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.


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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

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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, Birkhäuser 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.

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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 ( 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

O3 -Response Genes from Arabidopsis thaliana

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.

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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 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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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

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