months. 6. The digests can be stored at 4ºC for several days before isotope labeling; do not freeze. 7. Carefully examine the gel for air bubbles. If any are present in the gel, it may be necessary to prepare another gel. 8. The final volume of the reaction is 21.9 μL in theory, but the volume is decreased to approx 21 μL or less in practice. 9. Examine any air bubbles in the buffer on the gel; if present, carefully remove by drawing up with a 90º-cut long needle fitted to 1-mL syringe. 10. The higher the temperature, the faster the dye and DNA fragments will move, and thus there are variations in the duration of electrophoresis. In a rush experiment, electrophoresis for short fragments can be done at 200–250 V after running at 130 V for 2 h. For long fragments, electrophoresis at higher voltage, up to 150 V, can be used after electrophoresis at 100 V for 2–3 h. In general, electrophoresis at lower voltage results in better resolution and sharper spots. 11. To protect from dangerous ultraviolet radiation, wear a full safety mask. 12. Put an ID number on each tube. 13. The glass plates must be free of grease spots and dusts to avoid formation of air bubbles in the gel. 14. Adjust the amount of TEMED so that the gels polymerize about 10 min after the solution is poured.

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15. Spray gently from a distant place to prevent from disturbing the gel surface. Cover the top of the apparatus with Saran Wrap: plastic plate will crack after exposure to 2-butanol. 16. The position of the XC corresponds to approx 300-bp double-strand DNA fragments. If smaller fragments are the target, run a shorter gel.

References 1. Hatada, I., Hayashizaki, Y., Hirotsune, S., Komatsubara, H., and Mukai, T. (1991) A genomic scanning method for higher organisms using restriction sites as landmarks. Proc. Natl. Acad. Sci. USA 88, 9523–9527. 2. Asakawa, J., Kuick, R., Neel, J. V., Kodaira, M., Satoh, C., and Hanash, S. M. (1994) Genetic variation detected by quantitative analysis of end-labeled genomic DNA fragments. Proc. Natl. Acad. Sci. USA 91, 9052–9056. 3. Asakawa, J., Kuick, R., Neel, J. V., Kodaira, M., Satoh, C., and Hanash, S. M. (1995) Quantitative and qualitative genetic variation in two-dimensional DNA gels of human lymphocytoid cell lines. Electrophoresis 16, 241–252. 4. Kuick, R., Asakawa, J., Neel, J. V., Kodaira, M., Satoh, C., and Hanash, S. M. (1995), High yield of restriction fragment length polymorphisms in twodimensional separations of human genomic DNA. Genomics 25, 345–353. 5. Thoraval, D., Asakawa, J., Kodaira, M., Chang, C., Radany, E., Kuick, R., Lamb, B., Richardson, B., Neel, J. V., Glover, T., and Hanash, S. (1996) A methylated human 9-kb repetitive sequence on acrocentric chromosomes is homologous to a subtelomeric repeat in chimpanzees. Proc. Natl. Acad. Sci. USA 93, 4442–4447. 6. Hayashizaki, Y., Hirotsune, S., Okazaki, Y., Muramatsu, M., and Asakawa, J. (1997) Restriction Landmark Genomic Scanning Method. In Encyclopedia of Molecular Biology and Molecular Medicine, Vol. 5. Editor-in-chief: Meyers, R. A., VCH publisher, Weinheim, New York, Basel, Cambridge, pp. 304–319. 7. Asakawa, J., Kuick, R., Kodaira, M., Nakamura, N., Katayama, H., Pierce, D., Funamoto, S., Preston, D., Satoh, C., Neel, J. V., and Hanash, S. M. (2004) A genome scanning approach to assess the effects of radiation in mice and humans. Radiat. Res.; 161, 380–390.

11 Use of the Comet Assay in Environmental Toxicology Loren D. Knopper and James P. McNamee

Summary The comet assay, also known as the single-cell gel electrophoresis assay, is a method of detecting DNA damage in virtually any nucleated cell. The comet assay has significant advantages over other genotoxicity tests, but it is very sensitive to subtle changes that can yield appreciable variability in results. The purpose of our chapter is to present background information and detailed standard operating procedures for the use of the alkaline comet assay in environmental genotoxicity assessment. We address pitfalls and concerns associated with conducting the comet assay, and briefly discuss modifications of the general alkaline procedure that can be used to address different issues relevant to environmental toxicology. Key Words: Alkaline comet assay; DNA damage; environmental toxicology; genotoxicity.

1. Introduction The comet assay, also known as the single-cell gel electrophoresis assay, is a method of detecting DNA damage (including DNA single- and double-strand breaks, DNA–DNA crosslinks, DNA–protein crosslinks, and alkali-labile DNA damage) in virtually any nucleated cell. Significant advantages of the comet assay over other genotoxicity tests include its fairly simple methodology, sensitivity, requirement for small numbers of cells, and rapid production of data (1). However, researchers need to be aware that the technique is very sensitive to subtle changes (e.g., temperature, buffer volume, and pH) that can yield appreciable variability in results. From: Methods in Molecular Biology, vol. 410: Environmental Genomics Edited by: C. Cristofre Martin © Humana Press, Totowa, NJ

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The general technique begins by obtaining a single-cell suspension of nucleated cells of interest. The cells should be assessed for viability, diluted to the appropriate cell concentration, and then mixed with molten agarose and “cast” onto frosted glass slides (1,2) or into individual plastic chambers affixed to a piece of Gelbond acetate film (Fig. 1) (3). We suggest the use of the Gelbond technique because it eliminates the main technical problems associated with using glass slides (e.g., lack of agarose adhesion to slides, shrinking of gels during storage) and results in increased productivity and efficiency without decreasing assay reliability. After the solidification of the

Fig. 1. Comet assay setup using the GelBond technique. (a) Lab Tek II chambers are affixed to Gelbond; (b) casting agarose-cell suspension mixture into chambers; (c) removing chambers from Gelbond; (d) placing GelBond-agarose complex into lysis buffer.

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cell-agarose suspension, the entire Gelbond–gel or glass slide–gel complex is placed in a lysis buffer consisting of salts and detergents to degrade the cellular membrane and expose the nucleus. This is followed by submersion of the complex in an alkaline or neutral buffer (see later) that allows for DNA unwinding before electrophoresis, which is conducted for a given time based on cell type and/or desired sensitivity. After electrophoresis, the complex is placed in a neutralizing buffer to stabilize the gel, and then dehydrated in ethanol. This technique is common to most researchers but cell dilutions, buffer pH, and electrophoresis conditions (voltage and time) are specific to cell type, desired sensitivity, and equipment being used. (See Notes 1–3.) The National Institute of Health maintains a Listserve for the Comet Assay Interest Group (www.cometassay.com), where ongoing discussion and debate takes place regarding all aspects of the comet assay. The Listserve is regularly updated and all correspondence is archived so past postings can be viewed.

2. Materials 2.1. Solutions 1. 10× Ca2+ - and Mg2+ -free phosphate-buffered saline (PBS): 1.31 M NaCl, 50 mMNa2 HPO4 , 16 mMKH2 PO4 . Store in the dark at room temperature. Dilute to 1× before use and adjust to pH 7.4. 2. 0.75% Agarose: 750 mg of low melting point (LMP) agarose, 1× Ca2+ -and Mg2+ free PBS. Stir together on stirring plate with heat on medium/high and then dispense 1.0-mL aliquots into 1.5-mL Eppendorf tubes. Store at 4°C until needed. On the day of experiment, melt in microwave for 10×20 s on high and place in a heater block or water bath at 42°C. 3. 1.0% Agarose: 1.0 g of agarose, 100 mL of Ca2+ - and Mg2+ -free PBS. Stir together on stirring plate with heat on medium/high and then dispense 4-mL aliquots into Falcon tubes and store at 4°C until use. On the day of the experiment, melt in a microwave for 10–20 s on high, then maintain in a heater block or water bath at 42°C. 4. Lysis buffer (See Note 4): 2.5 M NaCl, 100 mM tetrasodium EDTA, 10 mM Tris base, 1% n-lauryl sarcosine. Cover and shake to remove clumps. Slowly add 1 L of purified H2 0 and stir on a stir plate. Adjust the pH to 10.0 with concentrated NaOH or HCl. Store in the dark at room temperature. Add 1% Triton X-100 to the required volume on the day of the experiment, 30 min before use. Store at 4°C (see later). 5. Electrophoresis buffer A (See Note 5): 0.3 M NaOH, 10 mM tetrasodium EDTA, 0.1% 8-hydroxyquinoline, 2% dimethyl sulfoxide (DMSO). Slowly add purified H2 0 and stir on a stir plate. Add DMSO while mixing. Adjust to pH 13.1 ± 0.1 on a calibrated pH meter with concentrated NaOH or HCl. Prepare fresh on the day of the experiment.

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6. Neutralization buffer: 1 M ammonium acetate. Adjust the pH to 7.0 with concentrated NaOH or HCl 7. Dual stain for cell viability: stock fluorescein diacetate (5.0 mg/mL acetone), stock ethidium bromide (200 μg/mL Ca2+ - and Mg2+ -free PBS), adjust to pH 7.4. Store stock fluorescein diacetate at –20°C and stock ethidium bromide, wrapped with aluminum foil, at 4°C. For a working stain, mix in an Eppendorf: 1.2 mL of Ca2+ and Mg2+ -free PBS, 7.5 μL of stock fluorescein diacetate, and 50 μL of stock ethidium bromide. Make fresh on the day of use. 8. DNA stain: Dispense 5.0 μL of SYBR Gold stain (Molecular Probes cat. no. S-11494) into 50 mL of purified H2 O. Wrap in aluminum foil. Store in the dark at 4°C. (See Note 6.)

2.2. Glassware and Labware 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14.

Pipets. Pipet tips (10, 20, 100, 200, 1000μL). Volumetric flasks (100 mL, 500 mL, and 1 L). Graduated cylinders (100 mL, 500 mL, 1 L). Beakers (250 mL, 500 mL, 1 L). Petri dishes (120 mm diameter). Dispensing bottle (for purified water). Ice bucket. Cryovials (1.0 mL). Eppendorf tubes (0.5 mL, 1.5 mL). Centrifuge tubes with screw caps (50 mL). Plastic/glass trays (e.g., lids and bottoms of pipet tip containers). Microscope slides. Microscope cover slips (22 × 50 mm).

2.3. Instrumentation 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12.

pH meter. Analytical balance. Magnetic stirrer and magnetic bars. Power supply capable of low voltage and high milli-amperage (e.g., 300 V, 2000 mA). Horizontal gel electrophoresis units. Tweezers. Scissors. Gelbond Film (agarose gel support medium) (Mandel cat. no. 53740 GB 1638). Lab Tek II chambers (Nalge Nunc cat. no. 154461-B). Heater block or water bath. Computer with Comet assay imaging system (Kinetic Komet 5.5 or other). Fluorescence microscope with ×40 oil immersion objective (Zeiss AxioPlan II or other) and proper excitation/emission filters for dye (e.g., SYBR Gold: ex 300, em 495/537 nm, bound to nucleic acid).

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3. Methods 3.1. Calibration Calibration of the comet assay technique can be achieved by assessing DNA damage in cells that have been exposed to either ionizing radiation (X-ray or -ray) or chemical exposure (hydrogen peroxide, cyclophosphamide, or methyl methanesulfonate) (See Note 7) (1,4,5). Generating a dose–response curve can also be used to assess the sensitivity of the technique in a given laboratory (Fig. 2). Calibration should be conducted with each new species and cell type being assessed. Some laboratories incorporate a concurrent positive control (e.g., radiation or chemical exposure) for each comet assay experiment to ensure assay conditions are consistent across experiments. 3.2. Cell Viability Cells that are apoptotic or necrotic do not display the typical comet appearance. Rather, they exhibit very small or nonexistent heads and very large diffuse tails (Fig. 3 ). These cells are commonly referred to as tear-drops, ghost cells, or hedgehog cells (1,4). Such cells can be produced after exposure to cytotoxic agents and/or nongenotoxic agents and should be excluded from analysis. Cells that have been exposed to genotoxins can also show this type of appearance and should be used in data analysis. Thus, it is important to conduct a concomitant assessment of cytotoxicity in the cell suspension in

Fig. 2. An example of a dose–response curve generated by exposing whole blood from meadow voles (Microtus pennsylvanicus) to gradations of 137 Cs radiation. N = 4 at each dose. Notice the plateauing trend in tail length and the exponential trend in moment with dose. (Modified from Knopper et al., 2005.) (22)

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Fig. 3. Control (a), damaged (b), moderately damaged (c), and apoptotic (i.e., hedgehog, ghost-cell) or highly damaged nucleoids (d) as observed after 20 min of electrophoresis at 1.5 V/cm. Cells stained with SYBR Gold DNA dye.

order to determine the cause of highly damaged cells and determine whether they should be included in the data analysis. Cell viability is predominantly assessed using one of two methods. The first is with the dual stain viability assay (6). In this technique, equal volumes of the cell solution and ethidium bromide/fluorescein diacetate working stain (See Subheading 2.1., item 7) are mixed together, then loaded into both sides of a hemacytometer (10 μL per side). Viable and nonviable cells are then counted manually via fluorescence microscopy. Metabolically competent (viable) cells will fluoresce green as a result of the conversion of fluorescein diacetate to its fluorescent metabolite fluorescein by cellular esterases, while metabolically incompetent (nonviable) cells fluoresce red because of membrane leakage and staining of DNA by ethidium bromide. Another way to assess viability is by the trypan blue exclusion assay. In this assay, equal volumes of the cell solution and trypan blue are mixed together, loaded into both sides of a hemacytometer (10 μL per side), then viable and nonviable cells counted manually under light

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microscopy after a settling period (usually between 2 and 5 min). Cells that take up the dye are nonviable, whereas those that exclude the dye are viable. In general, samples with viability below 70–75% of that in the control samples should be discarded from further analysis (1). 3.3. Species Concerns The comet assay can be conducted using virtually any nucleated cell, with the caveat that those cells are viable. It has been found that in some species (e.g., avian), whole blood is not appropriate for use with the comet assay because more than 80% of the cells exhibit the “ghost cell” or “hedgehog” appearance, presumably owing to degraded and functionally inert DNA/RNA within nucleated, mature erythrocytes. In this situation, leukocytes need to be separated from the nucleated erythrocytes to be used for the comet assay. Whole blood from amphibians does not appear to display this phenomenon. 3.4. Metrics for Assessing Damage To quantify DNA damage, gels are stained with a DNA stain (e.g., ethidium bromide, SYBR Gold, SYBR Green) and the comets scored for DNA migration under a fluorescence microscope using an appropriate software package such as Komet 5.5 (Kinetic Imaging, Nottingham, UK), Comet Assay IV (Perceptive Instruments, Suffolk, UK), or others. Damaged cells have an appearance similar to astronomical comets, with long tails of DNA migrating from the center of the exposed nucleoid. Damage is generally quantified using three main values: comet tail length; tail moment (i.e., tail length multiplied by the % DNA in the tail) or Olive tail moment (i.e., distance between the center of gravity of the tail and the center of gravity of the head, multiplied by % DNA in the tail); and % DNA in the tail (7,8). The most informative metric is currently under debate, mainly because computerized imaging programs tend to compute these metrics differently. Tail length is expected to increase quickly with low levels of exposure to a genotoxin, but this metric will plateau at higher exposures (9). However, the amount of DNA in the tail region can continue to increase as the dose increases, theoretically from 0 to 100% (7). Thus, with increasing dose, it is tail intensity that continues to increase with increasing doses, not tail length (Fig. 2). Because tail moment is calculated based on length measurements, it has been argued that tail intensity, or % DNA in the tail, should be the best metric of genotoxicity (10). 3.5. Freezing Samples When the comet assay cannot be conducted quickly after sample collection, samples can be frozen in liquid nitrogen and stored until the appropriate time,

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as long as they have been placed in a proper cryopreservant (11–13). We have found when using blood samples, Ca2+ - and Mg2+ -free PBS plus 10% DMSO or Ca2+ - and Mg2+ -free PBS plus 10% DMSO and 20 mM EDTA (12) are acceptable cryopreservatives. Samples should be thawed in a room temperature water bath and processed immediately thereafter. However, comet assay data from frozen samples cannot be directly compared to data for unfrozen samples, as the freezing process elevates the background level of DNA migration. As such, all experimental controls should be handled (frozen) in a similar fashion. 3.6. Alkaline Comet Assay on Gelbond Film 1. Remove 1.5-mL Eppendorf tubes of 0.75% low melting point agarose (LMP) and melt in microwave for approx 10–20 s on high. 2. Once melted, place tubes in a prewarmed heating block or water-bath, set at 42°C. 3. Affix Lab-Tek II chambers to Gelbond by applying constant pressure for approx 30 s. (See Note 8.) 4. Remaining steps should be conducted under subdued lighting. 5. Remove a 30-μL aliquot from the diluted cell suspension and add to 270 μL of liquified 0.75% LMP agarose. Mix gently by pipetting. (See Note 9.) 6. Cast a 120-μL aliquot of the cell–agarose mixture into an individual well of the two-well chamber. 7. Cast another 120-μL aliquot into a different chamber affixed to a different Gelbond film. Repeat steps 5–6 with each sample. 8. An internal control should be run simultaneously. (See Note 10.) 9. Once the agarose has solidified, carefully remove the Lab-Tek II chambers and place each Gelbond film in a small plastic box filled with 75 mL of lysis buffer. 10. Place dishes at 4°C overnight. 11. The following day, calibrate the pH meter. 12. Make fresh electrophoresis buffer. 13. Remove gels from the fridge/incubator/water bath. 14. Place a Petri dish in the sink and fill with fresh water. Allow water to gently run into dish. 15. Using tweezers, remove Gelbond film from lysis buffer and repeatedly dip into the Petri dish for about 30 s, or until the foam from the lysis buffer has drained. (See Note 11 and 12.) 16. Place Gelbond film into the gel electrophoresis units, filled to a volume of electrophoresis buffer required to achieve a level approx. 1 cm above the gel (See Note 13). Allow the gels to stand for 30 min. 17. After 30 min, electrophorese gels using constant voltage (time and voltage will depend on sample type; See Note 3). 18. Place a Petri dish in the sink and fill with fresh water. Allow water to gently run into dish.

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19. Using tweezers, gently remove the Gelbond film from the electrophoresis unit and repeatedly dip into the Petri dish for about 30 s, or until the electrophoresis buffer has drained. 20. Transfer Gelbond film to another tray containing 75 mL of neutralization buffer and leave for 30 min. 21. After 30 min, rinse as described in the preceding text, and place Gelbond film into approx. 75 mL of 85% ethanol for a minimum of 2 h. 22. Remove film and then air-dry overnight. 23. Store dried gels in labeled manila envelopes. 24. To stain gels, make SYBR Gold solution in a 50-mL tube (See Subheading 2.2., item 10) and fill another 50-mL tube with water. 25. Cut one of the duplicate Gelbond films into strips, so each strip contains two samples. 26. Label and place in stain. (See Note 14.) 27. After the staining period, remove the Gelbond strip with tweezers, and dip in water two or three times. 28. Place Gelbond, gel side up, on microscope slide and cover with a 22 × 50 mm glass cover slip. 29. Gently press with a paper towel to remove excess water and to form a seal. 30. Add a drop of immersion oil to the cover slip and then view on the microscope. A minimum of 50 cells, on each of two duplicate slides, should be scored using appropriate software.

3.7. Other Comet Assay Techniques for Environmental Toxicology Three modifications to the general alkaline version on the comet assay can also be useful tools for studies in environmental toxicology and genomics. First, the comet assay can be combined with fluorescence in situ hybridization (FISH) to determine in which gene regions DNA strand breaks are occurring (14,15). Second, the comet-DNA diffusion assay, which involves precipitating DNA with ethanol during the comet assay without the use of electrophoresis, can be used to discriminate between the mechanisms of cellular death (i.e., apoptosis (programmed cell death) and necrosis (death of cells through injury or disease) (16). Finally, the comet assay can also be conducted using a neutral buffer rather than alkaline buffers in order to assess only double-strand DNA breaks (5,17). This technique is also useful for assessing DNA damage in germ cells, which possess naturally high levels of alkali-labile sites (5,18,19). (See Note 6.) 4. Notes 1. Because the basis of the comet assay is to measure DNA damage in individual cells (i.e., generally no fewer than 50 per sample), it is important that cell density in

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the gels is not too high; otherwise the comets may overlap, making measurements impractical or impossible. Generally, a dilution of a sample resulting in 2–4 × 105 cells/mL is sufficient. For example, mammalian whole blood requires a 1:10 dilution with phosphate Ca2+ - and Mg2+ -free PBS whereas amphibian whole blood requires a 1:100 dilution. 2. Under neutral pH conditions, only double-stranded DNA breaks can be revealed using the comet assay (17). Under alkaline pH conditions (pH > 13.1), double-and single-strand breaks as well as alkali-labile sites (expressed as single-strand breaks under alkaline conditions) can be detected (1,4). Compared to radiation exposure, chemical genotoxins cause orders of magnitude fewer double-strand breaks than single-strand breaks (1). Thus, the alkaline comet assay is a more useful tool for biomonitoring studies than the neutral version. However, in some cases cells (e.g., sperm) may contain inherently high levels of ambient alkali-labile sites (18). Alkaline buffers will not be useful when using these cell types because these sites will be expressed as DNA single-strand breaks under alkaline conditions. Thus, control cells will exhibit substantial DNA damage. In these circumstances, the use of neutral buffers is suggested (e.g., Tris–borate EDTA or Tris–acetate EDTA; pH 7–8). Some researchers have used electrophoresis buffers of pH 9.0 (20) and 12.5 (21) with no apparent increases in ambient DNA damage. 3. If the voltage or running time used during electrophoresis is too low, DNA will not migrate from the comet head. Conversely, if the voltage or running time is too high, DNA from undamaged cells will migrate extensively. Thus, conditions need to be optimized so only true damage is detected. Generally, control cells should exhibit no more than 5–10% DNA in the tail and have tail lengths of no more than 15–20 μm (10). The appropriate voltage and time of electrophoresis will vary depending on the species and cell type being assessed, the equipment being used, and on the volume of buffer required to cover the gels. Electrophoresis voltage (constant voltage) should be expressed as V/cm, as determined by the running voltage divided by the distance between the anode and cathode in the electrophoresis units. A good starting point for blood samples is 1.5 V/cm for between 16 and 20 min of electrophoresis. It is imperative that these conditions be optimized before an experiment is conducted and then strictly adhered to within and across experiments. 4. This recipe for lysis buffer is the standard for many cell types, but when using germ cells, a different lysis buffer is required to expose the nucleus. For these cells, prepare lysis buffer containing 2.5 M NaCl (146.1 g/L), 100 mM tetra-sodium EDTA (41.6 g/L), 10.0 mM Tris-HCl (0.61 g/L). Cover and shake to remove clumps. Slowly add 1 L of purified H2 0 and stir on a stir plate. Adjust the pH to 10.0 with concentrated NaOH or HCl. Store in the dark at roughly 21°C. Add 1% Triton X-100 and 4 mM DTT to required volume on the day of experiment and stir. DO NOT refrigerate. After 1 h, decant buffer and add fresh buffer containing 0.1 mg/mL of proteinase K, and incubate at 37°C overnight. The addition of DTT and proteinase K to this buffer is required to decondense sperm chromatin.

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5. This recipe for electrophoresis buffer is the standard for the alkaline version of the comet assay. Germ cells have naturally abundant alkali-labile sites (18) and require the use of a neutral buffer. Neutral buffers are required when only double-stranded DNA damage is being quantified. Tris–acetate EDTA or Tris–borate EDTA (pH 7–8) buffer is appropriate for use in these circumstances. 6. When stain intensity has diminished, another 5 μL of SYBR Gold can be added to replenish the solution. This should be replenished only one time, after which a new stain solution should be made. Dispose of the solution by pouring through activated charcoal. 7. When using germ cells, where the chromatin is very tightly packed, cells must undergo lysis before chemical exposure (e.g., H2 O2 ). If in vitro damage is induced by radiation exposure, however, lysis is not required before irradiation (5). 8. New chambers have an adhesive backing, but this adhesive backing will degrade with use. It is possible to reuse these old chambers by dipping the bottoms of the chambers in a 1% agarose solution. Gently dip a gloved finger in melted 1% agarose and run along the bottom of a Lab-Tek II chamber until the bottom is coated, then affix to the Gelbond film and allow it to solidify. The agarose on the bottom of the chamber will form a seal, and samples can then be cast as previously described. 9. If using frozen samples: remove samples from the –80°C freezer and place in a room temperature water bath. Once the samples have thawed (∼1–3 min), gently mix, and place on ice. 10. If available, use a control sample. If no control is available, human blood (dilution: 20 μL of human blood mixed with 180 μL of Ca2+ - and Mg2+ -free PBS), obtained from finger prick, is a suitable substitute. Control samples can be frozen ahead of time and thawed when needed, but the viability and DNA damage in these samples should be checked regularly for consistency as time spent frozen is related to increased DNA damage. 11. Salt from lysis buffer that remains in the gel can cause misshapen comets during electrophoresis, so proper rinsing should be ensured. 12. Some researchers who are interested in assessing for the presence of specific types of DNA damage (e.g., DNA–protein crosslinks and several forms of oxidative base damage) employ a second lysis step in which DNA specific endonucleases (such as endonuclease III) or proteases are added. These enzymes recognize and attempt to repair certain types of DNA modifications. However, these repair complexes are unstable, and when the gels are exposed to alkaline conditions, these repair sites degrade to yield single-strand DNA breaks. The increased level of DNA damage in the presence of such enzymes, relative to controls, indicates the presence of DNA lesions specific to that recognized by the enzyme added (10). 13. Because many different models of horizontal electrophoresis chambers can be used, each with its own relative dimensions, the optimal electrophoresis buffer volume needs to be determined. As a general practice, gels should be covered with approx 1 cm of buffer. It is very important to be consistent and accurate with the volume of buffer being used in the electrophoresis units, because differing buffer

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volumes will result in differing electric fields strengths for DNA migration and will yield inconsistency in results. This is particularly important for matched samples within an experiment, if more than one electrophoresis chamber is required. 14. When using leukocytes from mammals and birds, and mammalian sperm, leave gels in stain for 10–15 min. Whole blood from amphibians requires approx 30 min.

References 1. Tice, R. R., Agurell, E., Anderson, D., Burlinson, B., Hartmann, A., Kobayashi, H., Miyamae, Y., Rojas, E., Ryu, J.-C., and Sasaki, Y.F. (2000) Single cell gel/comet assay: guidelines for the in vitro and in vivo genetic toxicity testing. Environ. Mol. Mutagen. 35, 206–221. 2. Krause, G., Wolz, L., and Scherer, G. (2001) Performing the “comet assay” for genetic toxicology applications. Life Sci. News. 7:1–3. 3. McNamee, J. P., McLean, J. R. N., Ferrarotto, C., and Bellier, P. V. (2000) Comet assay: rapid processing of multiple samples. Mutat. Res. 466, 63–69. 4. Hartmann, A., Agurell, E., Beevers, C., Brendler-Schwaab, S., Burlinson, B., Clay, P., Collins, A. R., Smith, A., Speit, G., Thybaud, V., and Tice, R. R. (2003) Recommendations for conducting the in vivo alkaline comet assay. Mutagenesis 118, 45–51. 5. Singh, N. P., Muller, C. H., and Berger, R. E. (2003) Effects of age on DNA double-strand breaks and apoptosis in human sperm. Fertil. Steril. 80, 1420–1430. 6. Strauss, G. H. (1991) Non-random cell killing in cryopreservation: implications for performance of the battery of leukocyte tests (BLT), I. Toxic and immunotoxic effects. Mutat. Res. 252, 1–15. 7. Collins, A. R., Dobson, V. L., Dusinská, M., Kennedy, G., and Stetina, R. (1997) The comet assay: what can it really tell us? Mutat. Res. 375, 183–193. 8. Schnurstein, A., and Braunbeck, T. (2001) Tail moment versus tail lengthapplication of an in vitro version of the comet assay in biomonitoring for genotoxicity in native surface waters using primary hepatocytes and gill cells from zebrafish (Danio rerio). Ecotoxicol. Environ. Safe. 49, 187–196. 9. Fairbairn, D. W., Olive, P. L., and O’Neil, K. L. (1995) The comet assay: a comprehensive review. Mutat. Res. 339, 37–59. 10. Collins, A. R. (2004) The comet assay for DNA damage and repair: principles, applications, and limitations. Mol. Biotechnol. 26, 249–261. 11. Anderson, D., Yu, T.-W., Dobrzynska, M. M., Ribas, G., and Marcos, R. (1997) Effects in the comet assay of storage conditions on human blood. Teratogen., Carcinogen. Mutat. 17, 115–125. 12. Tice, R. R., and Vasquez, M. (1999) Protocol for the application of the pH>13 alkaline single cell gel (SCG) assay to the detection of DNA damage in mammalian cells, in Integrated Laboratory Systems. Research Triangle Park, NC. http://www.cometassay.com/Files/raytice.doc

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13. Duty, S. M., Singh, N. P., Ryan, L., Chen, Z., Lewis, C., Huang, T., and Hauser, R. (2002) Reliability of the comet assay in crypopreserved human sperm. Hum. Reprod. 17, 1274–1280. 14. McKelvey-Martin, V. J., Ho, E. T., McKeown, S. R., Johnston, S. R., McCarthy, P. J., Rajab, N. F., and Downes, C. S. (1998) Emerging applications of the single cell gel electrophoresis (Comet) assay. I. Management of invasive transitional cell human bladder carcinoma. II. Fluorescent in situ hybridization Comets for the identification of damaged and repaired DNA sequences in individual cells. Mutagenesis 13, 1–8. 15. Horváthová, E., Dušinská, M., Shaposhnikov, S., and Collins, A. R. (2004) DNA damage and repair measured in different genomic regions using the comet assay with fluorescent in situ hybridization. Mutagenesis 19, 269–276. 16. Singh, N. P. (2000) A simple method for accurate estimation of apoptotic Cells. Exp. Cell Res. 256, 328–337. 17. Olive, P. L., Wlodek, D., and Banáth, J. P. (1991) DNA double-strand breaks measured in individual cells subjected to gel electrophoresis. Cancer Res. 51, 4671–4676. 18. Singh, N. P., Danner, D. B., Tice, R. R., McCoy, M. T., Collins, G. D., and Schneider, E. L. (1989) Abundant alkali-sensitive sites in DNA of human and mouse sperm. Exp. Cell Res. 184, 461–470. 19. Haines, G. A., Hendry, J. H., Daniel, C. P., and Morris, I. D. (2001) Increased levels of comet-detected spermatozoa DNA damage following in vivo isotopic- or X-irradiation of spermatogonia. Mutat. Res. 495, 21–32. 20. Duty, S. M., Singh, N. P., Silva, M., Barr, D. B., Brock, J. W., Ryan, L., Herrick, R. F., Christiani, D. C., and Hauser, R. (2003) The relationship between environmental exposures to phthalates and DNA damage in human sperm using the neutral comet assay. Environ. Health Persp. 111, 1164–1169. 21. Migliore, L., Naccarati, A., Zanello, A., Scarpato, R., Bramanti, L., and Mariani, M. (2002) Assessment of sperm DNA integrity in workers exposed to styrene. Hum. Reprod. 17, 2912–2918. 22. Knopper, L. D., Mineau, P., McNamee, J. P., and Lean, D. R. S. (2005) Use of comet and micronucleus assays to measure genotoxicity in meadow voles (Microtus pennsylvanicus) living in golf course ecosystems exposed to pesticides. Ecotoxicology 14, 323–335.

12 The Micronucleus Assay Determination of Chromosomal Level DNA Damage Michael Fenech

Summary The study of DNA damage at the chromosome level is an essential part of genetic toxicology because chromosomal mutation is an important event in carcinogenesis. Micronucleus assays have emerged as one of the preferred methods for assessing chromosome damage because they enable both chromosome loss and chromosome breakage to be measured reliably. Because micronuclei can only be expressed in cells that complete nuclear division a special method was developed that identifies such cells by their binucleate appearance when blocked from performing cytokinesis by cytochalasin-B, a microfilament-assembly inhibitor. The cytokinesis-block micronucleus (CBMN) assay allows better precision because the data obtained are not confounded by altered cell division kinetics caused by cytotoxicity of agents tested or suboptimal cell culture conditions. The method is now applied to various cell types for population monitoring of genetic damage, screening of chemicals for genotoxic potential and for specific purposes such as the prediction of the radiosensitivity of tumors and the interindividual variation in radiosensitivity. In its current basic form the CBMN assay can provide, using simple morphological criteria, the following measures of genotoxicity and cytotoxicity: chromosome breakage, chromosome loss, chromosome rearrangement (nucleoplasmic bridges), gene amplification (nuclear buds), cell division inhibition, necrosis and apoptosis. The cytosine arabinoside modification of the CBMN assay allows for measurement of excision repairable lesions. The use of molecular probes enables chromosome loss to be distinguished from chromosome breakage and importantly nondisjunction in nonmicronucleated binucleated cells can be efficiently measured. The CBMN technique therefore provides multiple and complementary measures of genotoxicity and cytotoxicity which can be achieved with relative ease within one system. The basic principles and methods (including detailed scoring criteria for all the genotoxicity and cytotoxicity end points) of the CBMN assay are described and areas for future development identified. From: Methods in Molecular Biology, vol. 410: Environmental Genomics Edited by: C. Cristofre Martin © Humana Press, Totowa, NJ

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Key Words: Chromosome breakage; chromosome loss; cytokinesis-block; micronuclei; micronucleus; nucleoplasmic bridges.

1. Introduction The observation that chromosome damage can be caused by exposure to ionizing radiation or carcinogenic chemicals was among the first reliable evidence that physical and chemical agents can cause major alterations to the genetic material of eukaryotic cells (1). Although our understanding of chromosome structure is incomplete, evidence suggests that chromosome abnormalities are a direct consequence and manifestation of damage at the DNA level - for example chromosome breaks may result from unrepaired double strand breaks in DNA and chromosome rearrangements may result from misrepair of strand breaks in DNA (2). It is also recognized that chromosome loss and malsegregation of chromosomes (nondisjunction) are an important event in cancer and ageing and that they are probably caused by defects in the spindle or the centromere or as a consequence of undercondensation of chromosome structure before metaphase (3–5). In the classical cytogenetic techniques chromosomes are studied directly by observing and counting aberrations in metaphases (6). This approach provides the most detailed analysis but the complexity and laboriousness of enumerating aberrations in metaphase and the confounding effect of artefactual loss of chromosomes from metaphase preparations has stimulated the development of a simpler system of measuring chromosome damage. It was proposed independently by Schmid (7) and Heddle (8) that an alternative and simpler approach to assess chromosome damage in vivo was to measure micronuclei (MNi), also known as Howell–Jolly bodies to haematologists, in dividing cell populations such as the bone marrow. The micronucleus (MN) assay in bone marrow and peripheral blood erythrocytes is now one of the best established in vivo cytogenetic assays in the field of genetic toxicology, however, it is not a technique that is applicable to other cell populations in vivo or in vitro and methods have since been developed for measuring MNi in a variety of nucleated cells in vitro. MNi are expressed in dividing cells that either contain chromosome breaks lacking centromeres (acentric fragments) and/or whole chromosomes that are unable to travel to the spindle poles during mitosis. At telophase, a nuclear envelope forms around the lagging chromosomes and fragments, which then uncoil and gradually assume the morphology of an interphase nucleus with the exception that they are smaller than the main nuclei in the cell, hence the term “micronucleus” (Fig. 1). MNi, therefore, provide a convenient and reliable index of both chromosome breakage and chromosome loss. Because MNi are

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expressed in cells that have completed nuclear division they are ideally scored in the binucleated stage of the cell cycle (9,10). Occasionally nucleoplasmic bridges between nuclei in a binucleated cell are observed. These are probably dicentric chromosomes in which the two centromeres were pulled to opposite poles of the cell and the DNA in the resulting bridge covered by nuclear membrane (Fig. 1). Thus nucleoplasmic bridges in binucleated cells provide an additional and complementary measure of chromosome rearrangement which can be scored together with the MN count. It is evident from the above that MNi can only be expressed in dividing eukaryotic cells. In other words, the assay cannot be used efficiently or quantitatively in nondividing cell populations or in dividing cell populations in which the kinetics of cell division is not well understood or controlled. Consequently,

Fig. 1. (A)The origin of micronuclei from lagging whole chromosomes and acentric chromosome fragments at anaphase. (B) The formation of a nucleoplasmic bridge from a dicentric chromosome in which the centromeres are pulled to opposite poles of the cell; the formation of an MN from the accompanying acentric chromosome fragment is also illustrated. The critical role of Cyt-B in blocking dividing cells at the binucleate stage is also indicated in this diagram. The example shown is for a hypothetical cell with two pairs of chromosomes only.

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there was a need to develop a method that could distinguish between cells that are not dividing and cells that are undergoing mitosis within a cell population. Furthermore, because of the uncertainty of the fate of MNi after more than one nuclear division it is important to identify cells that have completed one nuclear division only. These requirements are also necessary because cells divide at different rates in vivo and in vitro depending on the various physiological, genetic and micronutrient conditions. Several methods have been proposed based on stathmokinetic, flow cytometric and DNA labeling approaches but the method that has found most favor owing to its simplicity and lack of uncertainty regarding its effect on baseline genetic damage is the cytokinesis-block micronucleus (CBMN) assay (9–11). In the CBMN assay, cells that have completed one nuclear division are blocked from undergoing cytokinesis by using cytochalasin-B (Cyt-B) and are consequently readily identified by their binucleated appearance (Fig. 1). Cyt-B is an inhibitor of actin polymerization required for the formation of the microfilament ring that constricts the cytoplasm between the daughter nuclei during cytokinesis (12). The use of Cyt-B enables the accumulation of virtually all dividing cells at the binucleate stage in dividing cell populations regardless of their degree of synchrony and the proportion of dividing cells. MNi are then scored in binucleated cells only, which enables reliable comparisons of chromosome damage between cell populations that may differ in their cell division kinetics. The method was initially developed for use with cultured human lymphocytes (9,10) but has now been adapted to various cell types such as solid tumor and bone marrow cells (13,14). Furthermore, new developments have also occurred that allow (1) MNi originating from whole chromosomes to be distinguished from MNi originating from chromosome fragments (15–20), (2) the conversion of excision-repaired sites to MNi within one cell division (21), (3) the use of molecular probes to identify nondisjunction events in binucleated cells (22–24), and (4) the integration of necrotic and apoptotic cells within the CBMN assay (25,26). It has recently been proposed that the MN assay be used instead of metaphase analysis for genotoxicity testing of new chemicals. A recent special issue of Mutation Research has been dedicated to this topic (27). The current methodologies and data for the in vitro MN test were reviewed at the Washington International Workshop on Genotoxicity Test Procedures held in 1999 (28). The standard CBMN assay and its various modifications are described in detail in the next subheadings. The methods described are mainly applicable to cultured human lymphocytes; however, modifications of the assay for application to other cell types are included.

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2. Materials 2.1. Cytokinesis-Block Micronucleus Assay 1. Cytochalasin-B stock solution in 2 mg/mL of dimethyl sulfoxide (DMSO). 2. Ficoll Paque. 3. Hank’s balanced salt solution (HBSS): 0.137 M NaCl, 5.4 mM KCl, 0.25 mM Na2 HPO4 , 0.44 mM KH2 PO2 , 1.3 mM CaCl2 , 1.0 mM MgSO4 , 4.2 mM NaHCO3 . 4. RPMI 1640 culture medium + 10–15% heat inactivated fetal calf serum. 5. Phytohemagglutinin (PHA) (Glaxo Wellcome) stock: 2.25 mg/mL in H2 O (not required for transformed cell lines or tumor cell cultures). 6. Diff Quik (Lab-Aids, Australia) or Giemsa stain. 7. Depex (DPX) mounting medium (Electron Microscopy Sciences). 8. Sorensen’s phosphate buffer: 10 mM NaH2 PO4 ·H2 O/Na2 HPO4 .2H2 O, pH 6.9. 9. Acridine orange: 40 μg/mL in Sorensen’s phosphate buffer, pH 6.9.

2.2. Kinetochore Detection in Micronuclei 1. Serum samples from scleroderma patients of the CREST subtype. 2. Rabbit fluorescein isothiocyanate (FITC)-conjugated secondary anti-human IgG antibody. 3. Peroxidase-labeled rabbit anti-human IgG. 4. Diaminobenzidine (DAB) solution: 1 mg/mL in 0.5 M Tris base buffer stock, pH 7.6. 5. NiCl2 solution: 8% solution in 0.5 M Tris-base buffer stock, pH 7.6. Prepare immediately before use. 6. DAB reaction mixture: 1 mL of DAB solution, 3 mL of 0.5 M Tris base, pH 7.6, 25 μL of NiCl2 solution, 40 μL of 0.1M imidazole aqueous solution, and 10 μL of 30% hydrogen peroxide. Prepare immediately before use. 7. Neutral red solution: 0.1% in distilled water.

3. Methods 3.1. Standard Cytokinesis-Block Micronucleus Assay for Isolated Human Lymphocytes In this technique MNi are scored only in those cells that have completed one nuclear division after PHA stimulation. These cells are recognized by their binucleated appearance after they are blocked from undergoing cytokinesis by Cyt-B, which should be added before the first mitotic wave. Optimal culture conditions should yield 35–60% or more binucleates as a proportion of viable cells (i.e., all cells excluding necrotic and apoptotic cells) at 72 h after PHA stimulation. All equipment should have biosafety features to protect the operator, and solutions used in this procedure should be filter sterilized.

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3.1.1. Lymphocyte Isolation, Cell Culture, and Cell Harvesting 1. Fresh blood is collected by venepuncture in tubes with heparin as anticoagulant and stored at 22°C for less than 4 h before lymphocyte isolation. 2. The blood is then diluted 1:1 with sterile 0.85% NaCl solution and gently inverted to mix. 3. The diluted blood is overlaid gently on Ficoll Paque (Pharmacia) density gradients using a ratio of approx 1:3 (e.g., 2 mL of Ficoll Paque to 6 mL of diluted blood), being very careful not to disturb the interface. 4. The gradient is then centrifuged at 400g for 25–40 min at 22°C after carefully balancing the tubes. 5. The lymphocyte layer at the interface of Ficoll Paque and diluted plasma is collected with a sterile plugged Pasteur pipet and added to 3–5 times volume of HBSS at 22°C. The resulting cell suspension is centrifuged at 280–400g for 5–10 min depending on the volume. 6. The supernatant is discarded, and the cells are resuspended in 2–5 times volume HBSS and centrifuged at 180–400g for 5 min depending on the volume. 7. The supernatant is discarded and the cells resuspended in 1ml RPMI 1640 culture medium. 8. Cell concentration is then measured using a Coulter Counter or hemocytometer (according to the manufacturer’s instructions) and the concentration adjusted by the percentage of viable cells measured via trypan blue exclusion assay. 9. The cells are resuspended in RPMI 1640 medium containing 10–15% heatinactivated foetal calf serum at 0.5–1.0 × 106 cells/mL and cultured in a 0.75–1.0 mL volume in round-bottom tissue culture tubes (10 mm width). 10. Lymphocytes are then stimulated to divide by adding PHA (Glaxo Wellcome HA15) to each culture tube at 10 μLmL and incubated at 37°C with loose lids in a humidified atmosphere containing 5% CO2 . The concentration of PHA used has to be optimized depending on the purity and source of the reagent to ensure maximum number of binucleated cells after Cyt-B block. 11. Forty-four hours after PHA stimulation, 4.5 μg Cyt-B is added to each mL of culture [USE GLOVES AND FUME HOOD]: a 100-μL aliquot of Cyt-B stock solution is thawed, and 900 μL of culture medium added and mixed. Seventyfive microliters of the mixture is added to each 1 mL of culture to give a final concentration of 4.5 μg of Cyt-B/mL (other laboratories have successfully used 6.0 μg of Cyt-B/mL in their cultures). Culture tubes are then reincubated with loose lids. 12. Twenty-eight hours after addition of Cyt-B, cells are harvested by cytocentrifugation (Shandon Elliot). One hundred microliters of the culture medium is removed without disturbing the cells and then cells are gently resuspended in their tubes. One hundred to one hundred and twenty microliters of cell suspension is transferred to cytocentrifuge cups (Shandon Elliot) and centrifuged to produce two spots per slide (see Note 1). [Set the cytocentrifuge as follows: time: 5 min, speed: 100 g. Slides are removed from the cytocentrifuge and allowed to air dry for 10–2 min only and then fixed for 10 min in absolute methanol.

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13. The cells can be stained using a variety of techniques that can clearly identify nuclear and cytoplasmic boundaries. In our experience the use of “Diff Quik,” a commercial ready-to-use product, provides rapid and optimal results. (See Note 1.) 14. After staining, the slides are air-dried and cover slips placed over the cells using DPX mounting medium. This procedure is carried out in the fume hood and the slides are left to set in the fume hood and then stored indefinitely until required.

Duplicate cultures of control or genotoxin-treated cells should be set up and slides from each culture should be prepared. This is essential to obtain a measure of experimental variation, i.e., coefficient of variation, which should be quoted with each set of duplicate cultures (see Notes 2 and 3). This experimental design is summarized in Fig. 2. For fluorescence microscopy staining with acridine orange (40 μg/mL in Sorensen’s phosphate buffer, pH 6.9) is recommended. If a cytocentrifuge is not available slides can be prepared using the procedure, described below, for whole blood cultures. 3.1.2. Examination of Slides and Assessment of MN Frequency Slides are best examined at ×1000 magnification using a light or fluorescence microscope. Slides should be coded before analysis so that the scorer is not aware of the identity of the slide. A score should be obtained for slides from each duplicate culture ideally from two different scorers using identical

Fig. 2. An optimal sampling schedule for the in vitro MN assay that enables an estimation of experimental variation (results for A + C vs. B + D) as well as the effect of scorer bias (results for A + B vs. C + D).

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microscopes (see Notes 4 and 5). The number of cells scored (see Note 6) should be determined depending on the level of change in the MN index that the experiment is intended to detect and the expected standard deviation of the estimate. For each slide the following information should be obtained: 1. The number of micronuclei (MNi) in at least 1000 binucleate (BN) cells should be scored and the frequency of MNi per 1000 BN cells calculated. The criteria for scoring MNi in BN cells are detailed below. 2. The distribution of BN cells with zero, one, or more MNi; the number of MNi in a single binucleated cell normally ranges from 0 to 3 in lymphocytes of healthy individuals but can be greater than 3 on occasion depending on genotoxin exposure and age. 3. The frequency of micronucleated BN cells in at least 1000 BN cells. 4. The frequency of nucleoplasmic bridges in 1000 BN cells. Scoring criteria for nucleoplasmic bridges are described below. 5. The proportion of mono-, bi-, tri-, and tetranucleated cells per 500 cells scored. From this information the Nuclear Division Index (explained later) can be derived. 6. The number of dead or dying cells due to apoptosis or necrosis per 500 cells may also be scored on the same slide (scoring criteria for these cells are detailed later) while scoring the frequency of viable mono-, bi-, and multinucleated cells. (See Note 7.)

It is important to note that it is best to skip scoring a cell if one is uncertain on how to classify it. The basic elements of a typical score sheet are listed in Table 1. 3.1.3. Criteria for Selecting Binucleated Cells and Scoring Micronuclei, Nucleoplasmic Bridges, Nuclear Buds, Apoptotic and Necrotic Cells 1. Criteria for selecting binucleated cells: The cytokinesis-blocked cells that may be scored for MN frequency should have the following characteristics: a. The cells should be binucleated. b. The two nuclei in a binucleated cell should have intact nuclear membranes and be situated within the same cytoplasmic boundary. c. The two nuclei in a binucleated cell should be approximately equal in size, staining pattern and staining intensity. d. The two nuclei within a BN cell may be attached by a fine nucleoplasmic bridge that is no wider than 1/4th of the nuclear diameter. e. The two main nuclei in a BN cell may touch but ideally should not overlap each other. A cell with two overlapping nuclei can be scored only if the nuclear boundaries of each nucleus are distinguishable. f. The cytoplasmic boundary or membrane of a binucleated cell should be intact and clearly distinguishable from the cytoplasmic boundary of adjacent cells.

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Table 1 Information that Should Be Included on a Score Sheet for the Cytokinesis-Block Micronucleus Assay 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14.

Code number of each slide. Number of BN cells scored. The distribution of BN cells with 0, 1, 2, 3, or more MNi in at least 1000 BN cells. Total number of MNi in BN cells. The frequency of MNi in 1000 BN cells. The frequency of micronucleated BN cells in 1000 BN cells. Proportion of BN cells with nucleoplasmic bridges. Proportion of BN cells with nuclear buds. The proportion of mono-, bi-, tri-, and tetranucleated cells in 500 viable cells. The frequency of BN cells in a total of 500 viable cells. The nuclear division index. The proportion of cells that are undergoing apoptosis or necrosis in 500 cells. The nuclear division cytotoxicity index. Coefficient of variation for duplicate estimates of above parameters.

BN, binucleate; MNi, micronuclei.

Examples of the type of binucleated cells that may or may not be scored are illustrated diagrammatically in Fig. 3. The cell types that should not be scored for MN frequency include mono-, tri-, quadr-, and multinucleated cells, and cells that are necrotic or apoptotic (illustrated in Fig. 4). 2. Criteria for scoring micronuclei: MNi are morphologically identical to but smaller than nuclei. They also have the following characteristics: a. The diameter of MNi in human lymphocytes usually varies between 1/16th and 1/3rd of the mean diameter of the main nuclei which corresponds to 1/256th and 1/9th of the area of one of the main nuclei in a BN cell, respectively. b. MNi are nonrefractile and they can therefore be readily distinguished from artefact such as staining particles. c. MNi are not linked or connected to the main nuclei. d. MNi may touch but not overlap the main nuclei and the micronuclear boundary should be distinguishable from the nuclear boundary. e. MNi usually have the same staining intensity as the main nuclei but occasionally staining may be more intense.

Examples of typical MNi that meet the criteria set above are shown in Fig. 5. Examples of cellular structures that resemble MNi but should not be classified as MNi originating from chromosome breakage or loss are illustrated in Fig. 6.

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Fig. 3. Criteria for choosing binucleate cells in the CBMN assay. (A) Ideal binucleate cell. (B) Binucleate cell with touching nuclei. (C) Binucleate cell with narrow nucleoplasmic bridge between nuclei. (D) Binucleate cell with relatively wide nucleoplasmic bridge. Cells with two overlapping nuclei may be considered suitable to score as binucleated cells if the nuclear boundaries are distinguishable. Occasionally binucleated cells with more than one nucleoplasmic bridge are observed.

3. Criteria for scoring nuclear buds: Induction of gene amplification may lead to extrusion of amplified genes into nuclear buds (e.g., Fig. 6C, D) during S phase that are eventually detached from the nucleus to form a MN (Shimizu et al., 1998); it may be necessary to quantify the frequency of nuclei with nuclear bud formation if gene amplification is suspected (see Note 8). Nuclear buds are morphologically similar to micronuclei with the exception that they are clearly joined to the nucleus and having a continuous connection between the nucleoplasmic material in the nucleus and the nuclear bud. 4. Criteria for scoring nucleoplasmic bridges: Nucleoplasmic bridges are sometimes observed in binucleated cells following exposure to clastogens. a. They are a continuous link between the nuclei in a binucleated cell and are thought to be due to dicentric chromosomes in which the centromeres were pulled to opposite poles during anaphase. b. The width of a nucleoplasmic bridge may vary considerably but usually does not exceed 1/4th of the diameter of the nuclei within the cell. c. The nucleoplasmic bridge should have the same staining characteristics of the main nuclei. d. On very rare occasions more than one nucleoplasmic bridge may be observed within one binucleated cell. A binucleated cell with a nucleoplasmic bridge often contains one or more micronuclei. Examples of binucleated cells with nucleoplasmic bridges are illustrated in Figs. 1 and 5 (see Note 9).

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Fig. 4. The various types of cells that may be observed in the in vitro CBMN assay excluding binucleated cells. These cell types shown should not be scored for MN frequency. (A) Viable mono-, tri-, and quadrinuclear cells. (B) Mono- and binucleated cells at early stage of apoptosis when chromatin condensation has occurred but nuclear membrane has not disintegrated and late stage apoptotic cells with intact cytoplasm, no nucleus and apoptotic chromatin bodies within the cytoplasm. (C) Cells at the various stages of necrosis including early stages showing vacuolisation, disintegration of cytoplasmic membrane and loss of cytoplasm with an intact nucleus and late stages in which cytoplasm is partially or completely lost and nuclear membrane is visibly damaged and nuclear material is commencing to leak from the remnant nucleus.

5. Criteria for scoring apoptotic and necrotic cells: The various pathways and events that may be expected to occur in cultured lymphocytes exposed to a toxic agent are shown in Fig. 7. Cytogenetic genotoxicity assays that require hypotonic treatment for the preparation of interphase cells (for whole blood MN assay) or metaphase plates for chromosome analysis are not usable for cytotoxicity assays because hypotonic treatment may destroy necrotic cells and apoptotic cells making them unavailable for assay. Inclusion of necrosis and apoptosis is important for the accurate description of mechanism of action and measurement of cellular sensitivity to a chemical or radiation. Isolated lymphocyte culture assay or culture of cell lines does not require hypotonic treatment of cells for slide preparation, thus making it is possible to preserve the morphology of both necrotic and apoptotic cells. The use of Cyt-B, should make it easier to score apoptotic cells because it is expected to inhibit the disintegration of apoptotic cells into smaller apoptotic

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Fig. 5. Typical appearance and relative size of micronuclei in binucleated cells. (A) Cell with two micronuclei one with 1/3rd and the other 1/9th the diameter of one of the main nuclei within the cell. (B) Micronuclei touching but not overlapping the main nuclei. (C) A binucleated cell with nucleoplasmic bridge between main nuclei and two micronuclei. (D) A binucleated cell with six micronuclei of various sizes; this type of cell is rarely seen.

Fig. 6. Occasionally binucleated cells (or cells that resemble binucleated cells) may contain structures that resemble micronuclei but should not be scored as micronuclei originating from chromosome loss or chromosome breakage. These situations include (A) a trinucleated cell in which one of the nuclei is relatively small but has a diameter greater than 1/3 the diameter of the other nuclei, (B) dense stippling in a specific region of the cytoplasm, (C) extruded nuclear material that appears like a MN with a narrow nucleoplasmic connection to the main nucleus, and (D) nuclear blebs that have an obvious nucleoplasmic connection with the main nucleus.

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bodies. The latter process requires microfilament assembly (29) which is readily inhibited by Cyt-B (12). The following guidelines for scoring necrotic and apoptotic cells are recommended: a. Cells showing chromatin condensation with intact cytoplasmic and nuclear boundaries or cells exhibiting nuclear fragmentation into smaller nuclear bodies within an intact cytoplasm/cytoplasmic membrane are classified as apoptotic. b. Cells exhibiting a pale cytoplasm with numerous vacuoles and damaged cytoplasmic membrane with a fairly intact nucleus or cells exhibiting loss of cytoplasm and damaged/irregular nuclear membrane with a partially intact nuclear structure are classified as necrotic. These criteria and results for these measures with hydrogen peroxide have been recently reported elsewhere (26).

Figures 4 and 7 illustrate typical examples of necrotic and apoptotic cells. 3.1.4. Calculation of Nuclear Division Index (NDI) and Nuclear Division Cytotoxicity index (NDCI) 1. NDI is often calculated according to the method of Eastmond and Tucker (30). 500 viable cells are scored to determine the frequency of cells with one, two, three, or four nuclei and calculate the NDI using the formula: NDI = M1 + 2xM2 + 3xM3 + 4xM4/N where M1–M4 represent the number of cells with one to four nuclei and N is the total number of viable cells scored. The NDI and the proportion of binucleated cells are useful parameters for comparing the mitogenic response of lymphocytes and cytostatic effects of agents examined in the assay. 2. A more accurate assessment of nuclear division status is obtained if necrotic and apoptotic cells are included in the total number of cells scored because at higher toxic doses of chemicals tested one can expect a very large proportion of cells becoming nonviable. It is therefore important to note that both binucleate ratio and the NDI are overestimated if necrotic and apoptotic cells are not included when scoring cells. 3. A more accurate estimate of nuclear division status and cell division kinetics can be obtained using the following modified equation which takes account of viable as well as necrotic and apoptotic cells: NDCI = AP + Nec + M1 + 2xM2 + 3xM3 + 4xM4/N ∗

where NDCI = nuclear division cytotoxicity index, Ap = number of apoptotic cells, Nec = number of necrotic cells, M1–M4 = number of viable cells with one to four nuclei, and N* = total number of cells scored (viable and nonviable).

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Fig. 7. Genome damage and cell death biomarkers scored in the comprehensive CBMN assay. The CBMN assay allows the measurement of all possible outcomes after a genome damage event. In this assay a cell with genome damage may either undergo cell death via apoptosis or necrosis or alternatively may survive and undergo further nuclear division. In the latter case dividing cells are recognized as binucleated cells (BNCs) by blocking cytokinesis with Cyt-B. BNCs are then scored for the following genome damage events: (A) micronuclei (MNi) that originate from lagging whole chromosomes or broken chromosome fragments and are therefore a marker of chromosome breakage and chromosome loss events, the latter being due to defects in centromere or spindle structure, (B) nucleoplasmic bridges (NPB) that originate from dicentric chromosomes caused by misrepair of chromosome breaks and are therefore a marker of chromosome rearrangement, and (C) nuclear buds (NBUD) that are the mechanism by which the nucleus eliminates amplified DNA resulting from breakage– fusion–bridge cycles generated by NPB. NBUD therefore provide a measure of gene amplification. An increase in MNi, NPB, and NBUD is indicative of an increase in genome instability commonly seen in cancer.

3.2. Measurement of Excision-Repaired DNA Lesions in G0 /G1 HumanLymphocytes Using the Cytosine Arabinoside Micronucleus Assay in Human Lymphocytes After assessing the MN response in human G0 lymphocytes after exposure to a variety of genotoxins, it became evident that the extent of MN formation in relation to cytotoxicity was low for chemicals and ultraviolet radiation which

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mainly induce base-lesions and adducts on DNA rather than strand breakage or spindle damage (21). We hypothesized that this was due to either efficient repair of the lesions or that such sites, if left unrepaired, do not convert to a double stranded break in DNA after one round of DNA synthesis. Furthermore, we reasoned that inhibition of excision repair by cytosine arabinoside (ARA) would result in the conversion of such base lesions to a single-stranded break that would become a double-stranded break after DNA synthesis, leading to the production of an acentric fragment which would then be expressed as a MN within one division cycle (21,31). 1. Using this concept (illustrated in Fig. 8), we showed that addition of ARA (1 μg/mL in culture medium) during the first 16 h of lymphocyte culture (i.e., before DNA synthesis) did result in a dramatic increase (10-fold or greater) in the MN dose– response after ultraviolet or MNU treatment. However, the ARA-induced increase after X-ray exposure was only 1.8-fold, as would be expected from the proportion of DNA adducts or base lesions relative to the induction of DNA strand breaks. This method has since been used to identify pesticides that induce excision repair and to distinguish between genotoxic agents that do or do not induce excision repair (32).

Fig. 8. A schematic diagram explaining the mechanism for the conversion by ARA of an excision-repairable DNA lesion to a MN within one division cycle.

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2. The ARA protocol is an important adjunct to the basic CBMN assay and should be attempted, particularly if strong cytotoxic effects are observed in conjunction with weak MN induction. 3. Precise measurement of excision-repaired DNA lesions using the ARA method is possible only using the CBMN assay because (a) the conversion of excisionrepaired DNA lesions to MN occurs only in cells that have completed nuclear division and (b) the addition of ARA may also result in significantly altered cell division kinetics, which could confound results in MN assays without Cyt-B. 4. ARA inhibition of DNA polymerase may cause DNA strand breaks in cells undergoing replicative DNA synthesis. Therefore, it is only possible to use this method in PHA-stimulated G0 lymphocytes with ARA exposure occurring during the G1 phase and before the S-phase, because excision repair is activated during G1 . 5. In practice this means that cells are cultured in the presence of ARA during the first 16–20 h after PHA stimulation, after which the cells are washed to remove ARA and incubated in culture medium containing deoxycytidine to reverse ARA inhibition of DNA polymerase. 6. After these steps, the standard CBMN protocol (described earlier) is followed. For more procedure details and typical results refer to Fenech and Neville (21) and Surrales et al. (32).

3.3. CBMN Assay in Other Cell Culture Systems 3.3.1. Whole Blood Cultures for Human Lymphocytes 1. The CBMN assay in human lymphocytes can also be performed using whole blood cultures. 2. Typically 0.4–0.5 mL of whole blood is added to 4.5 mL of culture medium (e.g., RPMI 1640) supplemented with fetal calf serum containing l-glutamine, antibiotics (optional) and PHA. 3. Cyt-B is added at 44 h post PHA stimulation. The recommended optimal concentration of Cyt-B for accumulating binucleated cells in whole blood cultures is 6 μg/mL (33). 4. The binucleated lymphocytes are harvested 28 h after addition of Cyt-B as follows: a. The cells are centrifuged gently (300g) for 5 min and the supernatant culture medium is removed. b. The cells are hypotonically treated with 7 mL of cold (4°C) 0.075M KCl to lyse red blood cells and centrifuged immediately (300g) for 8 min. c. The supernatant is removed and replaced with 5 mL of fixative consisting of methanol–acetic acid (3:1) (the fixative should be added whilst agitating the cells to prevent clumps forming).

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d. The cells are then centrifuged again at 300g for 8 min and washed with two further changes of fixative. e. The cells are resuspended gently, and the suspension is dropped onto clean glass slides and allowed to dry. As an alternative it is also possible to isolate the binucleated lymphocytes directly from the whole blood culture using Ficoll gradients and then transfer cells to slides by centricentrifugation before fixation and staining (unpublished observation) which precludes the requirement for hypotonic treatment and enables optimal preservation of the cytoplasm. 5. Staining of cells can be done using either 10% Giemsa in 0.1 M potassium phosphate buffer, pH 7.3, for light microscopy or acridine orange (10 μg/mL in 0.1 M phosphate- buffered saline, pH 6.9, for fluorescence microscopy).

3.3.2. Murine Lymphocyte Cultures 1. Lymphocytes are isolated either from the spleen or peripheral blood and cultured according to the procedures described by Fenech et al. (34). 2. Because murine lymphocytes have shorter cell division cycles than human lymphocytes, it is essential to add Cyt-B no later than 18 h after stimulation by mitogen and to harvest the cells 20 h later. Depending on the culture conditions it is possible to obtain good binucleate ratios even at 72 h post mitogen stimulation.

3.3.3. Other Primary Cell Cultures Including Tumor Cell Cultures The CBMN assay can be readily adapted to other primary cell types to assess DNA damage induced in vitro, in vivo or ex vivo. The most important points to remember are to (1) ensure that MNi are scored in the first nuclear division following the genotoxic insult and (2) perform preliminary experiments to determine the concentration of Cyt-B and incubation time at which the maximum number of dividing cells will be blocked at the binucleate stage (see Note 10). It is also important to remember that Cyt-B may take up to 6 h before it starts to exert its cytokinesis-blocking action (unpublished observation). 1. When using established or primary cell lines from dividing cell populations it is usual to add Cyt-B shortly after exposure to genotoxin to capture all cells undergoing their first nuclear division as binucleated cells—this usually requires an incubation period of about 24–48 h, depending on the cell cycle time, before harvesting the cells. 2. Attached cells can be trypsinized and then prepared by cytocentrifugation as described for human lymphocytes. Specific methods have been described for use with nucleated bone-marrow cells (14), lung fibroblasts (35), skin keratinocytes (36), and primary tumor cell cultures (13). 3. It is generally more practical to assess in vivo induction of micronuclei by blocking cytokinesis in dividing cells after the cells have been isolated from the animal and

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placed in culture medium in the presence of Cyt-B; this approach has proven to be successful with a variety of cell types including fibroblasts, keratinocytes, and nucleated bone marrow cells.

3.4. Micronucleus Assay in Cell Lines or Cultured Tumor Cells with or without Cytokinesis Block 1. There is some debate that Cyt-B, used to accumulate binucleated cells, may interfere with the expression of MN (28). Studies with normal cells do not show an induction of MNi by Cyt-B or a dose–response effect of Cyt-B with MN frequency in binucleated cells at doses that are usually used to block cells in cytokinesis (10,37–39). A recent study suggests that MN expression induced by

Fig. 9. (A) Comparison of the MN dose–response in human lymphocytes exposed in vitro in G1 /S/G2 to mitomycin-C (MMC) measured either in mononucleated cells in cultures without Cyt-B (solid black bars) or in binucleated cells in cultures with Cyt-B (white bars). (B) The level of dividing cells assessed by measuring the percentage of binucleated cell in the cytokinesis-blocked cultures. It is evident that the assay without Cyt-B underestimates the extent of genetic damage induced by MMC, particularly at doses that inhibit nuclear division. The data represent the mean ± 1 SE of three replicate cultures.

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3.

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spindle poisons may be less than expected in the cytokinesis-blocked BN cells because of pole-to-pole distance shortening which may increase the probability of reinclusion of lagging chromosome fragments or whole chromosomes back into a nucleus but this did not diminish the effectiveness of the CBMN assay (40). There has been an increased interest in exploring further the possibility of performing the in vitro MN assay without Cyt-B to minimise the possible confounding effect of Cyt-B while running the potential risk of obtaining a false-negative result because of inadequate control of cell division kinetics (i.e., inhibition of nuclear division inhibits MN expression). Whilst the evidence of obtaining a false-positive result with the CBMN assay in normal cells is lacking, there is already adequate evidence that performing the MN assay in a manner that does not account for inhibition of nuclear division can lead to false-negative results or an underestimate of MN induction in human lymphocyte cultures (10,11,41) and an example of this defect of MN assays without Cyt-B is shown in Fig. 9. Nevertheless, recent studies comparing the MN assay with or without Cyt-B suggest that if cell lines with good growth characteristics are used and culture and nuclear division conditions are optimal it is possible to obtain comparable results between the CBMN assay and the MN assay without Cyt-B when strong clastogens are tested (42,43). A mathematical model of MN expression predicts that (1) scoring MN in BN cells is the most reliable way of determining MN frequency and (2) scoring MN in mononucleated cells in cultures without cytokinesis block is likely to generate false-negative results when nuclear division is significantly inhibited by the chemical tested or the culture conditions do not allow an optimal number of dividing cells (44). Consequently, results for MN frequency obtained by scoring micronuclei in mononucleated cells in cultures without Cyt-B cannot be considered conclusive and that a negative result with this system should be confirmed using the CBMN assay.

3.5. Molecular Techniques for Measuring Chromosome Loss in Micronuclei and Nondisjunction To take full advantage of the ability of the CBMN assay it is essential to distinguish between MNi originating from whole chromosomes or acentric fragments. This is best achieved by using probes that are specific for the centromeric DNA or antibodies that bind to the kinetochore proteins that are assembled at the centromeric regions of active chromosomes. The use of MN size as a discriminant is not recommended for human cells or other cell types in which the size of chromosomes is heterogenous because a small MN may contain either a fragment of a large chromosome or a whole small chromosome. The simplest and least expensive technique to use is the anti-kinetochore antibody method (45) but this approach does not distinguish between unique chromosomes and may not detect chromosome loss occurring as a result of

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absence of kinetochores on inactive centromeres (46). The use of in situ hybridization (ISH) to identify centromeric regions is more expensive and laborious but it can provide greater specificity; for example, centromeric probes for unique chromosomes can be used which also enables the detection of nondisjunctional events (i.e., unequal distribution of homologous chromosomes in daughter nuclei) in binucleated cells (17). In this chapter only the kinetochore antibody method is described. For details on the use of centromere detection by ISH refer to the papers by Farooqi et al. (17), Hando et al. (18), Ehajouji et al., (23,47), Schuler et al. (24), and the chapter by Pacchierotti and Sgura in this volume. The types of results that can be expected with the various techniques are illustrated in Fig. 10. 3.5.1. Kinetochore Detection in MNi in the CBMN Assay 3.5.1.1. Slide Preparation. 1. In this technique BN cells are accumulated as described in the standard CBMN assay, transferred to a slide using a cytocentrifuge, air-dried for 5 min, and fixed in methanol for 10 min and air-dried again. 2. At this stage slides may either be processed immediately or stored for a maximum of 3 months in a sealed desiccated box in a nitrogen atmosphere above liquid nitrogen. 3. For detection of kinetochores the stored slides are removed from the nitrogen atmosphere and allowed to equilibrate at room temperature within the sealed box.

3.5.1.2. Kinetochore Detection 1. The anti-kinetochore sera may either be obtained commercially or from an immunology clinic that has serum samples from scleroderma patients of the CREST subtype (48). Use of the latter sera would require Human Ethics approval and consent from the donor patient. 2. The sera should be tested on slides of metaphase spreads of cultured cells using a rabbit FITC-conjugated secondary anti-human IgG antibody and examined by fluorescence microscopy. Only sera that appear to react exclusively with kinetochores on metaphase chromosomes should be selected for the assay. 3. The use of FITC-conjugated secondary antibody to visualise kinetochores is a direct technique but requires the use of a fluorescence microscope and nonpermanent slide preparations; the fluorescence technique has been described in detail elsewhere (45). An alternative procedure is to use an immunoperoxidase staining method that allows permanent slide preparations to be obtained (49), which is more practical for routine screening and is described in the next paragraph. 4. In the immunoperoxidase technique, fixed slides are incubated overnight at 20°C in a humidity chamber with the primary anti-kinetochore antibody diluted 1:40 in Tris–saline buffer, pH 7.6 (6.0 g of Tris base/L saline). 5. Negative control slides are exposed to the diluted serum of a normal healthy individual.

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Fig. 10. The use of molecular techniques for identifying (A) a MN originating from a lagging acentric chromosome fragment, (B) an MN originating from a lagging whole chromosome, and (C) nondisjunction of a chromosome leading to aneuploid daughter nuclei. The white spots in the nuclei and micronuclei of the binucleated cells on the left of each panel show the centromeric or kinetochore pattern of staining when pancentromeric probes or kinetochore antibodies are used. The white spots in the nuclei and micronuclei of the binucleated cells on the right of each panel show the pattern of centromeric staining when a centromeric probe specific to the chromosomes involved in MN formation or nondisjunction events is used. The example shown is for a hypothetical cell with only two pairs of chromosomes. 6. The following day the slides are washed by dipping them for 30 s in the same Tris–saline buffer used to dilute the antibody. 7. Slides are then drained without drying, and incubated for 3 h with peroxidaselabeled rabbit anti-human IgG. 8. Again, slides are then drained without drying in preparation for the peroxidase histochemical reaction.

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9. The histochemical method that gives best contrast is the nickel chloride–imidazole modification of the standard DAB reaction, which produces a black precipitate (50,51). 10. The DAB reaction mixture is prepared just before use and applied immediately to slides through a 0.22-μm filter to minimize nonspecific precipitation on the slides. 11. Slides should be stained in batches including a slide with the negative control serum. 12. The reaction is allowed to proceed for 1 min at 20°C and then stopped by draining the slides and rinsing in water. 13. The slides are then air-dried, counterstained with the nuclear stain neutral red (0.1% in distilled water) for 30 s, washed in water, air-dried, and mounted to give permanent preparations.

3.5.1.3. Scoring Procedure. 1. Scoring of kinetochore status of MNi is restricted to those binucleated cells in which a minimum of 20 kinetochores within each nucleus is observed. 2. A minimum of 100 MNi should be classified according to whether they contain kinetochores or not and the number of kinetochores within each MN should be noted. 3. The final value for the proportion of MNi with kinetochores is determined by the following formula: [Ps – Pc ]/[1 – Pc ], where Pc is the proportion of MNi that has a positive peroxidase reaction in slides exposed to normal control serum and Ps is the proportion of MNi that have a positive peroxidase reaction in slides exposed to anti-kinetochore serum.

3.6. Treatment Schedules for In Vitro Chemosensitivity Testing 1. Ideally each chemical should be tested for its genotoxic potential at the various stages of the cell cycle. Because human peripheral blood lymphocytes are in the G0 phase when collected they are ideal for assessing damage at this stage. 2. However, cells are expected to be more sensitive to genotoxic effects during S phase, G2 phase, and M phase and for this purpose it essential to expose cell cultures when most cells are dividing. Because MN expression requires one nuclear division to be completed, the period between treatment and harvest time needs to allow for this. 3. With human peripheral blood lymphocytes treated in G0 it is necessary to accumulate binucleated cells as early as possible and for as long as possible to ensure that even cells experiencing mitotic delay are examined. Typically the standard protocol of adding Cyt-B at 44 h and harvesting cells at 72 h should suffice for this purpose. However, it is equally practical to add Cyt-B at 24 h and harvest cells at 96 h, maximizing the number of late dividing cells available for analysis. 4. If treatment of cells in S, G2 , and M phases is required, as would be the case with tumor cell cultures, then exposure to the chemical should occur during logarithmic

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growth phase of the culture, followed shortly afterward with Cyt-B to accumulate dividing cells, and cells are then harvested between 6 h and 24 h later depending on the stage of the cell cycle that is being examined. 5. At the very early harvest times mainly cells exposed in G2 or late S phase are accumulated as binucleated cells while at the later harvest time cells exposed in all stages of the cell cycle are blocked in the binucleate stage. Thus the harvest time relative to Cyt-B addition would affect the type of cell examined. 6. Typical schedules for use of the CBMN assay for in vitro genotoxicity testing are summarized in Table 2. 7. The use of a metabolic activation system such as S9 mix should be considered as an option when testing new chemicals but this could limit the exposure period because of the possible cytotoxicity of S9 to the target cells. A better option may be the use of metabolically competent cells such as genetically modified MCL-5 cells (52).

3.7. Future Developments It is evident that the MN assay has evolved into a robust assay for genetic damage with applications in ecotoxicology (53), nutrition (54), radiation sensitivity testing both for cancer risk assessment (55) and optimization of radiotherapy (13,56), biomonitoring of human populations (57), and, importantly, testing of new pharmaceuticals and agrichemicals (27,28). There is little doubt that there is a need for an automated scoring system for quicker and more reliable data acquisition that would ideally be based on the scoring of slides also prepared for visual scoring—this should enable consistent results to be obtained that are not influenced by the interindividual and temporal variability of human scorers. For this goal to be achieved it is essential that scoring criteria are well developed and that a robust slide preparation protocol be put in place and that slide preparations be permanent so that they can be reexamined visually if necessary. Currently image analysis systems have been developed for automated scoring of micronuclei in mammalian cells (58–62), but these systems do not take account of other important events such as nucleoplasmic bridges, nuclear buds, necrosis, apoptosis, and cytostasis, which are essential for the correct interpretation of the result obtained (26). In the future we should expect to have an automated system that can score reliably the various end points possible with the cytokinesis block MN assay outlined in this chapter. Finally, it is also essential to keep abreast of more recent developments in our understanding of MN formation and events that may alter expression of this end point. Some notable examples are (1) the formation of micronuclei as a result of gene amplification in which the cell eliminates excess amplified DNA directly from the nucleus, during S phase, into a MN produced by nuclear budding

Harvest cells Harvest cells

Add Cyt-Ba

Add Cyt-Ba Add Cyt-B Harvest cells Harvest cells

(1) Wash out ARA(2) Fresh medium with IL-2 and DC

Add PHA

CBMN AssayG1 /S Exposure

Harvest cells Harvest cells

Harvest cells Add test agent

Add test agent Add PHA

CBMN AssayG1 /S/G2 /M Exposure

Harvest cells Harvest cells

Add Cyt-Ba

Add test agent (1) Add PHA(2) Add ARA

CBMN/ARA Assay, G0 Exposure

Harvest cells

Add Cyt-B

CBMN AssayG1 /S/G2 /M Exposure

Cell Lines in Log Phaseb

The proposed protocols assume that the test agent is retained in the culture medium even after Cyt-B is added. However, it may also be desirable to remove test chemical by replacing culture medium (1) after a brief exposure period to test chemical or (2) just prior to addition of Cyt-B. In the latter case IL-2 should be added to fresh medium for lymphocyte cultures. ARA, cytosine arabinoside; Cyt-B, cytochalasin-B; DC, deoxycytidine; IL-2, interleukin-2; PHA, phytohemagglutinin. a Alternatively Cyt-B could be added at 24 h. b This treatment schedule is the most appropriate for testing chemosensitivity in tumor cell cultures.

20 24 44 48 72 96

Add test agent

Add test agent Add PHA

–4 0

16

CBMN Assay G0 Exposure

Culture Time (hours)

Peripheral Blood Human Lymphocytes

Table 2 Typical Protocols Used for Testing Micronucleus Induction by a Chemical or Radiation

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(63,64); (2) the use of the cytokinesis block MN assay to measure breakagefusion-bridge cycles that are the one of the hallmarks of genomic instability in preneoplastic cells and folate-deficient cells (64,65); (3) the observation that treatment with specific mitotic spindle inhibitors may cause mitotic slippage leading to polyploid nuclei and micronuclei and therefore implicating that it may be useful to score not only MNi in binucleated cells but also MNi in mononucleated cells in cytokinesis-blocked cultures (47); and (4) the possible elimination of micronucleated cells and micronuclei by apoptosis (66,67). Furthermore, scoring criteria in the cytokinesis-block MN assay are continually being reviewed as part of the activity of the HUMN project (www.humn.org), and a more comprehensive and recommended description of scoring criteria with photomicrographs has been published recently (68). All of the preceding point to the fact that the full potential of the cytokinesisblock MN assay as a “cytome” assay is readily achievable once all the morphological end points of cytotoxicity, cytostasis, and DNA damage are integrated into the system. Finally it is important to note that micronuclei can also be scored directly in cells originating from other tissues that are actively dividing in vivo such as in erythrocytes, colonocytes, and oral mucosa cells and that MN assays are being also successfully applied to determine optimal dietary requirements for genome health maintenance (69,70). 4. Notes 1. In our experience, most of the problems in the CBMN assay arise during slide preparation and staining because the quality of the score depends on the quality of the slide. Main points to note: (a) avoid cell clumps by gently resuspending cells before harvest and transfer to slides; (b) maintain a moderate cell density so that it is relatively easy to identify cytoplasmic boundaries; (c) stain only one slide initially to ensure that staining is optimal before staining the whole batch. 2. The use of duplicate cultures is critical for producing robust results also because it allows the measurement of the intraexperimental coefficient of variation. Cytogenetic assays should be subject to the same rigor as analytical assays that typically reject duplicate results with a CV greater than 10%. Owing to the visual scoring, greater latitude in the acceptable CV is understandable. In our experience and the results of international inter-laboratory scoring comparison (69) CVs greater than 40% are not acceptable for baseline data and with radiation exposed cultures in which more than 100 MN per 1000 BN cells are induced, CVs less than 20% are expected. 3. Scores from inexperienced personnel (e.g., students, new staff) should not be relied on until they are able to achieve acceptable CVs (no greater than 40%) for repeat scores of standard control slides. 4. Inter-scorer variability is one of the key sources of variation in the MN assay (71). It is therefore essential that the same scorers are maintained throughout a

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6.

7.

8.

9.

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single study and ideally two scorers are used, each providing a count from each of the duplicate cultures and their mean values calculated as indicated in Fig. 2. An alternative approach is to calibrate scorers by using a common set of standard slides with “low,” “medium,” and “high” MN frequencies. The scores of each scorer on the standard slides can then be used to calculate a corrected value. The latter approach is still in development but worth noting as an option because it can take account of differences in the visual capacity of scorers within the same laboratory and between laboratories. Another important source of variability between scorers and between laboratories is the quality of the microscopes and their optics. In our experience, scoring of nucleoplasmic bridges is influenced by the quality of the microscope because fine bridges can be missed with low-quality optics. The main issue here is for scorers to avoid switching microscopes during experiments and for the laboratory manager to upgrade the optics of the microscopes to a uniform and high level whenever possible. We recommend a system that will provide the best clarity and contrast of image and the flattest image possible. The system should be capable of up to ×1000 magnification. One of the most common questions is the number of BN cells to be scored in the CBMN assay. The accepted protocol is to score a minimum of 1000 BN cells per treatment or time point, although reports vary from between 500 BN cells and 2000 BN cells. An alternative approach is to keep on scoring BN cells until a fixed number of micronuclei are observed (e.g., 45 micronuclei). The latter has the advantage that more BN cells are scored when fewer MNi are induced, thus maintaining similar statistical power across different treatments. The main disadvantage is that more than 2000 cells may have to be scored in cultures with low MN frequency. In our experience scoring 1000 BN cells from each of the duplicate cultures always yields robust results. With respect to scoring slides it is best to first score the frequency of mononucleated, binucleated, multinucleated, apoptotic, and necrotic cells to determine the NDI and NDCI indeces. Then focus on scoring binucleated cells for the presence of micronuclei, nucleoplasmic bridges, and nuclear buds to determine the genome damage rate. It should be noted that the use of nuclear buds within the CBMN assay is expected to increase because of the consistent significant relationship of this biomarker of gene amplification with nucleoplasmic bridges and micronuclei (64). It is therefore recommended that nuclear buds be scored. In our experience the expression of this biomarker may be more prevalent after long-term exposure (>3 days) to a genotoxic agent, which is consistent with the notion that it may take 3 or more nuclear divisions for breakage–fusion–bridge cycles to generate sufficient amplified DNA to be eliminated by nuclear budding. When scoring nucleoplasmic bridges in binucleated cells it is important to note that the score may depend on the frequency of binucleated cells with nuclei that touch or overlap. This is because nucleoplasmic bridges are more likely to be visible in binucleated cells in which the nuclei are clearly separated from each other.

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10. In maximizing the number of cytokinesis-blocked binucleated cells by increasing exposure time to Cyt-B, there is the risk of also increasing the proportion of cytokinesis-blocked multinucleated cells that arise from binucleated cells that attempt another nuclear division while cytokinesis-blocked. Ideally the proportion of binucleated among cytokinesis-blocked cell should be in excess of 80%. The proportion of binucleated cells among all cells will depend on the proportion of dividing cells in the culture. The latter depends on the cell line or tumor cells and the culture conditions.

Acknowledgments The development of the CBMN assay was the result of research I performed at the Medical School of the Flinders University of South Australia in Prof. Alec Morley’s laboratory and CSIRO Health Sciences and Nutrition with the support of the Anti-Cancer Foundation of the Universities of South Australia. I also acknowledge the important role of Ms. J. Rinaldi, Ms. C. Aitken, Ms. S. Neville, Ms. J. Turner, Ms. F. Bulman, Ms. C. Salisbury, Mr. P. Thomas, Mr. J.Crott, Ms. S. Brown, and Mr. W. Greenrod, who have contributed significantly to the more recent research effort. I thank Prof. Micheline KirschVolders and Prof. Wushou P. Chang for critically reading the manuscript and for their constructive suggestions. References 1. Evans, H. J. (1977) Molecular mechanisms in the induction of chromosome aberrations, in: Progress in Genetic Toxicology (Scott, D., Bridges, B. A., and Sobels, F. H., eds.), Elsevier North Holland Biomedical Press, Amsterdam, pp.57–74. 2. Savage, J. R. K. (1993) Update on target theory as applied to chromosomal aberrations. Environ. Mol. Mutagen.22, 198–207. 3. Evans, H. J. (1990) Cytogenetics: overview. Prog. Clin Biol. Res. 340B, 301–323. 4. Dellarco, V. L., Mavournin, K. H., and Tice, R. R. (1985) Aneuploidy and health risk assessment: current status and future directions. Environ. Mutagen. 7, 405–424. 5. Guttenbach, M., and Schmid, M. (1994) Exclusion of specific human chromosomes into micronuclei by 5-azacytidine treatment of lymphocyte cultures. Exp. Cell Res. 211, 127–132. 6. Natarajan, A. T., and Obe, G. (1982) Mutagenicity testing with cultured mammalian cells: cytogenetic assays, in Mutagenicity: New Horizons in Genetic Toxicology (Heddle J. A. ed.), Academic Press, New York, pp.17–213. 7. Schmid, W. (1975) The micronucleus test. Mutat. Res. 31, 9–15. 8. Heddle, J. A. (1973) A rapid in vivo test for chromosome damage. Mutat. Res. 18, 187–192.

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68. Fenech, M.; Chang, W. P., Kirsch-Volders, M., Holland, N., Bonassi, S., and Zeiger, E. (2003) HUMN Project: Detailed description of the scoring criteria for the cytokinesis-block micronucleus assay using isolated human lymphocyte cultures. Mutat. Res. 534, 65–75. 69. Fenech, M., Baghurst, P., Luderer, W., Turner, J., Record, S., Ceppi, M., and Bonassi, S. (2005) Low intake of calcium, folate, nicotinic acid, vitamin E, retinol, -carotene and high intake of pantothenic acid, biotin and riboflavin are significantly associated with increased genome instability – results from a dietary intake and micronucleus index survey in South Australia. Carcinogenesis 26, 991–999. 70. Fenech, M. (2005) The Genome Health Clinic and Genome Health Nutrigenomics concepts: diagnosis and nutritional treatment of genome and epigenome damage on an individual basis. Mutagenesis 20, 255–269. 71. Fenech, M., Bonassi, S., Turner, J., Lando, C., Ceppi, M., Chang, W. P., Holland, N., Kirsch-Volders, M., Zeiger, E., Bigatti, M. P., Bolognesi, C., Cao, J., De Luca, G., Di Giorgio, M., Ferguson, L. R., Fucic, A., Garcia Lima, O., Hadjidekova, V. V., Hrelia, P., Jaworska, A., Joksic, G., Krishnaja, A. P., Lee, T.-K., Martelli, A., McKay, M. J., Migliore, L., Mirkova, E., Müller, W.-U., Odagiri, Y., Orsiere, T., Scarfì, M. R., Silva, M. J., Sofuni, T., Suralles, J., Trenta, G., Vorobtsova, I., Vral, A., and Zijno, A. (2003) Intra- and inter-laboratory variation in the scoring of micronuclei and nucleoplasmic bridges in binucleated human lymphocytes. Results of an international slide-scoring exercise by the HUMN project. Mutat. Res. 534, 45–64.

13 Fluorescence In Situ Hybridization for the Detection of Chromosome Aberrations and Aneuploidy Induced by Environmental Toxicants Francesca Pacchierotti and Antonella Sgura

Summary Numerous chemicals as well as ionizing radiations of different qualities can induce damage to chromosome integrity and/or chromosome distribution at mitosis and meiosis. Fluorescence in situ hybridization with many kinds of probes complementary to different DNA sequences has been developed to detect and quantify specific types of structural and numerical aberrations in metaphase and interphase cells. Probes for the whole sequence of specific chromosomes are applied to metaphase cells to detect stable rearrangements, which can be relevant for cell transformation and tumor development. Probes that recognize the pericentromeric sequence of all chromosomes of a species are used to distinguish micronuclei that contain centromeres from those that do not, and, on this basis, infer whether they were induced by chromosome loss or chromosome break. Conversely, probes that recognize the pericentromeric sequence of specific chromosomes, if available, can be used to count the number of chromosomes in interphase nuclei to detect hyperploid cells possibly produced by chromosome nondisjunction. Finally, probes that hybridize to the telomeric sequence common to all chromosomes can be used to label telomeres and quantify their individual and mean length, a cell parameter that has been recently related to genomic instability. All these different techniques share the basic principles of fluorescence hybridization, with some specific adjustments. This chapter provides a general protocol useful for any of the above applications and comments to specific requirements or modifications. Key Words: Aneuploidy; charge-coupled-device camera; chromosome painting; environmental mutagens; fluorescence in situ hybridization; telomeres; translocations.

From: Methods in Molecular Biology, vol. 410: Environmental Genomics Edited by: C. Cristofre Martin © Humana Press, Totowa, NJ

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1. Introduction Numerous synthetic chemicals and industrial by-products are introduced into the environment that possess genotoxic activity. These chemicals, or their metabolites produced by biotransformation reactions occurring in most eukaryotes, can react with DNA, forming covalently bound adducts that distort the double helix and eventually activate repair mechanisms that cause singleor double-strand breaks in the helix backbone. Ionizing radiations transferring their energy into a living organism are also well known to induce gene mutations and chromosome aberrations by a free radical mediated mechanism capable of directly causing DNA strand breaks (1). Other chemicals do not react with DNA, but may bind to proteins forming microtubules, including those of the metaphase spindle that organize chromosome segregation at mitosis and meiosis. These chemicals, which include antineoplastic drugs but also widely used pesticides, may induce errors of chromosome segregation and produce cells with missing or extra chromosomes (2). The characterization of the genotoxicity of chemical and physical agents is one of the main areas of environmental mutagenesis research, together with the determination of dose–effect relationships and of the numerous variables that affect the response of a given cell/organism to a certain mutagen exposure. The ultimate goal of this discipline is to prevent the occurrence of genetic changes in somatic and germ cells, additional to those produced by endogenous mechanisms, that can lead to severe degenerative or heritable diseases primarily in humans, but also in wild and domestic animals and plants. An important contribution of environmental mutagenesis is also to generate knowledge needed to set scientifically sound chemical and radiation exposure limits to human populations and to regulate labeling and classification of commercial products. To achieve its goals, environmental mutagenesis makes use of in vitro and in vivo laboratory models and field, or so-called biomonitoring studies, carried out mostly in human populations and more rarely in other natural living organisms. In all of these studies, the frequencies of appropriate biomarkers of genotoxic effects are measured in exposed cells or individuals and compared with those of unexposed control groups. Gene mutations, chromosome aberrations, and aneuploidy are all genetic changes causally related to cancer development, and high frequencies of chromosome aberrations in human blood lymphocytes were retrospectively shown to correlate with an increased risk of cancer (3). Structural aberrations essentially comprise fragments of broken chromosomes and various types of rearrangements between nonhomologous chromosomes. Aneuploidy includes loss and gain of whole intact chromosomes. In most cases, these aberrations do not persist for long times either because, at cell division, they are intrinsically unstable, such as are acentric fragments or dicentric chromosomes, or produce gene dosage effects leading to cell

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death. Nevertheless they represent an early biomarker of genotoxic exposure or genetic instability. Conversely, some structural and numerical chromosome changes may be stable through cell division and compatible with cell survival, such as reciprocal translocations and particular aneuploidies; such genetic alterations may directly contribute to the multistep process of cell transformation and tumor development (4). Structural and numerical chromosome aberrations can be detected by optical microscopy in metaphase and, more recently, also interphase cells. Traditionally, metaphase chromosomes stained by DNA specific dyes were scored for numerical and structural changes. Chromosome banding techniques could be applied to help detection of aberrations, such as centromeric- (C-) banding, in the case of mouse acrocentric chromosomes. Reciprocal translocations could not be identified by this approach unless they produced gross karyotypic changes. Numerous techniques, all based on the principles of fluorescence in situ hybridization (FISH), were developed in the past few years to improve the cost-effectiveness, speed, and sensitivity of chromosome aberration scoring. Chromosome painting denotes the technique based on metaphase chromosome FISH with whole chromosome specific probes. Some chromosomes are hybridized with fluorescent probes, and all of them are counterstained with DNA specific fluorescent dyes such as 4,6-diamidino2-phenylindole (DAPI). Chromosome rearrangements are detected as color switches between painted and unpainted sequences. Chromosome painting was initially developed to detect stable aberrations as long-term biomarkers of past radiation exposure in humans (5). Later on it was applied to experimental studies on mechanisms of aberration induction and, with the development of mouse chromosome probes, to the quantification and characterization of persistent aberrations experimentally induced by radiation and chemical mutagens (6). Further developments of chromosome painting were based on a cocktail of probes specific for each chromosome conjugated with different fluorescent dyes and computerized image analysis capable of recognizing each one of the 24 human or the 21 mouse different chromosomes. By this technique, complex rearrangements involving more than two chromosomes could be detected (7). Detection of target sequences is achieved not only on metaphase chromosomes, but also in interphase nuclei. The possibility of detecting specific sequences in interphase is of particular interest for cases in which chromosomes are difficult if at all possible to prepare, such as sperm (see Marchetti, this volume) micronuclei, or low proliferation tissues. A useful application of interphase FISH is that with pan-centromeric probes that hybridize close to the functional centromeric sequences of all human or

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mouse chromosomes (8). This FISH technique has been extensively applied to characterize the chromosome content of micronuclei, small chromatincontaining bodies surrounded by a nuclear membrane that are produced after chromosome breaks or chromosome loss at anaphase (see Fenech, this volume). Centromeric FISH-positive micronuclei are, in fact, interpreted as containing whole chromosomes lost at anaphase because of spindle damage or other types of mitotic defects, while negative micronuclei represent acentric chromosome fragments produced by chromosome breakage (9). Alternatively, whole-chromosome containing micronuclei were identified via immunofluorescent detection of kinetochore proteins with CREST antibodies (10). The simultaneous application of CREST and FISH labeling to micronuclei allowed evaluation of possible detachment of kinetochore proteins from the centromeric DNA sequences (11). Human chromosomes are characterized by chromosome-specific pericentromeric sequences. Probes for these sequences mark specific chromosomes in interphase nuclei. These probes can be used to trace chemically induce chromosome nondisjunction in pairs of nuclei of binucleate cells: when damage to any component of the mitotic machinery affects chromosome segregation, in fact, two sister chromatids may segregate to the same daughter nucleus. This event will give rise, at the same time, to one cell with a missing and one cell with an extra chromosome that can be easily detected by counting the number of specific probe signals in each nucleus (12). Finally, telomeres are specialized functional elements of eukaryotic chromosomes required for chromosome stability maintenance (13). Telomeres consist of repetitive DNA sequences and specific proteins. All vertebrate species have the same basic telomeric DNA sequence. Telomere function is impaired when telomere length changes either below or above the species-specific range. Nonfunctional telomeres cause end-to-end chromosome fusion producing various types of stable and unstable structural aberrations (14). Quantitative FISH (Q-FISH) is a FISH-based approach developed to measure telomere length with high resolution on a single chromosome basis (15). 2. Basic Principles of FISH FISH is a method for localizing and detecting specific nucleotide sequences in morphologically preserved tissue sections or cell preparations by hybridizing the complementary strand of a nucleotide sequence (the probe) with the DNA of interest (also called target). There are essentially five types of probe that can be used in performing in situ hybridization (16). 1. Oligonucleotide probes. These are produced synthetically by an automated chemical synthesis. These probes have the advantages of being resistant to RNases

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4.

5.

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and are small, generally around 40–50 base pairs (bp). Their small size allows for easy penetration into the cells or tissue of interest, and the single strand excludes the possibility of renaturation. This oligonucleotide probe allows the hybridization of short sequences, especially single-copy genes. Single-stranded DNA probes. These have similar advantages of the oligonucleotide probes except they are much larger, probably in the 200–500 bp size range. Double-stranded DNA probes. These are the common commercial probes used. They are longer than the oligonucleotide probes and single-stranded probes allowing hybridization of repeated sequences or entire chromosomes. Because the probe is double stranded, it means that denaturation has to be carried out before hybridization in order for one strand to hybridize with the target. RNA probes (cRNA probes or riboprobes). RNA probes have the advantage that RNA-DNA hybrids are very thermostable and are resistant to digestion by RNases. This allows the possibility of posthybridization digestion with RNase to remove non-hybridized RNA and therefore reduces the possibility of background staining. PNA probes. The peptide nucleic acid probes (PNAs) constitute a remarkable new class of synthetic nucleic acid analogues, based on their peptide-like backbone. This structure gives to PNAs the capacity to hybridize with high affinity and specificity to complementary RNA and DNA sequences and a great resistance to nucleases and proteinases (17).

The probes are synthesized using different existing protocols. Basically, one resynthesizes one strand of a double-stranded DNA molecule while incorporating modified nucleotides. A DNA probe could be labeled with a fluorescent dye (direct labeling) or a haptene (indirect labeling), usually in the form of fluor-dUTP or haptene-dUTP, that is incorporated into the DNA using enzymatic reactions, such as nick translation, random-primed or polymerase chain reaction (PCR). If the probe has been labeled indirectly, an extra step is required for visualization of the non-fluorescent haptene that uses an enzymatic or immunological detection system. Whereas FISH is faster with directly labeled probes, indirect labeling offers the advantage of signal amplification by using several layers of antibodies, and might therefore produce a signal that is brighter compared with background levels. A large variety of commercial probes are now available, so as to allow hybridization even in less optimal conditions. There are almost as many methods for carrying out in situ hybridization. The basic principles for in situ hybridization are the same. Here we report a brief outline of the common procedural cytogenetic steps. 2.1. Slide Preparation and Fixation Proper technique for preparing slides is considered essential for both classical cytogenetics (banding) and molecular cytogenetic procedures (FISH). Different protocols were specifically designed for use with particular types

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of biological material or cells or for particular purposes such as detection of structural aberrations, numerical aberrations, or sequence amplifications. To preserve morphology, the biological material must be fixed. For metaphase chromosome spreads, methanol–acetic acid fixation is usually sufficient. In the basic protocol, cells are resuspended in 3:1 methanol–acetic acid fixative(s) and dropped or smeared on commercially precleaned slides, which are dried at various temperatures. For paraffin-embedded tissue sections, use formalin fixation. Cryostat-sliced sections fixed for 30 min with 4% formaldehyde have been used successfully. Unfortunately, a fixation protocol that can be used for all substrates has not yet been described. The fixation must be optimized for different applications. 2.2. Aging Slides are subjected to dry heat and/or ethanol to remove water and fixative from the preparations, and to enhance the adherence of the material to the glass. 2.3. RNase Treatment RNase treatment serves to remove endogenous RNA and may improve the signal-to-noise ratio in DNA–DNA hybridization. 2.4. Permeabilization and Pretreatment The act of fixation results in crosslinking of proteins, which may present an obstacle to good accessing of the probe. Pretreatment serves to increase target accessibility by digesting the protein that surrounds the target nucleic acid. There are a number of different procedures for permeabilization. Three common reagents used are HCl, detergents (Triton or sodium dodecyl sulfate [SDS]) and enzymes such as proteinase K, trypsin, or pepsin. The most common enzyme used is pepsin because it has the advantage that it can be easily inactivated by pH changes, and the reaction is easier to control. 2.5. Denaturation Denaturation of DNA, which separates the two strands, is obtained by alkaline or heat treatment. Heat denaturation has become popular because of its experimental simplicity and greater effectiveness. Slide and probe denaturation can be performed together (called “simultaneous or direct denaturing”) or separately (called “separate or indirect denaturing”). Numerous experiments with both commercial and custom-prepared probes showed that results were

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similar with both procedures. Variations in time and temperature should be evaluated to find the best conditions for denaturation. 2.6. Hybridization The hybridization step consists in simply mixing the single-strand probes with the denatured target DNA. The denaturation of DNA allows access of the single-strand probes to their respective complementary single-strand DNA. Provided that different probes are conjugated with fluorochromes with nonoverlapping emission wavelengths, multiple targets can be simultaneously hybridized in the same cells. 2.7. Posthybridization Washes Labeled probe can hybridize nonspecifically to sequences that are partially but not entirely homologous to the probe sequence. Such hybrids are less stable than perfectly matched hybrids. They can be dissociated by performing washes of various stringencies. The stringency of the washes can be manipulated by varying the formamide concentration, salt concentration, and temperature. Often a wash in 2× saline sodium citrate (SSC) containing 50% formamide suffices. For some applications the stringency of the washes should be higher. 2.8. Detection and Ampliﬁcation of FISH Signal The detection of the probes is the final step of FISH. When a DNA probe is labeled by a fluor-dUTP the probe-target hybrids can be visualized under a microscope immediately after the hybridization reaction. If the probe is labeled with haptene-dUTP, the fluorescent signal is usually provided by a fluor-labeled molecule (primary detection reagent) that binds the haptene with high affinity. To amplify the signal, another fluorescent-labeled molecule (secondary detection reagent) that recognizes the primary reagent is used. Although these detection reactions sometimes are not based on the immunological principles of antigen– antibody recognition, but on protein binding with strong affinity (biotin/avidin), usually the term antibody is used to indicate the detection reagent. When a DNA probe is labeled by a fluor-dUTP, signals are usually weaker than the signal yielded by the same probe if it were detected with an antibody. The reason is that an antibody molecule carries three or four fluors attached to it. The weak signal of a fluorescent-labeled probe can sometimes be amplified using antibodies raised against that fluor molecule. Antibodies against fluorescein, rhodamine, Texas Red, Cascade Blue, and other fluors are available commercially. These primary antibodies will bind to the respective fluor, and a secondary antibody labeled with the same fluor will be used to bind to the primary antibody and amplify the signal.

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2.9. Counterstaining Fluorescent DNA counterstaining is usually performed with red fluorescent propidium iodide (PI) or with blue fluorescent DAPI. An antifade agent should be mixed with the counterstaining dye solution to retard fading. 2.10. Microscopy Finally the signals are evaluated by fluorescence microscopy. A fluorescent microscope contains a lamp for excitation of the fluorescent dye and a special filter that transmits a high percentage of light emitted by the fluorescent dye. There has been considerable improvement in both the hardware and software that are used for the analysis of FISH images. Cooled charge-coupled-device (CCD) cameras and fluorescence filter sets for microscopy that are more specific and efficient have improved the sensitivity and resolution of imaging, and sophisticated software facilitates the acquisition and processing of images. The general FISH procedure is illustrated in Fig. 1. In the following chapters, materials and methods are described that are common to the application of FISH to detection of chromosome aberrations in metaphase cells, detection of centromeric sequences in interphase nuclei and

Fig. 1. The basic principles of fluorescence in situ hybridization.

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micronuclei, and estimation of telomeric sequence length. Whenever necessary specific aspects of these techniques are presented; in the end, criteria of slide analysis are separately described. 3. Materials 3.1. Preparation of Slides and Fixation of Material 1. Cleaned slides. 2. Methanol–acetic acid (3:1).

3.2. Pretreatments of Material on Slides 1. 20× SSC: Dissolve 175.3 g of sodium chloride and 88.23 g of trisodium citrate in 900 mL of water. Adjust the pH to 7.0 with a few drops of concentrated HCl. Store at room temperature for up to 1 year. Dilute in water to make appropriate solutions for this protocol. 2. RNase A DNase-free (Roche, Mannheim, Germany) stock solution: 10 mg/mL. Store at –20°C. Prepare 1 mL of RNase working solution, 100 μg/mL, by adding 10 μL of RNase 10 mg/mL, 890 μL water, and 100 μL of 20× SSC. Prepare fresh each time. 3. Pepsin (Sigma-Aldrich) stock solution: 100 mg/mL in water. Store at –20°C. 4. 1× PBS + 50 mM MgCl2 solution: Add 50 mL of MgCl2 1 M to 950 mL of 1× phosphate-buffered saline (PBS). 5. 1% Formaldehyde: Add 2.7 mL of 37% formaldehyde to 100 mL of 1× PBS + 50 mM MgCl2 solution. 6. 1× PBS solution: Dissolve 8 g of NaCl, 0.2 g of KCl, 1.44 g of Na2 HPO4 , and 0.24 g of KH2 PO4 in 800 mL of water. Adjust the pH to 7.4 and adjust the volume to 1 L. Autoclave and store at room temperature. 7. Alcohol series: Prepare fresh 70%, 85%, and 100% ethanol and store at –20°C. 8. Plastic cover slips. (See Note 1.)

3.3. Denaturation of Target and Probe 1. Denaturation solution (70% formamide–2 SSC): for 50 mL, add 35 mL of deionized formamide (Kodak, Rochester, NY), 10 mL of distilled water, 5 mL of 20× SSC. Adjust the pH to 7.0 with HCl. 2. Hot plate.

3.4. In Situ Hybridization 1. Probe. 2. Humidified chamber. Humidified chamber can be prepared using a clean plastic box with a lid. Soak several paper towels in water and place them at the bottom of the tray. Put the lid on top and place chamber in the 37°C incubator. 3. Glass cover slips. 4. Glass cover slip sealant (rubber cement).

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3.5. Posthybridization Washes 1. Wash solution: 50% formamide (Fluka, Buchs, SG, Switzerland) 100 mL add 50 mL of formamide, 10 mL of 20× SSC, and 40 water. Adjust pH to 7.0. 2. 4× SSC + 0.1% Tween-20 solution: Add 100 mL of 20× SSC Tween-20 to a total of 500 mL of distilled water, mix well on plate, and store at room temperature (RT) for up to 1 month.

in 2× SSC. For mL of distilled and 0.5 mL of a magnetic stir

3.6. Detection and Ampliﬁcation 1. Plastic cover slips. (See Note 1.) 2. Blocking solution: 3% bovine serum albumin (BSA) in 4× SSC + 0.1% Tween-20. Prepare 5 mL with 150 mg of BSA, 1 mL of 20× SSC, 3.95 mL of water, and 50 μL of 10% Tween-20. 3. Detection solution: 1% BSA in 4× SSC + 0.1% Tween-20. Prepare 5 mL with 50 mg of BSA, 1 mL 20× SSC, 3.95 mL of water, and 50 μL of 10% Tween-20. 4. Antibodies: To detect the haptene-dUTP labeled probe or to amplify the weak signal of fluorescent probe, primary, secondary, and sometimes tertiary detection reagents can be used (Roche, Mannheim, Germany). Digoxigenin-labeled or biotinlabeled DNA probe can be detected with a variety of labeled antibodies (using different fluors) as shown in Table 1. Antibody combinatorial schemes allow, for example, triple color detection and signal amplification for two or more probes labeled with different haptene-dUTPs.

Table 1 Different Strategies to Label the Target and Amplify the Primary Fluorescent Signal

Haptene Digoxigenin

Primary detection reagent (Fluors) Mouse anti-DIG

FITC

Secondary detection reagent Anti-mouse Ig-DIG

Tertiary detection reagent (Fluors) Anti-DIG FITC Cy3

Cy3 Texas Red

Texas Red Rhodamine

Rhodamine Biotin

(Fluors) Avidin

Biotinylated anti-avidin

(Fluors) Avidin

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3.7. Counterstaining 3.7.1. With Propidium Iodide (for FITC-Labeled Probe) 1. Propidium iodide (Sigma-Aldrich) solution. Stock solution, 10 μg/mL in distilled water or PBS, can be stored at 4°C for several months. Working solution (PI/antifade): for each slide prepare in a microfuge tube 30 μL of solution with 6 μL of propidium iodide stock solution and 24 μL of mounting medium/antifade (Vector Laboratories, Burlingame, CA), mixing well (final concentration: 2 μg/mL). 2. Nail varnish.

3.7.2. With DAPI (for Cy3, Texas Red and Rhodamine-Labeled Probe) 1. DAPI (Sigma-Aldrich) solution. Stock solution, 100 μg/mL in PBS or 2× SSC, can be stored at 4°C for several months. Working solution: Prepare 50 mL of a solution with 100 μL of DAPI stock solution in 2× SSC (final concentration: 0.2 μg/mL). Put the DAPI solution in a glass Coplin jar. The jar with DAPI solution is wrapped in aluminum foil or is taped with black tape (to protect it from light). 2. Mounting medium/antifade Vectashield (Vector Laboratories). 3. Nail varnish.

4. Methods 4.1. Preparation of Slides and Fixation of Material Prepare metaphase chromosome spreads or interphase nuclei on a glass microscope slide according to standard cytogenetic procedure currently in use in your laboratory (see Notes 2 and 3). More details about metaphase and interphase cell preparation for subsequent FISH can be found in refs. 18 and 19. 4.2. Slide Aging The purpose of aging is to fix the biological material to the glass surface and to make the chromosome structure resistant to the subsequent DNA denaturing steps. Very fresh slides, if not aged, will either lose most of the nuclei/chromosomes during denaturation or the shape of the chromosomes will become very distorted. Too long aging will decrease the efficiency of hybridization (see Note 4). Slides are usually aged either by keeping them 2–3 days at RT, or overnight at 65°C or by heating 30 min at 90°C. 4.3. Pretreatments of Material on Slides (see Note 5) 1. Apply 100–200 μL of RNase stock solution (see Subheading 3.2.2. for details) to each slide and cover with plastic cover slip. Incubate for 1 h in humidified chamber at 37°C. Carefully peel back cover slip with forceps.

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2. Rinse slides 3 times in 2× SSC at RT, 5 min each rinse. 3. Prewarm 0.01 M HCl in Coplin jars to 37°C in a water bath. Add 25 μL of pepsin 100 mg/mL per 50 mL of 0.01 M HCl (final concentration: 50 μg/mL). Place slides in Coplin jars for 10 min. 4. Wash slides in a glass Coplin jar 2 times in 50 mL of 1× PBS at RT for 5 min. 5. Wash slides in 50 mL of 1× PBS + 50 mM MgCl2 at RT for 5 min. 6. Incubate slides for 10 min in 1% formaldehyde in PBS + MgCl2 at RT. 7. Wash slides for 5 min in PBS at RT. 8. Dehydrate slides in a series of three cold ethanol washes: 70%, 85%, and 100%. All washes are done in Coplin jars for 2 min each wash, starting with the 70% wash. (See Note 6.) 9. Allow slides to air-dry or dry under an air jet.

4.4. Denaturation of Target and Probe 1. Preset the hot plate to 70°C. (See Note 7.) 2. Prepare 100 μL of denaturation solution for each slide into a 1.5-mL microfuge tube. Denaturation solution must be prewarmed to 70°C. 3. Apply 100 μL of denaturation solution to each slide and cover with a glass cover slip. (See Note 8.) 4. Denature slides (no more than three slides at a time) for 2 min at 70°C on the prewarmed hot plate. (See Note 9.) 5. Immediately remove cover slip by soaking slides in a Coplin jar containing cold 70% ethanol and rinse for 2 min. Remove the cover slips from the Coplin jar with forceps. Repeat rinse in cold 85% and 100% ethanol solutions. 6. Allow slides to air-dry or dry under an air jet. 7. Prewarm probe at 37°C for 5 min. (See Note 10.) 8. Vortex-mix gently and centrifuge 2–3 s to collect contents on the bottom of the tube. 9. Place 30 μL of commercial probe for entire slide or 10 μL of probe for a 22 × 22 mm area (see Note 11) in a 0.5-mL microfuge tube (see Note 12) and denature for 5 min in a 70°C water bath. (See Note 13.) 10. Immediately place the tube on ice until ready to use.

4.5. In Situ Hybridization 1. Apply 30 μL of probe on each slide and cover with 22 × 50 mm glass cover slip. (See Notes 8 and 11.) 2. Seal the cover slip with glass cover slip sealant by applying sealant along the perimeter of the cover slip. 3. Incubate at 37°C for 12–16 h at 37°C in a humidified chamber.

4.6. Posthybridization Washes 1. Carefully remove the cover slip sealant with forceps. Do not remove the cover slip. This will fall off during the washing step. Take out the cover slips from the Coplin

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jar with forceps. Different probes require different stringency hybridization and postwashing conditions, according to the probe data sheet specifications (see Note 14). For this reason we can distinguish two separate post-hybridization washing protocols: low-stringency and high-stringency protocols (see Note 15).

Low-stringency posthybridization washing (prewarm in water bath all the following solutions): 2. Wash slides 3 times in a glass Coplin jar containing 50 mL of wash solution at 37°C for 5 min with intermittent agitation. 3. Wash slides 3 times in 50 mL of 2× SSC, pH 7, at 42°C for 5 min with intermittent agitation. 4. Rinse slides in 4× SSC + 0.1% Tween-20 at RT in a Coplin jar. Do not allow slides to dry beyond this step (see Note 16).

High-stringency posthybridization washing (prewarm in water bath all the following solutions): 2 . Wash slides 3 times in a glass Coplin jar containing 50 mL of wash solution at 42°C for 5 min with intermittent agitation. 3 . Wash slides 3 times in 50 mL of 0.1× SSC, pH 7, at 60°C for 5 min with intermittent agitation. 4 . Rinse slides in 4× SSC + 0.1% Tween-20 at RT in a Coplin jar. Do not allow slides to dry beyond this step (see Note 16).

If the DNA probe is labeled by a fluor-dUTP proceed immediately to counterstaining (see Subheading 4.9.). If the probe is labeled with haptenedUTP, proceed to the following step (detection). 4.7. Detection (see Note 12) 1. Remove slides from 4× SSC + 0.1% Tween-20 and briefly blot excess fluid from the edge. 2. Apply 100 μL of blocking solution to each slide (see Notes 17 and 18). Place a plastic cover slip over the solution. Incubate for 30 min in a humidified chamber at 37°C. 3. Carefully peel back the cover slip with forceps, tilt slide and allow fluid to drain briefly.

It is possible to detect the probe with individual antibodies or antibody combination. Antibody combination allows detection of two or more probes labeled with different haptene-dUTPs. 4. Apply 100 μL of primary antibody (fluors-labeled avidin and/or fluors-mouse antidigoxigenin [anti-DIG]) to each slide and replace plastic cover slip (see Notes 19 and 20). Incubate 30–60 min in humidified chamber at 37°C. 5. Wash slides 3 times in 50 mL of 4× SSC + 0.1% Tween-20 at 37°C for 5 min each wash with intermittent agitation (see Note 21).

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4.8. Ampliﬁcation 1. Apply 100 μL of secondary antibody (biotinylated anti-avidin and/or anti-mouse Ig DIG) to each slide and replace plastic cover slip over the solution (see Notes 19 and 20). Incubate 30–60 min in a humidified chamber at 37°C. 2. Wash slides 3 times in 50 mL of 4× SSC + 0.1% Tween-20 at 37°C for 5 min each wash with intermittent agitation. (See Note 21.) 3. Apply 100 μL of tertiary antibody (fluors-labeled avidin and/or fluors-labeled antiDIG) to each slide and replace plastic cover slip over the solution (see Notes 19 and 20). Incubate 30–60 min in humidified chamber at 37°C. 4. Wash slides 3 times in 50 mL of 4× SSC + 0.1% Tween-20 at 37°C for 5 min each wash with intermittent agitation (see Note 21).

4.9. Counterstaining 4.9.1. With Propidium Iodide 1. Apply 30 μL PI/Antifade (2 μg/mL) to each slide and cover with a glass cover slip. 2. Seal the edges of the cover slip with nail varnish (see Note 22). The signal keeps well for several weeks if slides are stored at 4°C.

4.9.2. With DAPI 1. 2. 3. 4. 5.

Slides are incubated 10 min in the staining solution. Rinse slides 10 min in 2× SSC to wash off excess dye. Wash slides for a few seconds in distilled water to remove any salt traces. Air-dry at room temperature and apply 30 μL of antifade solution. Cover with a glass cover slip and seal the edges of the cover slip with nail varnish (see Note 22).The signal keeps well for several weeks if slides are stored at 4°C.

4.10. Analysis of Slides 4.10.1. Chromosome Painting Chromosome painting (Fig. 2a) was mainly developed to allow detection of stable exchange-type aberrations, such as reciprocal translocations, for biological dosimetry purposes (5). In fact, the analysis of dicentrics (unstable aberrations) in solid-stained chromosome preparations is very reliable to estimate recent and acute radiation exposures, but not for chronic or past exposures because the yield of dicentric chromosomes decreases with time after irradiation (20). For a correct analysis of aberrations in painted (as well as in solid-stained) metaphases, the detection of centromeres may be critical, especially in the case of mouse acrocentric chromosomes: it is obtained either by DAPI counterstaining that, by itself, after alkaline/heat denaturation, gives a bright signal to centromeric heterochromatin or by centromeric FISH staining.

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Fig. 2. (a) Mouse bone marrow metaphase hybridized with probes specific for chromosomes 2 and 7. The products of a reciprocal translocation are indicated by arrows; (b) two sister nuclei of a cytokinesis-blocked human male lymphocyte hybridized with centromeric probes for chromosome X (in green) and 8 (in red). Nondisjunction of the X chromosome is shown by the 2+0 pattern of green signals (courtesy of Dr. A. Zijno). (c) Mouse bone marrow metaphase hybridized with telomere specific PNA probes for Q-FISH analysis. (d-g) Photomicrographs of normal and abnormal sperm nuclei (d, e: human ACM assay; f, g: mouse CT8 assay) taken under ×1000 magnification using fluorescence microscopy to visualize the probes and with phase-contrast imaging to visualize the sperm head and tail; (d) normal human haploid sperm showing one red (1cen), one blue (1q12), and one green (1p36.3) fluorescence domain. The 1cen and 1q12 regions are contiguous on chromosome 1, and their fluorescent domains are adjacent to each other in normal sperm; (e) human sperm with a break within 1q12, indicated by two 1q12 (blue) domains, one 1cen (red) domain, and one 1p36.3 (green) domain; (f) normal mouse sperm showing one red (2cen), one green (2tel), and one yellow (chromosome 8) fluorescence domain; (g) mouse sperm with a duplication of 2ter. There are two green signals, but only one red and one yellow signal.

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In solid-stained metaphases, the conventional nomenclature classifies each simple interchange as a single event (e.g., one dicentric plus an acentric fragment), and distinguishes between complete (if no unrejoined breaks are found) and incomplete exchanges (e.g., a dicentric chromosome without any acentric fragment). The use of painting techniques has led to the development of new nomenclature systems. These are mainly based on the recognition of color switches between the fluor-conjugated probe and the counterstaining dye. According to the Protocol for Aberration Identification and Nomenclature Terminology (PAINT) (21), each abnormal painted chromosome or fragment is described individually using the letters “A” and “a” to indicate counterstained chromosomal material, the letters “B” and “b” to indicate painted material, capital letters to designate centromere containing regions, lower case letters to indicate acentric regions. Thus, typical PAINT-classified aberrations are t(Ab), dic(AB) or ace(ab), where t stands for translocation, dic for dicentric, and ace for acentric fragment. Another independently developed nomenclature system is that of Savage and Simpson (S&S) (22), which is the method of choice for an interpretation of the mechanistic aspects of aberration formation. When FISH painting is applied to the analysis of aberrations induced by chemical mutagens (6,23) a further type of aberration, not encountered in radiation studies, must be considered for the correct assessment of induced effects: it is the presence of one or more supernumerary fully painted chromosomes in a metaphase with the euploid chromosome number for the specific species. These supernumerary chromosomes derive from chromatid-type exchanges in the pericentromeric region, which are induced by radiation only when G2 cells are targeted. Unless all chromosomes are hybridized with their specific painting probes conjugated with different fluorochromes and a computerized image analysis system is used to assign a different color to each of them, only a portion of the whole karyotype is usually painted. Aberrations involving painted chromosomes thus represent only a subset of all induced aberrations. It may be of importance to estimate the total number of aberrations induced to predict the fate of exposed cells or individuals. Assuming that aberrations are randomly distributed over different chromosomes, to convert the frequency of observed aberrations into an estimate of the frequency of total aberrations/genome, one has to know the proportion of the genome covered by each color probe (or probe cocktail). Then, if p represents the proportion of genome painted in a given color, q the proportion of genome painted in a second color and r the unpainted fraction of the genome, the fraction of detectable exchanges is calculated as S = 2pq + 2pr + 2qr. Multiplying the number of scored metaphases by S, one obtains the number of cell (genome) equivalents. The ratio between the number of observed aberrations and the number of cell equivalents gives

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the estimated frequency of aberrations/genome. Also, assuming that n is the recommended number of metaphases to be scored when the whole genome can be scanned for aberrations, n/S gives the number of painted metaphases to score to obtain the amount of information equivalent to that provided by n fully analyzed metaphases (24). 4.10.2. Centromeric FISH Staining Centromeric FISH staining allows to detect potential aneugenic agents by measuring the frequency of chromosome loss and chromosome nondisjunction events. To estimate chromosome loss events, probes specific for the centromeric sequence of all chromosomes are used to detect the presence of centromere(s) in micronuclei. Micronuclei labeled by the probe are classified as centromere positive (C+), assumed to contain one or more chromosomes, and ascribed to chromosome loss occurred at the preceding anaphase. To estimate nondisjunction events, probes that detect the (peri)centromeric region of specific chromosomes are used to assess the number of chromosomes present in each sister interphase nucleus of a binucleate cell. CytochalasinB-blocked cells are used that retain sister nuclei of a single mitotic division within the same cytoplasm. Two spots for each probe are expected for any autosome in euploid nuclei. Nondisjunction events occurred at the preceding mitotic division are detected via deviations from the expected 2+2 pattern of signals in sister nuclei, e.g., 3+1 or 4+0 combinations; nondisjunction of the X or the Y chromosome in male cells gives rise to a 2+0 pattern of signals (Fig. 2b). Only cells with the correct total number of hybridization signals with similar fluorescence intensity are analyzed. It has been demonstrated that nondisjunction events may be more sensitive indicators of aneugenic activity than chromosome losses (25). This emphasizes the importance of molecular cytogenetics in interphase cells as a fast and reliable approach to assess the frequency of spontaneous and induced nondisjunction events. 4.10.3. Quantitative FISH for Telomere Length Estimation 4.10.3.1. Quantitative Image Analysis.

Telomeres protect the end of chromosomes from end-to-end fusion. Shortening of telomeres beyond a critical length drives the cell into replicative senescence or is responsible of chromosome instability. Thus, the measurement of telomere length is a useful means for assessing the lifespan of a cell or for studying chromosome instability induced by physical or chemical agents. Recently, a new in situ technique has been developed, the quantitative fluorescence in situ hybridization (Q-FISH). This technique, which can be applied

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to metaphase and interphase nuclei, is based on the use of a peptide nucleic acid (PNA) telomere oligonucleotide probe that generates stronger and more specific hybridization signals than standard DNA oligonucleotide probes (17). Presently, Q-FISH is able to detect differences in individual telomere lengths below 1 kb (26). The assumption is made that the length of a telomere is directly related to its integrated fluorescence intensity (IFI) value, as the fluorescence probes used are assumed to hybridize quantitatively to telomere repeat sequences, allowing detection of short telomere length variations. Digital images are recorded with a camera on a fluorescence microscope equipped with a multiple filter wheel. Image acquisition must be performed with dedicated software. Digital images are acquired from DAPI-stained chromosomes and from Cy-3-stained telomeres (Fig. 2c). A dedicated computer program was developed for image analysis (TFL-Telo program) (27). In short, chromosomes and telomeres are identified through segmentation of the DAPI image and the Cy-3 image, respectively. Both images are combined and corrected for pixel shifts. To prevent a possible selection bias, metaphases are chosen solely on the basis of a good chromosome spread. TFL-Telo program allows to make contours from individual chromosomes and individual telomere spots and the two images are successively combined. Finally, the fluorescence intensity of telomeres on all individual chromosomes is expressed in arbitrary units of fluorescence (a.u.f.). 4.10.3.2. Calibration.

Two levels of calibration have to be used to ensure a reliable quantitative estimation of telomere length in various samples. First, to correct for daily variations in lamp intensity and alignment, images of fluorescent beads (orange beads, size 0.2 μm; Molecular Probes, Eugene, OR) are acquired and analyzed with the computer program in parallel to cell samples. Second, to relate fluorescence intensity to number of T2 AG3 repeats, mammalian cells with a defined (T2 AG3 )n length of 10 and 80 kb (LY-S and LY-R lymphoma cell lines respectively) (28) may be hybridized and analyzed. Based on the slope of the linear relationship between IFI values and telomere lengths of LY-S and LY-R cells, the telomere lengths of cell samples are estimated after their IFI values are acquired under the same conditions. Alternatively to cell lines with defined telomere length, plasmids with telomere sequence inserts of known size can be used (27). 4.11. Microscopy Filter Setting A very important part of FISH is the visualization and recording of the results with a fluorescence microscope and camera system. A fluorescence

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Table 2 Some of the Most Common Fluorochromes Used for the Detection of Chromosome Structural and Numerical Aberrations Fluorochrome

Color

Excitation (nm)

Emission (nm)

Propidium iodide DAPI Rhodamine CY3 Fluorescein

Red Blue Red Red Green

535 359 560 552 494

605 441 590 565 523

microscope contains a lamp for excitation of the fluorescent dye and a special filter that transmits a high percentage of light emitted by fluorescent dye. The filters allow certain wavelengths of light through, while blocking others. Therefore, the filters are individually designed for specific fluors and must be chosen properly. The propidium iodide or DAPI filter should be used to scan the slide for cells or metaphase spreads, while the filters specific for probe-conjugated fluors are used for visualization of the target(s) of interest. Table 2 gives general guidelines for choosing filters from your microscope manufacturer. 5. Notes 1. Plastic cover slips can be prepared by cutting a piece of parafilm. They can be removed from the slide more easily than glass cover slips and should be preferred except in those steps in which high temperatures or sealing requirements are not compatible with plastics. 2. To prepare metaphase chromosome spreads, cultured cells are treated with colcemid (10 μg/mL-Invitrogen, Paisley, UK) for 3 h. This protocol can be adapted for many cell cultures. For cells with a short cell cycle, such as mouse cells, 2 h of colcemid treatment could be enough. For cells that grow more slowly, the colcemid time could be from 3 h to overnight. The colcemid concentration should be adequate to the treatment time: long treatment time requires lower colcemid concentration. To obtain metaphases from in vivo experiments, 0.3 mL per mouse of colchicine 10−3 M (Sigma-Aldrich) is used. To prepare binucleate cells cytochalasin B (3 μg/mL, Sigma-Aldrich) is added to the cultures for the duration of one cell cycle. 3. This protocol is recommended for cultured cells and cell suspensions obtained from experimental animals or humans, fixed in methanol–acetic acid (3:1). 4. The slide aging conditions are always related to the probe used. For this reason we suggest to age slides according to the probe data sheet specifications.

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5. Many incubations are performed in 50-mL glass Coplin jars. To obtain optimal results, verify the solution temperatures, placing a clean thermometer directly into the Coplin jar. 6. Ethanol should be stored at –20°C before use. 7. If multiple slides are processed simultaneously, each slide will cause the hot plate temperature to drop 1°C. Therefore, the temperature of the hot plate must be set up 1°C higher for each slide to be denatured. 8. Care should be taken to avoid air bubbles. 9. Time and temperature are very important to maintain chromosome and nuclei morphology. Incomplete denaturation of target DNA can result in lack of signal. The suggested protocol has been verified for both metaphase and interphase human cells. Mouse cells may require higher denaturation temperatures up to 80°C. 10. Usually the probe is diluted in glycerol or formamide and the solution is very viscous. For this reason we suggest to prewarm the commercial probe at 37°C to decrease the solution viscosity. 11. For economy, only a target area of slides containing good quality and quantity of metaphase spreads or interphase nuclei could be hybridized. The selected area can be labeled under phase contrast with diamond pen. 12. We recommend wrapping in aluminum foil the tubes with the fluors labeled probe or fluors labeled antibodies, to protect them from light. 13. If two or more probes are to be used together, in the same hybridization, the final volume may be higher than the 10–12 μL accommodated under a standard 22 × 22 mm cover slip. To solve this problem, centrifuge the total volume of the mixed probes 10 min at 8400g and discard the supernatant. Allow the pellet to dry under air jet or by speed vacuum centrifuge, resuspend it in 10–12 μL of hybridization buffer (Hybrisol VI, Roche), and apply the mix to slides. 14. Temperature, time, and buffer concentration (stringency) of hybridization and postwashing solutions are important, as lower stringency can result in nonspecific binding of the probe to other sequences, and higher stringency can result in a lack of signal. 15. This step is used primarily to remove the nonspecific and/or repetitive DNA hybridized to the cells and chromosomes. It is impossible to present a standard procedure, but there are some guidelines that can help. The less concentrated the salt solution and the longer the duration of the wash and the higher the temperature, the higher will be the stringency and the more unbound or nonspecifically bound probe will be removed. For very short DNA probes (0.5–3 kb) or very complex probes (chromosome paint probes), the washing temperature should be lower (up to 45°C) and the stringency lower as well (1–2× SSC). When using single-locus, large probes, the temperature should be around 65°C and the stringency high (below 0.5× SSC). The temperature and stringency should be the highest for repetitive probes (such as -satellite repeats). 16. If necessary, slides can be stored at 4°C in 50 mL of 4× SSC + 0.1% Tween-20 up to 2 wk before detection.

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17. Unless specified otherwise, the volumes of detection solutions are given for the whole slide area of 22 × 50. If only a 22 × 22 area has been hybridized, reduce the volumes to 1/3. 18. BSA blocks some of the unspecific binding of the antibodies to the glass and this is important to decrease background, but may also partially block antibodies from reaching the labeled DNA. 19. With commercially available, fluorescent-labeled antibodies, the manufacturer’s instructions regarding antibody concentration for in situ procedures should be followed. As a general guideline, a 1:100 dilution in detection solution of a 1 mg/mL antibody stock solution should work well, after 30 min of incubation at 37°C. Every time a new antibody is purchased, it should be tested at different dilutions, to find the optimal range for that antibody. In our experience, we have found the following antibodies at the following concentration to be good for in situ hybridization: a. For biotin-labeled probes Fluorescein avidin DCS (Vector Laboratories) and rhodamine avidin DCS (Vector Laboratories) 1:300 in detection solution. Biotinylated anti-avidin D (Vector Laboratories) 1:100 in detection solution b. For DIG-labeled probes Mouse anti-DIG (Roche) 1:100–1:200 in detection solution. Anti-mouse Ig-DIG (Roche) 1:100 in detection solution. Anti-DIG– rhodamine or anti-DIG–fluorescein (Roche) 1:50 in detection solution. 20. If using an antibody combination prepare a double concentration of antibody solutions and mix 50 μL of each solution on the slides. 21. This wash could be performed at 37°C or 42°C. The optimal temperature should be empirically determined to remove excess and unbound detection reagents. 22. The nail varnish may increase the shelf-life of FISH slides.

Acknowledgments The authors thank Dr. Andrea Zijno (Rome, Italy) for the permission to reproduce one of his pictures of a nondisjunction event in a binucleate cell.

References 1. Friedberg, E. C., Walker, G. C., Siede, W., Wood, R. D., Schultz, R. A., and Ellenferger, T. (2005) DNA Repair and Mutagenesis, ASM Press, Washington, DC. 2. Parry, E. M., Parry, J. M., Corso, C., Doherty, A., Haddad, F., Hermine, T. F., Johnson, G., Kayani, M., Quick, E., Warr, T., and Williamson, J. (2002) Detection and characterization of mechanisms of action of aneugenic chemicals. Mutagenesis 17, 509–521. 3. Rossner, P., Boffetta, P., Ceppi, M., Bonassi, S., Smerhovsky, Z., Landa, K., Juzova, D., and Sram, R. J. (2005) Chromosomal aberrations in lymphocytes of healthy subjects and risk of cancer. Environ. Health Perspect. 113, 517–520.

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4. Albertson, D. G., Collins, C., McCormick, F., and Gray, J. W. (2003) Chromosome aberrations in solid tumors. Nat. Genet. 34, 369–376. 5. Tucker, J. D. (2001) FISH cytogenetics and the future of radiation biodosimetry. Radiat. Prot. Dosimetry 97, 55–60. 6. Stronati, L., Farris, A., and Pacchierotti, F. (2004) Evaluation of chromosome painting to assess the induction and persistence of chromosome aberrations in bone marrow cells of mice treated with benzene. Mutat. Res. 545, 1–9. 7. Dorritie, K., Montagna, C., Difilippantonio, M. J., and Ried, T. (2004) Advanced molecular cytogenetics in human and mouse. Expert. Rev. Mol. Diagn. 4, 663–676. 8. Mitchell, A. R. (1996) The mammalian centromere: its molecular architecture. Mutat. Res. 372, 153–162. 9. Huber, R., Salassidis, K., Kulka, U., Braselmann, H., and Bauchinger, M. (1996) Detection of centromeres in vinblastine- and radiation-induced micronuclei of human lymphocytes using FISH with an alpha satellite pancentromeric DNA probe. Environ. Mol. Mutagen. 27, 105–109. 10. Degrassi, F., and Tanzarella, C. (1988) Immunofluorescent staining of kinetochores in micronuclei: a new assay for the detection of aneuploidy. Mutat. Res. 203, 339–345. 11. Sgura, A., Antoccia, A., Cherubini, R., and Tanzarella, C. (2001) Chromosome nondisjunction and loss induced by protons and X rays in primary human fibroblasts: role of centromeres in aneuploidy. Radiat. Res. 156, 225–231. 12. Zijno, A., Marcon, F., Leopardi, P., and Crebelli, R. (1996) Analysis of chromosome segregation in cytokinesis-blocked human lymphocytes: non-disjunction is the prevalent damage resulting from low dose exposure to spindle poisons. Mutagenesis 11, 335–340. 13. de Lange, T. (2002) Protection of mammalian telomeres. Oncogene 21, 532–540. 14. Blasco, M. A., Lee, H. W., Hande, M. P., Samper, E., Lansdorp, P. M., DePinho, R. A., and Greider, C.W. (1997) Telomere shortening and tumor formation by mouse cells lacking telomerase RNA. Cell 91, 25–34. 15. Slijepcevic, P. (2001) Telomere length measurement by Q-FISH. Methods Cell Sci. 23, 17–22. 16. Read, A., and Strachan, T. (2003) Human Molecular Genetics, Garland, New York. 17. Nielsen, P. E., Egholm, M., Berg, R. H., and Buchardt, O. (1991) Sequenceselective recognition of DNA by strand displacement with a thymine-substituted polyamide. Science 254, 1497–1500. 18. Adler, I. D. (1984) Cytogenetic Tests in Mammals, in Mutagenicity Testing. A Practical Approach (Venitt, S. and Parry J. M., eds.), IRL Press, Oxford, pp. 275–306. 19. Fenech, M. (2000) The in vitro micronucleus technique. Mutat. Res. 455, 81–95. 20. Bauchinger, M. (1995) Quantification of low-level radiation exposure by conventional chromosome aberration analysis. Mutat. Res. 339, 177–189. 21. Tucker, J. D., Morgan, W. F., Awa, A. A., Bauchinger, M., Blakey, D., Cornforth, M. N., Littlefield, L. G., Natarajan, A. T., and Shasserre, C. (1995) A proposed system for scoring structural aberrations detected by chromosome painting. Cytogenet. Cell Genet. 68, 211–221.

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22. Savage, J. R., and Simpson, P. J. (1994) FISH “painting” patterns resulting from complex exchanges. Mutat. Res. 312, 51–60. 23. Sgura, A., Stronati, L., Gullotta, F., Pecis, A., Cinelli, S., Lascialfari, A., Tanzarella, C.., and Pacchierotti, F. (2005) Use of chromosome painting for detecting stable chromosome aberrations induced by melphalan in mice. Environ. Mol. Mutagen. 45, 419–426. 24. Tucker, J. D., Breneman, J. W., Briner, J. F., Eveleth, G. G., Langlois, R. G., and Moore, D. H. 2nd. (1997) Persistence of radiation-induced translocations in rat peripheral blood determined by chromosome painting. Environ. Mol. Mutagen. 30, 264–272. 25. Kirsch-Volders, M., Tallon, I., Tanzarella, C., Sgura, A., Hermine, T., Parry, E. M., and Parry, J. M. (1996) Mitotic non-disjunction as a mechanism for in vitro aneuploidy induction by X-rays in primary human cells. Mutagenesis 11, 307–313. 26. Slijepcevic, P. (1998) Telomere length and telomere-centromere reltionships? Mutat. Res. 404, 215–220. 27. Poon, S. S. S., Martens, U. M., Ward, R. K., and Lansdorp, P. M. (1999) Telomere length measurements using digital fluorescence microscopy. Cytometry 36, 267–278. 28. McIlrath, J., Bouffler, S., Samper, E., Cuthbert, A., Wojcik, A., Szumiel, I., Bryant, P. E., Riches, A. C., Thompson, A., Blasco, M. A., Newbold, R. F., and Slijepcevic, P. (2001) Telomere length abnormalities in mammalian radiosentive cells. Cancer Res. 61, 912–915.

14 Laboratory Methods for the Detection of Chromosomal Structural Aberrations in Human and Mouse Sperm by Fluorescence In Situ Hybridization Francesco Marchetti, Debby Cabreros, and Andrew J. Wyrobek

Summary The father, like the mother, can transmit genetic defects that are detrimental for development and genetic health for his children, but the mechanisms for paternally mediated abnormal reproductive outcomes remain poorly understood. A battery of sensitive methods has been developed for detecting genetic damage associated with infertility, spontaneous abortions, as well as inherited defects in children such as aneuploidy syndromes, translocation carriers, and certain genetic diseases directly in sperm. Among these, fluorescence in situ hybridization (FISH) sperm-based assays for measuring numerical abnormalities and structural chromosomal aberrations are now available for an expanding number of species including humans, rodents, and several domesticated animals. This new generation of sperm FISH methods has identified several paternal risk factors such as age, various drugs, lifestyles, and various environmental and occupational exposures. These sperm FISH assays provide new opportunities to identify and characterize male reproductive risks associated with genetic, lifestyle, and environmental factors. This chapter outlines the laboratory methods for the detection of sperm with chromosomal structural aberrations in humans (ACM assay) and mice (CT8 assay) that have been validated for detecting environmental germ cell mutagens. Key Words: Breaks; chromosomal damage, duplications; deletions; fluorescence in situ hybridization.

1. Introduction Paternally transmitted chromosomal abnormalities may lead to birth defects and genetic diseases in offspring (1–3). However, the etiologies of numerical From: Methods in Molecular Biology, vol. 410: Environmental Genomics Edited by: C. Cristofre Martin © Humana Press, Totowa, NJ

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and chromosomal defects in sperm and their association with subsequent abnormal reproductive outcomes remain generally unknown. Extensive testing in rodent models has shown that various environmental toxicants, when given to males or females before or after mating, can have profoundly deleterious effects on reproduction, such as infertility, lethality during development, malformations, as well as cancer among offspring (4,5). This has raised concern that certain environmental, occupational, or medical exposures can have detrimental effects on the genetic integrity of human male germ cells. Of special concern are long-term chronic low-dose exposures to environmental mutagens such as smoking and air pollution that affect large numbers of individuals, or short exposures of more limited populations to very high doses of mutagens such as cancer chemotherapies (1). Paternally mediated abnormal reproductive outcomes may be a consequence of abnormal reproductive physiology, predisposing genetic factors (6), past and present male environmental exposures (7,8), or random errors that occur during spermatogenesis (9). Elucidating the relative contribution of these factors from epidemiological surveys of affected offspring is extremely difficult because the sample sizes of offspring with specific defects are generally too small and prenatal selection against defective embryos vary among different types of genetic and chromosomal defects. This has provided an incentive for developing effective biomarkers to detect genomic damage directly in sperm where greater statistical power can be attained by detecting changes in the frequencies of defective sperm of only a few well characterized individuals. Fluorescence in situ hybridization (FISH) is an efficient approach for labeling DNA of chromosomes in interphase cells including sperm. Its effectiveness has improved with the availability of chromosome-specific DNA probes for every human chromosome and with the increased emphasis on the importance of scoring criteria (10). Since its introduction, FISH assays evolved from a one- to a two-, three-, and four-chromosome assay using multiple probes specific for each chromosome (11,12). Emphasis has also shifted from using any chromosome for which an effective DNA probe was available to selected use of chromosomes with clinical relevance in human aneuploidy syndromes (13). Human sperm FISH has identified several potential risk factors for chromosomally abnormal sperm including certain lifestyles factors and environmental/occupational exposures (8,14,15). Sperm FISH also has the intrinsic advantage of being broadly applicable to any laboratory and domestic species for which chromosome-region-specific probes are available and several multicolor FISH assays have also been developed to detect numerical abnormalities in laboratory animals (16–18). Sperm FISH assays for laboratory animals may provide a platform for systematic tests of the genetic damage to germ cells of the myriads of chemicals present in the environment, and for

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prioritizing human epidemiological studies of paternally mediated abnormal reproductive outcomes. An important recent extension of sperm FISH methods has been the development of assays for detecting human (ACM assay) and rodent (CT8 assay) sperm carrying chromosomal structural aberrations (19,20). Unlike transmitted germinal aneuploidy, which originates from meiotic nondisjunction

Fig. 1. Sperm FISH labeling strategies for detecting chromosomal structural aberrations in sperm of humans and mice. For both assays the approximate locations of each probe is indicated. (A) ACM assay for the detection of deletions and duplications of chromosome 1p, breaks in the classical satellite region of 1q, as well as numerical abnormalities in human sperm. (B) CT8 assay for detection of deletions and duplications of chromosome 2 and numerical abnormalities of chromosomes 2 and 8.

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predominantly during oogenesis (6), transmitted de novo structural aberrations tend to originate predominantly during spermatogenesis (9,21,22). Methods for detecting chromosomal aberrations in sperm have in common that two or more loci on a single target chromosome are interrogated simultaneously using two or more fluorescent DNA probes. As shown in Fig. 1A, the ACM assay uses three probes for specific regions of chromosome 1: 1cen (alpha satellite or A), 1q.12 (classical satellite or C), and 1p36.3 (midisatellite or M) to allow the simultaneous detections of three classes of chromosomal defects associated with abnormal reproductive outcomes: (1) numerical abnormalities, (2) segmental duplications and deletions, and (3) chromosomal breaks. The CT8 assay (Fig. 1B) uses a combination of probes specific for the centromeric (C) and telomeric (T) regions of chromosome 2 together with a subtelomeric probe for chromosome 8 to detect three types of damage: (1) duplications and deletions involving chromosome 2, (2) aneuploidies involving chromosomes 2 and 8, and (3) sperm diploidy. The specific types of chromosomal aberrations and their associated sperm FISH phenotypes are shown in Tables 1 and 2 for the ACM and CT8 assay, respectively. This chapter describes the laboratory protocols for these two FISH methods for the detection of chromosomal structural aberrations in sperm of humans and mice together with a detailed description of the criteria used for collecting data. A description of the basic principles of FISH can be found in the chapter by Pacchierotti and Sgura (this volume).

2. The Human Sperm ACM Assay The ACM method provides an important new approach for measuring exposure to chromosome-breaking agents and assessing genetic predisposition to such damage. As shown in Table 1, analyses of sperm from healthy men indicated that (1) the spontaneous frequencies of chromosomal structural abnormalities are higher than those of numerical aberrations; (2) chromosomal breaks are more prevalent than duplications and deletions; duplications and deletions of 1q36.3 are significantly higher (5-fold) than duplications and deletions of 1cen; and (3) within each chromosomal region, duplications and deletions tend to occur at the same rate (19). The ACM assay has been used recently to show that oligozoospermic men have higher frequencies of sperm with chromosomal structural aberrations than normozoospermic sperm (23) and that there is a gradual increase in the frequencies of sperm with chromosomal structural

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Table 1 Chromosomally Abnormal Sperm Detected by the Human ACM Assay Sperm chromosomal defects Segmental aneuploidies 1pter duplication 1pter deletion 1cen-1q12 duplication 1cen-1q12 deletion Chromosomal breaks Breaks between 1cen and 1q12 Breaks within 1q12 Total structural aberrations Numerical abnormalities Disomy 1 or diploidy Nullisomy 1

FISH phenotypea

Baseline frequenciesb

ACMM ACO ACACM OOM

1.0 0.5 2.2 1.6

± ± ± ±

0.3 0.2 0.4 0.3

A.CM AC.CM

5.4 ± 0.7 5.5 ± 0.9 16.2 ± 1.3

ACACMM OOO

12.9 ± 2.1 0.3 ± 0.2

a Each chromosomal region is represented by a letter code: A indicates the red signal representative of the alpha satellite; C indicates the blue signal representative of the classical satellite; M indicates the green signal representative of the Midi probe; O indictates the absence of the signal for a chromosomal region; A.C indicates a break between the a and classical regions; C.C indicates a break within the classical region. b Frequencies per 10,000 sperm ± standard deviation. Data from Sloter et al. (2006).

aberrations as men age (24). The following subheadings describe the materials, reagents, and protocols for performing the ACM assay. 2.1. Materials and Reagents 2.1.1. Preparation of Semen Smears 1. 3–9 μL of fresh or frozen semen. (See Note 1.) 2. Glass slides with frosted end. 3. 100% Ethanol–200 proof.

2.1.2. Decondensation of Sperm 1. 40 mL of dithiothreitol (DTT, Sigma-Aldrich, St. Louis, MO) solution in a Coplin jar placed in ice. 2. 40 mL of lithium 3,5-diiodosalicylate (LIS, Sigma-Aldrich) in a Coplin jar at room temperature (RT). 3. Tris (hydroxymethyl) aminomethane hydrochloride (Tris–HCl, Sigma); adjusted to pH 7.8. 4. Autoclaved distilled water (dH2 O). 5. Circulating water bath set to 77–78°C.

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6. 70% formamide/2× SSC: 315 mL of formamide (Shelton Scientific, Peosta, IA), 45 mL 20× SSC (87.65 g of NaCl, 44.1 g of Na citrate, 400 mL of dH2 O; adjust pH to 7.0 with HCl or NaOH; adjust volume to 500 mL with dH2 O), 60 mL autoclaved dH2 O, then adjusted to pH 7.0 with 2 N HCl and to volume 450 mL with extra autoclaved dH2 O. 7. 70%, 85%, 100% Ethanol in Coplin jars.

2.1.3. Random Priming of Probes 1. Template DNA: human 1cen unlabeled pSDZ1-1; human 1 classical satellite pUC1.77; 1midi D1Z2. 2. 2.5× Random primers (Invitrogen). 3. 10× Buffer: 10 μL of Tris-HCl, pH 7.5, 5 μL of Na2 EDTA (stop buffer), 485 μL of sterilized H2 O. 4. Bioprime labeling kit with Klenow, sterilized H2 O, dNTP mix (Invitrogen).

2.1.4. Probe Mixture Preparation 1. Master mix: 5.5. mL of formamide, 0.5 mL of 20× SSC, 1 dextran sulfate, heat at 72°C until completely dissolved, adjust pH to 7.0, and adjust the volume to 7 mL with dH2 O. Aliquot in 1.5-mL Eppendorf vials and store at −20°C. 2. Herring sperm (Invitrogen). 3. Labeled probes generated from random priming reactions.

2.1.5. Hybridization 1. 2. 3. 4. 5.

Slide warmer set to 37°–42°C. Glass cover slips 22 × 22 mm, No. 2 (Corning). Rubber cement (Starkey, La Grange, IL). Prewarmed humidity chamber. Incubator with temperature set to 37°C.

2.1.6. Posthybridization Washes 1. Circulating water bath set to 77–78°C. 2. 60% Formamide/2× SSC: 30 mL of formamide, 5 mL of 20× SSC, 10 mL of autoclaved dH2 0, then adjust pH to 7.0 with 2 N HCl and adjust volume to 50 mL with dH2 0. 3. 2× SSC: Dilute 100 mL of 20× SSC with 800 mL of autoclaved dH2 O. Adjust the pH to 7.0 with 2 N HCl. Add more water until the volume reaches 1000 mL. Filter sterilize. Store at RT.

2.1.7. Antibody Staining (per slide) 1. 40 μL of PNM: Add 5 g of nonfat dry milk to approx 100 mL of PN buffer and add 20 μL of 0.02% sodium azide (Sigma-Aldrich). Incubate at 37°C for 60 min. Leave on a lab bench overnight. Remove supernatant in two 50-mL centrifuge tubes and aliquot into 1.5-mL Eppendorf tubes. Store at 4°C.

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2. 1 μL of Pacific Blue–streptavidin (stock concentration 2.5 μg/mL; Molecular Probes, Eugene, OR). 3. 0.5 μL of anti-digoxigenin (DIG) –fluorescein isothiocyanate (FITC) (stock concentration 0.2 mg/mL; Boehringer Mannheim, Indianapolis, IN).

2.1.8. Counterstaining 1. Vectashield (Vector, Burlingame, CA). 2. Cover slips 22 × 22 mm, No. 1 (Goldseal).

2.2. Laboratory Methods for the Human ACM Assay 2.2.1. Semen Smear Slide Preparation 1. Soak slides in 100% ethanol for at least 2 days. Layer the slides in a jar or a container with a lid in a criss-cross fashion to expose as much of the slide as possible to the alcohol. Fill the jar with ethanol until all slides are covered. When removing the slides, wear appropriate personal protective equipment (PPE), wipe all slides vigorously with a Kimwipe, and place them in a slide box. 2. Using a diamond stylus, cut a piece from another clean slide in which the width is slightly less than the slide’s width (using another slide for measuring); after marking the surface, break the slide by bending inwards allowing the piece to break off carefully. Be careful not to touch the edge that will be in contact with the sample. As an alternative, cover slips may be used instead; however, they can break more easily during smearing. Wear appropriate PPE. 3. Apply 7 μL of semen sample onto a clean glass slide using a pipet. This volume may vary (i.e., 3–9 μL) depending on the sample’s sperm concentration. (See Note 2 for low sperm count donors.) 4. Smear the sample using a cut piece of a previously prepared slide. Holding the frosted end in one hand so that the slide is slightly inclined and the other end resting on the workbench, place the cut slide piece (or cover slip) directly on the sample drop; the clean untouched factory edge only should be in contact with the slide at this point. Slowly move the edge of the cut piece/cover slip so it comes in contact with the sample; the sample will automatically distribute along the length of the edge. After the sample is distributed along the edge, slide the edge toward the frosted end, allowing the sample to smear about 2.5–4 cm. 5. Lay the smeared slide flat on a clean paper towel. When the sample dries, store the smeared slides into a slide box with the cover slightly off to allow air-drying for at least 24 h. It is highly recommended to leave slides out for several days. (See Note 3.)

2.2.2. ACM Probe Preparation This process uses random priming to generate labeled probes for hybridization. The random octamers are annealed to denatured DNA template

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and extended by Klenow fragments. During the process, fluorescently labeled11-dUTP is incorporated during a 10- to 40-fold amplification to produce probes for use in FISH. (See Note 4.) 2.2.2.1. Alpha-Rhodamine 1. Prepare rhodamine dNTP mix (for five reactions, prepare the following: 2 μL of 25 mM dCTP, 2 μL of 25 mM dGTP, 2 μL of 25 mM dATP, 1.4 μL of 25 mM dTTP, 15 μL of 1 mM rhodamine-6-dUTP). 2. Mix 20 μL of random primer, 0.2 μL of template DNA (pSDZ1-1 1 cen rhodamine), and 23.8 μL of sterilized water in a polymerase chain reaction (PCR) tube. 3. Program thermal cycler to denature for 10 min at 99.9°C, remove the vial, and place immediately in ice for 5 min. 4. Add 5 μL of 10× buffer, 5 μL of dNTP mix, and 1 μL of Klenow (see Note 5) from Bioprime Kit to denatured template. Total volume = 55 μL. 5. Gently mix and centrifuge for 30 s. 6. Program thermal cycler to incubate at 37°C for 3 h, 70°C for 10 min, and maintain at 4°C. 7. Transfer probe to a 1.5-mL Eppendorf tube, label, and store in a –20°C freezer.

2.2.2.2. Classical Biotin 1. Mix 20 μL of random primer, 0.3 μL of template DNA (pUC1.77—classical biotin), and 23.7 μL of sterilized water in a PCR tube. 2. Program thermal cycler to denature for 10 min at 99.9°C, remove the vial, and place immediately in ice for 5 min. 3. Add 5 μL of 10× buffer, 5 μL of dNTP mix from the Bioprime Labeling Kit, and 1 μL of Klenow (see Note 5) from Bioprime Kit to denatured template. Total volume = 55 μL. 4. Gently mix and centrifuge for 30 s. 5. Program thermal cycler to incubate at 37°C for 3 h, 70°C for 10 min, and maintain it at 4°C. 6. Transfer the probe into a 1.5-mL Eppendorf tube, label, and store in a –20°C freezer.

2.2.2.3. Midi-Digoxigenin 1. Prepare DIG dNTP mix (for five reactions, prepare the following: 2 μL of 25 mM dCTP, 2 μL of 25 mM dGTP, 2 μL of 25 mM dATP, 1.4 μL of 25 mM dTTP, 15 μL of 1 mM DIG-11-dUTP). 2. Mix 20 μL of random primer, 0.1 μL of template DNA (D1Z2 midi-digoxigenin), and 23.9 μL of sterilized water in a PCR tube. 3. Program thermal cycler to denature for 10 min at 99.9°C, remove the vial, and place immediately in ice for 5 min. 4. Add 5 μL of 10× buffer, 5 μL of dNTP mix, and 1 μL of Klenow (see Note 5) from the Bioprime Kit to denatured template. Total volume = 55μL.

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5. Gently mix and centrifuge for 30 s. 6. Program the thermal cycler to incubate at 37°C for 3 h, 70°C for 10 min, and maintain it at 4°C. 7. Transfer the probe into a 1.5-mL Eppendorf tube, label, and store at –20°C.

2.2.3. Probe Mixture 1. Prepare probes using the following amounts per slide: 10.5 μL of master mix; 1.0 μL of herring sperm DNA; 1.5 μL of D1Z5 A. rhodamine; 1.0 μL of pUC1.77-C. biotin; 1.0 μL of D1Z2 M. digoxigenin. 2. Mix by gentle pipetting and centrifuge briefly (2 s) using a tabletop centrifuge. 3. Keep probe mix in –20°C until ready for denaturation.

2.2.4. Sperm Nuclei and Probe Decondensation 1. Place slide smears in DTT (4 mL of 1 M Tris-HCl, pH 7.8, + 36 mL of dH2 O + 0.0617 g of DTT) on ice for 30 min. DTT should be prepared fresh every time. Do not denature more than four slides per Coplin jar. DTT disrupts disulfide bridges between protamine molecules. 2. Transfer slides to LIS (4 mL of 1 M Tris-HCl pH 7.8 + 36 mL of dH2 O + 0.0633 g of LIS) at RT for 75–90 min. LIS penetrates sperm membranes and swells nuclei. If slides are kept in LIS for less than 75 min, dimmer signals are produced. If slides are kept in LIS for more than 90 min, diffuse signals are produced. 3. Dry slides upright up at RT on absorbent paper for 2–3 h. Over-drying (3+ h) may allow spontaneous reformation of disulphide bonds and recondensation of sperm. 4. To prepare for hybridization, warm denaturation solution (70% formamide/2× SSC) in circulating water bath set to 77–78°C (20–30 min before use). 5. Denature slides for 2 min in 70% formamide/2× SSC at 77–78°C in a circulating water bath. (See Note 6.) 6. Keep slides in ice-cold ethanol series of 70%, 85%, and 100% for 2 min each. (See Note 7.) 7. Air dry slides upright on absorbent paper at RT for 30 min. (See Note 8.) 8. After drying, check sperm density under microscope using phase contrast and etch an area with a consistent sperm density to hybridize. Choose an area big enough for a cover slip. Continue hybridization only if you have enough cells. 9. Immerse the cover slip in ethanol for a few seconds and clean dry with a Kimwipe. 10. Set the slide warmer to 37°C. 11. Retrieve probes from freezer, tap and centrifuge a few seconds using a benchtop centrifuge. (See Note 9.) 12. Denature probe mix for 6 min (it can be done for up to 10 min) at 77–78°C in circulating water bath and plunge immediately into ice for no more than 5 min.

2.2.5. Hybridization 1. Place the slide on a slide warmer and immediately drop denatured probe mix onto the etched area and add a cover slip. Avoid air bubbles. (See Note 10.) 2. Seal the cover slip with rubber cement.

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3. Put slides in a prewarmed humidity chamber. A plastic tip box with half an inch of water at the bottom works perfectly for this task. (See Note 11.) 4. Allow the slides to incubate at 37°C over two nights.

2.2.6. Posthybridization Washes and Detection (see Note 12) 1. Peel the rubber cement off carefully with forceps. Do not allow the cover slip to move around. 2. Place the slide in a Coplin jar containing 2× SSC at RT to allow cover slips to fall off. (See Note 13.) 3. Wash slides in two changes of fresh 2× SSC at RT for 3 min each. 4. Drain slides but do not allow them to completely dry. 5. Add 1 μL of Pacific Blue–streptavidin to 40 μL of PNM. Mix well and transfer to the slide. 6. Apply a plastic cover slip and incubate for 30 min in the dark at RT. (See Note 14.) 7. Drain excess liquid and wash two times in Coplin jars containing 2× SSC at RT for 3 min each. 8. Warm circulating water bath and washing solution (60% formamide/2× SSC) to 45°C. Thirty minutes is enough to time to allow solution to warm up to the desired temperature. (See Note 15.) 9. Wash slides (no more than two at a time) in 60% formamide–2× SSC at 45°C for 4 min. 10. Wash slides two more times in Coplin jars containing 2× SSC solution at RT for 5 min each. 11. Add 0.25 μL of Pacific Blue–streptavidin and 0.5 μL of anti-DIG FITC to 100 μL of PNM. Mix well and aliquot 40 μL of the mixture to each slide. 12. Apply a plastic cover slip and incubate using the humidity chamber in the dark 30 min at RT. 13. Drain excess liquid and wash again two times in the same Coplin jars containing 2× SSC solution for 3 min each.

2.2.7. Counterstaining 1. Drain the slides well but do not allow them to dry completely. Apply 10 μL of Vectashield antifade on the etched area and place a No. 1 thickness 22 × 22-mm cover slip on the slide. 2. Keep at 4°C until ready for analysis (see Subheading 4.).

3. The Mouse Sperm CT8 Assay The mouse CT8 assay is the first robust rodent screen for potential male germ cell aneugens and clastogens and it has been used to show that (20): (1) the baseline frequencies of sperm carrying structural aberrations involving chromosome 2 are more common than sperm aneuploidy for chromosomes 2 and 8 combined; (2) diploid sperm were the most common anomaly found in mouse sperm;

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Table 2 Chromosomally Abnormal Sperm Detected by the Mouse CT8 Assay Sperm chromosomal defects Segmental aneuploidies 2pter duplication 2pter deletion 2cen duplication 2cen deletion Numerical abnormalities Disomy 2 Nullisomy 2 Disomy 8 Nullisomy 8 Diploidy

FISH phenotypea

Baseline frequenciesb

CTT8 CO8 CCT8 OT8

0.1 0.1 0.1 0.3

± ± ± ±

0.3 0.3 0.3 0.5

CCTT8 OO8 CT88 CTO CCTT88

0.3 0.1 0.2 0.6 2.4

± ± ± ± ±

0.7 0.3 0.4 0.8 1.8

a

Each chromosomal region is represented by a letter code: “C” indicates the red signal representative of the 2cen region; “T” indicates the green signal representative of the 2tel region; “8” indicates the yellow signal representative of the 8 probe; “O” indicates the absence of the signal for a chromosomal region. b Frequencies per 5000 sperm ± standard deviation. Data from Marchetti et al.(2006).

and (3) sperm with duplications or deletions of the centromeric and telomeric region of chromosome 2 occurred at similar frequencies (Table 2). Comparison of the spontaneous frequencies of sperm with chromosomal structural aberrations between humans and mice showed that healthy human males appear to produce approx 6 times higher frequencies of sperm with chromosomal aberrations than mice (2). Recently, the CT8 assay was used to show that exposure of male mice to chemotherapeutic relevant doses of etoposide resulted in major increases in the frequencies of sperm carrying chromosomal aberrations in both meiotic pachytene (27- to 578-fold) and spermatogonial stem-cells (8- to 16fold), while aneuploid sperm were induced only after treatment of meiotic cells (27-fold) with no persistent effects in stem cells (25). The following sections describe the materials, reagents, and protocols for performing the CT8 assay. 3.1. Materials and Reagents 3.1.1. Epididymal Sperm Smear Preparations 1. 2. 3. 4.

2.2% Sodium citrate (isotonic solution). 1.5-mL microcentrifuge tubes. Incubator, Ethanol-cleaned glass microscope slides.

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3.1.2. Decondensation of Mouse Sperm 1. 40 mL of DTT (Sigma-Aldrich, St. Louis, MO) solution in Coplin jar placed in ice. 2. 40 mL of Tris-HCl (Sigma-Aldrich); adjust pH to 7.8. 3. Autoclaved dH2 O. 4. Circulating water bath set to 77–78°C. 5. 70% Formamide/2× SSC: 315 mL of formamide (Shelton Scientific, Peosta, IA), 45 mL of 20× SSC, 60 mL of autoclaved dH2 O, then adjust pH to 7.0 with 2 N HCl and adjust volume to 450 mL with extra autoclaved dH2 O. 6. 70%, 85%, 100% Ethanol in Coplin jars.

3.1.3. Random Priming of Probes 1. Template DNA: mouse chromosome 2 centromere probe (Research Genetics, 460H-4), mouse chromosome 2 telomere probe (Research Genetics, 121-E-1), clones 4a and 5e form mouse chromosome 8 (26). 2. 2.5× Random primers (Gibco, Bethesda, MD). 3. 10× buffer. 4. Bioprime Labeling Kit with Klenow, Sterilized H2 O, dNTP mix (Invitrogen).

3.1.4. Probe Mixture Preparation The following quantities are for preparing probes necessary to hybridize 4 slides. These probe quantities may need to be adjusted depending on the quality of each batch of probe (see Subheading 3.2.3. for probe generation). 1. 2. 3. 4. 5. 6. 7. 8. 9. 10.

30 μL of mouse Cot-1 DNA (Invitrogen). 4 μL of mouse 2cent-DIG. 2 μL of mouse 2tel-bio. 1 μL of mouse 8-bio (clone 4a). 1 μL of mouse 8-bio (clone 5e). 0.5 μL of mouse 8-DIG (clone 4a). 0.5 μL of mouse 8-DIG (clone 5e). 2 μL of herring sperm DNA (Invitrogen). 4.1 μL of 3 M sodium acetate. 112.75 μL of ice-cold 200 proof ethanol.

3.1.5. Hybridization Same as Subheading 2.1.5. 3.1.6. Posthybridization Washes and Antibody Staining 1. Circulating water bath set to 45o C. 2. 50% formamide/2× SSC: 25 mL of formamide (Shelton Scientific), 5 mL of 20× SSC, 10 mL of autoclaved dH20, then adjust pH to 7.0 with 2N HCL and adjust volume to 50 mL with dH2 0.

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3. 2× SSC: Dilute 10 mL of 20× SSC with 800 mL of autoclaved dH2 O. Adjust the pH to 7.0 with 2 N HCl. Add more water until volume reaches 1000 mL. Filter sterilize. Store at room temperature. 4. 100 mL of PN buffer. 5. Dual detection reagent (1 μL of fluorescein avidin DCS; 1 μL of anti-digoxigeninrhodamine, 498 μL of PN, store at 4° C).

3.1.7. Counterstaining Same as Subheading 2.1.8. 3.2. Laboratory Methods for the Mouse CT8 Assay 3.2.1. Epididymis Sperm Smear Preparation 1. Set the incubator to 32°C. 2. Fill 1.5-mL microcentrifuge tube with 300 μL of 2.2% sodium citrate, and prewarm to 32°C. 3. Euthanize mice with CO2 following the guidelines for the care and use of laboratory animals for research. Isolate the cauda epididymides from both testes. Hold each cauda with tweezers and cut small slits into the cauda using iris scissors. Be careful to keep the cauda as one piece. 4. Place both cauda in the microcentrifuge tube with 300 μL of 2.2% sodium citrate. 5. Incubate at 32°C for 10 min to allow sperm to swim out of the cauda. 6. Remove the cauda. The sperm suspension can be used immediately for making smears or stored at –20°C. Pipet 5 μL of the sperm suspension onto a clean glass slide. Gently smear the sperm suspension over an approx 22 × 22 mm2 area of the glass slide using the side of a pipet tip. Alternately, a glass slide may be used to make a smear as long as an adequate cell density is attained (see Subheading 2.2.1.). Allow the smears to air dry at room temperature for at least 24 h and store the slides at –20°C in nitrogen gas.

3.2.2. Decondensation of Mouse Sperm 1. Make 1 M Tris-HCl stock solution: 157.6 g of Tris-HCl and dissolve in approx. 800 mL of dH2 O; adjust pH to 7.8 by adding NaOH. Bring up to total volume of 1 L. Autoclave before use. 2. Make 10 mM DTT (made fresh for each use): a. Make 0.1 M Tris-HCl: Measure 36 mL of dH2 O, pour into a Coplin jar placed in ice. Add 4 mL of 1 M Tris-HCl stock (from step 1). Mix well. b. Make 10 mM DTT: 0.0617 g of DTT from Sigma and mix with 0.1 M Tris-HCl solution in the Coplin jar by stirring with spatula until the powder completely dissolves. Place the Coplin jar on ice. 3. Place the glass slides into the Coplin jar containing ice cold 10 mM DTT (step 2) for 30 min.

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4. Remove slides from Coplin jar and drain. Briefly dip slide in dH2 O at room temperature and drain. Dry slide at room temperature for at least 30 min. Slides are ready for hybridization when thoroughly dried.

3.2.3. Random Priming of Probes 3.2.3.1. Random Priming with Digoxigenin-Modiﬁed Nucleotides 1. Mix 25–500 ng of DNA template with dilute buffer or water to make a final volume of 19 μL in a microcentrifuge tube. Add 20 μL of the 2.5× random primer solution, mix well and denature for 5–10 min at 100°C. Immediately cool on ice. 2. Add 5 μL of reaction buffer, 5μL of DIG/dNTP mix, mix well and briefly centrifuge. 3. Add 1 μL of Klenow fragment. Mix gently but thoroughly and centrifuge for 30 s. 4. Incubate at 37°C for 3 h. 5. Heat to 70°C to deactivate the enzyme (it is not necessary to add stop buffer). Store at –20°C until use.

3.2.3.2. Random Priming with Biotin-Modiﬁed Nucleotides 1. Mix 25–500 ng of DNA template with dilute buffer or water to make a final volume of 19 μL in a microcentrifuge tube. Add 20 μL of the 2.5× random primer solution, mix well and denature for 5–10 min at 100°C. Immediately cool on ice. 2. Add 5 μL of the kit 10× dNTP mix, mix well, and briefly centrifuge. 3. Add 1 μL of Klenow fragment. Mix gently but thoroughly and centrifuge for 30 s. 4. Incubate at 37°C for 3 h. 5. Heat to 70°C to deactivate the enzyme (it is not necessary to add stop buffer). Store at –20°C until use.

3.2.4. Probe Mixture 1. Prepare the probe mixture with amounts of reagents as indicated in Subheading 3.1.4. These quantities (per slide to be hybridized) may need to be adjusted depending on the quality of each batch of probe. 2. Keep at –80°C for 1 h to overnight. 3. Centrifuge for 30 min at 180g, pour off the supernatant, and dry the pellet. 4. Reconstitute probe mix with 3 μL of water and 7 μL of CEP hybridization buffer (per slide). 5. Denature probe mix 10 min at 78°C, and preanneal at 37°C for 30 min.

3.2.5. Slide Pretreatment and Denaturation (see Note 15) 1. Flood slides with 3:1 methanol–acetic acid and air-dry. 2. Place slides in DTT on ice (4 mL of 1M Tris-HCl, pH 7.8, + 36 mL of dH2 O + 0.0617 g of DTT) for 30 min. 3. Dip in dH2 O and dry completely at RT (minimum of 30 min). 4. Denature smears in 70% formamide/2× SSC at 78°C for 6 min. (See Note 6.) 5. Dehydrate slides using a series of ice-cold 70%/85%/100% ethanol (2 min each).

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6. Dry completely at RT; check under a microscope for sperm density and mark area to be hybridized.

3.2.6. Hybridization 1. Clean glass cover slips with 100% ethanol. 2. Prewarm slides on a slide warmer set to 42°C for about 1 min. 3. Add 10 μL of the probe mixture and cover the slide with a 22 × 22 mm2 glass cover slip. 4. Leave the slide on the slide warmer for a few minutes and cover the edges with rubber cement. Make sure that all the edges are covered by rubber cement. Add more rubber cement if needed. 5. Incubate over 2 nights at 37°C in a moist prewarmed box. (See Note 11.)

3.2.7. Washing and Detection 1. 2. 3. 4. 5. 6.

Carefully remove the rubber cement with forceps. Wash for 5 min in 50% formamide/2× SSC at 45°C. Repeat two additional times. Wash for 5 min in 2× SSC at 45°C. Wash for 5 min in PN at 45°C. Wash for 5 min in PN at room temperature. Drain the slide and without allowing it to dry, add 30 μL of dual detection reagent, cover with a plastic cover slip, and keep at RT in a moist box for 40 min. 7. Final wash for 3 min in PN at room temperature (twice).

3.2.8. Counterstaining 1. Apply 10 μL of DAPI (0.01 μg/mL) in Vectashield on marked area and apply cover slip. 2. Keep at 4°C until scoring (see Subheading 4).

4. Microscope Data Collection Protocols for Sperm FISH Assays The previous subheadings have described the laboratory protocols for producing slides with good hybridization quality for both the ACM and CT8 assays. Hybridization quality, that is, high bright and compact FISH signals and low background, is probably the most important factor affecting the success of a sperm FISH study. However, equally important are the visualization of the FISH signals and protocols for data collection that were developed to reduce to a minimum the influence of technical factors on the experimental outcome (see Pacchierotti and Sgura, this volume, for information about fluorescence microscopes and filter settings). In the next few subheadings we will focus on several technical factors that are critical to the reliability of the sperm FISH assay such as: (1) the development of strict scoring criteria to determine whether a sperm has an abnormal number of spots; (2) blinding of scorers

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and procedures for data collection; and (3) harmonization of scoring criteria among different scorers. (See Note 16.) 4.1. Scoring Criteria for ACM and CT8 Sperm FISH Assays The subjective evaluation of FISH signal is an issue that can result in great variability in scoring results among different laboratories and within laboratories. Our group has invested significant effort in developing strict scoring criteria that reduce scorer-to-scorer variability. Strict adherence to these scoring criteria is essential for generating reproducible data. The first step in the scoring criteria is to determine whether a sperm should be scored. In order to score a sperm, it must meet all of the following criteria (these criteria apply to both human and mouse sperm): 1. The entire cell must be visible. Do not count a cell unless the total cell border can be seen. Do not count cells that are partially covered by other cells. Example: do not count cells that exist in large clumps where overlapping cells might obscure the fluorescence signals. 2. The cell must appear intact. The contents of the cell should remain within the cell borders. Sometimes overly decondensed cells will spill their nuclear content or their fluorescence domains and are not acceptable for scoring. Example: do not count cells where the fluorescence domains are outside the main body of the sperm head. 3. The cell should be in a well-hybridized area of the slide. Usually on hybridized slides, there are whole regions that hybridize efficiently, as well as areas that are less well hybridized. It may be necessary to pass over some areas if the hybridization is poor. 4. The cell size should be no less than 5 μm and no greater than 15 μm. An eye-piece with a reticle can be used for this task. Nondecondensed cells do not hybridize reliably and to keep from biasing the data, cells less than 5 μm on the reticle are not included in the denominator. A tally of such cells is kept but their fluorescence domains are not scored. Cells that overly decondense give very diffuse hybridization signals and cannot be reliably scored. 5. The cell should have no background.There may be some background signals on the slide. An area with high background makes it difficult to distinguish true signals from the background and should not be scored.

Only sperm that pass all these criteria can be scored for their FISH phenotype. Next, we describe the decision tree for the ACM assay, followed by the decision tree for the CT8 assay. 4.2. Decision Tree for the ACM Assay Before analyzing the FISH signals the scorer should determine whether the cell qualify for scoring by using the phase contrast to evaluate the cell’s

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outline (intact or spilled), presence of one or multiple tails. Also, it should be established that it is a single cell rather than two overlapping cells. To be scored, the following criteria must be met. 1. 2. 3. 4. 5. 6.

The entire cell is visible with no part hidden from view. The cell border appears intact. The cell has a tail or tail attachment site. (See Note 17.) All the hybridization signals are inside the cell The cell size is no less than 5 or greater than 15 notches by the reticle. The cell is in an area of the slide in which a majority of the cells have hybridized.

If any of these criteria are not met, this cell is not acceptable for scoring. If contents are spilled or the cell is too small or too large, this information should be tallied, but the hybridization signals are not scored. If the cell is acceptable for scoring, the next step in the decision tree is to determine the FISH phenotype. 4.2.1. Is There a Break in Between the Alpha (A) and Classical (C) Probes? To be defined as a break between the A and C probes the following criteria must be met: 1. The A and C signals must be at least one A signal domain width apart. 2. Both signals must be of normal size based on the hybridization signals seen for these probes over the same region of the slide. 3. The signals must be entirely separate and not attached by any threads. 4. The cell has a tail or a tail attachment site. 5. There are no hybridization signals either in surrounding cells or within this cell that would raise the question that this is an artifact.

If all of these criteria are met, the cell is scored as having a break between the A and C region; otherwise the cell is not scored. 4.2.2. Is There a Break Within the Classical (C) Probe? To be defined as a break within the C probe, the following criteria must be met: 1. Only one C signal is associated with the A signal. 2. If both C signals are of the same size they must be separated by a width as large as the individual signals; if they are of unequal size, they must be separated by at least the width of the larger C domain. 3. Both signals must have approximately the same intensity. 4. The signals must be entirely separate and not attached by any threads. 5. The cell has a tail or a tail attachment site. 6. There are no hybridization signals either in surrounding cells or within this cell that would raise the question that this is an artifact.

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If all of these criteria are met, the cell is scored as having a break within the C region; otherwise the cell is not scored. 4.2.3. Are There Duplications or Deletions of Any of the Signals? To be defined as a duplication of the C region, the following criteria must be met: 1. The two C signals within the cell must have approximately the same size. 2. Both signals must have approximately the same intensity. 3. Both signals should be approximately of normal size based on the hybridization signals seen for this probe over the same region of the slide. 4. The signals must be at least a full domain’s width apart. 5. The signals must be entirely separate and not attached by any threads. 6. The cell has a tail or a tail attachment site. 7. There are no hybridization signals either in surrounding cells or within this cell that raise the question that this is an artifact.

If all of these criteria are met, the cell is scored as having a duplication of the C region; otherwise the cell is not scored. To be defined as a deletion of the C region the following criteria must be met: 1. The absence of the C signal is confirmed by using the single filter specific for this signal. 2. There are no debris near or around cell. 3. The cell is not damaged. 4. The cell has a tail or a tail attachment site.

If all of these criteria are met, the cell is scored as having a deletion of the C region; otherwise the cell is not scored. A similar process is used to determine whether there are duplications and deletions of the A and M signals. To be defined as a duplication of the AC probes, the following criteria must be met: 1. The two signals of each probe must have approximately the same size. 2. The two signals of each probe must have approximately the same intensity. 3. The two signals of each probe must be approximately of normal size based on the hybridization signals seen for these probes over the same region of the slide. 4. The signals must be at least a full domain’s width apart. 5. The signals must be entirely separate and not attached by any threads. 6. The cell has a tail or a tail attachment site. 7. There are no hybridization signals either in surrounding cells or within this cell that raise the question that this is an artifact.

If all of these criteria are met, the cell is scored as having a duplication of the A and C regions; otherwise the cell is not scored.

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To be defined as a deletion of the AC regions, the following criteria must be met: 1. The absence of the A and C signals is confirmed by using triple, single, and DAPI filters. 2. There are no debris near or around cell. 3. The cell is not damaged. 4. The cell has a tail or a tail attachment site.

If all of these criteria are met, the cell is scored as having a deletion of the AC regions; otherwise the cell is not scored. 4.2.4. Is This Cell a Diploid Sperm? To be defined as a diploid sperm the following criteria must be met: 1. 2. 3. 4. 5. 6. 7. 8.

There are two A signals plus two C signals plus two M signals. The two signals of each probe must have approximately the same size. The two signals of each probe must have approximately the same intensity. The two signals of each probe must have approximately normal size based on the hybridization signals seen for this probe over the same region of the slide. The two signals of each probe must be at least a full domain’s width apart. The signals must be entirely separate and not attached by any threads. There are no hybridization signals either in surrounding cells or within this cell that raise the question that this is an artifact. The cell is really one cell and not two overlapped cells (this should be checked carefully by phase contrast).

If all of these criteria are met, the cell is scored as being diploid; otherwise the cell is not scored. 4.2.5. Are There No Signals Within the Cell? Occasionally sperm with no hybridization signals will be seen. In these cases, the absence of the signals should be confirmed using all filters. In addition, under phase contrast it should be checked that the cell is within size requirements, that there is no debris overlapping sperm, and that there are no cells overlapping. After checking the above criteria, this cell is scored as OOO (Table 1). 4.3. Decision Tree for the CT8 Assay Before analyzing the FISH signals the scorer should determine whether this cell qualifies for scoring by using phase contrast to evaluate the cell’s outline (intact or spilled), hooked shape of the head, and presence of one or multiplae tails. Also, it should be determined that it is a single cell rather than two overlapping cells. In order to be scored, the following criteria must be met:

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The entire cell is visible with no part hidden from view. The cell border appears intact. The cell has a hook. All the hybridization signals are inside the cell. The cell is in an area of the slide in which a majority of the cells have hybridized.

If any of these criteria are not met, this cell is not acceptable for scoring. If contents are spilled or the cell is too small or too large, this information should be tallied, but the hybridization signals are not scored. If the cell is acceptable for scoring, the next step in the decision tree is to determine the FISH phenotype. 4.3.1. Are There Duplications or Deletions of the Centromeric Region of Chromosome 2? To be defined as a duplication of the C region, the following criteria must be met: 1. The two C signals within the cell must have approximately the same size. 2. Both signals must have approximately the same intensity. 3. Both signals should be approximately of normal size based on the hybridization signals seen for this probe over the same region of the slide. 4. The signals must be at least a full domain’s width apart. 5. The signals must be entirely separate and not attached by any threads. 6. The cell has a hooked shape. 7. There are no hybridization signals either in the surrounding cells or within this cell that raise the question that this is an artifact.

If all of these criteria are met, the cell is scored as having a duplication of the C region; otherwise the cell is not scored. To be defined as a deletion of the C region the following criteria must be met: 1. The absence of the C signal is confirmed by using the single filter specific for this signal. 2. There are no debris near or around cell. 3. The cell is not damaged. 4. The cell has a hooked shape or a tail.

If all of these criteria are met, the cell is scored as having a deletion of the T region; otherwise the cell is not scored. 4.3.2. Are There Duplications or Deletions of the Telomeric Region of Chromosome 2? To be defined as a duplication of the T region, the following criteria must be met: 1. The two T signals within the cell must have approximately the same size. 2. Both signals must have approximately the same intensity.

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3. Both signals should be approximately of normal size based on the hybridization signals seen for this probe over the same region of the slide. 4. The signals must be at least a full domain’s width apart. 5. The signals must be entirely separate and not attached by any threads. 6. The cell has a hooked shape. 7. There are no hybridization signals either in surrounding cells or within this cell that raise the question that this is an artifact.

If all of these criteria are met, the cell is scored as having a duplication of the T region; otherwise the cell is not scored. To be defined as a deletion of the T region the following criteria must be met: 1. The absence of the T signal is confirmed by using the single filter specific for this signal. 2. There are no debris near or around cell. 3. The cell is not damaged. 4. The cell has a hooked shape or a tail.

If all of these criteria are met, the cell is scored as having a deletion of the T region; otherwise the cell is not scored. 4.3.3. Is This Cell Aneuploid for Chromosome 2? To be defined as being disomic for chromosome 2 the following criteria must be met: 1. The two C and two T signals within the cell have approximately the same size. 2. Both signals of each probe have approximately the same intensity. 3. Both signals of each probe are approximately of normal size based on the hybridization signals seen for these probes over the same region of the slide. 4. The two signals of each probe are at least a full domain’s width apart. 5. The two signals of each probe are entirely separate and not attached by any threads. 6. The cell has a hooked shape. 7. There are no other hybridization signals either in surrounding cells or within this cell that would raise the question that this is an artifact.

If all of these criteria are met, the cell is scored as being disomic for chromosome 2; otherwise the cell is not scored. To be defined as being nullisomic for chromosome 2 the following criteria must be met: 1. The absence of the C and T signals is confirmed by using the single filter specific for each signal. 2. There are no debris near or around cell. 3. The cell is not damaged. 4. The cell has a hooked shape or a tail.

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If all of these criteria are met, the cell is scored as being nullisomic for chromosome 2; otherwise the cell is not scored. 4.3.4. Is this Cell Aneuploid for Chromosome 8? To be defined as being disomic for chromosome 8 the following criteria must be met: 1. The two 8 signals within the cell have approximately the same size. 2. Both signals have approximately the same intensity. 3. Both signals are approximately of normal size based on the hybridization signals seen for these probes over the same region of the slide. 4. The two signals are at least a full domain’s width apart. 5. The two signals are entirely separate and not attached by any threads. 6. The cell has a hooked shape. 7. There are no other hybridization signals either in surrounding cells or within this cell that would raise the question that this is an artifact.

If all of these criteria are met, the cell is scored as being disomic for chromosome 8; otherwise the cell is not scored. To be defined as being nullisomic for chromosome 8 the following criteria must be met: 1. The absence of the 8 signal is confirmed by using the single filter specific for each signal. 2. There are no debris near or around cell. 3. The cell is not damaged. 4. The cell has a hooked shape or a tail.

If all of these criteria are met, the cell is scored as being nullisomic for chromosome 8; otherwise the cell is not scored. 4.4. Is this Cell a Diploid Sperm? To be defined as a diploid sperm the following criteria must be met: 1. 2. 3. 4.

There are two C signals plus two T signals plus two 8 signals. The two signals of each probe must have approximately the same size. The two signals of each probe must have approximately the same intensity. The two signals of each probe must have approximately normal size based on the hybridization signals seen for this probe over the same region of the slide. 5. The two signals of each probe must be at least a full domain’s width apart. 6. The signals must be entirely separate and not attached by any threads. 7. There are no hybridization signals either in surrounding cells or within this cell that would raise the question that this is an artifact.

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8. The cell is really one cell and not two overlapping cells (this should be checked carefully by phase contrast).

If all of these criteria are met, the cell is scored as being diploid; otherwise the cell is not scored. 4.5. Are There No Signals Within the Cell? Occasionally sperm with no hybridization signals will be seen. In these cases, the absence of the signals should be confirmed using all filters. In addition, under phase contrast it should be checked that the cell is within size requirements, there is no debris overlapping sperm, and there are no overlapping cells. After these criteria are checked, the cell is scored as OOO (Table 2). 4.6. Data Collection 4.6.1. Blinding of Scorers and Coding Procedures Despite the implementation of strict scoring criteria as described in the previous subheading, the scoring of sperm FISH signals still remains a subjective analysis and therefore subject to scoring biases. Extreme care should be taken to completely blind the scorers to the origin of the samples being scored. Several procedures can be used to achieve this goal. First, it is very important that the scorer is not able to identify the origin of the slide based solely on sperm concentration. Chemical exposure may result in a significant reduction in the number of sperm in the semen. In these situations, a third person should take particular care that a section of the slide with the highest concentration of sperm is chosen for hybridization so that slides from control and exposed samples do not differ greatly in sperm concentration. Second, the person coding the slides for scoring should ensure that all experimental groups and controls are represented in the set of five or six slides given to the scorer. Any marking on the slide should be removed and an adhesive sticker with the code should be applied on the frosted part of the slides. Third, the method used for tallying the number of sperm scored should provide information to the scorer only of the number of sperm scored and not of how many sperm with abnormal FISH phenotypes have been found. We routinely score 10,000 sperm per sample (in two scoring sessions of 5000 sperm each). The advantage of analyzing large numbers of sperm in a relatively short time by sperm FISH confers a relatively high level of sensitivity and statistical power to these assays, so that small increases can be detected by analyzing sperm from a small number of donors per experimental group. As previously mentioned, sets of five or six coded slides are given to the scorer

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(see Note 18). Once the scorer has finished analyzing 5000 sperm on each of the coded slides, these are returned to the coder, who removes the sticker, and applies a new sticker with a second code. Once the slides are recoded, they are given back to the scorer, who will score a second set of 5000 sperm. The scoring of the first set of 5000 sperm is routinely started on the left side of the cover slip, and the second set of 5000 starting from the right side of the cover slip. To make sure that two independent sections of the slide are scored, a mark with a permanent marker is made on the side of the slide to indicate where the scoring was stopped. In addition, coordinates of the location where scoring was stopped can be recorded. If during the scoring of the second set of 5000 sperm the scorer comes too close to the mark, the scoring should be stopped and another slide from the same donor/animal should be used to complete the second set of 5000 sperm. 4.6.2. Computer Software Cytoscore© for Data Collection Collection of data is done using the Cytoscore© program that was developed at Lawrence Livermore National Laboratory to keep track of the number of sperm scored and their FISH phenotype. The CytoScore© program allows the user to input, in addition to the FISH phenotype, other information, such as size and shape of sperm head, whether a second opinion was provided, and any additional information that the scorer may feel is necessary (Fig. 2). For each set of data, the CytoScore© creates three files: (1) total count, which shows the total data scored. At the beginning of the file, information about the date the data was collected, the scorer, microscope used, project, and additional microscope settings are reported. Following is the number of sperm analyzed and the number and types of abnormal cells; (2) split summary, which report the same information contained in total count; however, here the data are reported for each set of 500 sperm. Reporting the data in this way may provide information of the quality of the hybridization. In case of homogenous Fig. 2. Examples of Cytoscore© screenshots. (A) Cytoscore© set-up editor showing the menu listing and features that are stored for each sperm that is scored. (B) Cytscore© scoring data window showing the active keys that were selected during the set up of the CytoScore© generator (left column), the total number of sperm scored (center column), and the last 16 sperm that were scored with the most recent at the top (right column). Pressing the 4 key adds a CT8 sperm to the total count. Note that the number of anomalies found is not listed. (C) Cytoscore© anomaly page. Every time the zero key is pressed, this window pops-up and requires the scorer to provide information about the anomaly. In the example presented, we have a CCT8 sperm that was classified as possible because the 2 CC signals differ in size.

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hybridization, the abnormal cells should be divided evenly among the various splits. If this is not the case, it indicates that there may be regions of the slide that are better (or worse) than others. (3) Anomaly data, which report all cells that were called abnormal together with the location of the cell on the slide, plus a set of additional information including comments inserted by the scorer or the person giving a second opinion. The files generated by the CytoScore© software can be exported into Excel spreadsheets for statistical analyses. The first step in this process is to decode the samples and compare the two sets of 5000 sperm from the same slide using a 2 test. If the comparison is good the data become part of the study; if the comparison is not good, i.e., there are major variations between the two sets, the data are discarded and the slide must be rescored. 4.7. Scoring Harmonization Between Scorers, Experiments, and Collaborating Laboratories The goal of scoring harmonization is to establish standard scoring criteria among scorers, studies, and laboratories in order to maximize consistency of analyses and minimize technical variation. The approach of the harmonization process is to: (1) help the new scorer create visualization images on the situations described in the scoring criteria (see Subheadings 4.2. and 4.3.) and (2) conduct a scoring harmonization trial with samples with established frequencies of abnormal sperm. 4.7.1. Scoring Criteria Training The objectives of this training are to familiarize the new scorer with all components of the scoring criteria before observing slides, emphasizing the requirement to consistently use multiple filters, including phase contrast. In addition, the new scorer needs to work side by side with an experienced scorer who presents examples of sperm FISH phenotypes, especially cells that may introduce judgment difficulties, such as background noise, split signals, clusters of spots that remain connected by signal filaments, and cells with signals on the outside of the cell. Key aspects for the trainee to learn are to: 1. Recognize the difference between hybridization artifacts and abnormal cells. 2. Learn to recognize fields that are inappropriate for scoring because of: a. Unhybridized cells or uneven hybridization. b. Excessive clustering and overlapping of cells. c. Debris obscuring cells to be scored. 3. Keep track of areas that have already been scored in order to prevent duplication by documenting the starting and stopping coordinates.

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Active discussion among scorers is also a key aspect of the scoring training. When an atypical cell is observed, both scorers examine the sperm identified as having an abnormal phenotype and discuss any reason for suspecting an abnormality with respect to the established scoring criteria. It is beneficial to show examples of cells that are definitely normal versus normal cells that have various artifacts interfering with the detection of the hybridization signals, including background noise, spilled cells, and split signals. Finally, comparing slides and observing cells described by the other scorers helps to create visual images of the situations that are listed in the scoring criteria. 4.7.2. Scoring Harmonization Plan The objective of the scoring harmonization exercise is to test the new scorer’s ability to score sperm FISH samples using an established scoring criteria and replicate findings obtained by more experienced scorers. This is generally accomplished in three phases. During phase 1, the new scorer and at least one veteran scorer will score 10,000 cells on two reference samples. Scoring is conducted side by side, which each scorer showing to the other every abnormal cell encountered. Discussion of the reasons why each abnormal cell is classified as such is an important aspect of this training and should continue until a consensus is reached. After reaching 5000 sperm, the scorers will exchange slides and score 5000 cells in the same area that was already scored, with continued discussion of every abnormal cell encountered. Once the second set of 5000 sperm is analyzed, comparisons between the two sets of 10,000 sperm from each scorer are made using 2 analyses. If there are no statistically significant differences, the scorers proceed to phase 2; if statistically significant differences are found, the scorers review the scoring criteria and repeat phase 1. Phase 2 will be conducted in the same manner as phase 1, except that this time each scorer will show the other only those cells that he or she believes are questionable. At the end of the scoring, comparisons will be made between the results of the new scorer and those of the veteran scorer. If there are no significant differences, the scorers will proceed to phase 3. If there are significant differences the scorers will revert back to the side-by-side scoring exercise outlined in phase 1 using a new slide. In the last phase of the scoring harmonization plan, both scorers will score a common set of four coded slides. After a set of 5000 sperm on each slide are scored, the slides will be recoded and an additional 5000 cells will be scored. Finally, comparisons will be made between the first and second set of 5000 sperm scored for each slide by each scorer. Comparisons will also be made between the trainee’s data and data previously collected. If there are significant differences the scorers will revert to phase 2. If there are no

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significant differences, the scoring harmonization exercise was successful and the trainee will have generated data that are in line with historic data generated from those samples by experienced scorers and is ready to generate data from new samples. 5. Notes 1. Properly stored samples can be used for FISH for several years after collection. We have obtained good hybridization results from samples that had been stored at –80°C for more than 15 years. Fresh samples should be frozen as soon as possible to prevent sperm from settling at the bottom of the vial. 2. For low-density sperm count donors, a pipet tip can be used to make a spiral smear. The ideal volume is 8–10 μL depending on density observed microscopically after the smear is made. Pipet the appropriate volume, and then place the pipet at the center of the glass slide. From there, the pipet tip is moved in a circular motion while a small volume is pipetted. After the smear has been made, the smear will appear as a spiral similar to a snail’s shell. It is recommended to allow the smears to air-dry for at least 48 h before hybridizing or storing them in the freezer. If low sperm density is still a problem, the semen sample aliquot can be left in the refrigerator for a day or two to allow sperm to settle to the bottom before making smears. 3. If the slides are not immediately hybridized, place the slide box in a plastic bag and then fill the bag with nitrogen air. As much as possible of the oxygenated air should be removed. Seal the bag and place it in a freezer at –20°C. In preparation for hybridization, allow the bag to stand at room temperature for about 30 min before opening it and removing the slides. 4. Whenever a new probe is labeled it should be checked for efficiency and overall quality. A poor probe may be difficult to detect during a study, and it may not be noticed until several slides have been scored. a. Use a control slide to test the probe. Hybridize the probe alone or with other good probes. Some probes may behave differently when combined with others. b. Score at least 500 cells on the slide, looking only at the new probe, and only in regions that are well hybridized. The probe should be present in greater than 99% of the cells. c. Note the quality of the probe. It should be bright with minimal signal splitting and very little or no noise. If the probe is efficient, clean, bright, and tight, then the probe may be used for FISH. 5. Do not take the Klenow out of the freezer until ready to use. Once you have used the Klenow, return it immediately to the freezer. 6. Denature no more than two slides at a time in the 70% formamide–2× SSC solution. Each slide will drop the temperature of the solution about 1–1.5°C.

Detection of Chromosomal Aberrations by FISH 7. 8. 9. 10.

11.

12.

13.

14. 15.

16.

17.

18.

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Prepare fresh every time. Keep at –4°C for at least 30 min before using. On a humid day, 1 h of drying time provides better results. This step can be done while slides are drying. This should be done under yellow light or partial darkness because these stains are sensitive to white light. Also, prewarming cover slips on the warming plate a few minutes before adding the denaturing probes helps maintain the temperature at 37°C during the first few minutes of hybridization. Keep an empty pipet tip box with half an inch of water at the bottom in the incubator at all times so that it will be ready to serve as a humidity chamber any time it is needed. Periodically add water to replace that which has evaporated. The probe mixture for the ACM assay is a combination of directly labeled (A) and indirectly labeled (C and M) probes; therefore detections steps are needed for the C and M probes. Amplification of the C signal is done in two steps with an intermediate washing step with formamide. This step is necessary to reduce nonspecific staining for this probe. If cover slips do not fall off in less than 5 min, then the cover slips were not sealed adequately or there was not enough water in the humidity chamber. Leave cover slip in 2× SSC until cover slips fall off even if it takes longer than 5 min. If signals are not bright enough, this step can be done for 45 min. Small numbers of sperm may be removed from the slide during the denaturation and washing process and remain in the solution. For this reason, denaturing and washing solutions should be changed weekly if heavily used. They should also be changed whenever different treatment sets are hybridized. Equally important is the use of a fluorescence microscope that is equipped with the proper filter sets to detect the light emitted by each fluorochrome. See Table 2 in Pacchierotti and Sgura (this volume). This is especially critical when scoring human samples because of the occasional presence of somatic cells in the semen. The hooked shape of the mouse sperm is the main discriminating factor for identifying sperm from somatic cells. During the scoring of a large number of samples, it is encouraged to include periodically one slide already scored in the set of coded slides given to the scorer. This will help to keep track of the consistency of the scoring criteria used.

Acknowledgments We thank Eddie Sloter, Thomas Schmid, Xiu Lowe, Francesca Pearson, and other past and present laboratory members who helped in the development and validation of the ACM and CT8 assays. This work was performed under the auspices of the U.S. DOE by the University of California, LLNL under contract W-7405-ENG-48 with funding support from NIEHS IAG Y01-ES-8016-5 and from Superfund P42ES04705 from the National Institute of Environmental Health Sciences.

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References 1. Wyrobek, A. J., Schmid, T. E., and Marchetti, F. (2005) Relative susceptibilities of germ cells to genetic defects induced by cancer chemotherapies. J. Natl. Cancer Inst. Monogr. 34, 31–35. 2. Wyrobek, A. J., Schmid, T. S., and Marchetti, F. (2005) Cross-species spermFISH assays for chemical testing and assessing paternal risk for chromosomally abnormal pregnancies. Environ. Mol. Mutagen. 45, 271–283. 3. Marchetti, F., and Wyrobek, A. J. (2005) Mechanisms and consequences of paternally-transmitted chromosomal abnormalities. Birth Defects Res. C Embryo Today 75, 112–129. 4. Shelby, M. D. (1996) Selecting chemicals and assays for assessing mammalian germ cell mutagenicity. Mutat. Res. 325, 159–167. 5. Witt, K. L., and Bishop, J. B. (1996) Mutagenicity of anticancer drugs in mammalian germ cells. Mutat. Res. 355, 209–234. 6. Hassold, T., and Hunt, P. (2001) To err (meiotically) is human: the genesis of human aneuploidy. Nat. Rev. Genet. 2, 280–291. 7. Olshan, A. F., and van Wijngaarden, E. (2003) Paternal occupation and childhood cancer. Adv. Exp. Med. Biol. 518, 147–161. 8. Harkonen, K. (2005) Pesticides and the induction of aneuploidy in human sperm. Cytogenet. Genome Res. 111, 378–383. 9. Crow, J. F. (2001) The origins, patterns and implication of human spontaneous mutations. Nat. Rev. Genet. 1, 40–47. 10. Robbins, W. A., Segraves, R., Pinkel, D., and Wyrobekm, A. J. (1993) Detection of aneuploid human sperm by fluorescence in situ hybridization: evidence for a donor difference in frequency of sperm disomic for chromosomes 1 and Y. Am. J. Hum. Genet. 52, 799–807. 11. Wyrobek, A. J., Alhborn, T., Balhorn, R., Stanker, L., and Pinkel, D. (1990) Fluorescence in situ hybridization to Y chromosomes in decondensed human sperm nuclei. Mol. Reprod. Dev. 27, 200–208. 12. Wyrobek, A. J., and Adler, I. D. (1996) Detection of aneuploidy in human and rodent sperm using FISH and applications of sperm assays of genetic damage in heritable risk evaluation. Mutat. Res. 352, 173–179. 13. Frias, S., Van Hummelen, P., Meistrich, M. L., Lowe, X. R., Hagemeister, F. B., Shelby, M. D., Bishop, J. B., and Wyrobek, A. J. (2003) NOVP chemotherapy for Hodgkin’s disease transiently induces sperm aneuploidies associated with the major clinical aneuploidy syndromes involving chromosomes X, Y, 18, and 21. Cancer Res. 63, 44–51. 14. Rubes, J., Lowe, X., Moore, D., Perreault, S., Slott, V., Everson, D., Selevan, S., and Wyrobek, A. J. (1998) Smoking cigarettes is associated with increased sperm disomy in teenage men. Fertil. Steril. 70, 715–723. 15. Robbins, W. A., Vine, M. F., Truong, K. Y., and Everson, R. B. (1997) Use of fluorescence in situ hybridization (FISH) to assess effects of smoking, caffeine, and alcohol on aneuploidy load in sperm of healthy men. Environ. Mol. Mutagen. 30, 175–183.

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16. Lowe, X., O’Hogan, S., Moore, D., Bishop, J., and Wyrobek, A. (1996) Aneuploid epididymal sperm detected in chromosomally normal and Robertsonian translocation-bearing mice using a new three-chromosome FISH method. Chromosoma 105, 204–210. 17. Lowe, X., de Stoppelaar, J. M., Bishop, J., Cassel, M., Hoebee, B., Moore, D., and Wyrobek, A. J. (1998) Epididymal sperm aneuploidies in three strains of rats detected by multicolor fluorescence in situ hybridization. Environ. Mol. Mutagen. 31, 125–132. 18. Rubes, J., Vozdova, M., and Kubickova, S. (1999) Aneuploidy in pig sperm: multicolor fluorescence in situ hybridization using probes for chromosomes 1, 10, and Y. Cytogenet. Cell Genet. 85, 200–204. 19. Sloter, E., Lowe, X., Moore, D., Nath, J., and Wyrobek, A. J. (2000) Multicolor FISH analysis of chromosomal breaks, duplications, deletions, and numerical abnormalities in the sperm of healthy men. Am. J. Hum. Genet. 67, 862–872. 20. Hill, F. S., Marchetti, F., Liechty, M., Bishop, J., Hozier, J., and Wyrobek, A. J. (2003) A new FISH assay to simultaneously detect structural and numerical chromosomal abnormalities in mouse sperm. Mol. Reprod. Dev. 66, 172–180. 21. Shelby, M. D., Bishop, J. B., Mason, J. M., and Tindall, K. R. (1993) Fertility, reproduction, and genetic disease: studies on the mutagenic effects of environmental agents on mammalian germ cells. Environ. Health Persp. 100, 283–291. 22. Estop, A. M., Marquez, C., Munne, S., Navarro, J., Cieply, K., Van Kirk, V., Martorell, M. R., Benet, J., and Templado, C. (1995) An analysis of human sperm chromosome breakpoints. Am. J. Hum. Genet. 56, 4524–4560. 23. Schmid, T. E., Brinkworth, M. H., Hill, F., Sloter, E., Kamischke, A., Marchetti, F., Nieschlag, E., and Wyrobek, A. J. (2004) Detection of structural and numerical chromosomal abnormalities by ACM-FISH analysis in sperm of oligozoospermic infertility patients. Hum. Reprod. 19, 1395–1400. 24. Sloter, E., Marchetti, F., Eskenazi, B., Weldon, R. H., Nath, J., Cabreros, D., and Wyrobek, A. J. (2006) The frequencies of human sperm carrying chromosomal breaks and segmental duplications and deletions increases with advancing age. Fertil. Steril. 85, 1077–1086. 25. Marchetti, F., Pearson, F. S., Bishop, J. B., and Wyrobek, A. J. (2006) Etoposide induces chromosomal abnormalities in mouse spermatocytes and stem cell spermatogonia. Hum. Reprod. 21, 888–895. 26. Boyle, A. L., and Ward, D. (1992) Isolation and initial characterization of a large repeat sequence element specific to mouse chromosome 8. Genomics 12, 517–525.

III Determination of Species Diversity

15 Assembling DNA Barcodes Analytical Protocols Jeremy R. deWaard, Natalia V. Ivanova, Mehrdad Hajibabaei, and Paul D. N. Hebert

Summary The Barcode of Life initiative represents an ambitious effort to develop an identification system for eukaryotic life based on the analysis of sequence diversity in short, standardized gene regions. Work is furthest advanced for members of the animal kingdom. In this case, a target gene region has been selected (cytochrome c oxidase I) and pilot studies have validated its effectiveness in species discovery and identification. Based on these positive results, there is now a growing effort to both gather barcode records on a large-scale for members of this kingdom and to identify target barcode regions for the other kingdoms of eukaryotes. In this chapter, we detail the protocols involved in the assembly of DNA barcode records for members of the animal kingdom, but many of these approaches are of more general application. Key Words: Biodiversity; COI; cytochrome c oxidase I; DNA barcoding; DNA sequencing; mitochondria; species identification; taxonomy.

1. Introduction The DNA barcoding movement seeks to advance biodiversity science through the development of DNA-based systems that aid species identification and discovery (1–4). In particular, it aims to build these systems in a parsimonious fashion, by basing them, whenever possible, on sequence diversity in a short, standardized gene region. DNA barcoding is a young endeavor whose activation traces to a 2003 publication (1) that revealed the likelihood of developing an effective system for species identifications in the animal kingdom based on From: Methods in Molecular Biology, vol. 410: Environmental Genomics Edited by: C. Cristofre Martin © Humana Press, Totowa, NJ

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sequence variation in a 648 base pair (bp) region of the cytochrome c oxidase I (COI) gene. A number of studies have now validated this approach in varied taxonomic groups and geographic settings. Studies on North American birds have progressed to the point that barcode records are available for more than 95% of the species that breed on this continent (5,6). This work has revealed that DNA barcoding is effective; it provides species-level identifications for 94% of the 693 species analyzed and the few cases of compromised resolution regularly involve species pairs that are known to hybridize. The studies also affirm the value of DNA barcoding in species discovery; flagging 15 overlooked species of birds despite the intensity of prior taxonomic work on this continent. Fishes have also attracted substantial interest and barcode records are now available for more than 1000 species, with the most detailed study revealing 100% success in the identification of 250 Australian fish species (7). Work has also been carried out on a number of invertebrate lineages. Studies on molluscs have largely been restricted to members of a single family in which 94% success in species identification was reported (8). However, arthropods have been examined more intensively with studies on crustaceans (9), spiders (10), collembolans (11), and varied insect lineages including ants (12), mayflies (13), mosquitoes (14), and tachinid flies (15). Work on Lepidoptera has revealed the effectiveness of barcoding in both the discovery and identification of species in hyperdiverse tropical biotas (16,17). As well, wide-ranging studies on more than 1000 North American species of Lepidoptera revealed that regional variation in barcode sequences pose no difficulty for the development of an effective DNA-based identification system (18). The ultimate goal of the DNA barcode movement is the development of comprehensive barcode libraries for all lineages of eukaryotes. While past work has focused on animals, investigations are underway to develop protocols for protists, plants, and fungi. There is evidence that the region of COI targeted in animals may also be effective as a basis for identification systems in algae (19) and fungi (20). In plants, a different genic target will be required, but exploratory efforts are underway (21,22). While protocol development is the primary area of endeavor in other kingdoms of eukaryotes, work on animals is scaling up through major barcode campaigns (23). Some of these campaigns have a taxonomic focus, such as the efforts to gather barcode records for all species of birds and fishes. In other cases, barcode campaigns seek to build a comprehensive barcode library for the biota in a particular region. For example, efforts are underway to assemble barcode records for 10,000 Canadian animal species (10% of fauna) within 5 years. The progress toward the large-scale activation of barcoding is now coordinated by the Consortium for the Barcode of Life, headquartered at the Smithsonian’s National Museum of Natural History in Washington, but representing an alliance of more than 100 international organizations with interests in biodiversity science.

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The growing intensity of barcode research has created the need for simple, inexpensive protocols for barcode analysis (24). This chapter describes the protocols employed in a DNA Barcoding Centre that processes approx 100K specimens a year. However, because the primary considerations for protocol adoption by this facility were simplicity, cost minimization, and speed, these analytical approaches represent a generic solution to barcode acquisition. In this chapter, we consider the eight key steps in the transition from the collection of an organism to the injection of its sequence record into either the Barcode of Life Data System or one of the major genomics repositories. 2. Materials 2.1. Specimens and Tissue Handling 1. 96-Well format 2-D barcoded storage tubes (TrakMates, Matrix Technologies). 2. 99.9% Ethyl alcohol (Commercial Alcohols). Store in a flammable liquids cabinet. 3. Forceps and/or scalpel (FineScience Tools).

2.1.1. Genomic DNA Extraction/Puriﬁcation—Fresh or Frozen Tissue 1. Dry Release extraction buffer: 5–6 g of Chelex-100 (Bio-Rad Laboratories, Hercules, CA), 10 mL of 1% sodium azide (Sigma-Aldrich, St. Louis, MO), 1 mL of Tris-HCl, pH 8.3, ultrapure H2 O to 100 mL. Store at 4°C in 10-mL aliquots. 2. 20 mg/mL of proteinase K (Invitrogen, Carlsbad, CA): 100 mg of proteinase K, 5 mL of ultrapure H2 O. Store in 0.1- to 1-mL aliquots at −20°C. 3. DryRelease working solution: 10 μL of proteinase K, 100 μL of DryRelease extraction buffer. 4. Microplate (Eppendorf, New York, NY). 5. Cap strips (ABgene, Rochester, NY). 6. Thermocycler (Mastercycler EP Gradient, Eppendorf).

2.1.2. Genomic DNA Extraction/Puriﬁcation—Archival Specimens 1. 2. 3. 4. 5. 6. 7.

NucleoSpin 96 Tissue Kit (Macherey-Nagel). 99.9% Ethyl alcohol (Commercial Alcohols). Store in a flammable liquids cabinet. Microplate (Eppendorf). Cap strips (ABgene). Matrix ImpactII P1250 pipettor (Matrix Technologies). Centrifuge with deep-well plate rotor (25R, Beckman Coulter). Incubator (Fisher Scientific).

2.2. Polymerase Chain Reaction (PCR) Ampliﬁcation of the Barcode Region 1. 10% Trehalose: 5 g of d-(+)-trehalose dehydrate (Sigma-Aldrich), ultrapure H2 O to 50 mL. Store at −20°C in 1- to 2-mL aliquots.

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2. 10× PCR Buffer (New England Biolabs). Store at −20°C. 3. 50 mMMgCl2 : 2 mL of 1 MMgCl2 (Sigma-Aldrich), 38 mL of ultrapure H2 O. Store at −20°C in 1-mL aliquots. 4. 10 mM dNTP mix (New England Biolabs). Store at −20°C in 100-μL aliquots. 5. 100 μM primer stock: Dissolve desiccated primer (Invitrogen) in the amount of ultrapure H2 O indicated by the manufacturer to produce a final solution of 100 μM (i.e., add number of nmol × 10 μL of ultrapure H2 O). Store at −20°C. 6. 10 μM primer working solution: 20 μL of 100 μM primer stock, 180 μL of ultrapure H2 O. Store at −20°C. 7. Taq polymerase (New England Biolabs). Store at −20°C in 50-μL aliquots. 8. Microplate (Eppendorf). 9. Cap strips (ABgene). 10. Thermocycler (Mastercycler EP Gradient, Eppendorf).

2.3. PCR Product Check 1. Mother E-base (Invitrogen). 2. 2% E-gel 96 gels (Invitrogen). Store at room temperature prior to opening and at 4°C between reuse. Agarose is encased in glass but does contain the mutagen ethidium bromide; waste should be disposed of according to local regulations. 3. Gel documentation system (AlphaImager 3400, Alpha Innotech Corp.).

2.4. Sequencing Setup 1. BigDye v.3.1. Cycle Sequencing Kit (Applied Biosystems). Store at −20°C. 2. 5× Sequencing buffer (Applied Biosystems). Store at 4°C. 3. 10% Trehalose: 5 g of d-(+)-trehalose dehydrate (Sigma-Aldrich), ultrapure H2 O to 50 mL. Store at −20°C frozen in 1- to 2-mL aliquots. 4. 100 μM primer stock: Dissolve desiccated primer (Invitrogen) in the amount of ultrapure H2 O indicated by the manufacturer to produce a final solution of 100 μM (i.e., add number of nmol × 10 μL of ultrapure H2 O). Store at −20°C. 5. 10 μM primer working solution: 20 μL of 100 μM primer stock, 180 μL of ultrapure H2 O. Store at −20°C. 6. Microplate (Eppendorf). 7. Cap strips (ABgene). 8. Thermocycler (Eppendorf).

2.5. Sequencing Reaction Cleanup 1. Sephadex G-50 (Sigma-Aldrich). Skin exposure or inhalation may be harmful; wearing a face mask and gloves when handling is recommended. 2. AcroPrep 96 0.45 μM GHP filter plates (Pall Corp.). 3. Formamide (Applied Biosystems). Store at −20°C. This reagent is toxic; exposure should be minimized and waste should be disposed of according to local regulations.

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4. Column loader (Millipore). 5. Centrifuge alignment frame (Millipore). 6. 96-Well reaction plates (Applied Biosystems).

2.6. Sequence Analysis 1. 2. 3. 4. 5.

96-Well septa (Applied Biosystems). 96-Well plate bases (Applied Biosystems). 96-Well plate retainers (Applied Biosystems). 3730 Buffer (10×) with EDTA (Applied Biosystems). Store at 4°C. POP-7 polymer (Applied Biosystems). Stable at room temperature for 7–10 days. This reagent is toxic; exposure should be minimized and waste should be disposed of according to local regulations. 6. 48-Capillary (50 cm) array (Applied Biosystems). 7. 3730 DNA Analyzer capillary sequencer (Applied Biosystems).

2.7. Sequence Editing/Alignment 1. Sequence editing software: Sequencher (Gene Codes, Ann Arbor, MI), SeqScape (Applied Biosystems), or Lasergene (DNASTAR, Madison, WI).

3. Methods 3.1. Specimens and Tissue Handling Barcode analysis on most specimens is straightforward, but is highly dependent on the initial condition of the DNA. For this reason, care should be taken to ensure that specimens are killed in a DNA-friendly fashion, analysis should follow collection as quickly as possible (see Note 1), and precautions should be made to prevent contamination with DNA or other specimens (see Note 2). There are six key steps in ensuring the proper handling and documentation of each specimen: 1. Arrange specimens in batches of 94 (see Note 3). Laser-print small labels of unique specimen accession numbers for each of the 94 specimens. If a voucher specimen will be retained, affix a label to it or to the specimen container with this accession number. 2. Clean work surface with ethyl alcohol or a detergent for the removal of DNA and DNase contamination. 3. For each specimen, use acid- or flame-sterilized forceps and/or scalpel to remove a small tissue sample. Place tissue in an individual storage tube of the TrakMates box along with the corresponding specimen accession label. 4. Fill vials with ethyl alcohol for storage at room temperature. For dried tissue (e.g., insect legs) or tissue to be stored below freezing, the addition of ethanol can be omitted.

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5. Record specimen accession numbers and associated data in electronic spreadsheet (see www.barcodeoflife.org for spreadsheet and documentation). 6. Photograph each specimen.

3.2. Genomic DNA Extraction/Puriﬁcation Numerous options are available for the extraction and/or purification of genomic DNA (see Note 4). Several of these methods have been thoroughly tested for their efficacy in high-volume animal DNA barcoding (24); the two methods that demonstrated superior performance for fresh/frozen tissue and for archival tissue, respectively, are outlined in the following two subheadings. 3.2.1. Genomic DNA Extraction/Puriﬁcation—Fresh or Frozen Tissue The Chelex-based DryRelease method (24) (see Note 5) is a quick and cost-effective method of DNA isolation, particularly useful for fresh or frozen material. 1. Aliquot 30–110 μL of DryRelease working solution (see Note 6) into each well of a microplate using a multi-channel pipet, wide-bore tips, and a reservoir. Mix solution before and while aliquoting to ensure that the resin is equally dispersed between wells. Cover each row with cap strips. 2. Put a tiny amount of tissue (e.g., 1–2 mm of insect leg or 1–2 mm3 of ethanolpreserved tissue) into each well of the plate. To prevent cross-contamination, work with one row at a time. Close the lids and prevent shaking to ensure that the tissue fragment remains in the solution. 3. Incubate for 12–24 h at 55°C. 4. Centrifuge the plate at 1000g for 5 min. Incubate samples in a thermocycler at 95°C for 20 min to denature the proteinase K. 5. Store extractions at −20°C. 6. Before PCR set up, centrifuge the plate at 1000g for 5 min. 7. Use 1–2 μL of DNA sample for PCR. Ensure that Chelex resins are not transferred into the PCR reaction.

3.2.2. Genomic DNA Extraction/Puriﬁcation—Archival Specimens The NucleoSpin 96 Tissue Kit (Machery-Nagel) is a silica membrane-based method that performs well with both fresh and archival (older than 5 years) tissue (24) (see Note 7). The eluted DNA is highly purified, making this method ideal for isolating DNA for long-term storage. 1. Add a small amount of tissue (e.g., 2–4 mm of insect leg or 1–3 mm3 of ethanolpreserved tissue) (see Fig. 1) to each well of the round-well block supplied with the kit. Maceration of the tissue is optional, but dividing the tissue into smaller pieces improves the final yield of DNA.

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Fig. 1. Typical specimen sizes used for DNA extraction as compared to the head of a pencil. (A) Moth leg, (B) small crustacean (Daphnia), (C) bird feather, and (D) muscle tissue. (Reproduced from 24. )

2. Prepare a lysis working solution by combining 18 mL of buffer T1 with 2.5 mL of proteinase K in a reservoir. Using a multichannel pipet, transfer 200 μL of the working solution into each well of the round-well block (see Note 8). 3. Seal wells with the cap strips provided and shake vigorously for 10–15 s to mix. Centrifuge at 1500g for 15 s to collect samples at the bottom of the wells. 4. Incubate at 56°C for a minimum of 6 h (ideally 18–24 h) to allow digestion. Tape down cap strips to prevent them from popping off. 5. After digestion, centrifuge at 1500g for 15 s to remove any condensate from the cap strips. 6. Premix ethanol and buffer BQ1: mix 20 mL of ethanol with 20 mL of buffer BQ1 in a reservoir. Using a multichannel pipet, transfer 400 μL of the mixture into each well of the round-well block. Seal wells with cap strips. Shake vigorously for 10–15 s and centrifuge at 1500g for 10 s to remove any sample from the cap strips. 7. Remove cap strips and transfer lysate (about 600 μL) from the wells of the roundwell block into the wells of the tissue binding plate placed on top of a square-well block. Seal plate with a self-adhering PE foil supplied with the kit. 8. Centrifuge at 5600g for 10 min to bind DNA to the silica membrane. 9. Perform the first wash step: Add 500 μL of buffer BW to each well of the tissue binding plate using a reservoir and multichannel pipette. Use a new self-adhering PE foil to seal the plate and centrifuge at 5600g for 2 min. 10. To accommodate the volume of flow through, replace the current square-well block with a new square-well block, placing it underneath the tissue binding plate.

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11. Perform the second wash step: Add 700 μL of buffer B5 to each well of the tissue binding plate using a reservoir and multichannel pipet. Use a new self-adhering PE foil to seal the plate, then centrifuge at 5600g for 4 min. 12. Remove the self-adhering PE foil and place the tissue binding plate on a sterile microplate. Incubate at 70°C for 10 min to evaporate residual ethanol. 13. Dispense 30–100 μL ddH2 0 directly onto the membrane of each well of the tissue binding plate (see Note 9). Incubate at room temperature for 1 min and seal plate. 14. Place both the tissue binding plate and the microplate on an open rack of MN tube strips (see Note 10). Centrifuge at 5600g for 2 min. Rotate the plate/microplate/rack 180° and repeat the centrifugation. Carefully remove the tissue binding plate and rack. Seal the microplate with cap strips. 15. Keep DNA at 4°C for temporary storage or at −20°C for long-term storage. Use 1–2 μL of the DNA sample for PCR amplification.

3.3. PCR Ampliﬁcation of the Barcode Region Barcode markers were chosen in part for their ease of isolation, satisfying such criteria as high copy number and the presence of conserved flanking regions for primers. PCR amplification of these regions is therefore routine with compliant samples and well-designed primers (see Note 11). For recalcitrant cases, success is often achieved by targeting smaller fragments or making small modifications to the primer sequence. 1. Defrost your reagents and place in a cold block. 2. Mix reagents in a 1.5-mL tube following the recipe given in Table 1 (see Notes 12–14). Additives (see Note 15) or alternative enzymes (see Note 16) may increase the success, yield, and accuracy of the PCR. A list of primers to amplify the COI barcode region for a variety of taxonomic groups is given in Table 2. 3. Vortex-mix gently and aliquot 10.5 μL of the PCR mix into each well of the microplate. Time can be saved by aliquoting 1/8 (˜138 μL) of the total mix into an eight-tube PCR strip and dispensing to the microplate with a multichannel pipet. 4. Add 0.5–2 μL of DNA extract (see Notes 17 and 18) to each well using a multichannel pipet. 5. Seal the plate with cap strips. 6. Centrifuge at 1000g for 10 s. 7. Place in a thermal cycler (see Note 19) and select program (see Note 20); an example program is given in Fig. 2.

3.4. PCR Product Check For projects that are examining compliant samples, it is possible to proceed directly from the barcode PCR to a sequencing reaction. However, it is often critical to screen PCR reactions for successful amplification products when

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Table 1 Master-Mix Recipes for PCR Ampliﬁcation of the Barcode Region (in μL) Reagent 10% Trehalose ddH2 O 10× PCR buffer 50 mMMgCl2 10 mM dNTP 10 μM Primer 1 10 μM Primer 2 Taq polymerase Mix volume DNA template Total volume

1 Reaction

Plate

6.25 2 1.25 0.625 0.0625 0.125 0.125 0.0625 10.5 2 12.5

650 208 130 65 6.5 13 13 6.5 -

working with older specimens or with a new taxonomic group. This has traditionally been a laborious task involving gel casting and the loading of individual reaction products. The use of bufferless, precast agarose gels circumvents these and other problems (see Note 21). 1. Plug the Mother E-Base into an electrical outlet. Press and release the “pwg/prg” (power/program) button on the base to select program EG. Select a run time by pressing the “time” button (see Note 22).

Fig. 2. A sample thermocycler program for PCR amplification of the COI barcode region.

Sequence (5 -3 )

GGTCAACAAATCATAAAGATATTGG TAAACTTCAGGGTGACCAAAAAATCA TTCTCCAACCACAAAGACATTGGCAC ACGTGGGAGATAATTCCAAATCCTGG TCAACCAACCACAAAGACATTGGCAC TAGACTTCTGGGTGGCCAAAGAATCA TCGACTAATCATAAAGATATCGGCAC ACTTCAGGGTGACCGAAGAATCAGAA TTCTCAACCAACCACAAAGACATTGG TTCTCAACCAACCACAARGAYATYGG TTCTCAACCAACCAIAAIGAIATIGG TAGACTTCTGGGTGGCCAAAGAATCA TAGACTTCTGGGTGGCCRAARAAYCA TAGACTTCTGGGTGICCIAAIAAICA

ATTCAACCAATCATAAAGATATTGG TAAACTTCTGGATGTCCAAAAAATCA

TTTTCTACAAATCATAAAGACATTGG GGTTCTTCTCCACCAACCACAARGAYATHGG TAAACTTCAGGGTGACCAAAAAATCA TACTCTACTAATCATAAAGACATTGG CCTCCTCCTGAAGGGTCAAAAAATGA GGATGGCCAAAAAATCAAAATAAATG

Primer name

LCO1490 HCO2198 BirdF1 BirdR1 FishF1 FishR1 FishF2 FishR2 VF1 VF1d VF1i VR1 VR1d VR1i

LepF1 LepR1

CrustF1 CrustF2 HCO2198 Chel F1 Chel R1 Chel R2

Table 2 Primers for the PCR Ampliﬁcation of COI for Varied Taxonomic Groups

7

7, 33

Fishes

Mammals, reptiles, Amphibians

9, 32

10

Crustaceans

Chelicerates

17

5

Birds

Insects

32

Reference

Misc. Phyla

Taxonomic group

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2. Remove the gel from the package and remove the plastic comb from the gel. Slide the gel into the two electrode connections on the Mother E-Base. 3. Load 16 μL of ddH2 O with a multichannel pipettor. 4. Load appropriate DNA markers in the marker wells if necessary. 5. Load 4 μL of the sample with a multichannel pipettor. 6. To begin electrophoresis, press the “pwd/prg” button. The red light should change to green. 7. At the end of run (signaled with a flashing red light and rapid beeping), press and release the “pwr/prg” button. 8. Remove the gel cassette from the base and capture a digital image of the gel with a gel documentation system. 9. Arrange lanes and manipulate image as necessary using the Invitrogen E-Editor software (available at www.invitrogen.com/egels). 10. Incorporate the E-gel image into an “electronic lab book” spreadsheet for hit picking (see Note 23). (See www.barcodeoflife.org for lab spreadsheets and documentation.) 11. Gels can be stored at 4°C for at least one reuse.

3.5. Sequencing Setup Cycle sequencing reactions are set up directly from PCR products (see Note 24), minimized to cut costs (see Note 25), and can be premixed and stored frozen for convenience and quality control measures (see Note 26). 1. Defrost your reagents and place in a cold block. 2. Mix reagents in a 1.5-mL tube following the recipe given in Table 3 (see Note 27). Prepare a mix for both the forward and reverse reaction of each sample. 3. Vortex-mix gently and aliquot 9 μL of the sequencing mix into each well of the microplate. This can be done quickly by aliquoting 1/8 (∼117 μL) of the total mix into an eight-tube PCR strip and dispensing to the microplate with a multichannel pipet. Table 3 Master-Mix Recipes for Sequencing Reaction Setup (in μL) Reagent Dye terminator mix v3.1 5× Sequencing buffer 10% Trehalose 10 μM Primer ddH2 O Mix volume PCR product Total volume

1 Reaction

Plate

0.25 1.875 5 1 0.875 9 1 10

26 195 520 104 91 — — —

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4. Add 0.5–2 μL of PCR product (see Note 28) to each well using a multichannel pipet. 5. Seal the plate with cap strips. 6. Centrifuge at 1000g for 10 s. 7. Place in a thermal cycler and select the program (see Note 29); a sample program is given in Fig. 3.

3.6. Sequencing Reaction Cleanup The Sephadex column method of cycle sequencing reaction cleanup is a rapid, reliable, and cost-effective alternative to both traditional and newer cleanup methods (see Note 30). 1. Use the column loader to fill each well of the filter plate with Sephadex. 2. Hydrate the wells with 300 μL of ultrapure H2 O. 3. Let the Sephadex swell before use, either overnight in the refrigerator or for 3–4 h at room temperature. 4. Drain the excess water by using the centrifuge alignment frame to attach the Sephadex filter plate to an empty microplate. Rubber bands can be used to hold the plates in place. Centrifuge at 750g for 3 min and discard the water. 5. Using a multichannel pipet, add the sequencing reactions (∼10 μL) to the center of the Sephadex columns in each well. 6. Using a multichannel pipet and an eight-tube PCR strip, add 10 μL of formamide to each well of a sterile 96-well reaction plate. 7. To elute the purified sequencing reactions into the formamide, attach the reaction plate to the bottom of the Sephadex filter plate, securing it with rubber bands. Centrifuge at 750g for 3 min.

Fig. 3. Thermocycler program for sequencing reactions.

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3.7. Sequence Analysis The high production goals of a DNA barcoding facility make a multiple capillary instrument essential. Applied Biosystems and Amersham produce several highly reliable instruments with varied production capacities (see Note 31). The following protocol refers to an Applied Biosystems 3730 capillary sequencer. 1. 2. 3. 4. 5. 6.

Cover the reaction plate with septa. Place the reaction plate into the plate base and attach the plate retainer. Print and affix a barcode to the assembled plate. Stack assembled plate in the 3730. Perform routine 3730 maintenance as necessary. Using the Plate Manager of the Data Collection software (Applied Biosystems), import the plate record(s) for the plate being run. 7. Begin the run within Run Scheduler.

3.8. Sequence Editing/Alignment The minimum capabilities of a sequence editing and alignment software package for high-volume barcoding are the abilities to assemble bidirectional reads and edit trace files. Sequencher, SeqScape, and Lasergene are capable, plus provide other useful functions. 1. Using the sequence alignment editor of choice, open up the trace files of the finished run. Depending on the software being used, as meant samples can be analyzed at once. 2. Assemble the forward and reverse read of each sample. 3. Remove the primer sequences from the end of the read. 4. Review each base call, making adjustments to miscalled or uncalled bases as necessary. 5. Save sequence(s) in Fasta format and upload to the appropriate project in BOLD or to another online sequence repository.

4. Notes 1. Killing in a DNA-friendly fashion refers to freezing, cyanide exposure, or immersion in ethanol, and avoiding even brief exposure to killing/preservation agents such as ethyl acetate or formalin that damage DNA. DNA in dried specimens ordinarily remains in good condition for at least a year, but degradation becomes increasingly problematic as time passes. DNA in frozen specimens (especially those held in cryogenic conditions) remains stable indefinitely, but DNA in ethanolpreserved material often degrades as a result of acidification. As a result, barcode analysis should follow collection as quickly as possible. 2. All samples should be handled on a clean working surface and all tissue-handling instruments should be acid or flame sterilized between each sample. A Bunsen

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burner flame is convenient for sterilization; small propane tanks are ideal for settings where gas is not online. In any laboratory that seeks high production rates, it is critical to carry out all stages of barcode analysis in 96-well plates. Two wells should be left blank for positive and negative controls in the preceding steps, leaving 94 wells for samples. Care must be taken to avoid cross-contamination between wells when loading these plates with samples. In addition to the Chelex and membrane-based methods discussed here, there are many alternate kits and protocols for DNA release or isolation. The use of magnetic beads is becoming increasingly popular, primarily because of their amenability to automation. When speed is critical, several kits offer DNA extraction in minutes, such as the Extract-N-Amp PCR Kit (Sigma-Aldrich) and FTA cards (Whatman, Florham Park, NJ). These methods are currently being evaluated for their value in high-throughput DNA barcoding. DryRelease is a Chelex-based, DNA release protocol that rapidly liberates DNA into solution, making it accessible for downstream applications (24). It requires minute tissue samples and minimal technician time, but is not suitable for samples with high levels of PCR inhibitors (e.g., hemoglobin), for samples in which DNA is degraded, or where pure DNA for long-term storage is required. Working volumes can range from 30 to 110 μL depending on sample size. For example, small crustaceans or legs of small insects should be extracted in 30 μL, whereas small blocks of vertebrate tissue can be extracted in 100–110 μL. The membrane-based DNA extraction method of the Nucleospin 96 Tissue kit relies on DNA binding to the silica membrane in the presence of a high concentration of chaotropic salt. It results in highly pure DNA and is exceptionally sensitive (25), making it useful for studies on specimens with degraded DNA (24). The higher cost and large demands on technician time of this and similar methods can limit its utility. A manual or electronic multichannel pipet is required to effectively perform 96well DNA extractions with this and most other kits. Similarly, nearly all steps of 96-well kits and protocols can be automated on robotic liquid handling stations if available. As with DryRelease, elution for smaller specimens (or specimens where DNA is likely degraded) should be done with a small volume of water, whereas larger volumes should be employed for larger or fresh specimens. Centrifuging the plates on a square-well block or an open rack of MN tube strips prevents the wells of the microplate from being crushed or cracked by the force. Primer design is critical and minor adjustments can have large impacts on barcode recovery. The first phase of any study on a new group should involve a serious effort to identify optimal primers. To address mismatches between primers and target DNA, the use of degenerate or inosine containing primers is recommended (26). Primers with two to four degenerate positions or inosine bases will often rescue barcodes from recalcitrant specimens and may also protect from amplifying nuclear pseudogenes (27).

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12. The use of sterile barrier tips is recommended for all PCR reagents to avoid contamination. Clean the bench top with alcohol or detergent before setting up reactions. DNA templates (DNA extracts or PCR products) should be kept away from the PCR reagents while you are setting up the reaction mixes. Add DNA only after all of the reagents have been returned to the freezer. 13. The addition of trehalose is optional but may enhance PCR success (28) (see Fig. 4) and makes it possible to freeze aliquoted master mixes (29). Master mix can be stored in tubes at −20°C for 1–3 months or aliquoted directly into 96-well plates at −20°C for up to 1 month. 14. To reduce costs, reaction volumes can be significantly lowered. To dispense such small volumes accurately, it is useful to make up enough master mix for several plates. Note that it is necessary to include extra volume to allow for pipetting mistakes and dead volume in the multichannel pipet (e.g., to make ten 96-well plates with 12.5-μL reactions per well, include about 40 extra reactions). 15. The presence of PCR inhibitors can usually be overcome by incorporating amplification facilitators such as trehalose, bovine serum albumin (BSA), betaine, or dimethyl sulfoxide in the PCR mix (28,30,31). 16. There is a growing diversity of polymerases or polymerase cocktails. Some enable PCR to be executed much more quickly (e.g., Z Taq, Takara Bio, Otsu, Japan); others aid the amplification of damaged templates (Restorase, Sigma-Aldrich) or permit high-fidelity replication (Diamond Taq, Bioline, Randolph, MA). 17. The amount of DNA extract used will depend on the specimen and extraction method employed. Although this may take some adjustment, it is not usually

Fig. 4. Demonstration of PCR-enhancing ability of the additive trehalose. A dark band on the E-gel images (negative exposure) indicates a successful PCR amplification for that sample; the clear slots indicate the loading wells. A12 and B12 are negative controls; column M contains size markers. (A) Regular PCR master mix without trehalose, and (B) PCR master mix with 5% trehalose.

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necessary to quantify genomic DNA extracts because even a few copies of the target gene are sufficient for PCR amplification. It is best to keep the volume of DNA template as low as possible to avoid adding reaction inhibitors that may be present, and to avoid unspecific amplification. Always include a sample without template as a negative control to check for contamination of the reagents. Also include a positive control (a DNA sample that has amplified in the past) to test the effectiveness of the PCR reagents. The latest generation of thermal cyclers (e.g., Eppendorf MasterCycler EP Silver) have faster thermal ramping that allows PCR amplifications to be completed more quickly (1–2 vs. 3–4 h). The annealing temperature for a PCR reaction is generally the only variable that needs to be changed when using new primers. Setting the annealing temperature at 2–6°C lower than the melting temperature of your primers is a general rule of thumb. The Invitrogen E-gel 96 system is quick, sensitive, consistent, and minimizes exposure to ethidium bromide. The capital cost is low, the gels are moderately priced, and the technical time is negligible. Bio-Rad and Amersham Biosciences have similar products. For an approx 700-bp amplicon, it takes only 6 min to fully resolve the band. The use of electronic lab books with incorporated E-gel images facilitates the preparation for hit picking of successful PCR reactions. This is a difficult and time-consuming task; a liquid handling system should be used if available. PCR products are often purified to remove unincorporated nucleotides and residual primers. If this step is omitted, it leads to degradation in the sequencing results for the first 20–50 bp. Such degradation is of little concern when the PCR product is slated for bi-directional sequencing. However, when the PCR product is sequenced in just a single direction, several protocols (e.g., ethanol precipitation) and numerous kits are available such as MultiScreen Filter Technology (Milllipore). To reduce costs, sequencing reactions can be cut to 10 μL and prepared with just 0.25 μL of BigDye (1/16 concentration). Because BigDye is one of the costliest reagents in the barcoding protocol, lowering its usage is a critical step in minimizing costs. Sequencing reaction cocktails can be prepared in either 96-well plates or in larger volume tubes well in advance of use, then frozen for up to 3 months. Trehalose is added to ensure stability of the enzyme during freeze–thaw cycles. BigDye v.3.1. cycle sequencing chemistry provides a robust sequencing platform, consistently producing long (∼750 bp), high-quality reads, even on GC-rich templates. Amersham Biosciences provides a creditable alternative with the DYEnamic ET Terminator cycle sequencing kit that is fully compatible with Applied Biosystems instruments. The volume of PCR template to add should be estimated based on the intensity of bands on the PCR check gel.

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29. The annealing temperature of the program may be varied according to the primer specificity, but 55°C works well for most COI barcode sequencing reactions. 30. Many high-volume genomics facilities use either ethanol precipitation or magnetic bead protocols for sequencing reaction cleanup. Solid-phase reversible immobilization (SPRI) based methods may be particularly well suited to high-throughput barcoding labs. 31. The Applied Biosystems multicapillary sequencer models include the 3100, 3130, 3730, and 3730XL with 4, 16, 48, and 96 capillaries, respectively. Similarly, the Amersham Biosciences line includes the MegaBASE 500, 1000, and 4000 with up to 48, 96, and 384 capillaries, respectively.

Acknowledgments We thank Alex Borisenko, Rob Dooh, Teresa Crease, Robert Hanner, Angela Holliss, Stephanie Kirk, Paula Mackie, Pia Marquart, Erin Penton, Keith Pickthorn, Cadhla Ramsden, Sujeevan Ratnasingham, Alex Smith, Janet Topan, Taika von Konigslow, Adam Yule, and Tyler Zemlak for their aid in protocol development. Grants from the Gordon and Betty Moore Foundation, NSERC, the Canada Foundation for Innovation and the Ontario Innovation Trust aided preparation of this chapter. References 1. Hebert, P. D. N., Cywinska, A., Ball, S. L., and deWaard, J. R. (2003) Biological identifications through DNA barcodes. Proc. R. Soc. B 270, 313–322. 2. Blaxter, M. (2003) Counting angels with DNA. Nature 421, 122–124. 3. Hebert, P. D. N., and Gregory, T. R. (2005) The promise of DNA barcoding for taxonomy. Syst. Biol. 54, 852–859. 4. Savolainen V., Cowan, R. S., Vogler, A. P., Roderick, G. K., and Lane, R. (2005) Towards writing the encyclopaedia of life: an introduction to DNA barcoding. Philos. Trans. R. Soc. B 360, 1805–1811. 5. Hebert, P. D. N., Stoeckle, M. Y., Zemlak, T. S., and Francis, C.M. (2004) Identification of birds through DNA barcodes. PLoS Biol. 2, 1657–1663. 6. Kerr, K. A. , Stoeckle, M. Y. , Dove, C, Weigt, L. A. , Francis, C. M., and Hebert, P. D. N. (2007) Comprehensive DNA barcode coverage of North American birds. Mol. Ecol. Notes. 7, 535–543. 7. Ward, R. D., Zemlak, T. S., Innes, B. H., Last, P. R., and Hebert, P. D. N. (2005) DNA barcoding Australia’s fish species. Philos. Trans. R. Soc. B 360, 1847–1857. 8. Meyer C. P., and Paulay, G. (2005) DNA barcoding: Error rates based on comprehensive sampling. PLoS Biol. 3, 2229–2238. 9. Costa, F. O., deWaard, J. R., Boutillier, J., Ratnasingham, S., Dooh, R. T., Hajibabaei, M., and Hebert, P. D. N. (2006) Biological identifications through DNA barcodes: the case of the Crustacea. Can. J. Fish. Aquat. Sci. 64, 272–295.

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10. Barrett, R. D. H., and Hebert, P. D. N. (2005) Identifying arachnids through DNA sequences. Can. J. Zool. 83, 481–491. 11. Hogg, I. D., and Hebert, P. D. N. (2004) Biological identification of springtails (Collembola: Hexapoda) from the Canadian Arctic, using mitochondrial DNA. Can. J. Zool. 82, 749–754. 12. Smith, M. A., Fisher, B. L., and Hebert, P. D. N. (2005) DNA barcoding for effective biodiversity assessment of a hyperdiverse arthropod group: the ants of Madagascar. Philos. Trans. R. Soc. B 360, 1825–1834. 13. Ball, S. L., Hebert, P. D. N., Burian, S. K., and Webb, J. M. (2005) Biological identifications of mayflies (Ephemeroptera) using DNA barcodes. JNABS 24, 508–524. 14. Cywinska, A., Hunter, F., and Hebert, P. D. N. (2006) Identifying Canadian mosquito species through DNA barcodes. Med. Vet. Entomal. 20, 413–424. 15. Smith, M. A., Woodley, N. E., Janzen, D. H., Hallwachs, W., and Hebert, P. D. N. (2006) DNA barcodes reveal cryptic host-specificity within the presumed polyphagous members of a genus of parasitoid flies (Diptera: Tachinidae). Proc. Natl. Acad. Sci. USA 103, 3657–3662. 16. Hebert, P. D. N., Penton, E. H., Burns, J. M., Janzen, D. H., and Hallwachs, W. (2004) Ten species in one: DNA barcoding reveals cryptic species in the neotropical skipper butterfly Astraptes fulgerator. Proc. Natl. Acad. Sci. USA 101, 14812–14817. 17. Hajibabaei, M., Janzen, D. H., Burns, J. M., Hallwachs, W., and Hebert, P. D. N. (2006) DNA barcodes distinguish species of tropical Lepidoptera. Proc. Natl. Acad. Sci. USA 103, 968–971. 18. Hebert, P. D. N., deWaard, J. R., and Landry, J-F. (2007) DNA barcodes deliver: species identifications and revelations for 1/1000 of the Animal Kingdom. In preparation. 19. Saunders, G. W. (2005) Applying DNA barcoding to red macroalgae: a preliminary appraisal holds promise for future applications. Philos. Trans. R. Soc. B 360, 1879–1888. 20. Seifert, K. A., Sampson, R. A., deWaard, J. R., Houbracken, J. A., Levesque, C. A., Montcalvo, J.-M., Louis-Seize, G. and Hebert, P. D. N. (2007) Prospects for fungus identification using CO1 DNA barcodes, with Penicillium as a test case. Proc. Natl. Acad. Sci. USA 104, 3901–3906. 21. Kress, J. W., Wurdack, K. J., Zimmer, E. A., Weigt, L. A., and Janzen, D. H. (2005) Use of DNA barcodes to identify flowering plants. Proc. Natl. Acad. Sci. USA 102, 8369–8374. 22. Chase, M. W., Salamin, N., Wilkinson, M., Dunwell, J. M., Kesanakurthi, R. P., Haidar, N., and Savolainen, V. (2005) Land plants and DNA barcodes: short-term and long-term goals. Philos. Trans. R. Soc. B 360, 1889–1895. 23. Marshall, E. (2005) Will DNA barcodes breathe life into classification? Science 307, 1037. 24. Hajibabaei, M., de Waard, J. R., Ivanova, N. V., Ratnasingham, S., Dooh, R. T., Kirk, S. L., Mackie, P. M., and Hebert, P. D. N. (2005) Critical factors for

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assembling a high volume of DNA barcodes. Philos. Trans. R. Soc. B 360, 1959–1967. Hoss, M., and Paabo, S. (1993) DNA extraction from Pleistocene bones by a silica-based purification method. Nucleic Acids Res. 21, 3913–3914. Batzer, M. A., Carlton, J. E., and Deininger, P. L. (1991) Enhanced evolutionary PCR using oligonucleotides with inosine at the 3?-terminus. Nucleic Acids Res. 19, 5081. Sorenson, M. D., Ast, J. C., Dimcheff, D. E., Yuri, T., and Mindell, D. P. (1999) Primers for a PCR-based approach to mitochondrial genome sequencing in birds and other vertebrates. Mol. Phylogenet. Evol. 12, 105–114. Spiess, A. N., Mueller, N., and Ivell, R. (2004) Trehalose is a potent PCR enhancer: lowering of DNA melting temperature and thermal stabilization of Taq polymerase by the disaccharide trehalose. Clin. Chem. 50, 1256–1259. Franks, F. (1990) Freeze drying: from empiricism to predictability. Cryoletters 11, 93–110. Al-Soud, W. A., and Rådström, P. (2000) Effects of amplification facilitators on diagnostic PCR in the presence of blood, feces, and meat. J. Clin. Microbiol. 38, 4463–4470. Frankman S., Kobs G., Simpson D., and Storts D. (1998) Betaine and DMSO: enhancing agents for PCR. Promega Notes 65, 27. Folmer, O., Black, M., Hoeh, W., Lutz, R., and Vrijenhoek, R. (1994) DNA primers for amplification of mitochondrial cytochrome c oxidase subunit I from diverse metazoan invertebrates. Mol. Mar. Biol. Biotechnol. 3, 294–299. Ivanova, N. V., deWaard, J. R., and Hebert, P. D. N. (2006) An inexpensive, automation-friendly protocol for recovering high-quality DNA. Mol. Ecol. Notes. 6, 998–1002.

16 Application of Suppressive Subtractive Hybridization to Uncover the Metagenomic Diversity of Environmental Samples Elizabeth A. Galbraith, Dionysios A. Antonopoulos, and Bryan A. White

Summary Metagenomics addresses the collective genetic structure and functional composition of a microbial environmental sample without the bias or necessity for culturing the microorganisms from the community in question. Metagenomic studies are now beginning to take advantage of the plethora of complete genome sequences (1,2) and the associated tools, such as bacterial artificial chromosome (BAC) and fosmid vectors, to discover novel genes and survey the structure and function of microbial communities. Complementary and less expensive methods to compare genomes from individual microbes have been utilized in comparative genomic studies. Suppressive subtractive hybridization (SSH) is one such approach, which has been utilized to compare the genomic content of closely related species of bacteria (3–6). Recently, SSH has also been used as a comparative method to examine the microbial diversity (i.e., species composition) and functional differences (i.e., gene composition) in the genomic content of two different rumen environmental communities (7). Through a series of hybridizations and polymerase chain reaction (PCR) amplifications, metagenomic differences between two environmental samples can be isolated by SSH. Subsequent DNA sequencing and bioinformatic analyses allow the putative identification of these differences. Key Words: Metagenomics; subtractive hybridization.

1. Introduction Metagenomics addresses the collective genetic structure and functional composition of a microbial environmental sample without the bias or necessity for culturing the microorganisms from the community in question. An extensive From: Methods in Molecular Biology, vol. 410: Environmental Genomics Edited by: C. Cristofre Martin © Humana Press, Totowa, NJ

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suite of molecular based approaches developed over the past decade has enabled the study of these uncultured microorganisms and is the direct outcome of the use of small subunit ribosomal RNA (SSU rRNA) targets which have been widely utilized to study the bacterial diversity, community structure, and microbial interactions within these microbial ecosystems (8–10). Although these methods provide a wealth of information about microbial community structure, they are limited to a subset of identifying sequences. Thus, metagenomic studies are now beginning to take advantage of the plethora of complete genome sequences (1,2) and the associated tools to discover novel genes and survey the structure and function of microbial communities. Community genome complexity and size have been estimated by analyzing the reassociation of genomic DNA from environmental samples (11,12). However, construction of metagenomic libraries using bacterial artificial chromosome (BAC) and fosmid vectors (13,14) is a more direct route to gaining insight into the functional nature of mixed microbial samples. The sequencing and assembly of these large insert libraries have also been hypothesized to lead to reconstruction of a nearly complete microbial genome (15). Complementary and less expensive methods to compare genomes from individual microbes have been utilized in comparative genomic studies. Suppressive subtractive hybridization (SSH) is one such approach, which has been utilized to compare the genomic content of closely related species of bacteria (3–6). Recently, SSH has also been used as a comparative method to examine the microbial diversity (i.e., species composition) and functional differences (i.e., gene composition) in the genomic content of two different rumen environmental communities (7). The core concept of subtractive hybridization was first documented in a publication by Bautz and Reilly (1966) (16). Here, the authors demonstrated that a bacteriophage T4 deletion mutant identified by phenotypic methods could be utilized to isolate mRNA corresponding to the specific deleted gene from the wild-type counterpart without extensive knowledge of the gene and surrounding sequences. Denatured (single-stranded) DNA from the deletion mutant was bound to a nitrocellulose column and used as a template to bind mRNA from the wild type. The mRNA in common with the wild-type and deletion mutant bound its DNA-based template and adhered to the column while the mRNA species corresponding to the deleted (missing) DNA target of interest emerged from the column, resulting in an effective subtraction. Approximately 20 years later this basic idea was systematically applied to isolating differences in genomic DNA from eukaryotes, whose genomes are several orders of magnitude larger than that of a bacteriophage. A series of three papers (17–19) documented the identification of specific genomic DNA regions using an approach modified from that of Bautz and Reilly, and introduced the term “genomic subtraction” (as well as “difference cloning”) to describe the

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process. By 1990, the technique was formalized by the incorporation of three crucial concepts: (1) the introduction of the “tester” and “driver” nomenclature; (2) the use of an excess amount of “driver” DNA to “drive” the subtraction; and (3) the incorporation of polymerase chain reaction (PCR) to amplify the remaining “tester”-specific fragments following the subtraction process. Two papers demonstrated the utility of this streamlined approach (20,21). Straus and Ausubel (1990) applied “genomic subtraction” to the isolation of wildtype DNA that was absent in a yeast deletion mutant, while Wieland et al. (1990) utilized a similar “subtractive hybridization” procedure to compare phage DNA and placental DNA, as well as placental DNA compared with DNA from Epstein–Barr virus-immortalized lymphocytes. Both publications involved the use of an avidin–biotin-mediated approach to isolating DNA fractions. Southern blot analyses illustrated the success of both approaches, and indicated the power of the technique relative to the investment of time and finances. However, the widest use for subtractive hybridization at the time was in the normalization of cDNA libraries (22,23). Because cDNA libraries are based on extracted mRNA from a specific eukaryotic tissue, certain mRNA species may be under-represented in one tissue type versus another. Subtractive hybridization would allow the recovery of these low-expression mRNA species to produce a full representation of the possible mRNA species (24–26). An earnest description of the subtractive hybridization process utilizing computer-based modeling was soon developed (27–29). This line of research produced not only a series of computer utilities to predict the ideal parameters for a successful experiment but also a further refinement of the subtractive hybridization procedure—which came to be called suppressive subtractive hybridization (26). Suppressive subtractive hybridization (SSH; other descriptions in the literature also refer to it as “suppression subtractive hybridization”) introduced the use of specially designed adaptors (short double-stranded DNA segments). If these adaptors are ligated to themselves or to small DNA fragments then the secondary structure (a “panhandle”-like structure) introduced by their sequence content would effectively inhibit the PCR amplification in the final preparation (Fig. 1). This would efficiently decrease the detection of background fragments and allow the identification of larger, differential DNA of sufficient “informational” length. The amended version of subtractive hybridization was demonstrated in a comparison of the genomes of two strains of Helicobacter pylori (3). A patented kit, produced by Clontech, is based on this publication and provides a convenient collection of the adaptors and enzymes to successfully construct SSH libraries. Variations of the subtractive hybridization approach to comparative microbial genomics have been used to described specific strain differences for

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Fig. 1. An overview of the suppressive subtractive hybridization procedure. Note the inhibitory “pan-handle” secondary structure (“b -> b”’ bottom panel) formed in the

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many microbial species (6,30–40). The streamlining of genome sequencing methodologies and concomitant decrease in economic cost has allowed the adoption of the technique to screen specific strains relative to a known fully sequenced one. The most thorough description of this approach was described by Nesbo et al. (2002) in studying strains of Thermotoga (5). They followed up this work with screening of phage libraries with the SSH products recovered in the aforementioned paper in order to obtain large flanking regions for various Thermotoga strains (41). More recently Antonopoulos et al. (2004) have described the application of the technique in parallel with the sequencing of the R. flavefaciens FD-1 genome, emphasizing that the technique does not require full sequence data to be effective (42). Assembled contigs from a draft genome sequence (2× coverage) provide large sequence scaffolds that SSH products can be assembled with and demonstrated the clustering of SSH products to specific genomic regions. Subtractive hybridization can also be effectively used to direct the closure of a draft genome sequence, as described for Xylella fastidiosa—yet another extension of the technique (43). Recently, we have pioneered the application of the technique to compare metagenomes of microbial consortia from rumen samples (7). This study utilized specific rumen samples for which our group had amassed phylogenetic data from denaturing gradient gel electrophoresis and SSU rDNA libraries (44). Moreover, rumen samples were chosen because the Bacterial and Archaeal diversity and functionality within the microbial community of the rumen had long been the foci of rumen microbiologists (45,46) and there was a considerable knowledge base from both classical culture-based methods as well as current molecular approaches (47,48). Therefore, for this study, SSH was employed to isolate DNA fragments present in one rumen sample, but absent from another. DNA of these subtracted samples was cloned, and the DNA sequences determined and analyzed via computer-assisted DNA and amino acid comparisons. Subtractive hybridization fragments of both Bacterial and Archaeal origin were identified, demonstrating that SSH provided a powerful complementary approach to current methods for the metagenomic analysis of complex microbial environments. Based on simple quality control assessments of the SSH libraries constructed, the technique appears capable of discerning and identifying DNA fragments unique to one complex microbial community Fig. 1. final PCR amplification; this greatly decreases the recovery of “background noise” (i.e., short non-specific sequences). (Reproduced from the manual for CLONTECH’s PCR-Select Bacterial Genome Subtraction Kit. Courtesy of Clontech Laboratories, Inc., a Takara Bio Co., [2006] Clontech Laboratories Inc.).

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relative to another. In this chapter, detailed methods for performing SSH with environmental microbial consortia are presented, including quality control experiments to determine subtraction efficiency. 2. Materials 2.1. Extraction of Total Genomic DNA from Environmental Samples 1. 0.1 M sodium phosphate buffer, pH 8.0: to make 200 mL, mix 5.3 mL of solution A (27.6 g of NaHPO4 · H2 O/L [0.2 M]) with 94.7 mL of solution B (53.65 g of Na2 HPO4 · 7H2 O/L [0.2 M]) and then dilute with H2 O to 200 mL. 2. Lysis solution: 0.15 M NaCl, 0.1 M EDTA, pH 8.0. Autoclave and then add 15 mg/mL of lysozyme. Store at 4°C. 3. SDS solution: 0.1 M NaCl, 0.5 M Tris-HCl, pH 8.0, 10% sodium dodecyl sulfate. 4. Proteinase K (Sigma-Aldrich). 5. Buffered phenol, pH 8.0. Store at 4°C in a dark glass bottle. 6. Phenol-chloroform-isoamyl alcohol (25:24:1). 7. Chloroform-isoamyl alcohol (24:1). 8. 10.5 M ammonium acetate: dissolve 80.93 g of ammonium acetate per 100 mL of ddH2 O. Filter sterilize using a 0.22-μm filter. 9. Cold isopropanol (4°C). 10. 70% Ethanol in water. 11. TE buffer, pH 8.0: 10 mM Tris-HCl, pH 8.0, 1 mM EDTA, pH 8.0. 12. RNase One™ Ribonuclease (Promega). Store at –20°C.

2.2. Suppressive Subtractive Hybridization (SSH) The subtractive hybridization process utilizes the CLONTECH PCR-Select™ Bacterial Genome Subtraction Kit (Clontech Laboratories Inc. cat. no. 637404). The following listed items are supplied in the kit. Store 4× hybridization buffer at room temperature. All other reagents are to be stored at –20°C. 2.2.1. Endonuclease Digestion 1. 10× RsaI restriction buffer: 100 mM bis-tris-propane-HCl, pH 7.0, 100 mM MgCl2 , 10 mM dithiothreitol (DTT). 2. 12 μL of RsaI (10 U/μL).

2.2.2. Adaptor Ligation 1. 21 μL of T4 DNA Ligase (400 U/μL; contains 3 mM ATP). 2. 5× DNA ligation buffer: 250 mM Tris-HCl, pH 7.8, 50 mM MgCl2 , 10 mM DTT, 0.25 mg/mL of bovine serum albumin (BSA). 3. 30 μL of adaptor 1 (10 mM). 4. 30 μL of adaptor 2R (10 mM).

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2.2.3. Hybridization 1. 4× Hybridization buffer: 200 mM N-2-hydroxyethylpiperazine-N -2-ethanesulfonic acid (HEPES-HCl), pH 8.0, 2 M NaCl, 0.8 mM EDTA, pH 8.0. Store at room temperature. 2. Dilution buffer: 20 mM HEPES-HCl, pH 8.3, 50 mM NaCl, 0.2 mM EDTA, pH 8.0.

2.2.4. PCR Ampliﬁcation 1. 2. 3. 4.

50 μL of PCR primer 1 (10 mM). 100 μL of nested primer 1 (10 mM). 100 μL of nested primer 2R (10 mM). 10 μL pf PCR control subtracted cDNA.

2.2.5. Control Reagents 1. 2. 3. 4.

5 μL of E. coli genomic DNA (1 mg/mL). 10 μL of control DNA (3 ng/μL; øX174/HaeIII digest). 50 μL of 10 μM 23S RNA forward primer*. 50 μL of 10 μM 23S RNA reverse primer*.

2.3. Cloning and Sequencing of Library This process utilizes a TOPO TA Cloning Kit for Sequencing (with pCR® 4TOPO® vector) with One Shot® TOP10 Electrocomp™ E. coli (Invitrogen; cat. no. K4580-01). The following listed items are supplied in the kit. Store all reagents at –20 °C unless noted otherwise. 2.3.1. TOPO TA Cloning Kit for Sequencing (with pCR® 4-TOPO® Vector) 1. 25 μL of pCR® 4-TOPO™ (10 ng/μL of plasmid DNA in 50% glycerol, 50 mM TrisHCl, pH 7.4 [at 25°C], 1 mM EDTA, 2 mM DTT, 0.1% Triton X-100, 100 μg/mL of BSA, 30 μM phenol red). 2. 100 μL of 10× PCR buffer (100 mM Tris-HCl, pH 8.3 [at 42°C], 500 mM KCl, 25 mM MgCl2 , 0.01% gelatin). 3. 10 μL of dNTP mix (12.5 mM dATP, 12.5 mM dCTP, 12.5 mM dGTP, 12.5 mM dTTP neutralized at pH 8.0 in water). 4. 50 μL of salt solution (1.2 M NaCl, 0.06 M MgCl2 ). 5. 1 mL of sterile water. 6. 20 μL of M13 forward (–20) primer (0.1 μg/μL in TE buffer, pH 8 [407 pmol]). 7. 20 μL of M13 reverse primer (0.1 μg/μL in TE buffer, pH 8 [385 pmol]). 8. 20 μL of T3 primer (0.1 μg/μL in TE buffer, pH 8 [329 pmol]). 9. 20 μL of T7 primer (0.1 μg/μL in TE buffer, pH 8 [328 pmol]). 10. 10 μL of control PCR template (0.1 μg/μL in TE buffer, pH 8).

*These primers will amplify a 374-bp fragment of the E. coli 23S rRNA gene.

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11. 10 μL of control PCR primers (0.1 μg/μL each in TE buffer, pH 8). M13 forward (–20) M13 reverse T3 T7

5´-GTAAAACGACGGCCAG-3´ 5´-CAGGAAACAGCTATGAC-3´ 5´-ATTAACCCTCACTAAAGGGA-3´ 5´-TAATACGACTCACTATAGGG-3´

2.3.2. One Shot® TOP10 Electrocomp™ E. coli and Reagents 1. S.O.C. medium (2% tryptone, 0.5% yeast extract, 10 mM NaCl, 2.5 mM KCl, 10 mM MgCl2 , 10 mM MgSO4 , 20 mM glucose [6 mL]). Store at +4°C or room temperature. 2. 21× 50 μL of TOP10 cells Electrocomp™. Store at –80°C. 3. 50 μL of pUC19 control DNA (10 pg/μL in 5 mM Tris-HCl, 0.5 mM EDTA, pH 8). Store at –80°C.

2.3.3. Inventory of Other Supplies and Reagents 1. 2. 3. 4. 5. 6.

7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18. 19.

AluI restriction enzyme (includes NEBuffer2 from New England Biolabs). 1 kb DNA Ladder (Invitrogen). 1.5-mL tubes (Fisher Scientific siliconized/ low-retention microcentrifuge tubes). 0.2-mL thin-walled PCR reaction tubes (Fisher Scientific Costar PCR tubes). Sterile, deionized H2 O. 5× TBE electrophoresis buffer: dissolve 54 g of Tris base, 27.5 g of boric acid, 20 mL of 0.5 M EDTA, pH 8.0, in 1 L of ddH2 O. Working solution for agarose gel electrophoresis is 0.5× (45 mM Tris-borate, 1 mM EDTA). 0.2 M EDTA, pH 8.0. Phenol–chloroform–isoamyl alcohol (25:24:1) (Sigma-Aldrich). Chloroform–isoamyl alcohol (24:1) (Fluka via Sigma-Aldrich). 3 M sodium acetate, pH 5.2. 20 mg/mL of glycogen (Sigma-Aldrich). 100% Ice-cold ethanol. 80% Ice-cold ethanol. TaKaRa Ex Taq DNA polymerase (includes dNTPs and buffer) (Fisher Scientific). Electroporation cuvettes, 0.1 cm (Bio-Rad). LB agar plates (1.0% tryptone, 0.5% yeast extract, 1.0% NaCl, pH 7.0, 1.5% agar) supplemented with ampicillin (50–100 μg/mL). LB (Luria-Bertani) medium (1.0% tryptone, 0.5% yeast extract, 1.0% NaCl, pH 7.0). 96-Well cell culture cluster, flat bottom with low evaporation lid (Corning). Glycerol.

2.4. Conﬁrmation of Tester-Speciﬁc DNA Fragments by Dot-Blot and Southern Blot Hybridizations 2.4.1. Preparation of the Dot-Blot Membrane 1. QIAquick PCR Purification kit (QIAGEN). 2. Sterile ddH2 O.

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Whatman 3 MM filter paper. 1 M NaOH. 0.4 M NaOH. 200 mM EDTA, pH 8.2. Hybond N+ charged nylon membrane (Amersham). Dot-blot manifold such as the Bio-Dot Microfiltration Apparatus (Bio-Rad). Clean glass tray. 2× SSC. UV transparent plastic wrap.

2.4.2. Preparation of the Southern Blot Membrane 1. 2. 3. 4. 5. 6. 7.

QIAquick PCR Purification kit (QIAGEN). Sterile ddH2 O. Whatman 3 MM filter paper. Agarose and ethidium bromide stain. 0.25 M HCl. Denaturation solution: 1.5 M NaCl, 0.5 M NaOH, store at room temperature. Neutralization solution: 1.5 M NaCl, 0.5 M Tris·Cl, pH 7.0. Store at room temperature. 8. Two clean glass trays. 9. 20× SSC: 3 M NaCl, 0.3 M Na3 citrate·2H2 O, adjust to pH 7.0 with 1 M HCl. Store at room temperature. 10. 2× SSC. Store at room temperature.

2.4.3. Radiolabeling of DNA Probes for Dot and Southern Blots 1. Sterile ddH2 O. 2. Items supplied in the Random Primers DNA Labeling System (Invitrogen) dATP solution, dGTP solution, dTTP solution, random primers buffer mixture, Klenow fragment, stop buffer, store at –20°C. 3. [-32 P]dCTP, 3000 Ci/mmol, 10 μCi/μL. Store at –20°C.

2.4.4. Puriﬁcation of Radiolabeled DNA Probes for Dot and Southern Blots 1. Nensorb™ 20 Nucleic Acid Purification Cartridge (NEN Life Science Products, Boston, MA). 2. Disposable 5-mL syringe. 3. 100% Methanol. 4. Reagent A: 0.1 M Tris·HCl, 10 mM triethylamine (TEA), 1 mM dipotassium or disodium EDTA, pH 7.7, stable for 1 month at 4°C. 5. 50% Methanol in water. 6. Molecular biology grade water. 7. 1.5-mL Eppendorf tube.

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2.4.5. Hybridization of Dot and Southern Blot Membranes to Radiolabeled DNA Probes 1. 2. 3. 4.

Hybridization bottle. PerfectHyb Plus Hybridization Buffer (Sigma-Aldrich). Low-stringency wash solution: 2× SSC, 0.1% SDS. Store at room temperature. Moderate stringency wash solution: 0.2× SSC, 0.1% SDS. Store at room temperature. 5. 2× SSC. 6. Phosphor imaging plates or X-ray film. 7. Phosphor imaging system such as BAS-1800 II (Fujifilm) or X-ray film processor.

2.4.6. Stripping of Dot and Southern Blot Membranes 1. 0.1% SDS. 2. Glass dish.

2.5. Total RNA Extraction and 16S rRNA Dot-Blot Hybridization 2.5.1. Total RNA Extraction 1. 0.1 M sodium phosphate buffer, pH 6.0: to make 200 mL, mix 87.7 mL of solution A (27.6 g of NaHPO4 ·H2 O/L [0.2 M]) with 12.3 mL of solution B (53.65 g of Na2 HPO4 ·7H2 O/L [0.2 M]), then dilute with H2 O to 200 mL. 2. Acetate buffer, pH 5.1 (50 mM sodium acetate, 10 mM EDTA). 3. 20% SDS (w/v). 4. Buffered phenol, pH 5.1. Store at 4°C in a dark glass bottle. 5. 0.1-mm zirconium beads, autoclaved. 6. 2-mL Eppendorf tubes. 7. Phenol-chloroform-isoamyl alcohol (25:24:1). 8. Chloroform-isoamyl alcohol (24:1). 9. 3 M sodium acetate. 10. Absolute ethanol. 11. 80% Ethanol. 12. RNase-free water. 13. RQ1 RNase-free DNase (Promega). 14. Agarose and ethidium bromide stain.

2.5.2. Radiolabeling of DNA Probes for RNA Dot Blots 1. 2. 3. 4. 5. 6.

Polynucleotide kinase (Invitrogen). 10× Kinase buffer (Invitrogen). [-32 P]ATP. 1% IGEPAL CA-630 (Sigma-Aldrich). RNase-free water. Nensorb™ 20 Nucleic Acid Purification Cartridge (NEN Life Science Products, Boston, MA).

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7. Disposable 5-mL syringe. 8. 100% Methanol. 9. Reagent A: 0.1 M Tris·HCl, 10 mM triethylamine (TEA), 1 mM dipotassium or disodium EDTA, pH 7.7, stable for 1 month at 4°C. 10. 50% Methanol in water. 11. 1.5- mL Eppendorf tube.

2.5.3. Preparation of RNA Dot-Blot Membrane and Hybridization of Membrane to Radiolabeled DNA Probes 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13.

2% Glutaraldehyde. Hybond N+ charged nylon membrane (Amersham). Whatman 3 MM filter paper. Dot-blot manifold such as the Bio-Dot Microfiltration Apparatus (Bio-Rad). Clean glass tray. RNase free water. UV transparent plastic wrap. Hybridization bottle. PerfectHyb Plus Hybridization Buffer (Sigma-Aldrich). RNA dot-blot washing solution (1× SSC, 1% SDS). Phosphor imaging plates or X-ray film. Phosphor imaging system such as BAS-1800 II (Fujifilm) or X-ray film processor. ImageQuant (Fujifilm) or other densitometry software.

3. Methods 3.1. Extraction of Total Genomic DNA from Environmental Samples This procedure has been adapted from a method for the extraction of DNA from sediments (49) and optimized for use with rumen samples. Before DNA isolation, fresh rumen contents can be strained through two layers of cheesecloth to remove large particulate matter. Alternate methods of DNA isolation may be preferable for other types of environmental samples. 1. Wash the sample by mixing 2 g of the environmental sample with 10 mL of 0.1 M sodium phosphate buffer, pH 8.0, shaking briefly. Centrifuge the sample at 6000g for 10 min at 4°C and discard the supernatant. 2. Suspend the sample in 10 mL of sodium phosphate buffer, pH 8.0, and repeat step 1. 3. Add 2 mL of lysis solution per gram of pelleted sample and incubate with gentle shaking (75 rpm on a rotating shaker) for 30 min at 37°C. 4. Add an equal volume of SDS solution to the sample and incubate with gentle shaking (75 rpm on a rotating shaker) for 30 min at 37°C. 5. Perform three cycles of freezing at –80°C and thawing at room temperature. 6. Add proteinase K to a final concentration of 50 μg/mL and incubate for 30 min at 37°C.

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7. Add an equal volume of phenol and gently invert the sample tube for 4 min. Centrifuge at 10,000g for 30 min at 4°C. Pipet the aqueous phase into a clean tube. 8. Add an equal volume of phenol–chloroform–isoamyl alcohol mixture and gently invert the sample tube for 4 min. Centrifuge at 10,000g for 30 min at 4°C. Pipet the aqueous phase into a clean tube. 9. Add an equal volume of chloroform mixture and gently invert the sample tube for 4 min. Centrifuge at 10,000g for 15 min at 4°C. Pipet the aqueous phase into a clean tube. 10. Add 10.5 M ammonium acetate to a final sample concentration of 2.5 M. Add an equal volume of cold isopropanol. Place the sample at –80°C for 30 min to precipitate the nucleic acids. 11. Centrifuge the sample at 12,000g for 30 min at 4°C. Discard the supernatant and wash the nucleic acid pellet with 2 mL of 70% EtOH. 12. Dry the pellet and suspend in 200 μL of TE buffer. 13. Add 1 μL of RNase One™ to remove RNA from sample. 14. Store DNA at 4°C or –20°C.

3.2. Suppressive Subtractive Hybridization The described protocol is based on the manufacturer’s protocol for CLONTECH’s PCR-Select™ Bacterial Genome Subtraction Kit (courtesy of Clontech Laboratories, Inc., a Takara Bio Co. [2006] Clontech Laboratories, Inc.). Specific steps have been amended due to metagenomic DNA being used instead of genomic DNA from pure cultures. Further information can additionally be obtained from the “Suppression Subtractive Hybridization” chapter in the Methods in Molecular Biology volume on “Gene Expression Profiling” (50). 3.2.1. Restriction Digestion of Metagenomic DNA Perform the following procedure with each experimental tester and driver metagenomic DNA, and with the control E. coli genomic DNA. The driver DNA will be used to “subtract” the DNA common with the tester DNA, thereby enriching for fragments unique to the tester DNA. Testing the efficacy of the restriction enzyme used is crucial to the success of the technique. Specifically, it serves as an indicator for the presence of confounding inhibitors of latter steps if digestion does not occur. In our experiments we used AluI as the restriction enzyme; for the E. coli control you must use the included RsaI restriction enzyme. If you do not, then you will not be able to conduct the associated control experiments using the E. coli DNA. 1. Label three 0.5-mL microcentrifuge tubes with a Sharpie as follows: T (this will contain the metagenomic tester DNA), D (this will contain the metagenomic driver

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DNA), and C (this will contain the E. coli control DNA). These designations will be used throughout the procedure. 2. Add the following reagents into the appropriately labeled 0.5-mL microcentrifuge tubes (see Note 1):

T-labeled tube (metagenomic tester DNA): Metagenomic tester DNA (2 μg in X μL) 10× AluI restriction buffer AluI (10 U/μL) Sterile, deionized H2 O Total reaction volume:

X μL 5.0 μL 1.5 μL X μL 50 μL

D-labeled tube (metagenomic driver DNA): Metagenomic driver DNA (2 μg in X μL) 10× AluI restriction buffer AluI (10 U/μL) Sterile, deionized H2 O Total reaction volume:

X μL 5.0 μL 1.5 μL X μL 50 μL

C-labeled tube (E. coli control DNA): E. coli control DNA (2 μg in X μL) 10× RsaI restriction buffer RsaI (10 U/μL) Sterile, deionized H2 O Total reaction volume:

X μL 5.0 μL 1.5 μL X μL 50 μL

3. Mix each tube by gently flicking the end of the tube and centrifuge briefly. Vortexmixing may give the appearance of digestion as opposed to specific restriction enzyme digestion of the DNA. 4. Incubate at 37°C for 5–16 h. 5. Use 5 μL of the digest mixture to analyze the efficiency of digestion. Electrophorese 0.2 μg of uncut metagenomic DNA and 5 μL of AluI-digested metagenomic DNA on a 1% agarose gel run in 0.5× TBE buffer. Compare the results side by side. The undigested genomic DNA should appear as a high-molecular weight band at the top of the gel running higher than the 1 Kb DNA Ladder. (This will also be dependent on the method used to extract DNA from the environmental sample. Some methods are more prone to DNA shearing than others, e.g. those using bead-beating). The AluI-digested DNA, in contrast, will be drastically decreased in size. (See Note 2 if this is not the case.) 6. Add 2.5 μL of 0.2 M EDTA, pH 8.0, to terminate the reaction and transfer the volume to a 1.5-mL tube.

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7. Add 450 μL of sterile, deionized H2 O to bring volume up to 500 μL. Then add 500 μL of phenol–chloroform–isoamyl alcohol (25:24:1). (This minimizes the loss of material associated with removing the aqueous layer in step 10.) 8. Vortex-mix thoroughly. 9. Centrifuge the tubes at 10,000 g for 10 min to separate phases. 10. Remove the top aqueous layer and place in a fresh 1.5-mL tube. 11. Add 500 μL of chloroform–isoamyl alcohol (24:1) and vortex-mix thoroughly. 12. Centrifuge the tubes at 10,000 g for 10 min to separate phases. 13. Remove the top aqueous layer and place in a clean 1.5-mL tube. 14. Add 1 μL of glycogen (20 mg/mL), 0.1 volume of 5 M sodium acetate, pH 5.2, and 2 volumes of 100% ice-cold ethanol. (Therefore, if the volume of the aqueous layer is 500 μL, add 1 μL of glycogen [20 mg/mL], 50 μL of 5 M sodium acetate [pH 5.2], and 1000 μL of 100% ice-cold ethanol.) 15. Vortex-mix the mixture thoroughly. 16. Centrifuge the tubes for 15 min at 10,000 g at room temperature. A small white pellet of precipitated DNA should appear at the bottom. (See Note 3 if it does not.) 17. Remove the supernatant carefully. 18. Gently overlay the pellets with 200 μL of 80% ethanol. 19. Centrifuge at 10,000 g for 5 min. 20. Carefully remove the supernatant. 21. Air-dry the pellets for 5–10 min. 22. Dissolve the pellet in 6.5 μL of H2 O and store at –20°C. The final concentration of driver DNA should be approx 300 ng/μL.

3.2.2. Adaptor Ligation Three adaptor ligations must be performed for each metagenomic tester DNA, as well as for the control tester DNA. Each DNA is aliquotted into two separate tubes: one aliquot is ligated with adaptor 1 (tester T1 and C1, according to the “T” and “C” labeling convention from Subheading 3.2.1.), and the second is ligated with Adaptor 2R (tester T2 and C2). In a third tube, portions of each aliquot are combined so that the DNA is ligated with both adaptors (unsubtracted tester control T-c and C-c). Each unsubtracted tester control DNA serves as a negative control for subtraction. An additional tester DNA for the positive control subtraction is prepared by mixing the RsaIdigested control E. coli genomic DNA with the Control DNA (øX174/HaeIII digest) provided in the kit. 1. Label two tubes as “T-diluted” and “C-diluted”. Dilute 1.2 μL of each corresponding restriction enzyme-digested experimental DNA with 1.8 μL of sterile, deionized H2 O. The digested metagenomic tester DNA, tube T from Subheading 3.2.1., will go into “T-diluted,” while digested control E. coli DNA, tube C from Subheading 3.2.1., will go into “C-diluted.” 2. Prepare positive control tester DNA:

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a. Briefly centrifuge the tube containing the control DNA (øX174/HaeIII digest, provided in the kit). b. Dilute 3 μL of the control DNA with 42 μL of sterile, deionized H2 O (up to 0.2 ng/μL). c. Mix 1.2 μL of the RsaI-digested control E. coli genomic DNA (300 ng/uL) with 1.8 μL of the diluted øX174/HaeIII DNA. This is your control tester DNA (see Note 4). 3. Prepare a ligation master mix by combining the following reagents in a 0.5-mL microcentrifuge tube. To ensure that you have sufficient master mix, prepare enough for all ligations plus one additional reaction.

Sterile, deionized H2 O 5× Ligation buffer T4 DNA ligase (400 U/μL)

per reaction 4 μL 2 μL 1 μL

4. For the metagenomic tester DNA (two tubes labeled T1 and T2) and for the control tester DNA (two tubes labeled C1 and C2), combine the reagents below in 0.5mL microcentrifuge tubes in the order shown. Pipet mixture up and down to mix thoroughly.

Tube no.: Component Diluted tester DNA* Adaptor 1 (10 mM) Adaptor 2R (10 mM) Master mix Final volume

T1 (μL) 1 2 — 7 10

T2 (μL) 1 — 2 7 10

C1 (μL) 1 2 — 7 10

C2 (μL) 1 — 2 7 10

5. In a fresh microcentrifuge tube labeled T-c, mix 1.5 μL of tester T1 and 1.5 μL of tester T2. This will be designated as the unsubtracted tester control T-c. Do the same for the control tester DNA (tester C1 and C2; label this fresh tube as C-c). After ligation, approx 1/3 of the DNA molecules in each unsubtracted tester control tube will have two different adaptors on their ends. 6. Centrifuge tubes briefly, and incubate at 16°C overnight. 7. Terminate ligation reactions by adding 1 μL of 0.2 M EDTA. 8. Heat samples at 72°C for 5 min to inactivate the ligase.

*Diluted tester DNA refers to either the "T-diluted" or "C-diluted" tube from step 1 above.

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9. Briefly centrifuge the tubes. Preparation of the experimental and control Adaptorligated tester DNAs (T1, T2, C1, and C2) is now complete. 10. Dilute 1 μL of each unsubtracted tester control (T-c and C-c) in separate tubes into 1 mL of H2 O. These samples will be used for PCR amplification later. 11. Store samples at –20°C.

3.2.3. Analysis of Ligation This subheading describes a quality control test for the ligation of adaptors and is specifically designed for the control E. coli genomic DNA included in the Clontech kit. The primers will amplify a 374-bp region of the 23S rRNA gene of E. coli and are used to verify that at least 25% of the control tester DNA fragments have adaptors on both ends. Amplified fragments span the adaptor/ DNA junctions of testers C1 and C2. (See Note 5.) 1. Dilute 1 μL of each E. coli control-ligated DNA (e.g., testers C1 and C2) into 200 μL of H2 O. 2. Set up four PCR reactions with the following template and primer combinations (volumes are in μL):

Tester C1 (ligated to adaptor 1) Tester C2 (ligated to adaptor 2R) 23S RNA forward primer (10 mM) 23S RNA reverse primer (10 mM) PCR primer 1 (10 mM) Total volume

1 1 — 1 — 1 3

2 1 — 1 1 — 3

3 — 1 1 — 1 3

4 — 1 1 1 — 3

3. Prepare a master mix for all of the reaction tubes plus one additional tube. For each reaction planned, combine the reagents below in the order shown:

Sterile H2 O 10× PCR reaction buffer dNTP mix (10 mM) Total volume

Amount per component reaction tube (μL) 18.75 2.5 0.5 21.75

Amount for 4 reactions (μL) 93.75 12.5 2.5 108.75

4. Mix well by vortex-mixing and then briefly centrifuge the tube. 5. Aliquot 21.75 μL of master mix into each of the reaction tubes from step 2 above.

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6. Mix well by vortex-mixing and briefly centrifuge the tube. 7. Incubate the reaction mix in a thermal cycler (using a Applied Biosystems GeneAmp® System 9700) at 72°C. As soon as the temperature has reached 72°C, pause the thermal cycler and add 0.25 μL of TaKaRa Ex Taq DNA polymerase (1.25 U) to each tube (see Note 6). Incubate the tubes for 2 min to extend the adaptors. This step “fills in” the missing strand of the adaptors (Fig. 1) and thus creates binding sites for PCR primer 1. At this stage, do not remove the samples from the thermal cycler. 8. Immediately commence thermal cycling (using a Applied Biosystems GeneAmp® System 9700): • 94°C 30 s 23 cycles: • 94°C 10 s • 65°C 30 s • 68°C 1 min Final extension at 72°C for 7 min, then hold at 4°C indefinitely. 9. Examine the products by electrophoresis on a 2% agarose gel run in 0.5× TBE buffer. If you cannot see a product after 23 cycles, perform four more cycles, and again analyze the product by agarose gel electrophoresis (see Note 7).

3.2.4. First Hybridization In the first hybridization, an excess of driver DNA is added to the tester DNA, and the samples are heat denatured and allowed to anneal. The concentration of fragments, present in single and multiple copies, is equalized among the type a molecules (Fig. 1) because reannealing is faster for the more abundant molecules due to the second-order kinetics of hybridization. 1. For each of the metagenomic (designated HT below) and control (HC) subtractions, combine the reagents in 0.2-mL microcentrifuge tubes in the order shown:

Component

Hybridization sample HT1 (μL)

Hybridization sample HT2 (μL)

AluI-digested driver DNA Adaptor 1-ligated tester T1 Adaptor 2R-ligated tester T2 4× Hybridization buffer Final volume

2.0 1.0 — 1.0 4.0

2.0 — 1.0 1.0 4.0

2. Incubate samples in a thermal cycler at 98°C for 1.5 min. 3. Incubate samples at 63°C for 1.5 h, then proceed immediately to the next section (see Note 8).

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3.2.5. Second Hybridization In the second hybridization the two samples from the first hybridization are mixed together and fresh denatured driver DNA is added to further enrich for tester-specific sequences. New hybrid molecules that correspond to testerspecific DNAs with different adaptors on each end are formed. Do not denature the primary hybridization samples at this stage. Also, do not remove the hybridization samples from the thermal cycler for longer than is necessary to add fresh driver. Repeat the following steps for your experimental tester DNA and for the control tester DNA. 1. Dilute the 4× hybridization buffer with an equal volume of deionized H2 O to make 2× hybridization buffer. 2. Combine the following reagents in a sterile 0.2-mL microcentrifuge tube:

Alul-digested driver DNA* 2× Hybridization buffer

1 μL 1 μL

3. Incubate in a thermal cycler at 98°C for 1.5 min. 4. Remove the tube of freshly denatured driver from the thermal cycler. Use the following procedure to simultaneously mix the driver with hybridization samples 1 and 2 (HT1 and HT2 for the metagenomic DNA, and HC1 and HC2 for the E. coli control DNA). This ensures that the two hybridization samples mix together only in the presence of driver. a. Set a micropipettor at 15 μL. b. Gently touch the pipet tip to the sample surface of the tube containing hybridization sample 2 (HT2 for the metagenomic sample, or HC2 for the E. coli control). c. Carefully draw the entire sample part way into the pipet tip. d. Remove the pipet tip from the tube, and draw a small amount of air into the tip, creating a slight air space below the droplet of sample. e. Repeat steps b–d with the tube containing the freshly denatured driver. The pipet tip should now contain both samples separated by a small pocket of air. f. Transfer the entire mixture to the tube containing hybridization sample 1 (HT1 for the metagenomic sample, or HC1 for the E. coli control). g. Mix by pipetting up and down. 5. 6. 7. 8. 9.

Briefly centrifuge the tube if necessary. Incubate reaction at 63°C overnight. Add 200 μL of dilution buffer to the tube and mix by pipetting. Heat in a thermal cycler at 63°C for 7 min to eliminate nonspecific hybridization. Store at –20°C. ∗

For the metagenomic DNA this would correspond to tube “D” from Subheading 3.2.1.

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3.2.6. PCR Ampliﬁcation In the following PCR, tester-specific DNA fragments are selectively amplified. The missing strands of the adaptors are first “filled in” again by a brief preincubation at 72°C which creates the binding sites for PCR primer 1 (Fig. 1). Following that, in the first round of PCR amplification, only dsDNA with different adaptor sequences on each end are exponentially amplified. Subsequently, in the second round, nested PCR is used to enrich for testerspecific sequences thereby reducing the background. At least five reactions should be prepared: 1. 2. 3. 4. 5.

The The The The The

subtracted metagenomic DNA (mix of HT1/HT2). unsubtracted metagenomic tester control (T-c). subtracted control DNA from E. coli (mix of HC1/HC2). unsubtracted tester control for the control subtraction (C-c). PCR control subtracted DNA (provided in the kit).

The PCR control subtracted DNA from the kit provides a positive PCR control that can be compared directly with the subtracted control DNA from E. coli and contains a successfully subtracted mixture of HaeIII-digested øX174 DNA fragments. In addition, you should also include a standard PCR control to ensure that the Taq DNA polymerase used in the PCR is performing efficiently. 1. Prepare the PCR templates as follows: a. Aliquot 1 μL of each diluted DNA (i.e., each subtracted sample and the corresponding diluted unsubtracted tester control) into an appropriately labeled tube. This is to be done for the prepared metagenomic subtracted sample (HT1/HT2), the diluted metagenomic unsubtracted sample (T-c), the prepared E. coli subtracted sample (HC1/HC2), and the diluted E. coli unsubtracted sample (C-c). b. Aliquot 1 μL of the PCR control subtracted DNA (provided in the kit) into an appropriately labeled tube. 2. Prepare a master mix for all of the primary PCR tubes plus one additional tube. Combine the reagents in the order shown below:

Component

Amount per reaction (μL)

For a 5-reaction experiment (μL)

Sterile H2 O 10× PCR reaction buffer dNTP mix (10 mM) PCR primer 1 (10 mM) Total volume

19.75 2.5 0.5 1.0 23.75

118.5 15.0 3.0 6.0 142.5

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3. Vortex-mix well and briefly centrifuge the tube. 4. Aliquot 23.75 μL of master mix into each of the reaction tubes prepared in step 1. 5. Incubate the reaction mix in a thermal cycler (using a Applied Biosystems GeneAmp® System 9700) at 72°C. As soon as the temperature has reached 72°C, pause the thermal cycler and add 0.25 μL of TaKaRa Ex Taq DNA polymerase (1.25 U) to each tube (see Note 6). Incubate the tubes for 2 min to extend the adaptors. At this stage, do not remove the samples from the thermal cycler. 6. Immediately commence thermal cycling (using a Applied Biosystems GeneAmp® System 9700): Initial hold at 94°C for 5 min 25 cycles: • 94°C 10 s • 66°C 30 s • 72°C 1.5 min Final extension at 72°C for 7 min, then hold at 4°C indefinitely. 7. Three to five more cycles may be needed depending on the Taq DNA polymerase used. In our case, with TaKaRa Ex Taq DNA polymerase, we obtained optimal results with 5 extra cycles (total of 30). 8. Analyze 7 μL from each tube on a 2% agarose gel run in 0.5× TBE buffer. You can set these 7-μL aliquots aside and run them on the same gel used to analyze the secondary PCR products (step 16 below). 9. Dilute 1 μL of each primary PCR mixture in 39 μL of H2 O. 10. Aliquot 1 μL of each diluted primary PCR mixture from the previous step into an appropriately labeled tube. 11. Prepare master mix for the secondary PCRs plus one additional reaction by combining the reagents in the order shown:

Reagent

Amount per reaction (μL)

For a 5-reaction experiment (μL)

Sterile H2 O 10× PCR reaction buffer Nested primer 1 (10 mM) Nested primer 2R (10 mM) dNTP mix (10 mM) Total volume

18.75 2.5 1.0 1.0 0.5 23.75

112.5 15.0 6.0 6.0 3.0 142.5

12. Vortex-mix well and briefly centrifuge the tube. 13. Aliquot 23.75 μL of master mix into each reaction tube and incubate the reactions in a thermal cycler (using a Applied Biosystems GeneAmp® System 9700) at 94°C for 5 min. As soon as the temperature has reached 94°C, pause the thermal cycler and add 0.25 μL of TaKaRa Ex Taq DNA polymerase (1.25 U) to each tube (see Note 6).

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14. Immediately commence thermal cycling (using a Applied Biosystems GeneAmp® System 9700): 10–12 cycles: • 94°C 10 s • 68°C 30 s • 72°C 1.5 min Final extension at 72°C for 7 min, then hold at 4°C indefinitely. 15. Three to five more cycles may be needed depending on the Taq DNA polymerase used. In our case, with TaKaRa Ex Taq DNA polymerase, we obtained optimal results without extra cycles for the secondary PCR (total of 12 cycles only). 16. Analyze 7 μL from each reaction on a 2.0% agarose gel run in 0.5× TBE buffer. 17. Store reaction products at –20°C (see Note 9).

3.2.7. Analysis of PCR Products 1. Agarose gel analysis of primary PCR. With the PCR control subtracted DNA, the major bands that appear after 25 cycles should correspond to the øX174/HaeIII fragments. This result should look similar to the control subtraction performed. The øX174/HaeIII bands may appear only after the secondary PCR—this is normal. At this stage, success with the control reactions is the only indication that the subtraction with metagenomic DNA may work. The primary PCR products from the experimental subtraction with metagenomic DNA should appear as a smear of amplified products from 0.2–2 kb. Confirmation of successful subtraction with metagenomic DNA is described in the subsequent sections using Southern blots. What follows here are guidelines on how to interpret the results to this point now and troubleshoot some basic problems. a. If you cannot see any products after 25 cycles, perform three to five more cycles, and analyze the products by gel electrophoresis. (As noted in Subheading 3.2.6., our experiment required a total of 30 cycles). This variable is usually the most responsive and easiest to compare when products from increasing cycles are analyzed side-by-side on the same agarose gel. b. If PCR products are not detected for either the subtracted (prepared E. coli subtracted sample [HC1/HC2]) or unsubtracted (unsubtracted tester control Cc) samples or the PCR control subtracted DNA, make sure that the Taq DNA polymerase is working. If the Taq DNA polymerase is working optimally, then try optimizing the PCR cycling parameters. Two variables to modify independently include decreasing the annealing or the extension temperature in small increments. Lowering these temperatures can dramatically increase the background. Initially, try reducing the annealing temperature from 66°C to 64°C. The extension temperature should be lowered in smaller increments (e.g., from 72°C to 71°C).

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2. Agarose gel analysis of secondary PCR. The patterns of the secondary PCR products from the PCR control subtracted DNA and from the E. coli control subtraction should resemble each other (a few additional background bands may appear). Because of the increased complexity of metagenomes relative to pure cultures the experimental subtracted samples usually look like smears. In contrast, for subtraction experiments with DNA from pure cultures, a number of distinct bands may also appear in addition to the smeared products. a. If the expected øX174/HaeIII bands appear in the PCR Control Subtracted DNA, but not in the control subtraction (i.e., either a smear of products or randomly sized bands appear), then there was probably an issue during the subtraction. Poor ligation efficiency is likely. If the ligation efficiency is not a problem, then be sure that the protocol was followed correctly. b. If you cannot see a product after 12 cycles, perform three more cycles, and again check the product by gel electrophoresis. Be conservative when adding cycles; too many cycles will increase background. c. If you do not observe øX174/HaeIII bands in the PCR Control Subtracted DNA, you will need to optimize the PCR conditions. Again, because of the variation in Taq DNA polymerases and thermal cycler models, many of these conditions need to be determined empirically. d. In a successful subtraction, the banding pattern of your unsubtracted DNA ligated with both adaptors (unsubtracted tester control T-c) should be different from the banding pattern of your experimental subtracted DNA samples. With metagenomic DNA this can sometimes be detected as a shift in the size of the smear of amplified products.

3.2.8. Clone Library of Enriched Fragments After Subtraction In the following steps, 4 μL of secondary PCR products from the metagenomic subtraction (HT1/HT2 from Subheading 3.2.6.) will be cloned into the pCR® 4-TOPO® vector (Invitrogen) and transformed into E. coli TOP10 cells (Invitrogen) by electroporation. The described protocol is based on the manufacturers’ protocol for the TOPO TA Cloning® Kit for Sequencing (courtesy of Invitrogen, © [2006]). 1. Set up the following reaction: Secondary PCR product Dilute salt solution TOPO® vector Final volume: 6 μL

4 μL 1 μL 1 μL

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4. 5. 6. 7.

8. 9.

10.

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Addition of the dilute salt solution in the TOPO® cloning reaction brings the final concentration of NaCl and MgCl2 in the TOPO® cloning reaction to 50 mM and 2.5 mM, respectively. Mix reaction gently and incubate for 5 min at room temperature (22–23°C). Place the reaction on ice. Add 2 μL of the TOPO® Cloning reaction into a vial of One Shot® Electrocompetent E. coli and mix gently. Do not mix by pipetting up and down; gently stir with the pipet. Store remaining cloning reaction volume at –20°C when finished. Carefully transfer solution to a 0.1-cm gapped cuvette, avoiding the formation of bubbles. Electroporate your samples. Electroporation conditions are 2.5 kV, 25 μF, and 200 using a 0.1-cm gapped cuvet and a Bio-Rad GenePulser™ (see Note 10). Immediately add 250 μL of room temperature SOC medium. Immediately plate transformants onto LB agar plates supplemented with 100 μg/mL of ampicillin and grow overnight at 37°C. Plate two different volumes to ensure that at least one plate will have well-spaced colonies. An efficient TOPO® cloning reaction should produce several hundred colonies. Pick 96 random transformants and grow at 37°C in 200 μL of Luria–Bertani medium supplemented with ampicillin for 2 h. Use the cycling conditions from the primary PCR, but the primers from secondary PCR to verify the presence of clone inserts and for use in dot and Southern blot hybridizations (see Subheading 3.2.6.). Use 1 μL of each cell culture as a template for PCR screening. Stock cultures of transformants should be maintained at –80°C in 96-well plates containing LB media with glycerol (1:1 ratio).

3.3. Conﬁrmation of Tester-Speciﬁc DNA Fragments by Dot-Blot and Southern Blot Hybridizations The efficiency of SSH can be tested by performing dot-blot and Southern blot hybridizations. In the dot-blot procedure, cloned subtraction fragments are spotted onto a membrane and hybridized to a tester metagenomic DNA probe and later to a driver metagenomic DNA probe (Fig. 2A). The Southern blot approach is similar, but uses the cloned subtraction fragments to probe a membrane containing the restriction digests of both tester and driver metagenomic DNA (Fig. 2B). If the SSH procedure was successful, there should be little or no hybridization of subtraction fragments to the driver metagenomic DNA on either type of blot. The method described here utilizes radiolabeled DNA probes; however, other methods of labeling are also suitable. All membrane manipulations should be performed using sterile forceps. (For further reference information on Southern blotting consult Current Protocols in Molecular Biology (51)).

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Fig. 2. Hybridization analysis of randomly selected clones from the subtracted library of the tester rumen metagenome sample. (A) PCR amplicons from 96 randomly selected clones were transferred to nylon membranes with a dot blot manifold and hybridized with 32 P–labeled genomic DNA from the tester metagenome (a) or the driver metagenome (b). (B) Southern blot hybridizations of five clones. Subtracted clones and AluI-digested genomic DNA from both tester (T) and driver (D) were electrophoresed using agarose gels, transferred to nylon membranes, and hybridized with respective 32 P–labeled subtracted clones. A 1 Kb DNA Ladder (L) (Invitrogen) is also shown. Clones H10 (a), C08 (b), E08 (c), and A04 (d) were randomly chosen. Clone C10 (e) was chosen as a positive control probe to confirm the dot blot hybridization results. (Reproduced from Galbraith et al. [2004] (7) with permission from Blackwell Publishing.)

3.3.1. Preparation of the Dot-Blot Membrane 1. Purify PCR products from randomly selected transformants using the QIAquick PCR Purification kit (QIAGEN) as per the manufacturer’s instructions. This results in purified DNA in a final volume of approx 50 μL per sample. 2. Dilute 10 μL of each PCR product with 40 μL of 1 M NaOH, 5 μL of 200 mM EDTA, pH 8.2, and 45 μL of sterile water. Denature the samples by heating at 99.9°C for 10 min in a thermocycler.

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3. Prepare a dot-blot manifold as per the manufacturer’s instructions. A charged nylon membrane (Hybond N+ ) cut to fit the manifold is laid on the surface of a tray of distilled water to allow the membrane to wet by capillary action. The membrane is submerged and soaked for 10 min. 4. Cut filter paper to fit the manifold, soak the paper in sterile distilled water, and place it on the manifold. 5. Lay the wet membrane on top of the filter paper and assemble the manifold, ensuring that no bubbles are trapped in the apparatus. 6. Prewash the membrane by adding 500 μL of sterile ddH2 O per well. A small amount of bromphenol blue dye can be added to the prewash water if desired. (See Note 11.) 7. Turn on the vacuum and allow the water to be slowly pulled through the membrane. Then, turn the vacuum off and slowly break the vacuum seal. 8. Apply denatured DNA samples to the membrane (one sample per well). 9. Turn the vacuum on once more, allowing the DNA to be pulled onto the membrane and the liquid from the samples to be pulled through. 10. Rinse each well with 500 μL of 0.4 M NaOH and dismantle the manifold. 11. Rinse the membrane briefly in 2× SSC and air-dry for 10 min. 12. Crosslink the DNA to the membrane by wrapping the membrane in UV transparent plastic wrap and irradiating in a UV crosslinker (Stratalinker, Stratagene) using the Autocrosslink function as per the manufacturer’s instructions. 13. Store the membrane dry at room temperature for up to 2–3 months, or at 4°C for longer periods of time.

3.3.2. Preparation of the Southern Blot Membrane 1. Purify PCR products from randomly selected transformants using the QIAquick PCR Purification kit (QIAGEN) as per the manufacturer’s instructions. This results in purified DNA in a final volume of approx 50 μL per sample. 2. Load approx 50 ng of a purified subtraction fragment, 1 μg of tester metagenomic DNA digest (digested to completion with the same restriction enzyme used in the SSH) and 1 μg of driver metagenomic DNA digest (also digested to completion) in separate lanes on a 0.8% agarose gel run in 0.5× TBE buffer. Electrophorese the gel for 3 h at 80 V, stain in 0.1 μg/mL of EtBr and visualize using a UV transilluminator. 3. Prepare the gel for DNA transfer by performing a series of washes at room temperature. All washes should be performed by slowly shaking the gel in a clean glass dish in approx 10 gel volumes of liquid. Between each of the washes, the gel should be rinsed with distilled water. Wash the gel once 0.25 M HCl for 30 min, twice in denaturation solution for 20 min, and twice in neutralization solution for 20 min. 4. Fill a glass dish with approx 1 L of 20× SSC. Balance a solid support large enough to hold the gel over the dish, leaving space on both sides of the support to allow a filter paper wick to hang down into the 20× SSC.

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5. Cut a Whatman 3 MM filter paper wick so that its width is the same size as the length of the gel and its length is long enough to lay across the solid support and hang into the 20× SSC solution on both sides of the support. Wet this wick with 20× SSC and place it on the support. 6. Place the gel on the filter paper and squeeze out any air bubbles. 7. Cut a charged nylon membrane (Hybond N+ ) to fit the agarose gel and lay it on the surface of a tray of distilled water to allow the membrane to wet by capillary action. The membrane is submerged and soaked for 10 min. 8. Place the wet membrane on the surface of the gel. Remove any air bubbles by slowly rolling a glass pipet over the surface. Flood the surface of the membrane with 20× SSC. 9. Cut five pieces of Whatman 3 MM filter paper to the size of the membrane, and stack on top of the membrane. Fold or cut paper towels to the size of the membrane and place on top of the filter paper. 10. Lay a solid support over the paper towels and place a weight on top of the support to hold the transfer pyramid in place. 11. Allow the transfer to proceed overnight at room temperature. 12. Disassemble the transfer pyramid, and using pencil, mark the position of gel wells and gel orientation on the membrane. 13. Rinse the membrane briefly in 2× SSC and air-dry for 10 min. 14. Cross link the DNA to the membrane by wrapping the membrane in UV transparent plastic wrap and irradiating in a UV crosslinker (Stratalinker, Stratagene) using the Autocrosslink function as per the manufacturer’s instructions. 15. Store the membrane dry at room temperature for up to 2–3 months, or at 4°C for longer periods of time.

3.3.3. Radiolabeling of DNA Probes for Dot and Southern Blots Labeling of several probes can be done simultaneously provided that the purified, labeled probes are stored at –20°C and used within a week. Dot blots should be probed with restriction enzyme digested metagenomic driver DNA first and subsequently with restriction enzyme digested metagenomic tester DNA. Hybridizing the dot blot in this order reduces the likelihood of residual signal present on the membrane after stripping. The restriction enzymes used for digestion of metagenomic probes should be identical to those used in the subtraction procedure. For Southern blots, a probe should consist of a single purified subtraction fragment. 1. Dilute purified probe DNA to 10 ng/μL in distilled water. Denature 100 ng of purified probe DNA by heating for 5 min in a boiling water bath, then immediately cooling on ice. 2. To the denatured probe, make the following additions on ice:

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2 μL dATP solution 2 μL dGTP solution 2 μL dTTP solution 15 μL random primers buffer mixture 5 μL (∼50 μCi) [-32 P]dCTP, 3000 Ci/mmol, 10 μCi/μL Distilled water to a total volume of 49 μL. Mix briefly. 3. Add 1 μL of Klenow fragment and mix thoroughly. Briefly centrifuge to deposit contents at the bottom of the tube. 4. Incubate at 25°C for 3 h. 5. Add 5 μL of stop buffer then proceed to probe purification or store probe at –20°C for up to 1 wk.

3.3.4. Puriﬁcation of Radiolabeled DNA Probes for Dot and Southern Blots Unincorporated dNTPs, salts, and other impurities should be removed from the DNA probes before hybridization. Purification using a Nensorb™ 20 Nucleic Acid Purification Cartridge (NEN Life Science Products, Boston, MA) is described here; however other purification methods may also be used. In the following steps, whenever pressure is applied to the cartridge following the application of a reagent or a sample, the flow rate of the solvent should not exceed one drop per 2 s. In addition, drying of the cartridge resin should be avoided until the final elution step. 1. Clamp one Nensorb 20 cartridge to a support, remove the protective cap from the cartridge, and place a waste collection beaker below the cartridge. 2. Pipet 2 mL of 100% methanol into the top of the cartridge. 3. Securely attach an adapter to the top of the cartridge forming an airtight seal. 4. Attach a disposable 5-mL syringe to the adaptor and gently push the methanol through the cartridge until the meniscus reaches the top of the cartridge resin bed. 5. Disconnect the syringe and the adapter and add 2 mL of reagent A to the top of the cartridge. 6. Reconnect the adapter and the syringe and gently push the reagent A through the cartridge until the meniscus reaches the top of the cartridge resin bed. 7. Disconnect the syringe and the adapter. 8. Add 200 μL of reagent A to the labeled probe and gently mix. Pipet this mixture into the top of the cartridge. Reattach the adapter and syringe and gently push the sample into the cartridge resin. 9. Disconnect the syringe and the adapter. 10. To wash the cartridge, pipet 3 mL of reagent A into the top of the cartridge, reattach the adapter and syringe, and gently push the liquid until the meniscus reaches the top of the cartridge resin bed.

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11. Disconnect the syringe and the adapter and wash the cartridge with 3 mL of molecular biology grade water. Reattach the adapter and syringe and gently push the liquid until the meniscus reaches the top of the cartridge resin bed. 12. Remove the waste beaker and place a sterile 1.5-mL Eppendorf tube below the cartridge to collect the purified probe. 13. Disconnect the syringe and adapter. 14. Pipet 500 μL of 50% methanol in water into the top of the cartridge. Reattach the adapter and the syringe and elute the purified probe into the Eppendorf tube. (Continue eluting until no further liquid escapes the cartridge.) 15. Use the purified probe immediately for a dot blot or Southern blot hybridization or store at –20°C for up to 1 wk. All probes should have specific activity measurements of at least 107 cpm mg−1 .

3.3.5. Hybridization of Dot and Southern Blot Membranes to Radiolabeled DNA Probes Hybridization of multiple membranes to a single probe can be performed simultaneously in a single hybridization bottle, provided that the quantity of probe and hybridization solution are adjusted accordingly and a mesh sheet is used to separate the membranes. (See Note 12.) 1. Carefully place the membrane in a hybridization bottle, add 7 mL of warm (68°C) PerfectHyb Plus Hybridization Buffer (Sigma-Aldrich), and place the bottle in a rotating hybridization oven for 15 min at 68°C. This is the prehybridization step. 2. Pour the prehybridization solution off of the membrane and discard. 3. Add 100 ng of radiolabeled probe to 7 mL of warm (68 °C) PerfectHyb Plus and mix gently. 4. Add the PerfectHyb Plus containing the probe to the hybridization bottle containing the membrane. 5. Place the hybridization bottle in a rotating hybridization oven for 16 h at 68 °C. 6. Pour off the hybridization solution and add 50 mL of low-stringency washing solution. Wash the membrane for 10 min at room temperature by gently shaking the hybridization bottle so that the wash solution flows gently over the membrane. Discard the wash solution and repeat this step with fresh low-stringency wash solution. 7. Discard the low-stringency wash solution and add 50 mL of moderate stringency wash solution to the hybridization bottle. Place the bottle in the rotating hybridization oven for 15 min at 42°C. Discard the moderate-stringency wash solution and repeat this step. 8. Rinse the membrane briefly with 100 mL of 2× SSC. 9. Gently remove the membrane from the hybridization bottle, wrap it tightly in transparent plastic wrap, and expose the membrane to a phosphor imaging plate for 20 min. Longer exposures may be performed if desired. 10. Use a phosphor imaging system to capture the blot image from the phosphor imaging plate. Alternatively, X-ray film may be used for imaging, however

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exposure time must be increased to 1–3 h and an X-ray developer must be used. Examples of the results produced from both dot blots and Southern blots are shown in Fig. 2.

3.3.6. Stripping of Dot and Southern Blot Membranes 1. Place the wet membrane in a glass dish and pour 300–400 mL of boiling 0.1% SDS over it. Gently shake the membrane until the 0.1% SDS reaches room temperature. 2. Remove the membrane and wrap it tightly in plastic wrap. The membrane is now ready to be rehybridized, or it can be stored wet at 4°C for a few days.

3.4. Total RNA Extraction and 16S rRNA Dot-Blot Hybridization After an SSH experiment, it can be useful to confirm the presence of specific microorganisms in the environmental sample and determine the proportion of their 16S rRNA relative to the total 16S rRNA in the tester and driver samples. For this purpose, DNA oligonucleotide probes are designed to target the populations of interest. In the study exemplified here, it was desirable to determine the proportions of Bacteria and Archaea in the tester and driver rumen samples, and therefore domain-specific 16S rRNA probes were used. The rRNAs from reference cultures must be used as controls to ensure that each probe is hybridizing properly and to assist with quantification. Reference cultures should be chosen appropriately for the environmental metagenomes under scrutiny. One reference culture should be obtained for each probe and defined quantities of RNA should be spotted on the blot. Measuring the intensity of hybridization signal from reference rRNA allows estimates to be made of the quantity of rRNA from environmental samples. The procedure for extraction of total RNA described below is slightly modified from the methods of Stahl et al. (1988) and Lin et al. (1997) (52,53). (For further reference information on Northern blotting consult Current Protocols in Molecular Biology (51)). 3.4.1. Total RNA Extraction 1. Combine 0.8 g of each environmental sample of interest with 400 μL of 0.1 M sodium phosphate buffer, pH 6.0, 400 μL of acetate buffer pH 5.1, 100 μL of 20% SDS (w/v), 500 μL of phenol (buffered to pH 5.1), and 0.1 g of sterile zirconium beads (0.1 mm diameter) in a 2.2-mL Eppendorf tube. 2. For reference microorganisms, centrifuge to obtain pellets from 40-mL cultures (E. coli BW13711 or M. acetivorans C2A in this example) and combine 0.8 g from each with 400 μL of 0.1 M sodium phosphate buffer, pH 6.0, 400 μL of acetate buffer pH 5.1, 100 μL of 20% SDS (w/v), 500 μL of phenol (buffered to pH 5.1), and 0.1 g of sterile zirconium beads (0.1 mm diameter) in a 2.2-mL Eppendorf tube. 3. Incubate all samples for 2 min at 60°C.

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4. Homogenize all samples in a mini-beadbeater reciprocating shaker (Biospec Products, Bartlesville, OK). 5. Repeat steps 3 and 4 twice. 6. Centrifuge samples at 4°C at 10,000g for 5 min. Transfer supernatants to clean tubes. 7. To each sample, add 1 mL of phenol (buffered at pH 5.1), and shake in the minibeadbeater for 2 min. Centrifuge samples for 5 min at 4°C and 10,000g. Transfer the aqueous phases to clean tubes. 8. Add 1 mL of phenol–chloroform–isoamyl alcohol (125:24:1) to each sample and shake in the mini-beadbeater for 2 min. Centrifuge samples for 5 min at 4°C and 10,000g. Transfer the aqueous phases to clean tubes. Repeat this step once. 9. Add 1 mL of chloroform–isoamyl alcohol (24:1) to each sample and shake in the mini-beadbeater for 2 min. Centrifuge samples for 5 min at 4°C and 10,000g. Transfer the supernatant from each sample to clean tubes. 10. Precipitate total nucleic acids from samples by addition of 50 μL of sodium acetate (3 M) and 1 mL of cold absolute ethanol to the supernatant followed by chilling at –80 C for 30 min. 11. Centrifuge samples at 4°C for 15 min at 10,000g to pellet nucleic acids. 12. Wash pellets with 100 μL of 80% ethanol, dry pellets, and suspend them in 100 μL of RNase-free water. 13. Treat samples with 1 U/μg of RQ1 RNase-free DNase (Promega) for 30 min at 37 C followed by a second extraction with phenol, phenol–chloroform–isoamyl alcohol, and chloroform–isoamyl alcohol performed in the same manner as steps 7–9. 14. Nucleic acids were precipitated, pelleted, and washed again in the same manner as steps 10–12 but suspend in 40 μL of RNase-free water. 15. Estimate the concentration of RNA in each sample by measuring the absorbance at 260 nm and confirm the concentration by electrophoresing small aliquots on 1% agarose gels run in 0.5× TBE buffer. Adjust the concentration of each sample to 100 ng/uL−1 .

3.4.2. Radiolabeling of DNA Probes for RNA Dot Blots End labeling of DNA oligonucleotide probes can be performed using polynucleotide kinase (Invitrogen) and [-32 P]ATP. The amount of polynucleotide kinase used in the labeling experiment should be calculated considering a 1:1 molar ratio of probe to [-32 P]ATP label. 1. In a 1.5-mL Eppendorf tube, combine the following: 3.0 μL 1.5 μL Calculated amount Calculated amount 1.0 μL

10× Kinase buffer 1% IGEPAL Oligonucleotide probe [-32 P]ATP Polynucleotide kinase

Add RNase-free water to bring the total volume up to 30 μL.

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2. Incubate the probe at 37°C for 30 min. 3. Inactivate polynucleotide kinase by heating probe to 65°C for 2 min. 4. Place probe on ice and purify using a Nensorb column as described for probes in Subheading 3.3.4. 5. Use the purified probe immediately for a dot-blot hybridization or store at –20°C for up to 1 wk. All probes should have specific activity measurements of at least 107 cpm mg−1 .

3.4.3. Preparation of RNA Dot-Blot Membrane and Hybridization of Membrane to Radiolabeled DNA Probes Hybridization temperature and the temperature for the final membrane wash should be determined by the sequence and length of the probe used. Calculations for determining Tm of probes and appropriate hybridization and wash temperatures can be found in Current Protocols in Molecular Biology (51). For hybridization of multiple membranes simultaneously to the same probe see Note 12. 1. Denature reference and environmental sample RNA by adding 3 volumes of 2% glutaraldehyde to 1 volume of RNA sample and incubate at room temperature for 10 min. Place samples on ice and dilute samples to desired concentrations with RNase- free water. 2. Prepare nylon membrane and dot blot manifold as described above in Subheading 3.3.2. Prewash the membrane by adding 500 μL of sterile RNase free water to each well (see Note 11) and allow the vacuum to pull the water slowly through the membrane. 3. Spot denatured RNA samples directly onto the nylon membrane, remove membrane from manifold, dry, and crosslink as described in Subheading 3.3.2. RNA from each sample should be spotted onto the membrane in triplicate, in amounts of 100 ng, 50 ng, 20 ng, 5 ng, 2 ng, 0.5 ng, and 0.2 ng of RNA per spot (each spotted in a volume of 100 μL). 4. Carefully place the membrane in a hybridization bottle, add 7 mL of warm (40°C) PerfectHyb Plus Hybridization Buffer (Sigma-Aldrich), and place the bottle in a rotating hybridization oven for 2 h at 40°C. This is the prehybridization step. 5. During the last 10 min of prehybridization, denature probe by heating for 10 min to 100°C in a boiling water bath or a heat block. 6. Pour the prehybridization solution off of the membrane and discard. 7. Add 100 ng of denatured, radiolabeled probe to 7 mL of warm (40°C) Perfect-Hyb Plus and mix gently. 8. Add the PerfectHyb Plus containing the probe to the hybridization bottle containing the membrane. 9. Place the hybridization bottle in a rotating hybridization oven for 16 h at 40°C or appropriate hybridization temperature calculated based on the sequence and length of the probe.

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10. Pour off the hybridization solution and add 50 mL of warm (40°C) RNA dot blot washing solution. Place the bottle in the rotating hybridization oven for 1 h at 40°C. Discard the wash solution and repeat this step. 11. Pour off the wash solution and add 50 mL of fresh warm (40°C) wash solution. Place the bottle in the rotating hybridization oven for 30 min at an appropriate temperature calculated for the specific probe. Discard the wash solution, wrap wet membrane tightly in transparent plastic wrap, and expose the membrane to a phosphor imaging plate as described in Subheading 3.3.5. 12. Capture an image of the blot using a phosphor imaging system or X-ray film processor as described for dot blots in Subheading 3.3.5. (Fig. 3). 13. Utilize a software program such as ImageQuant (Fujifilm) to determine densitometry measurements on each dot blot.

3.5. DNA Sequencing and Bioinformatic Analsyses Following Southern blot confirmation of the uniqueness of the cloned subtracted products relative to one metagenome versus another, the suppressive subtractive hybridization library should be sequenced. Descriptive protocols for DNA sequencing and subsequent bioinformatic analyses is beyond the scope of this chapter and are better described in other volumesof the Methods in Molecular Biology series. 4. Notes 1. To suspend precipitated DNA and mix reactions, gently pipet them up and down and then centrifuge the tube briefly to deposit contents at the bottom. For phenol– chloroform extractions vortex-mix the reactions. Always add enzymes to reaction mixtures last, and thoroughly incorporate the enzyme by gently pipetting the reaction mixture up and down. Do not increase the amount of enzyme added or the concentration of DNA in the reactions. The amounts and concentrations have been carefully optimized. 2. The choice of the appropriate restriction enzyme should depend on its ability to digest metagenomic DNA to produce fragments of optimal length. To determine if a sample is completely digested, remove small aliquots of DNA after 1 and 3 h of digestion and compare the samples on an agarose gel. If the DNA size distribution for both samples is the same (including relative to the E. coli positive control), then the digestion is complete. If the size distribution is not reduced after AluI digestion or is longer than 2 kb, your metagenomic DNA cannot be efficiently cut by (a) any restriction enzyme because of some inhibiting impurities in the DNA sample, or (b) the restriction enzyme because of the DNA’s nucleotide composition (unlikely with metagenomic DNA). The described protocol here uses AluI, but there are other four-base, blunt-end producing restriction enzymes: HaeIII and RsaI. A combination of several sixbase, blunt-cutting restriction enzymes can also be used to obtain the desired level

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Fig. 3. 16S rRNA dot-blot hybridization analysis of rumen samples from hay fed steers 218 and 256. Total RNA from rumen samples 218 and 256, E. coli, and Methanosarcina acetivorans was spotted on nylon membranes in triplicate in concentrations of 100 ng, 50 ng, 20 ng, 10 ng, 5 ng, 0.5 ng and 0.2 ng. Membranes shown in (A) and (B) were hybridized with a universal 32 P–labeled probe specific for all 16S rRNA (S-*-Univ-1392-a-A15). Bacterial and Archaeal reference organisms, E. coli and M. acetivorans were used to normalize the concentrations of rRNA from each rumen sample. Blots were then probed with Bacterial-specific (S-D-Bact-0338-a-A-18) and Archaealspecific (S-D-Arch-0915-a-A-20) 32 P–labeled probes shown in (C) and (D), respectively.

of DNA digestion. In this case, perform several standard digestions with different restriction enzymes: a. Add the following reagents into each reaction tube: Metagenomic DNA (0.2 μg) 10× restriction enzyme buffer Restriction enzyme (1 U) Sterile, deionized H2 O Total volume

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3. If a white pellet is not visible, try placing the tubes at –80°C for 15 min and then centrifuging again. Alternately, addition of a small amount of ethanol (in 100-μL increments) can also assist in the precipitation. 4. The tester control DNA contains one copy of HaeIII digested øX174 DNA per E. coli genome (the E. coli genome is about 5 × 106 base pairs). After subtraction of the tester DNA against the driver DNA, the primary bands produced in the final PCR should correspond to these control fragments of the øX174/HaeIII digest. 5. Successful adaptor ligation with the control E. coli genomic DNA does not guarantee a successful ligation with your metagenomic DNA; impurities in the DNA preparation may inhibit the ligase. The experiment only allows an indication that the basic steps of the adaptor ligation were successfully conducted. In addition, you should also include a standard PCR control to ensure that the Taq DNA polymerase used in the PCR is performing efficiently. 6. The cycling parameters and incubations in this protocol were performed on a Applied Biosystems GeneAmp® System 9700 using TaKaRa Ex Taq DNA polymerase. Following the procedures described here using different thermal cyclers, Taq DNA polymerases, and templates may require optimization. A hot start MUST be used to reduce nonspecific DNA synthesis during the set-up of the PCR (54) but other approaches can achieve the desired effect (e.g., TaqStart Antibody (55)). We used a manual hot start in our work with metagenomic DNA by addition of the Taq DNA polymerase during the initial denaturation step. 7. The PCR product generated using one gene-specific primer (23S RNA forward primer) and PCR primer 1 should be about the same intensity as the PCR product amplified using two gene specific primers (23S RNA forward and reverse primers). If the intensity of bands corresponding to these PCR products differs by more than 4-fold, then the efficiency of your ligation is less than 25%. This will significantly reduce your subtraction efficiency, and we recommend that you repeat the ligation reaction. If the PCR product amplified using the 23S RNA forward primer and PCR Primer 1 contains a 574-bp band in addition to the expected 374-bp band, your DNA is not fully digested. If there is a significant amount of this 574-bp product, we recommend that you repeat the RsaI digestion. If your ligation did not work well, you should repeat the ligation before performing the hybridizations. 8. The CLONTECH PCR-Select™ Bacterial Genome Subtraction Kit was configured to work with genomic DNA obtained from pure cultures of microorganisms. Therefore, troubleshooting the procedure when working with pure cultures is much more

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straightforward, as in the case of optimizing the hybridization temperature used. This variable may warrant investigation if information is available as to the composition of the metagenome being studied. The hybridization temperature, for example, is dependent on the GC content of a particular genome. The optimal temperature for genomic DNA with average GC content (40–51%) is 63°C. If the GC content of the genomic DNA is unusually low (e.g., the genome of Clostridium perfringens has a GC content of 26.5%), then the hybridization temperature should be lowered. The PCR mixture should now be enriched for metagenome tester-specific DNA. The products should be cloned to make a subtracted DNA library. The amplified products can be directly inserted into a T/A cloning vector. If you experience arcing during electroporation then precipitate the remainder of the TOPO® Cloning reaction and suspend in water before trying again. Another factor in arcing of your samples during electroporation may be the volume of cells. They should be between 50 and 80 μL (0.1 cm cuvettes) or 100-200 μL (0.2 cm cuvettes). A solution of 0.004% bromphenol blue in water can be used as the prewash water in dot blotting. The dye will slightly color the membrane in the areas where samples will be spotted. If the manifold is assembled and working properly the blue color will remain in areas where it is spotted. Bleeding of the color across the membrane indicates that the manifold is not assembled properly. In addition, the colored dye spots will allow you to easily discern which side of the membrane contains your samples after the dot blot manifold is disassembled. When hybridizing multiple membranes simultaneously, increase the amount of probe and hybridization solution proportionally. To prevent the membranes from overlapping within the hybridization bottle, cut a piece of mesh such as Denville Scientific mesh (Denville Scientific, Metuchen, NJ) to the appropriate size and lay it between the membranes.

Acknowledgments This work was supported by USDA NRI grant no. 970–326, and by the Agricultural Experimental Station of the University of Illinois. The authors gratefully acknowledge Professor Adrian Egan of the University of Melbourne for spawning this idea at the RRI-INRA meeting in Aberdeen, Scotland. References 1. Doolittle, R. F. (1997) Microbial genomes opened up. Nature 392, 339–42. 2. Fraser, C. M., Eisen, J. A., and Salzberg, S. L. (2000) Microbial genome sequencing. Nature 406, 799–803. 3. Akopyants, N. S., Fradkov, A., Diatchenko, L., Hill, J. E., Siebert, P. D., Lukyanov, S. A., Sverdlov, E. D., and Berg, D. E. (1998) PCR-based subtractive hybridization and differences in gene content among strains of Helicobacter pylori. Proc. Natl. Acad. Sci. USA 95, 13108–13113.

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18. Lamar, E. E., and Palmer, E. (1984) Y-encoded, species-specific DNA in mice: evidence that the Y chromosome exists in two polymorphic forms in inbred strains. Cell 37, 171–177. 19. Nussbaum, R. L., Lesko, J. G., Lewis, R. A., Ledbetter, S. A., and Ledbetter, D. H. (1987) Isolation of anonymous DNA sequences from within a submicroscopic X chromosomal deletion in a patient with choroideremia, deafness, and mental retardation. Proc. Natl. Acad. Sci. USA 84, 6521–6525. 20. Straus, D. and Ausubel, F. M. (1990) Genomic subtraction for cloning DNA corresponding to deletion mutations. Proc. Natl. Acad. Sci. USA 87, 1889–1893. 21. Wieland, I., Bolger, G., Asouline, G., and Wigler, M. (1990) A method for difference cloning: gene amplification following subtractive hybridization. Proc. Natl. Acad. Sci. USA 87, 2720–2724. 22. Duguid, J. R., and Dinauer, M. C. (1990) Library subtraction of in vitro cDNA libraries to identify differentially expressed genes in scrapie infection. Nucleic Acids Res. 18, 2789–2792. 23. Cecchini, E., Dominy, P. J., Geri, C., Kaiser, K., Sentry, J., and Milner, J. J. (1993) Identification of genes up-regulated in dedifferentiating Nicotania glauca pith tissue, using an improved method for constructing a subtractive cDNA library. Nucleic Acids Res. 21, 5742–5747. 24. Hubank, M., and Schatz, D. G. (1999) cDNA representational difference analysis: a sensitive and flexible method for identification of differentially expressed genes. Methods Enzymol. 303, 325–349. 25. Ji, W., Wright, M. B., Cai, L., Flament, A., and Lindpaintner, K. (2002) Efficacy of SSH PCR in isolating differentially expressed genes. BMC Genomics 3, 12. 26. Diatchenko, L., Lau, Y. F., Campbell, A. P., Chenchik, A., Moqadam, F., Huang, B., Lukyanov, S., Lukyanov, K., Gurskaya, N., Sverdlov, E. D., and Siebert, P. D. (1996) Suppression subtractive hybridization: a method for generating differentially regulated or tissue-specific cDNA probes and libraries. Proc. Natl. Acad. Sci. USA 93, 6025–6030. 27. Ermolaeva, O. D. and Wagner, M. C. (1995) SUBTRACT: a computer program for modeling the process of subtractive hybridization. CABIOS 11, 457–462. 28. Ermolaeva, O. D., Lukyanov, S. A., and Sverdlov, E. D. (1996) The mathematical model of subtractive hybridization and its practical application. Proc. Int. Conf. Intell. Syst. Mol. Biol. 4, 52–58. 29. Ermolaeva, O. D., and Sverdlov, E. D. (1996) Subtractive hybridization, a technique for extraction of DNA sequences distinguishing two closely related genomes: critical analysis. Genet. Anal. 13, 49–58. 30. Emmerth, M., Goebel, W., Miller, S. I., and Hueck, C. J. (1999) Genomic subtraction identifies Salmonella typhimurium prophages, F-related plasmid sequences, and a novel fimbrial operon, stf, which are absent in Salmonella typhi. J. Bacteriol. 181, 5652–5661. 31. Ferreira, H., Rodrigues Neto, J., Goncalves, E. R., and Rosato, Y. B. (1999) A simplified subtractive hybridization protocol used to isolate DNA sequences specific to Xylella fastidiosa. Microbiology 145, 1967–1975.

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32. Sawada, K., Kokeguchi, S., Hongyo, H., Sawada, S., Miyamoto, M., Maeda, H., Nishimura, F., Takashiba, S., and Murayama, Y. (1999) Identification by subtractive hybridization of a novel insertion sequence specific for virulent strains of Porphyromonas gingivalis. Infect. Immun. 67, 5621–5625. 33. Vinnemeier, J., and Hagemann, M. (1999) Identification of salt-regulated genes in the genome of the cyanobacterium Synechocystis sp. strain PCC 6803 by subtractive RNA hybridization. Arch. Microbiol. 172, 377–386. 34. Lai, Y. C., Yang, S. L., Peng, H. L., and Chang, H. Y. (2000) Identification of genes present specifically in a virulent strain of Klebsiella pneumoniae. Infect. Immun. 68, 7149–7151. 35. Agron, P. G., Walker, R. L., Kinde, H., Sawyer, S. J., Hayes, D. C., Wollard, J., and Andersen, G. L. (2001) Identification by subtractive hybridization of sequences specific for Salmonella enterica serovar enteritidis. Appl. Environ. Microbiol. 67, 4984–4991. 36. Ahmed, I. H., Manning, G., Wassenaar, T. M., Cawthraw, S., and Newell, D. G. (2002) Identification of genetic differences between two Campylobacter jejuni strains with different colonization potentials. Microbiology 148, 1203–1212. 37. Herd, M., and Kocks, C. (2001) Gene fragments distinguishing an epidemicassociated strain from a virulent prototype strain of Listeria monocytogenes belong to a distinct functional subset of genes and partially cross-hybridize with other Listeria species. Infect Immun 69, 3972–3979. 38. Pettigrew, M. M., Foxman, B., Marrs, C. F., and Gilsdorf, J. R. (2002) Identification of the lipooligosaccharide biosynthesis gene lic2B as a putative virulence factor in strains of nontypeable Haemophilus influenzae that cause otitis media. Infect. Immun. 70, 3551–3556. 39. Pradel, N., Leroy-Setrin, S., Joly, B., and Livrelli, V. (2002) Genomic subtraction to identify and characterize sequences of Shiga toxin-producing Escherichia coli O91:H21. Appl. Environ. Microbiol. 68, 2316–2325. 40. Parsons, Y. N., Banasko, R., Detsika, M. G., Duangsonk, K., Rainbow, L., Hart, C. A., and Winstanley, C. (2003) Suppression-subtractive hybridisation reveals variations in gene distribution amongst the Burkholderia cepacia complex, including the presence in some strains of a genomic island containing putative polysaccharide production genes. Arch. Microbiol. 179, 214–223. 41. Nesbø, C. L., and Doolittle, W. F. (2003) Targeting clusters of transferred genes in Thermotoga maritima. Environ. Microbiol. 5, 1144–1154. 42. Antonopoulos, D. A., Nelson, K. E., Morrison, M., and White, B. A. (2004) Strain specific genomic regions of Ruminococcus flavefaciens FD-1 as revealed by combinatorial random-phase genome sequencing and suppressive subtractive hybridization. Environ. Microbiol. 6, 335–346. 43. Frohme, M., Camargo, A. A., Czink, C., Matsukuma, A. Y., Simpson, A. J., Hoheisel, J. D., and Verjovski-Almeida, S. (2001) Directed gap closure in largescale sequencing projects. Genome Research 11, 901–903. 44. Kocherginskaya, S. A., Aminov, R. I., and White, B. A. (2001) Analysis of the rumen bacterial diversity under two different diet conditions using

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17 16S rRNA Targeted DGGE Fingerprinting of Microbial Communities Vesela A. Tzeneva, Hans G. H. J. Heilig, Wilma Akkermans van Vliet, Antoon D. L. Akkermans, Willem M. de Vos, and Hauke Smidt

Summary The past decades have seen the staggering development of molecular microbial ecology as a discipline that uses the detection of so-called biomarkers to monitor microbial communities in environment samples. A variety of molecules can be used as biomarkers, including cell-wall components, proteins, lipids, DNA or RNA. Especially, the application of small subunit ribosomal RNA (rRNA) and the corresponding genes have proven invaluable for advances in microbial ecology. Several types of fingerprinting methods have been developed for the description of microbial communities in environmental samples. Among the most commonly used approaches is denaturing gradient gel electrophoresis (DGGE) of PCR-amplified fragments. DGGE allows separation of DNA fragment mixtures of equal length depending on their sequence. The separation is based on their sequence-specific melting point in a polyacrylamide gel with a gradient of a denaturant chemical (generally a combination of urea and formamide). DGGE allows for a rapid analysis and comparison of microbial communities. Compositional diversity can be visualized using DGGE where each band in principle represents a bacterial phylotype. After staining bands are visualized at each position in the gel where DNA molecules stopped migration. In principle, DGGE fingerprinting can resolve single base pair differences. Key Words: Bacillus benzoevorans-relatives; DGGE; fingerprinting; 16S rRNA genes; soil microorganisms.

From: Methods in Molecular Biology, vol. 410: Environmental Genomics Edited by: C. Cristofre Martin © Humana Press, Totowa, NJ

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1. Introduction Microbial communities are pivotal to development and function of practically any environment on and even beyond the planet Earth, and the astounding development of molecular microbial ecological approaches targeting a variety of cellular biomarkers now allows for monitoring of community composition and functionality independent of the ability to cultivate microorganisms (1,2). Targeted biomarkers can be any biological component that indicates the presence of a particular group of microorganisms at different levels of taxonomic resolution, and include cellular components such as cell-wall constituents, proteins, lipids, DNA, or RNA. These molecules can be detected even when the cell is not alive. This enables us to study microbial communities and to follow their development under changing environmental conditions, even from historical samples, archived, e.g., as air-dried specimen in the past for future analyses, or retrieved from the paleome, completely inaccessible to cultivation-based efforts (3–5). Convenient for microbial ecology is the application of small subunit ribosomal RNA (SSU rRNA; 16S rRNA for Bacteria and Archaea, 18S rRNA for Eukarya) and the corresponding genes as biomarkers. Targeting these molecules, a variety of complementary molecular techniques can be applied for monitoring of the total bacterial, archaeal, or eukaryal communities, as well as the specific groups of interest, in different environments. Molecular fingerprinting methods, mostly targeting the SSU rRNA gene as a universal phylogenetic marker, are frequently used in molecular microbial ecology to rapidly monitor differences in microbial community composition in space and/or time. In combination with statistical analyses, these techniques are powerful tools to investigate how changing environmental factors can influence microbial community composition (6,7). Several types of gradient gel electrophoresis have been validated for the description of microbial communities in environmental samples. Most commonly used are denaturing gradient gel electrophoresis (DGGE) and temperature gradient gel electrophoresis (TGGE), but recently also temporal temperature gradient gel electrophoresis (TTGE), which combines the principles of both DGGE and TGGE (8). The first application of DGGE in microbial ecology was used for the profiling of bacterial diversity in a marine ecosystem (9). The gel electrophoresis techniques are useful for rapidly comparing microbial communities from different environments and monitoring changes in the composition of abundant members of a specific community over time. DGGE is a technique that allows separation of DNA fragment mixtures of equal length depending on their sequence (10). The separation is based on their sequence-specific melting point in a polyacrylamide gel with a gradient of a denaturant chemical (generally urea and formamide). Complete denaturation

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of the DNA fragments is prevented by addition of a so-called GC- clamp to the 5 -end of the molecule. This GC- clamp is 30–50 base pairs (bp) long, and is added to one of the primers used for PCR amplification of the target gene fragment prior to DGGE analysis. In principle, a single base pair difference at any given position throughout the amplified fragment can be separated for PCR products not longer than 500 bp. Fragments with different sequences will stop migration at different positions in the denaturing gradient. 2. Materials 2.1. DNA Isolation Using the Fast DNA® SPIN® Kit The Fast DNA® SPIN® Kit (For Soil) (Q BIOgene, Cambs, United Kingdom) is a commercially available DNA isolation kit and commonly used for a wide variety of environmental matrices, including soil, sediment, and feces, at our laboratory. However a large number of other DNA isolation methods and protocols are available. 2.2. Polymerase Chain Reaction (PCR) PCR can be performed with Taq polymerase (Invitrogen) in a Whatman Biometra Thermocycler (Göttingen, Germany). (See Note 1.) Prepare a master mix with the following components, without the template (DNA): 1. 2. 3. 4. 5. 6. 7.

5 μL of 10× PCR buffer (10× buffer: 200 mM Tris-HCl, pH 8.4, 500 mM KCl). 3 μL of MgCl2 (50 mM). 1 μL of forward primer (10 μM). 1 μL of reverse primer (10 μM). 1 μL of dNTP (10 mM). 0.25 μL of Taq DNA polymerase (5 U/μL). 37.75 μL of MilliQ 49 μL of reaction volume

Multiply the amounts mentioned above by the number of reactions needed + 1 extra, and aliquot the master mix in 0.2-mL PCR microtubes (49 μL in every tube). Finally add the template DNA (1 μL). The final amount of template DNA in each reaction should be 10–100 ng. Place the tubes in the PCR machine and start the required program. 2.3. DGGE 2.3.1. DGGE Technical Components 1. Large and small glass plate (20-cm system; Bio-Rad). 2. Gelbond (Tebu-bio, www.tebu-bio.com)

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3. Bio-Rad DGGE technical equipment, including: a. The temperature control module containing the heater, stirrer, pump and electrophoresis leads that operate the system. b. The electrophoresis tank—a reservoir for the running buffer. c. The sandwich core—holds one gel assembly on each side. When attached each gel assembly forms one side of the upper buffer chamber. The inner plate is clamped against a rubber gasket on the core to provide a greaseless seal for the upper chamber. d. The sandwich clamps (20-cm system). e. Spacer set. f. Combs. 4. Power supply (Bio-Rad). 5. Pump and mixing chamber (Bio-Rad).

2.3.2. DGGE Stock Solutions 1. 100% denaturant, 8% PAG solution: 200 mL of 40% acryl–bis-acrylamide 37.5:1, 400 mL of formamide, 10 mL of 50× TAE buffer, 20 mL of glycerol (see Note 2), 421.6 g of urea. Carefully heat to hand-warm, add stirrer, and dissolve. Adjust to final volume of 1 liter with demi-water. Store in the dark at room temperature. 2. 0% Denaturant, 8% PAG solution: 200 mL of 40% acryl–bis-acrylamide 37.5:1, 10 mL of 50× TAE buffer, 20 mL of glycerol (see Note 2). Adjust to final volume of 1 L with demi-water. Store in the dark at room temperature. 3. 50× TAE solution: 242 g of Tris base, 57.1 mL of glacial acetic acid, 100 ml of 0.5 M EDTA, pH 8.0.

2.3.3. DGGE Staining Solutions 1. Cairns’ 8× fixation solution: 200 mL of 96% ethanol, 10 mL of acetic acid, 40 mL of demi-water. Add 50 mL of 8× Cairns’ fixing solution to 350 mL of demi-water. 2. Silver staining solution: 0.4 g of AgNO3 in 1× Cairns’ fixing solution (200 mL). 3. Developer solution: A spatula tip of NaBH4 (∼10 mg), 250 mL 1.5 % NaOH solution, 750 μL of formaldehyde. 4. Cairn‘s preservation solution: 250 mL of 96% ethanol, 100 mL of glycerol, 650 mL of demi-water.

3. Methods DGGE is a well-established molecular fingerprinting technique in molecular microbial ecology (11,12). DGGE is a technique by which microbial communities can be analysed in a rapid way. Complex microbial communities can be visualized using DGGE where each band in principle represents a bacterial phylotype. One of the limitations of the technique is, however, that only a

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restricted amount of genetic information is being used (e.g., 16S rRNA genes). The DGGE principle consists of the separation of PCR amplicons of the same length, based on their sequence. Double-stranded DNA dissociates under the influence of denaturing chemicals. DNA migrates to the positive pole in an electrical field and in the same direction the percentage of denaturing compounds of the polyacrylamide gel increases linearly (Fig. 1). During electrophoresis PCR products migrate to the positive pole, and are melted at a specific position, which is sequence-specific (dissociation of the two nucleotide strands). A GC-clamp introduced to either the 5 - or 3 -end of the PCR product during amplification prevents complete denaturation, resulting in a fork-like structure that practically stops migrating. After staining bands are visualized at the position in the gel where DNA molecules stopped migration. In principle, DGGE fingerprinting can resolve single base pair differences. 3.1. DNA Isolation DNA is isolated directly from soils (or other environmental samples) (1 g) using, e.g., the Fast DNA® SPIN® Kit (For Soil) (Q BIOgene, Cambs, UK) according to the manufacturer‘s instructions. The kit has been designed for efficient lysis of all microorganisms including historically difficult sources such as bacterial spores and endospores, gram-positive bacteria, yeast, algae, nematodes, and fungi.

Fig. 1. General principle of DGGE.

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3.2. PCR Programs Indicated below are a PCR cycling parameters that have been optimized for the primer sets provided in Table 1. Choose the appropriate program for the group of organisms that you are working with. 1. Bacterial community DGGE-PCR: Predenaturation

2 min

95ºC

Denaturation Hybridization Elongation

30 s 40 s 1 min

95ºC 56ºC 72ºC

Postelongation

5 min

72ºC

2. Archaeal DGGE-PCR: Predenaturation 5 min

94ºC

Denaturation Hybridization Elongation

30 s 40 s 60 s

94ºC 52ºC 68ºC

Postelongation

5 min

68ºC

35 cycles

35 cycles

3. Eukaryotic community DGGE-PCR: Predenaturation

5 min

94ºC

Denaturation Hybridization Elongation

30 s 45 s 130 s

94ºC 56ºC 72ºC

Postelongation

10 min

72ºC

35 cycles

3.3. DGGE-PCR Primers A selection of general and group-specific DGGE-PCR primers is provided in Table 1. (See Note 3.) 3.4. DGGE 3.4.1. Gel Sandwich Preparation (See Note 4) 1. Clean one large and a smaller glass plate with soap, dry them, and clean them again with 96% ethanol. (See Note 5.) 2. Cut the gel bond to the size of the larger glass plate. (See Note 6.) 3. Add some water to the surface of the large glass plate. 4. Place the gel bond hydrophobic side down on this glass plate (you can easily check this by adding a drop of water on the gelbond, it will roll off easily).

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Table 1 Most Commonly Used DGGE-PCR Primers for Different Groups of Microorganisms Primers Bacteria Bact-968-GC-F

L1401-R Bact-0357-F-GC

Bact-0518-R Bact-0954-F-GC

Bact-1369-R Archaea Arch-109-F Uni-515-GC-R

Eukaryotes Euk1A Euk516r-GC

Sequence

Reference

5 - CGC CCG GGG CGC GCC CCG GGC GGG GCG GGG GCA CGG GGG GAA CGC GAA CCT TAC-3 5 - GCG TGT GTA CAA GAC CC-3 5 -CCG GGG GCG CGC CCC GGG CGG GGC GGG GGC ACG GGG GGC CTA CGG GAG GCA GCA G-3 5 -ATT ACC GCG GCT GCT GG-3 5 -CGC CGG GGG CGC GCC CCG GGC GGG GCG GGG GCA CGG GGG GGC ACA AGC GGT GGA GCA TGT GG-3 5 -GCC CGG GAA CGT ATT CAC CG-3

(17)

5 -ACT GCT CAG TAA CAC GT-3 5 -CGC CCG GGG CGC GCC CCG GGC GGG GCG GGG GCA CGG GGG GAT CGT ATT ACC GCG GCT GCT GGC AC-3

(19) (20)

5 -CTG GTT GAT CCT GCC AG-3 5 - CGC CCG GGG CGC GCC CCG GGC GGG GCG GGG GCA CGG GGG GAC CAG ACT TGC CCT CC-3

(21) (21)

(17) (18)

(18) (18)

(18)

5. Fix the gel bond, without removing the paper sheet, with a roller, then take of the paper sheet. 6. Dry the gel bond carefully with some tissues. 7. Clean a set of spacers with 96% ethanol, and place them on left and right site on the gel bond. 8. Place the smaller glass plate on top. 9. Add the clamps to the sides of the glass sandwich, and place in the sandwichholder. For proper distance between the spacers slide the plastic card between the glass plates. 10. Press the spacers down and fasten the screws on the clamps. 11. Place the sandwich (plate cassette) on top of the rubber gasket and press down the handles. 12. Apply the gel “plug” using a pipet.

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3.4.2. Gel “Plug” Preparation A 0% denaturant PAG gel “plug” is cast at the bottom of the DGGE plate cassette prior to making the main resolving gradient gel—preventing leakage from the bottom of the plates. The method for making the gel “plug” is as follows: 1. Make sure that the casting stand is level; adjust screw feet on the stands if necessary. 2. Prepare the “plug” solution as follows (volumes for one gel): 1.5 mL 4.5 μL 15 μL

0% Denaturant, 8% PAG solution TEMED 10% (w/v) ammonium persulfate (APS)

3. Add 1 mL of the “plug” solution to the plate cassette, by pipetting the volume using the side of the spacer. (See Note 7.) 4. Leave the plug for approx 10 min to set. 5. Cast the gradient gel and stacking gel.

3.4.3. Preparing the Gel 1. Prepare on ice the gel solutions required, being high, low, and a stacking gel according to the mixing Table 2. (See Note 8.) 2. Rinse the gradient maker and tubes with demi-water, switch on the pump at running speed (19 mL/min) and drain the system. 3. Close the screw between the compartments of the gradient maker. 4. Dry the compartments with a tissue. 5. When the gel solutions are cooled, add 10% APS to the high and low percentage denaturant solutions. 6. Pour the high solution in the right compartment, and the low in the left. 7. Start the stirrer, open the screw and immediately start the pump at 4.5 mL/min. 8. Place the needle between the glass plates. 9. Remove the needle when the gel is poured, switch off the pump and transfer to the Erlenmeyer flask. 10. Rinse the compartments with demi-water and switch on the pump and drain the system. 11. Add the 10% APS to the stacking gel. 12. Close the screw between the compartments and add the stacking gel to the right compartment. 13. Place the needle between the glass plates and start the pump set at 3 mL/min. 14. When the stacking gel is poured place the comb that will form the slots carefully in the stacking gel. Avoid air bubbles because they will appear as dents in all your bands. 15. Leave the gel to polymerize for 1 h.

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Table 2 DGGE Gradient Mixing Table Gradient Comp. (%)

0% (Ml)

100% (mL)

FinalVolume (mL)

TEMED (μL)

10%APS (μL)

0 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

9 9.1 8.97 8.84 8.71 8.58 8.45 8.32 8.19 8.06 7.93 7.8 7.67 7.54 7.41 7.28 7.15 7.02 6.89 7.76 6.63 6.5 6.37 6.24 6.11 5.98 5.85 5.72 5.59 5.46 5.33 5.2

— 3.9 4.03 4.16 4.29 4.42 4.55 4.68 4.81 4.94 5.07 5.2 5.33 5.46 5.59 5.72 5.85 5.98 6.11 6.24 6.37 6.5 6.63 6.76 6.89 7.02 7.15 7.28 7.41 7.54 7.67 7.8

9 13 13 13 13 13 13 13 13 13 13 13 13 13 13 13 13 13 13 13 13 13 13 13 13 13 13 13 13 13 13 13

13 13 13 13 13 13 13 13 13 13 13 13 13 13 13 13 13 13 13 13 13 13 13 13 13 13 13 13 13 13 13 13

50 50 50 50 50 50 50 50 50 50 50 50 50 50 50 50 50 50 50 50 50 50 50 50 50 50 50 50 50 50 50 50

3.4.4. Running a Gel 1. Add fresh 0.5× TAE buffer to the buffer tank. 2. Switch on the Dcode at least 90 min before electrophoresis, so that the buffer can equilibrate at 60ºC.

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3. After 1 h of polymerization remove the comb carefully. 4. Rinse off nonpolymerized gel in and above the slots with demi-water and click the sandwich in the sandwich-holder. (See Note 9.) 5. Switch off the Dcode and remove the lid. 6. Take this sandwich cassette and place it into the buffer tank. 7. Switch on the Dcode until the upper buffer compartment is filled with buffer. 8. Switch off the Dcode and remove the lid. 9. Rinse all slots with syringe and needle filled with buffer. 10. Apply your samples in the slots. (See Note 10.) 11. Switch on the Dcode after returning the lid. 12. Switch on the power supply for 10 min at 200 V and lower to 85 V for 16 h.

3.5. Gel Staining (see Notes 11 and 12) 1. Switch off power. 2. Remove gel from the plates and put the gel in a stainless steel tray. 3. Add 200 mL of 1× Cairn‘s fixation solution and rock the tray for 3 min. Remove the solution and store for later application. 4. Add 200 mL of silver staining solution (caution: use gloves) to the tray and rock 10 min. 5. Discard the solution. (See Note 13.) 6. Rinse the tray with demi-water. 7. Add fresh demi-water and rock for 2 min. 8. Discard water. 9. Rinse the gel and gel bond with demi-water. 10. Place the gel in a stainless steel developer tray. 11. Add a small part of developer solution and rock slightly; discard this part and replace by the remaining developer solution. 12. Rock until the gel has developed well. 13. Discard the developer solution. 14. Add the previously used 200 mL of 1× Cairns’ solution to the box and rock for 5 min. 15. Discard the fixation solution. 16. Add some demi-water and rock for 2 min. 17. Replace the demi-water with Cairns’ preservation solution and rock for 7 min. 18. Place the gel on a glass plate (gelbond faced down). 19. Prewet a convenient sized piece of cellophane foil in the preservation solution and place it on the gel. (See Note 14.) 20. Dry the gel overnight at 60ºC.

To illustrate the value of the approach described here for the characterization and comparison of complex microbial communities at different levels of taxonomic resolution, Figures 2 and 3 show examples of DGGE profiles of the total bacterial and Bacillus benzoevorans-related communities in different fresh and air-dried soil specimen.

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Fig. 2. DGGE profiles of the total bacterial and Bacillus benzoevorans-related communities in Drentse A (A) and Friesian (B) soil samples. Bands were identified by cloning and sequence analysis, and are indicated with numbers and explained in the table below. Lane I, Total bacterial fingerprint (universal DGGE primers GC-0968f/1401r); lane II, B. benzoevorans-related fingerprint (specific primers REX460f/REX1466r and REX576f/REX1446r, and universal DGGE primers GC-0968f/1401r). (Reproduced from ref. 3 with permission of American Society for Microbiology.)

3.6. Statistical Analysis of DGGE Gels After staining and overnight drying the DGGE gels are scanned at 400DPI, and analyzed using e.g. the software Bionumerics 4.0 (Applied Maths BVBA, Belgium). A variety of approaches has been described to assess the similarity between DGGE profiles, which in general can be determined by calculating similarity indices from the presence or absence of bands, or based on the densitometric curves of the profiles compared using the Pearson product-moment correlation (13,14). The UPGMA algorithm can be used as implemented in the analysis software for the construction of dendrograms (Fig. 3b). Multivariate statistical analysis (e.g., CANOCO software) is also applicable for DGGE fingerprints interpretation, and a discussion of additional statistical approaches that have been applied to the analysis of microbial community fingerprints is given, e.g., by Fromin et al. (15).

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Fig. 3. DGGE profile of the B. benzoevorans related community in soil samples from different places (a) and corresponding dendogram (UPGMA clustering) (b). M, Marker; 1, Wageningen University experimental field (The Netherlands); 2, river Rhine shore (The Netherlands); 3, Drentse A (The Netherlands); 4, Friesland (The Netherlands); 5, Alnus glutinosa nodules (The Netherlands); 6, Alnus glutinosa rhizosphere (The Netherlands); 7, river site (Portugal); 8, grassland (Portugal); 9, pine forest (Portugal); 10, fruit garden (Portugal); 11, vegetable garden (Portugal); 12, Himalaya, bulk soil; 13, Himalaya, rhizosphere soil from Coriaria nepalensis. (Reproduced from ref. 3 with permission of American Society for Microbiology.)

4. Notes 1. Polymerase from other providers and other thermocycler machines are also suitable for use. 2. Glycerol is added only when silver staining is performed. By using other staining procedures no glycerol addition. 3. The GC-clamp can be attached to both forward and reverse primer but always at their 5 end.

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4. Caution: During preparation and development of DGGE gels, highly toxic and carcinogenic materials are used. Wear gloves and change them when they are contaminated. 5. The careful cleaning of the glass plates will provide good quality gels. Fat spots can cause disturbances in the polyacrylamide gel surface. 6. Gel bond bigger than the big glass plate will lead to further impossibility of the preparation of the gel. 7. Avoid making any air bubbles. 8. Use a flow hood. 9. Always put an empty sandwich (without spacers) at the other side of the cassette holder to get a closed upper buffer compartment. 10. Remember you have to load mirror-wise. 11. According to Sanguinetti et al. (16) with minor modifications. 12. The gels can also be stained with SYBR GOLD or ethidium bromide. 13. For the disposal of the solutions in steps 5 until 8 use the chemical waste! 14. Avoid air bubbles; press the edges of the gel to the glass plate.

Acknowledgment This work was supported by a grand from the European Communities EC project “Exploration of Genomic and Metabolite Diversity of a Novel Group of Abundant Soil Bacteria” (BACREX-project QLK3-2000-01678). References 1. Head, I. M., Saunders, J. R. and Pickup, R. W. (1998). Microbial evolution, diversity, and ecology: a decade of ribosomal RNA analysis of uncultivated microorganisms. Microbiol. Ecol. 35, 1. 2. La Duc, M. T., Kern, R. and Venkateswaran, K. (2004). Microbial monitoring of spacecraft and associated environments. Microbiol. Ecol. 47, 150. 3. Tzeneva, V. A., Li, Y., Felske, A. D. M., de Vos, W. M., Akkermans, A. D. L., Vaughan, E. E. and Smidt, H. (2004). Development and application of a selective PCR-denaturing gradient gel electrophoresis approach to detect a recently cultivated Bacillus group predominant in soil. Appl. Environ. Microbiol. 70, 5801–5809. 4. Dolfing, J., Vos, A., Bloem, J., Ehlert, P. A. I., Naumova, N. B. and Kuikman, P. J. (2004). Microbial diversity in archived soils. Science 306, 813. 5. Inagaki, F., Okada, H., Tsapin, A. I. and Nealson, K. H. (2005). Microbial survival: the paleome: a sedimentary genetic record of past microbial communities. Astrobiology 5, 141–153. 6. Noll, M., Matthies, D., Frenzel, P., Derakshani, M. and Liesack, W. (2005). Succession of bacterial community structure and diversity in a paddy soil oxygen gradient. Environ. Microbiol. 7, 382–395.

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7. Egert, M., Marhan, S., Wagner, B., Scheu, S. and Friedrich, M. W. (2004). Molecular profiling of 16S rRNA genes reveals diet-related differences of microbial communities in soil, gut, and casts of Lumbricus terrestris L. (Oligochaeta: Lumbricidae). FEMS Microbiol. Ecol. 48, 187–197. 8. Yoshino, K., Nishigaki, K. and Husimi, Y. (1991). Temperature sweep gel electrophoresis: a simple method to detect point mutations. Nucleic Acids Res. 19, 31–53. 9. Muyzer, G., de Waal, E. C. and Uitterlinden, A. G. (1993). Profiling of complex microbial populations by denaturing gradient gel electrophoresis analysis of polymerase chain reaction-amplified genes coding for 16S rRNA. Appl. Environ. Microbiol. 59, 695–700. 10. Muyzer, G. and Smalla, K. (1998). Application of denaturing gradient gel electrophoresis (DGGE) and temperature gradient gel electrophoresis (TGGE) in microbial ecology. Antonie Leeuwenhoek 73, 127–141. 11. Kowalchuk, G. A., Stephen, J. R., De Boer, W., Prosser, J. I., Embley, T. M. and Woldendorp, J. W. (1997). Analysis of ammonia-oxidizing bacteria of the beta subdivision of the class Proteobacteria in coastal sand dunes by denaturing gradient gel electrophoresis and sequencing of PCR-amplified 16S ribosomal DNA fragments. Appl. Environ. Microbiol. 63, 1489–1497. 12. Zoetendal, E. G., Akkermans, A. D. L., and De Vos, W. M. (1998). Temperature gradient gel electrophoresis analysis of 16S rRNA from human fecal samples reveals stable and host-specific communities of active bacteria. Appl. Environ. Microbiol. 64, 3854–3859. 13. Häne, B. G., Jäger, K., and Drexler, H. (1993). The pearson product-moment correlation coefficient is better suited for identification of DNA fingerprint profiles than band matching algorithms. Electrophoresis 14, 967–972. 14. Zoetendal, E. G., Akkermans, A. D. L., Akkermans - van Vliet, W. M., de Visser, J. A. G. M., and de Vos, W. M. (2001). The host genotype affects the bacterial community in the human gastrointestinal tract. Microbial Ecol. Health Dis. 13, 129–134. 15. Fromin, N., Hamelin, J., Tarnawski, S., Roesti, D., Jourdain-Miserez, K., Forestier, N., Teyssier-Cuvelle, S., Gillet, F., Aragno, M. and Rossi, P. (2002). Statistical analysis of denaturing gel electrophoresis (DGE) fingerprinting patterns. Environ. Microbiol. 4, 634–643. 16. Sanguinetti, C. J., Neto, D., and Simpson, A. J. G. (1994). Rapid silver staining and recovery of PCR products separated on polyacrylamide gels. Biotechniques 17, 915–919. 17. Nübel, U., Engelen, B., Felske, A., Snaidr, J., Wieshuber, A., Amann, R. I., Ludwig, W., and Backhaus, H. (1996). Sequence hetereogeneities of genes encoding 16S rRNAs in Paenibacillus polymyxa detected by temperature gradient gel electrophoresis of 16S rDNA fragments. J. Bacteriol. 178, 5636–5643. 18. Yu, Z. and Morrison, M. (2004). Comparisons of different hypervariable regions of rrs genes for use in fingerprinting of microbial communities by PCR-denaturing gradient gel electrophoresis. Appl. Environ. Microbiol. 70, 4800–4806.

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19. Grosskopf, R., Janssen, P. H. and Liesack, W. (1998). Diversity and structure of the methanogenic community in anoxic rice paddy soil microcosms as examined by cultivation and direct 16S rRNA gene sequence retrieval. Appl. Environ. Microbiol. 64, 960–969. 20. Lane, D. J. (1991) 16S/23S rRNA Sequencing. John Wiley & Sons, UK. 21. Diez, B., Pedros-Alio, C., Marsh, T. L. and Massana, R. (2001). Application of denaturing gradient gel electrophoresis (DGGE) to study the diversity of marine picoeukaryotic assemblages and comparison of DGGE with other molecular techniques. Appl. Environ. Microbiol. 67, 2942–2951.

18 An Emulsion Polymerase Chain Reaction–Based Method for Molecular Haplotyping James G. Wetmur and Jia Chen

Summary Genotypes are easily measured using a variety of experimental methods. However, experimental methods for measuring haplotypes, i.e., molecular haplotyping, are limited. Instead, haplotypes often are statistically inferred from genotype data with varying degrees of confidence, depending on the extent of linkage disequilibrium (LD) between markers. We have developed a method for molecular haplotyping, linking-emulsion polymerase chain reaction (LE-PCR), that should find application in studies where LD is limited, especially when the polymorphisms in question affect the function of a single gene product. We have illustrated this technology with the human paraoxonase 1 gene (PON1), where polymorphisms affecting transcription and enzymatic activity show incomplete LD. PON1 is an enzyme with multiple activities, including detoxification of organophosphates. Key Words: Emulsion PCR; genotype; haplotype; linking PCR; organophosphate; paraoxonase 1; pesticide; phenotype.

1. Introduction Linking-emulsion polymerase chain reaction (LE-PCR) (1) is carried out in an oil–water emulsion with two pairs of primers designed to amplify two linked polymorphic loci. The genomic DNA is diluted such that no more than a single template molecule is present in an aqueous droplet. The primers are designed to facilitate linking of the two amplicons in the droplet. Thus minichromosomes are formed preserving the phase of the polymorphic loci. In the method described below, the haplotypes of the minichromosomes are determined by quantitative allele-specific PCR (qASPCR). From: Methods in Molecular Biology, vol. 410: Environmental Genomics Edited by: C. Cristofre Martin © Humana Press, Totowa, NJ

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Other molecular haplotyping methods are, on the whole, quite complicated and thus not suitable for association studies. Long-range qASPCR (2,3) is limited to closely linked markers because of the inherent contradiction between optimizing for allele specificity and optimizing for long-range PCR. Sperm can be haplotyped, but are usually neither available nor appropriate for association studies (4). Haplotyping fosmid/cosmid pools (5) requires cloning, and haplotyping somatic cell hybrids (6) requires even more manipulation. Finally, there are methods based on PCR of single, isolated template molecules. The oldest approach was based on limiting dilution (7), and is still in use (8). Limiting dilution haplotyping, like sperm haplotyping, requires relatively efficient amplification from two loci from a single template molecule and extreme precautions to prevent contamination. Polony haplotyping (9), which uses a gel instead of the emulsion in LE-PCR to isolate the template molecules, is quite robust but requires unconventional instrumentation. LE-PCR demonstrated here requires a real-time PCR machine only for the qASPCR haplotype readout. We have demonstrated that LE-PCR is a powerful molecular haplotyping technology that can be applied in population studies (1). 2. Materials 2.1. Human Genomic DNA Any source of quality human DNA of high integrity may be used. For our example, the study population is from an ongoing study at the Mount Sinai Children’s Environmental Health Center to assess prospectively infant growth and neurodevelopment associated with pesticide exposure in urban New York City. The study protocol was approved by the Institutional Review Board. The study consists of pregnant women of multiethnic origin (Caucasian, African American, and Hispanic of Caribbean origin) at 26–30 weeks of gestational age. Leukocyte DNA was isolated from blood as previously described (10). 2.2. Oligonucleotides as Illustrated for PON1 -909g>c and Q192R 1. All primers were synthesized by IDT. 2. External primers for amplification across PON1 -909g>c and Q192R: CAAAATCAAATCCTTCTGCCACCACTCGAA and ACATGGAGCAAATCATTCACAGTAA, respectively. 3. Linking primers for PON1 -909g>c and Q192R (5 -biotinylated): Bio-AAAGTGCTCAGGTCCCACACTGATAATGGGGCATTTGAGTAA and Bio-GCCCCATTATCAGTGTGGGACCTGAGCACTTTTATGGCACAA, respectively.

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4. Capping oligonucleotides for PON1 -909g>c and Q192R (3 -phosphorylated): AAAAAAGCCCCATTATCAGTG-P and AAAAAAAAAGTGC TCAGGTCCCA-P, 5. qASPCR primers: 192T: CAAATACATCTCCCAGGATT 192C: CAAATACATCTCCCAGGATC -909C: GCAGACAGCAGAGAAGAGAC -909G: GCAGACAGCAGAGAAGAGAG

2.3. Buffers (All 1×) 1. 2. 3. 4.

Taq: 10 mM Tris-HCl, pH 8.0, 50 mM KCl. NX: 100 mM NaCl, 1% Triton X-100, 10 mM Tris-HCl, pH 7.5, 1 mM EDTA. B&W: 10 mM Tris-HCl, pH 7.5, 1 mM EDTA, 2 M NaCl. qASPCR: 1× Taq buffer, 1.5 mM MgCl2 , 200 μM each dNTP, 2% glycerol, 1× BSA (NEB), and 1× SYBR green (Molecular Probes).

2.4. Emulsion Constituents 1. Oil phase final concentrations: 4.5% Span 80 (cat. no. 85548, Fluka), 0.4% Tween80 (cat. no. S-8074, Sigma), 0.05% Triton X-100 (cat. no. T-9284, Sigma-Aldrich) made up to 100% with mineral oil (cat. no. M-3516, Sigma-Aldrich). 2. Aqueous phase final concentrations: 1× Taq buffer, 300 μM each dNTP, 2.5 mM MgCl2 , 50 μM Me4 NCl (tetramethylammonium chloride), 1 μM each external primer, 0.1 μM each linking primer, 100 mU/μL of Amplitaq Gold (Applied Biosystems), and 1 ng/μL o f human genomic DNA.

2.5. Capping Reaction Constituents (Final Concentrations) 1. 1× Taq buffer, 1.5 mM MgCl2 , 200 μM each dNTP. 2. 1 μM each capping oligonucleotide. 3. 5 U/40 μL of Taq DNA polymerase (Promega, not hot start).

2.6. qASPCR Constituents (Final Concentrations) 1. 1× qASPCR buffer. 2. 1 μM each qASPCR primer. 3. 2.5 U/20 μL of Amplitaq Gold DNA polymerase (Applied Biosystems).

2.7. Other Materials 1. PCR purification kit (Qiagen). 2. Dynabeads Myone Streptavidin (Dynal Biotech).

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3. Methods LE-PCR will be illustrated for two common polymorphisms in human paraoxonase 1 (PON1) for the common polymorphisms, -909g>c and Q192R, separated by about 15 kb on the genome. The -909g>c polymorphism is in nearly complete LD with a functional promoter polymorphism at -108 that affects the level of transcription. Q192R affects substrate specificity. In a total population of 378, 77 subjects had compound heterozygous genotypes at both loci, thus ambiguous haplotypes. We determined molecular haplotypes for these subjects and measured the enzymatic activity (phenotype) using two substrates. We also carried out molecular haplotyping for a third polymorphism, L55M, with -909g>c and with Q192R for subjects with compound heterozygous genotypes at L55M and a second locus. Only two molecular haplotype measurements were required to determine the haplotype for subjects who were heterozygous at all three loci. By comparing the predictive power of molecular vs. inferred haplotypes to the phenotype, we demonstrated the utility of molecular haplotyping by LE-PCR in population studies (1). This chapter deals only with LE-PCR illustrated for PON1 -909g>c and Q192R and not with measurements of phenotypes. The overall logic for LE-PCR is illustrated in Fig. 1. Two amplicons are produced spanning the linked polymorphisms within an aqueous droplet in an emulsion as illustrated under the template. Linking PCR connects these amplicons into a single minichromosomes preserving the phase information of the two polymorphisms.

Fig. 1. Linking emulsion-PCR with human PON1. The exons (horizontal stripes) of the 27-kb human PON1 gene are drawn to scale. PCR amplicons containing the promoter polymorphism -909g>c (vertical stripes) and the missense polymorphism Q192R in exon 6 (solid) are larger than scale. Linking PCR in an aqueous droplet in an emulsion leads to the minichromosome preserving phase (vertical stripes and solid).

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3.1. Primer Design 1. Emulsion PCR requires two primers for each of the two amplicons as illustrated in Fig. 2A. Amplicons should be limited to 200 nucleotides. 2. The external primers are typical PCR primers of about 25 nt. It is our experience that primers containing approx 50% GC and ending in AA are optimal. 3. The internal primers are the linking primers (11). The overlap of the partially complementary linking primers for PON1 -909g>c and Q192R is depicted in Fig. 2B. Thirty-two of 42 nucleotides are complementary beginning at the 5 end. The 26 nucleotides at the 3 end are complementary to the template and act as the primer. The 16 nucleotides at the 5 end of each linking primer are derived by complementarity to the other linking primer, and hence the template for the other

A

B

Fig. 2. Linking emulsion-PCR primers. (A) External and biotinylated linking primers for synthesis of two linked amplicons in an emulsion droplet. The dotted lines between the amplicons are 15 kb in the PON1 example in Fig. 1. All primers are shown as solid lines with arrows. Only the 3 ends of the linking primers are complementary to the template where positioned. (B) Design of linking primers illustrated for PON1 -909g>c and Q192R. The partially complementary sequences are shown. The vertical lines separate sequences derived from the two amplicons.

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amplicon. They must carry a 5 -biotin to allow subsequent separation of unlinked amplicons.

3.2. Emulsion Formation 1. The template between the amplicons must be intact in the aqueous droplets before PCR. We have shown that emulsification did not affect template integrity for PON1 -909g>c to Q192r (1). We have observed water droplets to be of the order of 10 μm and less. By limiting template concentration to 1 ng/μL (∼300 haploid genome equivalents/μL), the probability of having more than one template per droplet is less than 1%. 2. Oil–water emulsions are produced based on previously published methods (12–14). The oil phase is assembled from the constituents listed in Subheading 2.4., item 1. The aqueous phase is assembled from the constituents listed in Subheading 2.4., item 2. The external and linking primer sequences for PON1 -909g>c and Q192R are given in Subheadings 2.2., item 2 and 2.2, item 3, respectively. The human template DNAs to be haplotyped are heterozygous for both linked polymorphisms. 3. Emulsification involves vortex-mixing one part aqueous phase and two parts oil phase (typical volume 150 μL) for 5 min. A simple approach is illustrated in Fig. 3. (See Notes 1 and 2.)

3.3. LE-PCR 1. For the example used in this chapter, the PCR cycling conditions were 30 cycles for 1 min 67ºC, 1 min 60ºC, 30 s 94ºC following incubation at 95ºC for 9 min to activate the polymerase and followed by a final incubation for 7 min at 60ºC.

Fig. 3. Emulsion formation for several samples using a modified laboratory vortex apparatus.

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2. The external primers are present in 10-fold excess over the linking primers. The PCR conditions were chosen so that at later cycles, 1 min at 67ºC favored extension from longer primers, in this case first from the dilute linking primers, and later from the amplicon strands themselves acting as primers, as required for linking.

3.4. PCR Cleanup 1. Add 3 volumes of NX buffer to 5 volumes of emulsion. Vortex-mix for 20 s. Separate the phases in a microcentrifuge and remove most of the oil. 2. Transfer the aqueous phase to a Qiagen PCR purification kit and use according to the manufacturer’s instructions. Elute in 40 μL. The Qiagen PCR purification kit tolerates oil carryover.

3.5. Removal of Biotinylated Primers and Unlinked Amplicons 1. All of the products of LE-PCR are illustrated in Fig. 4A. These include singlestranded DNA runoff products from the two amplicons resulting from the excess of external primers over linking primers as well as the desired double-stranded minichromosomes, both lacking biotins. Removal of biotinylated nucleic acids is illustrated in Fig. 4A. Biotins are present both on the long and self-complementary linking primers and on amplicons that have failed to link. 2. Wash 3 μL of Dynabeads Myone streptavidin (Dynal Biotech) 3 times in B&W buffer and once in Taq buffer. Resuspend the beads in the 40-μL eluate from the PCR cleanup and add 4 μL of 10× Taq buffer. Incubate at room temperature for 30 min, magnetize, and retain the supernatant.

3.6. Capping of Runoff Products 1. The use of capping oligonucleotides is illustrated in Fig. 4B. Two products remain after step 3.5, the minichromosomes and any single-stranded runoff products that escaped PCR cleanup. The 3 -phosphate on capping oligonucleotides prevents their acting as primers. Instead, they are used as templates to extend the runoff products, thus preventing the runoff products from acting as primers and forming new minichromosomes. 2. It is our experience that capping the runoff products is essential to prevent formation of new minichromosomes from the runoff products. Because the emulsion is gone, any new minichromosomes will not preserve phase information of the two polymorphisms. 3. Capping reactions are assembled as described in Subheading 2.5. The oligonucleotides in Subheading 2.2., item 4 were used for PON1 -909g>c and Q192R runoff products as illustrated in the figure. Incubate at 55ºC for 30 min. This step completes LE-PCR.

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Fig. 4. After-emulsion PCR steps. (A) Removal of biotinylated primers and products (single amplicons) on streptavidin-coated magnetic beads. The top depicts single-stranded runoff products and double-stranded minichromosomes remaining after purification. Arrows depict incorporated primers. (B) Capping of single-stranded runoff products. Arrows depict incorporated primers or capping oligonucleotides. The 3 -phosphate on capping oligonucleotides prevents their acting as primers. The vertical striped sequence at the 3 end of the runoff products is partially complementary to the capping oligonucleotide. A DNA polymerase will extend the runoff products using the capping oligonucleotide as template.

3.7. Determination of Haplotypes by qASPCR 1. Each qASPCR reaction used 2 μL of capped product in a 20-μL PCR reaction. The reactions were assembled as noted in Subheading 2.6. All four possible reactions were assembled with one qASPCR primer per SNP. Ideally, the assays are carried

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out at least in duplicate. The primers for PON1 -909g>c and Q192R are given in Subheading 2.2., item 5. 2. PCR was carried out in a LightCycler (Roche) with cycling 1 min 55ºC, 1 min 72ºC, 30 s 94ºC after an initial incubation at 95ºC for 9 min to activate the polymerase. Ct values were determined by the LightCycler second derivative algorithm for each of the four primer pairs. 3. Any real-time PCR instrument should be equally effective.

3.8. Data Analysis 1. All LE-PCR reactions will lead to the formation of two minichromosomes preserving the phase information of the polymorphic alleles on the two template chromosomes. There are thus two possible pairs of minichromosomes for the two alleles. 2. The calculations assume all qASPCR primers are equally efficient when amplifying PCR matched templates, as was the case for our four examples. If not, appropriate corrections must be employed. 3. Calculate the average Ct for the first possible haplotype pair: here 192T/-909c plus 192C/-909g. 4. Calculate the average Ct for the second possible haplotype pair: here 192T/-909g plus 192C/-909c. 5. Subtract to obtain Ct . 6. It is required that BOTH Ct values for the first possible haplotype pair (e.g., both amplifications with allele-specific primers for 192T/-909c and with allelespecific primers for 192C/-909g) be less than BOTH Ct values for the second possible haplotype pair (e.g., 192T/-909g and 192C/-909c) OR BOTH Ct values for the first possible haplotype pair be greater than BOTH Ct values for the second possible haplotype pair. Thus the four qASPCR measurements consistently favor one haplotype pair. 7. It is required that Ct > 1 OR Ct < –1. Thus the extent to which the qASPCR measurements favor one haplotype pair is well above the experimental error of the technique. 8. If both conditions 6 and 7 are met, call the haplotype based on the lowest Ct values. 9. If both conditions 6 and 7 are not met, see Notes 3–6.

4. Notes 1. 2. 3. 4.

It is important that the tubes be in contact with the platform. Devices hanging the tubes around the sides produce poor emulsions. LE-PCR failures are characterized by observing Ct values close to zero. Early in the development of this method, LE-PCR failures occurred about 20–30% of the time, but after practice in forming the emulsions, the failure rate dropped to less than 5%.

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5. We found that all such failures occurred in the emulsion PCR step and could not be rescued by repeating the purification and assay steps. 6. If a haplotype cannot be called, repeat the entire LE-PCR experiment.

Acknowledgments This work was supported by Grants R21 ES011634 and P01 ES09584 from the National Institute of Environmental Health Sciences and RD831-711 from the Environmental Protection Administration.

References 1. Wetmur, J. G., Kumar, M., Zhang, L., Palomeque, C., Wallenstein, S. and Chen, J. (2005) Molecular haplotyping by linking-emulsion PCR: analysis of paraoxonase 1 haplotypes and phenotypes. Nucleic Acids Res. 33, 2615–2619. 2. Michalatos-Beloin, S., Tishkoff, S. A., Bentley, K. L., Kidd, K. K., and Ruano, G. (1996) Molecular haplotyping of genetic markers 10 kb apart by allele-specific long-range PCR. Nucleic Acids Res. 24, 4841–4843. 3. McDonald, O. G., Krynetski, E. Y., and Evans, W. E. (2002) Molecular haplotyping of genomic DNA for multiple single-nucleotide polymorphisms located kilobases apart using long-range polymerase chain reaction and intramolecular ligation. Pharmacogenetics 12, 93–99. 4. Li, H. H., Gyllensten, U. B., Cui, X. F., Saiki, R. K., Erlich, H. A., and Arnheim, N. (1988) Amplification and analysis of DNA sequences in single human sperm and diploid cells. Nature 335, 414–417. 5. Burgtorf, C., Kepper, P., Hoehe, M., Schmitt, C., Reinhardt, R., Lehrach, H., and Sauer, S. (2003) Clone-based systematic haplotyping (CSH): a procedure for physical haplotyping of whole genomes. Genome Res. 13, 2717–2724. 6. Douglas, J. A., Boehnke, M., Gillanders, E., Trent, J. M., and Gruber, S. B. (2001) Experimentally-derived haplotypes substantially increase the efficiency of linkage disequilibrium studies. Nature Genet. 28, 361–364. 7. Ruano, G., Kidd, K. K., and Stephens, J. C. (1990) Haplotype of multiple polymorphisms resolved by enzymatic amplification of single DNA molecules. Proc. Natl. Acad. Sci. USA 87, 6296–6300. 8. Vogelstein, B., and Kinzler, K. W. (1999) Digital PCR. Proc. Natl. Acad. Sci. USA 96, 9236–9241. 9. Mitra, R. D., Butty, V. L., Shendure, J., Williams, B. R., Housman, D. E., and Church, G. M. (2003) Digital genotyping and haplotyping with polymerase colonies. Proc. Natl. Acad. Sci. USA 100, 5926–5931. 10. Chen, J., Kumar, M., Chan, W., Berkowitz, G., and Wetmur, J. G. (2003) Increased influence of genetic variation on PON1 activity in neonates. Environ. Health Perspect. 111, 1403–1409.

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11. Horton, R. M., Hunt, H. D., Ho, S. N., Pullen, J. K., and Pease, L. R. (1989) Engineering hybrid genes without the use of restriction enzymes: gene splicing by overlap extension. Gene 77, 61–68. 12. Tawfik, D. S., and Griffiths, A. D. (1998) Man-made cell-like compartments for molecular evolution. Nat. Biotechnol. 16, 652–656. 13. Ghadessy, F. J., Ong, J. L., and Holliger, P. (2001) Directed evolution of polymerase function by compartmentalized self-replication. Proc. Natl. Acad. Sci. USA 98, 4552–4557. 14. Dressman, D., Yan, H., Traverso, G., Kinzler, K.W., and Vogelstein, B. (2003) Transforming single DNA molecules into fluorescent magnetic particles for detection and enumeration of genetic variations. Proc. Natl. Acad. Sci. USA 100, 8817–8822.

Index Adaptor ligation, 308–310 Antibody anti-DIG, 8, 229 anti-kinetochore, 204 FITC-conjugated, 204 HRP-conjugated, 118 in situ detection, 8 2D western blot, 118 Autoradiography, 165 CBMN assay, 188 kinetochore detection, 204–206 modifications, 200–203, 208 cDNA synthesis, 19, 62–63 Cell culture, 190–191 Cloning cDNA, 64 PCR products, 21 subtracted library, 316–317 Comet assay, 171–173 alkaline, 178–179 modifications, 179 Cytosine arabinoside assay, 198–200

FISH human sperm, 242–244 metaphase chromosomes, 217–225 Human sperm ACM assay, 244–245 Hybridization automated in situ, 9 dot blot, 317, 322–323, 325–326 FISH, 219–225, 228, 241–244, 249–250, 252, 255 macroarray, 36, 49 microarray, 94–95 suppressive subtraction, 296–300, 311–313 whole mount in situ, 4, 8 Image analysis pixel threshold method, 11 quantification of in situ signal, 11 quantitative FISH, 233–234 RLGS, 166–268 2D protein gels, 132–133 In situ Pro Automated System, 9 Interphase chromosome, 219

DGGE fingerprinting, 336–337 DNA bar coding, 275–277 DNA isolation, 280–282, 305, 339 DNA sequencing analysis of, 74–77, 287, 326 reaction, 34, 285–286 Dot blot, 318–319

Library cDNA amplification, 64 cDNA normalization, 33 construction, 32–33 screening, 65–67

Electrophoresis DGGE, 340–344 formaldehyde denaturing gel, 71, 91 Gelbond film, 178 PAGE, 20 PCR products, 282–283, 315–316 RLGS, 154 1st dimension (disc gel), 156, 159, 162 2nd dimension, 164–165 2D isoelectric focusing, 115, 130 SDS-PAGE, 115–116, 130–131 Excision of cDNA clones, 67–69

Macroarray automated printing, 48 cDNA, 30–32, 43 nylon membrane filter preparation, 34, 38 Metabolites biofluid isolation, 142 polar and lipophilic isolation, 142–143 Metabolomics, 138–140 Metaphase chromosomes, 219 Microarray heterologous probing, 83–88 Micronuclei, 186–188

363

364 Microscopy ACM assay, 255–263 assessment of MN frequency, 191–192 cell viability assay, 176 FISH, 224–225 fluorescent filter settings, 234–235 scoring kinetochore data, 206 scoring MN data, 192–197 stained zebrafish embryos, 11 Mouse sperm ACM assay, 250–251 Northern blotting probing, 73–74 production, 71–73 PCR bar code, 282 cloning, 21 DGGE, 340 differential display (DD-PCR), 16 fluorescent RNA arbitrarily primed (FRAP), 16, 20 product isolation, 21 product re-amplification, 21 quantitative, 104 semi-quantitative, 96–98, 104 tester-specific fragments, 313–315 Peroxisome, 123–124 Plasmid mini prep isolation, 69–70 preparation for in vitro synthesis, 6 removal of insert DNA, 70 Probe labeling ACM assay, 247–249 Cy5-labeled primer, 16 DIG-labeled RNA, 6 DNA, 324–325 dot and southern blot, 320–322 end-labeling, 159 FISH, 220–221, 228 FITC-labeled primer, 16 radio-labeled cDNA, 49, 70–71, 93–94 random primer, 252, 254 rhodamine-labeled primer, 16 Protein isolation from cells, 114–115 isolation of peroxisomes, 126–128 precipitation, 128–129 solubilization, 130 trypsin digestion, 116–118

Index Quantitative PCR, 104 Restriction endonuclease, 154 digestion, 159 in situ digestion, 157–158, 162–164 RLGS alternates, 168–169 SSH, 306–308 RNA isolation concentration determination, 62, 91 environmental samples, 323–324 from cells, 18 from tissues, 18, 61–62, 91 poly-A RNA, 32, 92 Southern blot, 319–320 Spectroscopy MALDI-TOF, 118 NMR, 139–140, 143–145 analysis of, 145–147 Staining alkaline phosphatase, 8 cell viability, 176 centromeric, 223 colloidal coomassie for protein, 132 comet assay gel, 177 counterstaining chromosomes, 224, 230, 250, 255 DGGE gel, 344–345 ethidium bromide, 176 NBT/BCIP, 9, 11 RNA gel, 71 silver staining for protein, 116 SYBR Gold, 179 Statistical analysis ACM assay, 264–268 DGGE gels, 345–346 macroarray, 36–38, 51–52 microarray, 95–96, 104 NDI and NDCI, 197–198 NMR, 145–147 Tissue fixation chromosomes, 221–222, 227 sperm cells, 247, 251 zebrafish embryos, 7, 9 Toxicology, 4, 16, 123–124, 218–219 Western blot 2D immunoblotting, 118