MicroRNA Expression Detection Methods

MicroRNA Expression Detection Methods Zhiguo Wang Baofeng Yang l MicroRNA Expression Detection Methods Dr. Zhiguo ...

Author: Zhiguo Wang | Baofeng Yang

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MicroRNA Expression Detection Methods

Zhiguo Wang Baofeng Yang l

MicroRNA Expression Detection Methods

Dr. Zhiguo Wang Montreal Heart Institute Research Center 5000 Belanger Street Montreal QC H1T 1C8 Canada [email protected] or [email protected]

Dr. Baofeng Yang Harbin Medical University Dept. Pharmacology 150086 Harbin China, People’s Republic [email protected]

ISBN: 978-3-642-04927-9 e-ISBN: 978-3-642-04928-6 DOI 10.1007/978-3-642-04928-6 Springer Heidelberg Dordrecht London New York Library of Congress Control Number: PCN Applied for # Springer-Verlag Berlin Heidelberg 2010 This work is subject to copyright. All rights are reserved, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilm or in any other way, and storage in data banks. Duplication of this publication or parts thereof is permitted only under the provisions of the German Copyright Law of September 9, 1965, in its current version, and permission for use must always be obtained from Springer. Violations are liable to prosecution under the German Copyright Law. The use of general descriptive names, registered names, trademarks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. Cover design: WMXDesign GmbH, Heidelberg, Germany Printed on acid-free paper Springer is part of Springer Science+Business Media (www.springer.com)

Preface

MicroRNAs (miRNAs), endogenous noncoding regulatory mRNAs of 22nucleotides, have rapidly emerged as the central players in gene expression regulation. Owing to their ever-increasing implications in the control of various biological and pathological processes, miRNAs have now been considered novel biomarkers of various human diseases including, cancer, viral disease, cardiovascular disorders, metabolic disturbances, etc. Particular expression profiles have been associated with particular pathological states. Expression profiling of miRNAs have therefore become extremely important not only for fundamentalists but also for clinicians. However, the methodologies used for detecting protein-coding mRNAs cannot be directly applied to miRNAs because of their small size. Over the past years, researchers have made great efforts to developing techniques suitable for miRNA detection and quantification; a wide spectrum of creative and innovative techniques (more than 30 different methods) have been invented and validated. It has come to the time now to summarize these methods and present them in an orderly manner for better understanding and utilization of these methods to miRNA research and applications. In particular, the development of methods for quantifying circulating miRNAs opens up a fascinating opportunity for realizing miRNA as diagnostic and prognostic biomarkers of human disease. A book on this subject may help boosting up the passion of researchers to further improve the existing techniques and develop more new methods to fit to new application needs. These considerations prompted us and urged us to undertake the work: writing a book focusing on miRNA expression detection methods. This book is aimed to target a wide range of readers from graduate students to post-doctoral fellow and senior researchers involving miRNA research of any fields in universities and research institutions. The contents of the book are also suitable for medical practitioners from residents to professors of various types of medical fields, who are interested in developing or utilizing miRNA profiling as a complementary and an alternative strategy for clinical diagnosis of human disease. It provides state-of-the art approaches, cutting-edge methods, and practical protocols as powerful, efficient tools for miRNA detection, profiling and quantification for

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Preface

both basic research and diagnostic analysis of miRNA-related diseases. The contents of chapters are organized essentially based on the hands-on laboratory experience from many outstanding investigators worldwide. In microRNA Expression Detection Methods, the authors provide comprehensive descriptions of the innovative strategies and methodologies for detecting miRNA expression, and their applications to miRNA research and their potential as tools for clinical diagnosis and prognosis. The book is divided into 11 sections that include a total of 33 chapters. The book begins with Sect. 1 introducing the overall concept and strategies of miRNA expression detection methods emphasizing the need of a wide variety of miRNA detection methods to suit specific requirements for research and clinical examination in the laboratories. From Sect. 2 to Sect. 11, each of the 32 chapters is focused on an independent, unique method of miRNA detection. Each single chapter contains five subsections: Summary, Introduction, Protocol (including Materials, Instrument, Reagent, and Procedure), Application and Limitation, and Reference. The development of the technique, ideas behind it, and mechanisms underlying the method are given in Introduction of each chapter. The step-by-step protocols are detailed in Protocol section. Then, the applications and limitations of the methods are discussed. Finally, the literature citations are listed in Reference section. Schematic diagrams are included where needed and appropriate for better illustrating the principle of the methodologies. In addition, flowcharts are also provided to outline the protocols for each of miRNA expression detection methods. Canada China

Zhiguo Wang, PhD Baofeng Yang, MD, PhD

Acknowledgements

There are old Chinese sayings, “everything is difficult to do at the very beginning” and “as soon as revives, two chapter of ripeness.” Yet despite that this is the second one we have written in the line of miRNA book series, we were still feeling nervous and sometimes unconfident. Thanks God! We have finally made it, after a mixture of the bitter and sweet. But we have been enjoying the process more than the harvest. We entirely credit to the actual contributors and inventors of the techniques and methodologies described in this book. Since the beginning of the writing process, we have been immediately inspired by the actual contributors by their ingenious, creative thoughts, strongly amazed by their serious efforts of searing scientific doctrine, and deeply touched by their non-conserved sharing-out of every detail of their experimental protocols. We were not only gathering the information from their studies for our book writing, but are also learning a great deal from these investigators for “great works are performed not by strength but by perseverance (Samuel Johnson)” and “the talent of success is nothing more than doing well whatever you do without a thought of fame (Longfellow)”. We give our most sincere and respectful thanks to all these people, whether we knew them in person or not; we know their names from their papers and their works by heart. Without their sweat and intellect, this book would have been impossible. We give our hearty gratitude to our wives who are standing behind us and selflessly supporting us, while sharing with us the hash times and the joyfulness as well. We also thank our kids for understanding our long-time unavailability of being with them for fun and our lack of care of them as a result of deprivation of our attention from them by the writing task.

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About the Authors

Wang, Zhiguo He received his PhD in cardiovascular pharmacology from McGill University, Canada. He is presently a Professor of Biomedical Sciences, University of Montreal (Canada) and Department of Phamacology, Harbin Medical University, Harbin, Heilongjiang, China. He is the Director of the Cardiovascular Research Institute of Harbin Medical University. He is also a ChangJiang Scholar Endowed Professor (China), the Ministry of Education of China and a Longjiang Scholar Endowed Professor, the Education Committee of Heilongjiang province, China. His current research interests include cardiovascular disease and gene therapy related to microRNAs and ion channels. Yang, Baofeng He received his PhD in pharmacology from Tongji Medical University, China. He is presently a Professor of Pharmacology and the President, Harbin Medical University, China. He is an Academician, the Academy of Engineering of China. He is a Chief Scientist for the National Program on Key Basic Research Project of China, the Vice President of Chinese Pharmacologic Society, a Visiting Professor of West Virginia University, University of Missouri-Kansas City and Nippon Medical University, and an Honorable Professor of Perm Pharmaceutical Academy and Shiga-University of Medical Science. His current research interests include cardiovascular and molecular pharmacology involving microRNAs and ion channels.

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Contents

Part I 1

Detection, Profiling, and Quantification of miRNA Expression . . . . . . . 1.1 miRNA Biology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.1.1 Biogenesis of miRNAs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.1.2 Actions of miRNAs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.2 Human Disease-Related Expression Profiles of miRNAs . . . . . . . . . . . . . 1.2.1 Spatiotemporal Expression Profiles . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1.2.2 miRNA Transcriptome and Human Physiology . . . . . . . . . . . . . . 1.2.3 miRNA Transcriptome in Diseased States . . . . . . . . . . . . . . . . . . . . 1.3 miRNAs as Biomarkers for Human Disease . . . . . . . . . . . . . . . . . . . . . . . . . 1.4 Methods for Analyzing miRNAs Expression . . . . . . . . . . . . . . . . . . . . . . . . 1.4.1 Ideal Methods for miRNA Detection . . . . . . . . . . . . . . . . . . . . . . . . . . 1.4.2 Classification of Methods for miRNA Detection . . . . . . . . . . . . . 1.4.3 Brief Introduction to the Currently Available miRNA Detection Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Part II 2

General Remarks 3 3 3 7 9 9 17 20 37 38 39 39 40 53

miRNA Microarray Methods

Microarray and Its Variants for miRNA Profiling . . . . . . . . . . . . . . . . . . . 2.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2 Protocol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2.1 Materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2.2 Instruments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2.3 Reagents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2.4 Procedures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3 Application and Limitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

67 68 70 70 71 72 73 77 78

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Part III 3

Northern Blotting and Its Variants for Detecting Expression and Analyzing Tissue Distribution of miRNAs . . . . . . . . . . . 3.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2 Protocol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2.1 Basic Protocols . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2.2 Protocols with Improved Sensitivity . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2.3 Protocols with Improved Specificity and Sensitivity . . . . . . . . . . 3.2.4 Protocols with Nonisotopic Detection . . . . . . . . . . . . . . . . . . . . . . . . . 3.3 Application and Limitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Part IV 4

83 83 85 85 89 91 97 99 99

In Situ Hybridization Methods

In Situ Hybridization and Its Variants for Detecting Expression and Analyzing Cellular Distribution of miRNAs . . . . . . . 4.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2 Protocol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2.1 Basic Protocol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2.2 LNA-Modified Protocol with Enhanced Sensitivity and Specificity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.2.3 Ultramer Extension Protocol with Reduced Stringency and Expense . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4.3 Application and Limitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Part V 5

Northern Blotting Methods

103 104 106 106 116 122 126 127

Real-Time RT-PCR Methods

End-Point Stem-Loop Real-Time RT-PCR for miRNA Quantification . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2 Protocol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2.1 Materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2.2 Instruments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2.3 Reagents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.2.4 Procedures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5.3 Application and Limitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

131 132 134 134 135 135 135 139 139

Contents

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6

miR-Q RT-PCR for miRNA Quantification . . . . . . . . . . . . . . . . . . . . . . . . . . 6.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.2 Protocol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.2.1 Materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.2.2 Instruments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.2.3 Reagents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.2.4 Procedures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.3 Application and Limitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

141 141 142 142 142 143 143 146 146

7

Poly(A)-Tailed Universal Reverse Transcription . . . . . . . . . . . . . . . . . . . . . 7.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.2 Protocol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.2.1 Materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.2.2 Instruments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.2.3 Reagents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.2.4 Procedures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7.3 Application and Limitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

147 147 148 148 149 149 149 151 151

8

Multiplexing RT-PCR for High-Throughput miRNA Profiling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.2 Protocol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.2.1 Materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.2.2 Instruments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.2.3 Reagents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.2.4 Procedures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8.3 Application and Limitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

153 154 155 155 155 155 155 156 157

miRNA Amplification Profiling (mRAP) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.2 Protocol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.2.1 Materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.2.2 Instruments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.2.3 Reagents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.2.4 Procedures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9.3 Application and Limitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

159 159 160 160 162 163 164 169 169

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Part VI 10

miRNA Serial Analysis of Gene Expression (miRAGE or SAGE) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10.2 Protocol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10.2.1 Materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10.2.2 Instruments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10.2.3 Reagnets . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10.2.4 Procedures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10.3 Application and Limitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Part VII 11

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13

Cloning Methods

173 173 174 174 176 176 177 187 188

Nanoparticle Methods

Electrocatalytic Nanoparticle Tags Technique for High-Sensitivity miRNA Expression Analysis . . . . . . . . . . . . . . . . . . . . 11.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.2 Protocol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.2.1 Materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.2.2 Instruments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.2.3 Reagents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.2.4 Procedures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11.3 Application and Limitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

191 191 194 194 194 194 195 197 197

Nanoparticle-Amplified SPR Imaging for High-Sensitivity miRNA Profiling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.2 Protocol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.2.1 Materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.2.2 Instruments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.2.3 Reagents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.2.4 Procedures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12.3 Application and Limitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

199 199 201 201 201 202 202 205 206

Conducting Polymer Nanowires Technique for High-Sensitivity miRNA Expression Analysis . . . . . . . . . . . . . . . . . . . . . . . . 13.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13.2 Protocol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13.2.1 Materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13.2.2 Instruments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

207 207 209 209 209

Contents

14

xv

13.2.3 Reagents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13.2.4 Procedures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13.3 Application and Limitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

209 211 214 215

Gold Nanoparticle Probe Method for miRNA Quantification . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 14.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 14.2 Protocol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 14.2.1 Materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 14.2.2 Instruments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 14.2.3 Reagents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 14.2.4 Procedures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 14.3 Application and Limitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

217 217 220 220 220 220 221 224 224

Part VIII

Other Methods

15

Splinted Ligation Method . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 15.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 15.2 Protocol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 15.2.1 Materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 15.2.2 Instruments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 15.2.3 Reagents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 15.2.4 Procedures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 15.3 Application and Limitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

229 230 230 230 230 232 233 238 239

16

Padlock-Probes and Rolling-Circle Amplification . . . . . . . . . . . . . . . . . . . 16.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 16.2 Protocol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 16.3 Materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 16.3.1 Instruments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 16.3.2 Reagents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 16.3.3 Procedures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 16.4 Application and Limitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 16.4.1 Applications . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 16.4.2 Limitations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

241 241 243 243 243 243 244 246 246 247 247

17

Invader Assay . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 17.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 17.2 Protocol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 17.2.1 Materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

249 249 250 250

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Contents

17.2.2 Instruments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 17.2.3 Reagents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 17.2.4 Procedures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 17.3 Application and Limitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

250 250 251 255 255

18

Single Molecule Method . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 18.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 18.2 Protocol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 18.2.1 Materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 18.2.2 Instruments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 18.2.3 Reagents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 18.2.4 Procedures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 18.3 Application and Limitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

257 257 258 258 259 259 259 263 264

19

Enzymatic Method . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 19.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 19.2 Protocol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 19.2.1 Materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 19.2.2 Instruments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 19.2.3 Reagents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 19.2.4 Procedures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 19.3 Application and Limitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

267 267 269 269 269 269 270 274 274

20

Surface-Enhanced Raman Spectroscopy Method . . . . . . . . . . . . . . . . . . . . 20.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 20.2 Protocol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 20.2.1 Materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 20.2.2 Instruments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 20.2.3 Reagents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 20.2.4 Procedures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 20.3 Application and Limitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

275 275 277 277 277 277 278 280 280

21

RAKE Assay . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 21.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 21.2 Protocol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 21.2.1 Materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 21.2.2 Instruments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 21.2.3 Reagents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 21.2.4 Procedures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

281 281 283 283 283 283 284

Contents

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21.3 Application and Limitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 287 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 287 22

Bead-Based Flow Cytometric miRNA Expression Profiling . . . . . . . . . 22.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 22.2 Protocol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 22.2.1 Materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 22.2.2 Instruments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 22.2.3 Reagents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 22.2.4 Procedures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 22.3 Application and Limitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

289 289 290 290 290 290 290 293 294

23

Bioluminescence miRNA Detection Method . . . . . . . . . . . . . . . . . . . . . . . . . . 23.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 23.2 Protocol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 23.2.1 Materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 23.2.2 Instruments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 23.2.3 Reagents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 23.2.4 Procedures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 23.3 Application and Limitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

295 295 296 296 297 297 298 301 302

24

Molecular Beacon Method . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 24.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 24.2 Protocol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 24.2.1 Materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 24.2.2 Instruments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 24.2.3 Reagents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 24.2.4 Procedures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 24.3 Application and Limitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

303 303 306 306 307 307 307 310 310

25

Ribozyme Method . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 25.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 25.2 Protocol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 25.2.1 Materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 25.2.2 Instruments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 25.2.3 Reagents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 25.2.4 Procedures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 25.3 Application and Limitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

313 313 315 315 315 315 316 318 319

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26

Contents

Electrocatalytic Moiety Labeling Technique for High-Sensitivity miRNA Expression Analysis . . . . . . . . . . . . . . . . . . . . . . . . 26.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 26.2 Protocol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 26.2.1 Materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 26.2.2 Instruments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 26.2.3 Reagents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 26.2.4 Procedures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 26.3 Application and Limitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

Part IX

321 321 322 322 323 324 324 327 328

Circulating miRNA Detection Methods

27

Serum and Plasma miRNA Detection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 27.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 27.2 Protocol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 27.2.1 Materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 27.2.2 Instruments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 27.2.3 Reagents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 27.2.4 Procedures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 27.3 Application and Limitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

331 331 332 332 333 333 333 337 337

28

miRNA Detection from Peripheral Blood Microvesicles . . . . . . . . . . . . 28.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 28.2 Protocol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 28.2.1 Materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 28.2.2 Instruments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 28.2.3 Reagents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 28.2.4 Procedures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 28.3 Application and Limitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

339 339 340 340 340 341 341 343 343

29

Detection of Placental miRNAs in Maternal Plasma . . . . . . . . . . . . . . . . . 29.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 29.2 Protocol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 29.2.1 Materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 29.2.2 Instruments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 29.2.3 Reagents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 29.2.4 Procedures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 29.3 Application and Limitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

345 345 346 346 346 346 347 348 349

Contents

Part X 30

xix

Single-cell miRNA Detection Methods

Quantitative LNA-ELF-FISH Method for miRNA Detection in Single Mammalian Cell . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 30.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 30.2 Protocol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 30.2.1 Materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 30.2.2 Instruments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 30.2.3 Reagents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 30.2.4 Procedures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 30.3 Application and Limitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

353 353 354 354 354 355 355 358 358

31

Single Cell Stem-Looped Real-Time PCR . . . . . . . . . . . . . . . . . . . . . . . . . . . . 31.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 31.2 Protocol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 31.2.1 Materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 31.2.2 Instruments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 31.2.3 Reagents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 31.2.4 Procedures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 31.3 Application and Limitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

361 361 363 363 363 364 364 367 367

32

miRNA Function-Reporter Expression Assay . . . . . . . . . . . . . . . . . . . . . . . . 32.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 32.2 Protocol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 32.2.1 Materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 32.2.2 Instruments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 32.2.3 Reagents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 32.2.4 Procedures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 32.3 Application and Limitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

369 369 370 370 371 371 371 373 374

Part XI 33

Whole Mount In Situ Analysis

Whole Mount In Situ Hybridization (WM-ISH) for miRNA Expression Profiling During Vertebrate Development . . . . . . . . . . . . . . 33.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 33.2 Protocol . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 33.2.1 Materials . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 33.2.2 Instruments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

377 377 379 379 379

xx

Contents

33.2.3 Reagents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 33.2.4 Procedures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 33.3 Application and Limitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

379 380 383 384

Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 385

Part I General Remarks

Chapter 1

Detection, Profiling, and Quantification of miRNA Expression

Abstract MicroRNAs (miRNAs), endogenous noncoding regulatory mRNAs of 22 nucleotides, have rapidly emerged as the central players in gene expression regulation. Owing to their ever-increasing implications in the control of various biological and pathological processes, miRNAs have now been considered novel biomarkers of various human diseases including, cancer, viral diseases, cardiovascular disorders, metabolic disturbances, etc. Particular expression profiles have been associated with particular pathological states. Expression profiling of miRNAs have therefore become extremely important not only for fundamentalists but also for clinicians. Over the past years, researchers have made great efforts to develop techniques suitable for miRNA detection and quantification; a wide spectrum of creative and innovative techniques (more than 30 different methods) have been invented and validated. While the aim of this book is to introduce these miRNA expression detection methods, this chapter serves to pave the way for better understanding of these techniques. The chapter begins with the basics of miRNAs in terms of the fundamental aspects of miRNA biology: the biogenesis and actions of miRNAs. Next, miRNA transcriptome related to human physiology and pathology is described. The concept of miRNAs as potential biomarkers of human disease is then introduced. And finally, a brief introduction to miRNA expression detection technologies is given to link to the following chapters.

1.1

miRNA Biology

1.1.1

Biogenesis of miRNAs

1.1.1.1

Transcriptional Regulation

Genes for miRNAs are located in the chromosomes, and many of them are identified in clusters that can be transcribed as polycistronic primary transcripts. Some miRNAs Z. Wang and B. Yang, MicroRNA Expression Detection Methods, DOI 10.1007/978-3-642-04928-6_1, # Springer-Verlag Berlin Heidelberg 2010

3

4

1 Detection, Profiling, and Quantification of miRNA Expression

are encoded by their own genes and others are encoded by the sequences as a part of the host protein-coding genes. On the basis of the genomic arrangement of their genes, miRNAs can be grouped into two classes (Wang 2009b): 1. Intergenic miRNAs (miRNA-coding genes located in between protein-coding genes) 2. Intragenic miRNAs (miRNA-coding genes located within their host proteincoding genes). Further, the intragenic miRNAs can be divided into the following subclasses: (a) Intronic miRNAs (miRNA-coding genes located within introns of their host protein-coding genes) (b) Exonic miRNAs (miRNA-coding genes located within exons of host protein-coding genes) (c) 30 UTR miRNAs (miRNA-coding genes located within 30 UTR of host protein-coding genes) (d) 50 UTR miRNAs (miRNA-coding genes located within 50 UTR of host protein-coding genes) According to our analysis, in the human miRNAs identified thus far, a majority of miRNAs belong to intergenic and intronic miRNAs comprising ~42 and ~44% of the total, respectively, and the other three categories are rare, with the exonic miRNAs being ~7%, 30 UTR miRNAs being 1.5%, and 50 UTR miRNAs being 1%. Clearly, miRNAs either have their own genes or are associated with their host genes; accordingly, miRNAs are generated by two different mechanisms. Biogenesis of miRNAs can be summarized as a five-step process as detailed below (see also Fig. 1.1). 1. Generation of primary miRNAs: transcription of miRNA genes. The intergenic miRNA genes are first transcribed as long transcripts, called primary miRNAs (pri-miRNAs), mostly by RNA polymerase II or RNA polymerase III (Ying and Lin 2005). The pri-miRNAs are capped and polyadenylated and can reach several kilobases (kb) in length (Cullen 2004; Kim 2005). The clustered miRNA genes in polycistronic transcripts are likely to be coordinately regulated (Bartel 2004). The intronic miRNAs are processed by sharing the same promoter and other regulatory elements of the host genes. They are first transcribed along with their host genes by RNA polymerase II and then processed by Droshaindependent pathway from excised introns by the RNA splicing machinery for their biogenesis in Drosophila, C elegans, and mammals (Berezikov et al. 2007; Okamura et al. 2007; Ruby et al. 2007).

1.1.1.2

Post-transcriptional Processing

2. Generation of precursor miRNAs: endonuclease processing of pri-miRNAs. The pri-miRNAs are processed to precursor miRNAs (pre-miRNAs) by the RNase

1.1 miRNA Biology

5

CTD TEFb

TFIIF

Pol II

Transcription

AAAA

m7G

Targeted mRNA cleavage

Cytoplasm

Nucleus

pri-miRNA

miRNA

pre-miRNA

degradation

miRISC passenger strand RISC

protein-coding mRNA m7G

guide strand RISC

RISC

translation inhibition

~ ~ ~ ~ ~ ~

RISC

AAAA

ORF

3’UTR

II Protein

Fig. 1.1 Illustrative diagram for the action of miRNAs. A protein-coding gene is transcribed to an mRNA that is subsequently translated to a protein. On the other hand, a small RNA-coding gene is initially transcribed into a long primary miRNA (pri-miRNA) with stem-loop structure which is rapidly processed to remove branches and become precursor miRNA (pre-miRNA). Pre-miRNA is then transported out of the nucleus to the cytoplasm where it is further processed to become a double-stranded mature miRNA. A mature miRNA is incorporated into a protein complex called RISC (RNA-induced silencing complex). One of the strands is then removed from the complex and gets degraded, being a passenger strand. The remaining strand called guide strand can guide the RISC to find its complementary sites in the 30 UTR of target genes. The binding of RISC then primarily causes translation inhibition of protein-coding mRNA and may cause mRNA degradation as well

endonuclease-III Drosha and its partner DGCR8/Pasha in the nucleus (Lee et al. 2002b; Denli et al. 2004; Gregory et al. 2004; Landthaler et al. 2004). These premiRNAs are ~60–100 nts with a stem-loop or hairpin secondary structure. Specific RNA cleavage by Drosha predetermines the mature miRNA sequence and provides the substrates for subsequent processing steps. Cleavage of a primiRNA by microprocessor begins with DGCR8 recognizing the single-stranded RNA (ssRNA)–double-stranded RNA (dsRNA) junction typical of a pri-miRNA (Han et al. 2006). Then, Drosha is brought close to its substrate through interaction with DGCR8 and cleaves the stem of a pri-miRNA ~11 nt away from the two single-stranded segments. miRNA precursor-containing introns have recently been designated “mirtrons” (Miranda et al. 2006). Mirtrons are derived from certain debranched introns that fold into hairpin structures with 50 monophosphates and 30 2-nt

6

1 Detection, Profiling, and Quantification of miRNA Expression

hydroxyl overhangs, which mimic the structural hallmarks of pre-miRNAs and enter the miRNA-processing pathway (Okamura et al. 2007; Ruby et al. 2007). The discovery of mirtrons suggests that any RNA, with a size comparable to a pre-miRNA and all the structural features of a pre-miRNA, can be utilized by the miRNA processing machinery and can potentially give rise to a functional miRNA. 3. Nucleus to cytoplasm translocation of pre-miRNAs. Pre-miRNAs then get exported to the cytoplasm from the nucleus through nuclear pores by RanGTP and exportin-5 (Bohnsack et al. 2004; Lund et al. 2004; Yi et al. 2003). After a pre-miRNA is exported to the cytoplasm, RanGTP is hydrolyzed by RanGAP to RanGDP, and the pre-miRNA is released from Exp-5. 4. Generation of mature miRNAs: endonuclease processing of pre-miRNAs. In the cytoplasm, pre-miRNAs are further processed by Dicer in animals, which is a highly conserved, cytoplasmic RNase III ribonuclease that chops pre-miRNAs into ~22-nt duplexes of mature miRNAs containing a guide strand and a passenger strand (miRNA/miRNA*), with 2-nt overhangs at the 30 termini (Kim 2005). Like other RNase III family proteins, Dicer interacts with doublestranded RNA-binding protein (dsRBP) partners. In mammalian cells, Dicer associates with transactivation-response element RNA-binding protein (TRBP) and protein activator of the interferon-induced protein kinase (PACT) (Chendrimada et al. 2005; Lee et al. 2006). In plants, miRNAs are cleaved into miRNA:miRNA* duplex, possibly by Dicer-like enzyme 1 (DCL1) in the nucleus rather than in the cytoplasm (Bartel 2004; Lee et al. 2002a), then the duplex is translocated into the cytoplasm by HASTY, the plant ortholog of exportin 5 (Bartel 2004). The strands of this duplex separate and release mature miRNA of 19–25 nts in length (Bartel 2004; Lee et al. 2002). Plant miRNAs undergo further modification by methylation at the 30 end by HEN1 (Yu et al. 2005). 5. Formation of miRISC. Mature miRNAs get integrated into a RNA-induced silencing complex (RISC) to form the miRNA:RISC complex (miRISC). Only one strand of miRNA/miRNA*, the guide strand, is successfully incorporated into RISC, while the other strand, the passenger strand, is eliminated. Strand selection may be determined by the relative thermodynamic stability of two ends of miRNA duplexes (Khvorova et al. 2003; Schwarz et al. 2003). The strand with less stability at the 50 end is favorably loaded onto RISC, whereas the passenger strand is released or destroyed. miRISC contains several proteins, such as Dicer, TRBP, PACT, and Gemin3, but the components directly associated with miRNAs are Argonaute proteins (Ago). These proteins contain four domains: the N-terminal, PAZ, middle, and Piwi domains. The PAZ domain binds to the 30 end of guide miRNA, while the other three domains form a unique structure, creating grooves for target mRNA and guide miRNA interactions (Liu et al. 2002b; Song et al. 2004; Ma et al. 2005; Parker et al. 2005). In mammalian cells, four Ago proteins have been identified, all of which can bind to endogenous miRNAs (Meister and Tuschl 2004). Despite the sequence similarity among these Ago proteins, only Ago2 exhibits endonuclease activity to slice

1.1 miRNA Biology

7

complementary mRNA sequences between positions 10 and 11 in the 50 end of guide strand miRNA. Therefore, human Ago2 is a component not only of miRISC but also of siRISC (siRNA-induced silencing complex), a RISC assembled with exogenously introduced siRNA. The roles of various Ago proteins in mammalian RISC are ambiguous, but the division of labor among Ago proteins in Drosophila is well defined. Drosophila Ago1 and Ago2 have been shown by biochemical and genetic evidence to participate in two separate pathways: Ago1 interacts with miRNA in translational repression, whereas Ago2 associates with siRNA for target cleavage (Carmell et al. 2002; Okamura et al. 2004).

1.1.2

Actions of miRNAs

1.1.2.1

Mechanisms of Actions

miRNAs exist in double-stranded form (duplex), activate in single-stranded form (simplex), and act in complex form miRISC. Mature miRNAs confer sequence specificities to the RISC complex. Subsequently, a miRNA in the miRISC binds to the 30 untranslated region (30 UTR) of its target mRNA through a WatsonCrick basepairing mechanism with its 50 -end 2–8 nts exactly complementary to recognition motif within the target (taking into account that an RNA– RNA hybrid can also contain G–U matches). This 50 -end 2–8 nt region is termed “seed sequence” or “seed site” as it is critical for miRNA actions (Lewis et al. 2003, 2005). Partial complementarity with the rest of the sequences of a miRNA also plays a role in producing post-transcriptional regulation of gene expression, presumably by stabilizing the miRNA:mRNA interaction. Moreover, the mid and 30 -end regions of a miRNA may also be important for forming miRISC. Studies have shown that in addition to 30 UTR, coding region and 50 UTR can also interact with miRNAs to induce gene silencing (Jopling et al. 2005; Luo et al. 2008; Tay et al. 2008). In mammalian species, the assembly of the miRNA/RISC on a 30 UTR can potentially influence protein production by enhancing de-adenylation with subsequent degradation of the mRNA or by repressing translation initiation or both (Yekta et al. 2004; Lim et al. 2005; Giraldez et al. 2005; Pillai et al. 2007), depending upon at least the following factors (see Fig. 1.1): 1. The overall degree of complementarity of the binding site 2. The number of recognition motif corresponding to 50 -end 2–8 nts of the miRNA, and 3. The accessibility of the bindings sites (as determined by free energy states) (Jackson and Standart 2007; Nilsen 2007; Pillai et al. 2007) The greater the degree of complementarity of accessible binding sites, the more likely a miRNA degrades its targeted mRNA. The loose binding constraints allow

8

1 Detection, Profiling, and Quantification of miRNA Expression

one miRNA to bind to several sites within one 30 UTR. Perfectly complementary targets (full miRNA:mRNA interaction) are efficiently silenced by the endonucleolytic cleavage activity of some Argonaute proteins (Hutva´gner and Zamore 2002; Yekta et al. 2004; Davis et al. 2005), but the vast majority of predicted targets in animals are only partially paired (Partial miRNA:mRNA interaction) (Lewis et al. 2003, 2005; Grun et al. 2005; Krek et al. 2005; Rajewsky and Socci 2004; Brennecke et al. 2005) and can hardly be cleaved (Haley and Zamore 2004). Some miRNAs have only seed-site complementarity (seed-site miRNA:mRNA) and this interaction primarily leads to translation inhibition. And those miRNAs that display imperfect sequence complementarities with target mRNAs primarily lead to translational inhibition (Lewis et al. 2003, 2005; Jackson and Standart 2007; Nilsen 2007; Pillai et al. 2007). The mechanisms for translational inhibition remain largely unkown, although inhibition of translation initiation has been identified as one such mechanism by several studies (Humphreys et al. 2005; Pillai et al. 2005). Greater actions may be elicited by a miRNA if it has more than one accessible binding sites in its targeted miRNA, presumably by the cooperative miRNA:mRNA interactions from different sites. mRNA degradation by miRISC is initiated by deadenylation and decapping of the targeted mRNAs (Pillai et al. 2007). A recent study demonstrated, however, that miRNAs can also act to enhance translation when AU-rich elements and miRNA target sites coexist at proximity in the target mRNA and when the cells are in the state of cell-cycle arrest (Vasudevan et al. 2007). In plants, miRNAs base pair with their mRNA targets by precise or nearly precise complementarity (Wang et al. 2006). The loose binding constraints also allow one miRNA to bind to multiple mRNA targets within the transcriptome. This multiplicity endows miRNAs in principle with the ability to inhibit several genes at once, leading to a much stronger biological response due to multiple effects on one pathway or coordinated effects on several pathways. It has been predicted that each single miRNA can have >1,000 target genes and each single protein-coding gene can be regulated by multiple miRNAs (Lewis et al. 2003, 2005; Jackson and Standart 2007; Nilsen 2007; Pillai et al. 2007; Alvarez-Garcia and Miska 2005; Ambros 2004). This is at least partially a result of a lax requirement of complementarity for miRNA:mRNA interaction (Lim et al. 2005). This implies that actions of miRNAs are sequence- or motif-specific, but not gene-specific; different genes can have same binding motifs for a given miRNA and a given gene can have multiple binding motifs for distinct miRNAs. On the downside, the relaxed stringency of miRNAs, with regard to their potential targets, enhances the possibility of disadvantageous off-target effects on inappropriate mRNAs. Another disadvantage is the difficulty it raises for researchers trying to interpret the biological significance of altered expression of a given miRNA by determination of its relevant downstream targets. On the basis of the characteristics of miRNA actions, we postulated that a miRNA should be viewed as a regulator of a cellular function or a cellular program, not of a single gene (Wang et al. 2008).

1.2 Human Disease-Related Expression Profiles of miRNAs

1.1.2.2

9

Cellular Functions of miRNAs

miRNAs are an abundant RNA species constituting >3% of the predicted human genes, which regulates ~30% of protein-coding genes (Lim et al. 2005). The high sequence conservation across metazoan species and the ability of individual miRNAs to regulate the expression of multiple genes confer strong evolutionary pressure and participation of miRNAs in essential biologic processes such as cell proliferation, differentiation, apoptosis, metabolism, stress, etc. (Alvarez-Garcia and Miska 2005; Ambros 2004; Wang and Blelloch 2009; Wu et al. 2009a; Yang et al. 2009).

1.2 1.2.1

Human Disease-Related Expression Profiles of miRNAs Spatiotemporal Expression Profiles

Expression of miRNAs in mammalian species under normal conditions is genetically programmed with certain spatial (depending on cell-, tissue-, or organ-type) and temporal (depending on developmental stage) patterns. On one hand, expression of miRNAs is not spatially uniform; instead, some miRNAs are expressed in cell/tissue/organ-restricted manners and others may be ubiquitously expressed. On the other hand, expression of miRNAs is dynamic, but not static, along with the lifespan of an organism from fetal development to aging process. These spatial heterogeneities and temporal differences are critical for the involvement of miRNAs in the fine regulation of versatile cellular functions and and cell lineage decisions with right timings in right places. Chromosomal location and genomic distribution of a miRNA gene are important determinants of its expression from at least three perspectives. (1) Approximately 60% of miRNA genes are located within introns of defined transcription units, and their expression is frequently correlated with the expression profiles of their host genes. (2) Many miRNA genes are distributed as clusters, and a microarray expression profiling of 175 miRNAs in 24 human tissues showed that proximally paired miRNA genes at a distance up to 50 kb are generally co-expressed. The best example may be the four miRNA genes (miR-196b, miR-10a, miR-196a-2, and miR-10b) that are embedded in the Hox gene clusters (Hox A, Hox B, Hox C, and Hox D, respectively). By histochemical staining and in situ hybridization, expression patterns of miR-10a and Hoxb4 mRNA are very similar, suggesting that they share regulatory control of transcription. (3) miRNA genes are frequently located at fragile sites, as well as in regions of loss of heterozygosity, regions of amplification, or common breakpoint regions. Expression of miRNA genes within the regions afflicted by chromosomal aberration, a hallmark characteristic of neoplastic cells, could also be directly affected. For example, miR-15a and miR-16-1 are located

10

1 Detection, Profiling, and Quantification of miRNA Expression

at a frequently deleted site in most of the B cell chronic lymphocytic leukemia (CLL) patients, and induce apoptosis in a leukemia cell line model.

1.2.1.1

Spatial Heterogeneity of miRNA Expression

miRNA Distribution in Mouse Tissues The first attempt to characterize tissue distribution of miRNAs in mammalian species was made by Lagos-Quintana et al. (2002) with mice. They used the cloning method to investigate the tissue-specific distribution of miRNAs in nine different mouse tissues including heart, liver, small intestine, colon, spleen, cortex, cerebellum, and midbrain. For the purpose of tissue-specificity of miRNA expression, cloning of miRNAs from specific tissues is preferred over whole organism-based cloning because low-abundance miRNAs that normally go undetected by Northern blot analysis are identified clonally. They demonstrated that miR-1 accounts for 45% of all mouse miRNAs found in heart, yet miR-1 was still expressed at a low level in liver and midbrain, even though it remained undetectable by Northern analysis. In liver, variants of miR-122 account for 72% of all cloned miRNAs, and miR-122 was undetected in all other tissues analyzed. In spleen, miR-143 appeared to be most abundant, at a frequency of ~30%. In colon, miR-142-as was cloned several times and also appeared at a frequency of 30%. Variants of a particular miRNA, miR-124, dominated and accounted for 25–48% of all brain miRNAs. miR-101, -127, -128, -131, and -132, also cloned from brain tissues, were further analyzed by Northern blotting and shown to be predominantly brain specific (Lagos-Quintana et al. 2002). A study also performed with mice examined miRNA expression profiles from 13 neuroanatomically distinct areas of the adult mouse central nervous system (CNS), employing microarray profiling (see Chap. 2) in combination with real-time RT-PCR (see Chap. 5) and LNA (locked nucleic acid)-based in situ hybridization (see Chap. 4) (Bak et al. 2008). The authors uncovered 44 miRNAs displaying more than threefold enrichment in the spinal cord, cerebellum, medulla oblongata, pons, hypothalamus, hippocampus, neocortex, olfactory bulb, eye, and pituitary gland. These include miR-9, miR-124a, miR-125b, miR-127, miR-128, and members of the let-7 family. This is consistent with some previopus studies (Babak et al. 2004; Barad et al. 2004; Miska et al. 2004; Sempere et al. 2004; Shingara et al. 2005; Thomson et al. 2004). More than 50% of the identified mouse CNS-enriched miRNAs showed different expression patterns compared to those reported in zebrafish, although the mature miRNA sequences are nearly 100% conserved between the two vertebrate species. Their data further revealed that miR-195, miR-497, and miR-30b are enriched in the cerebellum. The medulla oblongata displayed enrichment of miR-34a, miR-451, miR-219, miR-338, miR-10a, and miR-10b. miR-7 and miR-7b were enriched in the hypothalamus. The hippocampus showed accumulation of miR-218, miR-221, miR-222, miR-26a, miR-128a/b,

1.2 Human Disease-Related Expression Profiles of miRNAs

11

miR-138, and let-7c and enrichment of any miRNAs in the amygdala, mesencephalon, and thalamus. It was found that miR-7 and miR-7b were enriched in the pituitary and hypothalamus (Farth et al. 2005); miR-195 in the cerebellum (Hohjoh and Fukushima 2007); miR-375, miR-141, and miR-200a in the pituitary (Landgraf et al. 2007); whereas miR-10a and miR-10b were enriched in the spinal cord (Kloosterman and Plasterk 2006). These findings suggest that a large number of mouse CNS-expressed miRNAs may be associated with specific functions within these regions. Another investigation compared two different methodologies (linear amplification of miRNAs – labeled-aRNA or using a direct labeling strategy – labeled-cDNA) for the preparation of labeled miRNAs from mouse CNS tissue for microarray analysis (Saba and Booth 2006). The most abundant miRNAs, including brain specific or enriched miRNAs, miR-124a-1, -9-1, -9*-1, -127, -136, -138-1, 149, -154, -218-1, -219-1, -222, -125a, -125b-1, -128a, -26a-1, -29a, -29b-1, -30c-1, and -34a were equally identified by both types of microarray labeling techniques. In mouse heart, miR-1, miR-133, miR-125, miR-30, let-7, miR-23, miR-24, miR-26, miR-29, miR-99, and miR-143 are highly enriched. Many miRNAs are enriched in a tissue-/cell-specific manner (Landgraf et al. 2007): miR-1, miR-16, miR-27b, miR-30d, miR-126, miR-133, miR-143, and the let-7 family are abundantly but not exclusively expressed in adult cardiac tissue. In addition to cardiomyocytes, the heart contains many other “non-cardiomyocyte” cell types, such as endothelial cells, smooth muscle cells, fibroblasts, and immune cells, which may have completely distinct miRNA expression profiles. Indeed, (skin) fibroblasts mainly express miR-16, miR-21, miR-22, miR-23a, miR-24, miR-27a, and others, an expression pattern that is highly different from that of cardiomyocytes. In artery smooth muscle, the most abundant miRNAs are miR145, let-7, miR-125b, miR-125a, miR-23, and miR-143 (Ji et al. 2007), despite the fact that the “muscle-specific” miR-1 and miR-133 are also expressed in artery smooth muscle. Other miRNAs, such as the let-7 family, miR-126, miR-221, and miR-222, are highly expressed in human endothelial cells (Kuehbacher et al. 2007; Harris et al. 2008). In addition, miRNA expression profiles can change during cardiac development, and many miRNAs that are only normally expressed at significant levels in the fetal human heart are re-expressed in cardiac disease, such as heart failure (Landgraf et al. 2007; Bauersachs and Thum 2007).

miRNA Distribution in Human Tissues Tissue distribution of miRNAs has also been measured in humans. One report used a bead-based detection platform (see Chap. 21) to profile expression of 217 miRNAs in a broad spectrum of normal human tissues, but low sensitivity and specificity make the results problematic for miRNAs that are less abundant (Lu et al. 2007). Thus far, the study recently reported by Liang et al. (2007) has been the only one systematic, detailed miRNA expression profiling in human tissues. These authors

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1 Detection, Profiling, and Quantification of miRNA Expression

used real-time RT-PCR-based method (see Chap. 5) to examine global profiles of distribution and expression of 345 unique miRNAs in 40 normal human tissues, in combination with public datasets to systematically analyze the association between genomic locations of miRNAs and their expression, and the correlation of expression between miRNAs and their predicted target genes. This study provides a wealth of invaluable information regarding the tissue distribution of miRNAs and the pertinent genomic association, target gene correlation. A number of findings were revealed: 1. A considerable portion of miRNAs have tissue-specific expression patterns and the average miRNA copy numbers in all tissues are highly variable. 2. MicroRNA genes localized within a genomic cluster are preferentially co-expressed as a “transcription unit.” In general, normal human tissues derived from similar anatomical locations or with related physiological functions were primarily clustered together according to the clusters of miRNAs. For example, tissues derived from different parts of heart (atrium versus ventricle) were clustered with skeletal muscle. Hence, localization in the same genomic cluster is the most recognizable feature for miRNAs that have correlated abundance and expression patterns among tissues. 3. The clustering of tissues using the miRNA expression profiles is very similar to that obtained by the mRNA expression profiles. The authors explain this finding on the ground that miRNAs preserve more of the “cellular identity signature” compared to mRNAs under the genomic instability and heterogeneity that characterize neoplastic cells, whereas in normal tissues such a variable environment does not exist, so the performance of both miRNA and mRNA expression profiles on tissue classification is comparable. However, among some tissue types the clustering patterns by their mRNA and miRNA expression profiles are quite different. Lung is clustered together with female reproductive organs and esophagus by mRNA expression profile, but miRNA expression profile of lung is similar only to that of fallopian tube. Thyroid is similar to different parts of the heart in miRNA expression but not in mRNA expression. 4. A group of 15 miRNAs are universally expressed at similar levels in all 40 different normal tissues. These universal miRNAs include miR-15, miR-16b, miR-29a, miR-29aN, miR-29bN, miR-30e, miR-92, miR-92N, miR-93, miR103, miR-106b, miR-140, miR-152b, miR-324-3p, and miR-423. Such a feature characterizes these miRNAs as a candidate of universal reference to normalize miRNA expression in normal human tissues. 5. Seventy-seven percent of the miRNAs have coherent expression patterns with their host genes. This result further corroborates the hypothesis that expression of intronic miRNAs is co-regulated with their host genes, and it also identifies the host genes that could be surrogate markers for expression of their intronic miRNAs. 6. A large number (~100) of miRNAs have pronounced expression in placenta compared to most of the other tissues.

1.2 Human Disease-Related Expression Profiles of miRNAs

13

Important to note is that this human miRNA profiling confirms many of the results from murine studies described earlier. A few examples are given below (Readers are strongly referred to the original article by Liang et al. (2007a); see Table 1.1). 1. miR-1 and miR-133 show highest expression in different parts of the heart and skeletal muscle as well as in vena cava. Other muscle-preferential miRNAs include miR-30, miR-125, miR-26, miR-23, miR-126, miR-92, miR-99, miR100, let-7c, and let-7f. 2. The miRNA expression profile is quite distinct in brain and not shared by other tissue types. Brain expresses high levels of miR-124, miR-221, miR-222, miR514, miR-524, miR-299, miR-320, miR-196, miR-527, miR-34a, and let-7a. 3. Intriguingly, in addition to the expression of liver-specific miR-122a (Jopling et al. 2005), liver also shares similar high levels of expression of several miRNAs as in heart: miR-30, miR-125, miR-26, miR-23, miR-92, miR-148, and miR-126. In addition, miR-192, miR-321, and miR-19 are abundantly expressed in liver but sparsely in other tissues. 4. Strikingly, the gender-related organs/tissues, including testicle, prostate, ovary, fallopian tube, uterus, and breast, express a quite similar set of miRNAs: miR125, miR-26, miR-21, miR-24, miR-30, let-7c, miR-100, miR-99, and miR-92. 5. miR-126, miR-21, miR26, miR-30, miR-24, miR-223, miR-92, and miR-142-3p are among the most abundant miRNAs in human lung. Interestingly, these miRNAs are also highly expressed in spleen. 6. The characteristic set of miRNAs in pancreas includes miR-375, miR-21, miR200c, miR-148, miR-30, miR-26, miR-29, and miR-125. 7. Surprisingly, the muscle-preferential miRNAs miR-133, miR-1, and miR-206 are also among the most abundant miRNA species in thyroid. Moreover, other miRNAs (miR-30, miR-125, miR-26, miR-126, miR-92, let-7c, and let-7f) that are enriched in heart also demonstrate their predominance in thyroid. Apparently, the restriction of miRNA expression can be qualitative (some miRNAs are expressed exclusively in certain tissue or cell types but not in others) or quantitative (some miRNAs are abundantly expressed only in certain tissue or cell types and modestly in others). To be more appropriate, while each individual miRNA may not be expressed in a tissue/cell-specific manner, the expression profile of miRNAs (co-expression of miRNAs) appears to be tissue/cell-specific. The differential tissue distributions of miRNAs suggest tissue– or even cell type– specific functions of these molecules. For instance, the cell lineage-specific miRNA expression patterns may be required to control timing of development and tissue specification. Probably more important is the fact that the expression profile of miRNAs is disease-dependent. A particular pathological process may be associated with the expression of a particular group of miRNAs; this is what the signature expression pattern of miRNAs implies. This issue will be discussed in the following sections of this chapter.

Table 1.1 Top ten abundant miRNAs expressed in various normal human tissues 1 2 3 4 5 6 7 Brain miR-514 let-7 miR-524 miR-299 miR-233 miR-222 miR-124a Heart LV miR-133 miR-1 miR-26 miR-30 miR-24 miR-125 miR-126 RV miR-133 miR-1 miR-26 miR-30 miR-126 miR-24 miR-125 LA miR-133 miR-26 miR-1 miR-125 miR-30 miR-24 miR-126 RA miR-133 miR-26 miR-125 miR-30 miR-1 miR-24 miR-126 Pericardium miR-26 miR-21 miR-16 miR-126 let-7 miR-321 miR-125 Vena Cava miR-26 miR-16 miR-133 miR-125 miR-126 miR-30 let-7 Lung miR-26 miR-126 miR-21 miR-30 miR-223 miR-16 miR-24 Trachea miR-26 miR-16 miR-21 miR-125 miR-200 let-7 miR-24 Liver miR-30e miR-26 miR-30 miR-122 miR-192 miR-92 miR-19 Kidney miR-26 miR-30 miR-21 miR-125 miR-16 miR-29 miR-24 Spleen miR-26 miR-16 miR-223 miR-126 miR-142-3p miR-21 miR-150 Pancreas miR-21 miR-375 miR-26 miR-30 miR-200 miR-16 miR-148 Thyroid miR-133 miR-26 miR-1 miR-26 miR-16 miR-126 miR-30 Esophagus miR-26 miR-125 miR-21 miR-24 miR-16 miR-30 miR-145 Stomach miR-26 miR-200 miR-30 miR-21 miR-375 miR-16 miR-148 Small Intestine miR-26 miR-21 miR-192 miR-321 miR-194 miR-16 miR-92 Colon miR-192 miR-194 miR-26 miR-21 miR-200 miR-215 miR-16 Lymph Node miR-142-3p miR-150 miR-26 miR-16 miR-21 miR-126 miR-29 Thymus miR-26 miR-16 miR-142-3p miR-125 miR-92 miR-20 miR-21 Adrenal miR-26 miR-125 miR-16 let-7 miR-21 miR-126 miR-30 Adipose miR-26 miR-126 miR-125 let-7 miR-16 miR-30 miR-24 Ovary miR-26 miR-125 let-7 miR-92 miR-195 miR-100 miR-99 Uterus miR-26 miR-125 let-7 miR-16 miR-100 miR-24 miR-30 Cervix miR-26 miR-125 let-7 miR-99 miR-100 miR-29 miR-21 Breast miR-26 miR-126 miR-125 miR-30 miR-21 miR-16 let-7 Testicle miR-26 miR-125 let-7 miR-514 miR-16 miR-21 miR-30 Prostate miR-26 miR-125 miR-21 let-7 miR-24 miR-16 miR-27 Bladder miR-26 miR-125 miR-16 miR-21 let-7 miR-126 miR-24 Ileum miR-26 miR-21 miR-192 miR-16 miR-194 miR-30 miR-24 PBMC miR-125 miR-92 miR-29 miR-124 miR-150 miR-30 miR-26 Source: Liang et al. (2007) LV left ventricle; RV right ventricle; LA left atrium; RA right atrium; PBMC peripheral blood mononuclear cells 8 miR-15b miR-23 miR-23 let-7 let-7 miR-199* miR-24 let-7 miR-29 miR-125 let-7 miR-30 let-7 miR-29 miR-27 miR-24 miR-30 miR-321 let-7 miR-150 miR-24 miR-100 miR-30 miR-99 miR-24 miR-24 miR-92 miR-100 miR-30 miR-92 miR-9

9 miR-196b miR-16 miR-27 miR-23 miR-100 miR-92 miR-21 miR-92 miR-126 miR-126 miR-126 miR-92 miR-29 let-7 miR-23 miR-125 miR-200 miR-30 miR-30 miR-30 miR-321 miR-99 miR-199 miR-10 miR-195 miR-30 miR-20 miR-30 miR-321 miR-20 miR-16

10 miR-29bN let-7 let-7 miR-21 miR-99 miR-24 miR-1 miR-29 miR-223 miR-21 miR-192 miR-20 miR-125 miR-125 miR-126 miR-321 miR-20 miR-92 miR-20 miR-321 miR-29 miR-92 miR-29 miR-27 miR-199 miR-195 miR-24 miR-99 miR-100 miR-126 miR-21

14 1 Detection, Profiling, and Quantification of miRNA Expression

1.2 Human Disease-Related Expression Profiles of miRNAs

1.2.1.2

15

Temporal Difference of miRNA Expression

miRNAs in Normal Bone Marrow Cell Lineages Differentiation Hematopoietic stem cells, an example of adult stem cells, are undifferentiated cells that reside in specific niches of the bone marrow and have the capacity to differentiate into any type of blood cell. miRNAs play an important role in hematopoiesis, not only in regulation of hematopoietic stem cell self-renewal but also in hematopoietic differentiation (Lawrie 2007). In mouse BM were identified lineage specific miRNA: miR-223 for myeloid cells and miR-181 for B cells (Chen et al. 2004). miR-150 is usually expressed in mature B and T cells and a premature expression of miR-150 in B cell progenitors stops pro-B cell transition. An up-regulation of miR150 in early B cell development stages blocks the expression of genes that are crucial for B cell maturation and function (Xiao et al. 2007; Fazi et al. 2005). The miR-181a is a key regulator of lymphoid cells differentiation; it is expressed in BM B cells and promotes B cell differentiation possibly through repression of Notch signals (Chen et al. 2005). A screening in naive, effector, and memory T cells showed the importance of miR-21 in T cell differentiation and function (Wu et al. 2007).

miRNA Expression Signatures of Human Mesenchymal Stromal Cells Human mesenchymal stromal cells (MSCs) can produce osteocytes and chondrocytes for bone and cartilage development, and adipocytes for maintaining fat tissue as well. MSCs offer great hope for the treatment of tissue degenerative and immune diseases, but their phenotypic similarity to dermal fibroblasts may hinder robust cell identification and isolation from diverse tissue harvests. To identify genetic elements that can reliably discriminate MSCs from fibroblasts, Bae et al. (2009) performed comparative gene and miRNA expression profiling analyzes with genome-wide oligonucleotides microarrays. They observed similar miRNA expression profiles between MSCs and fibroblasts. A notable exception to the homologous miRNA pattern shared by MSCs and fibroblasts is miR-335 whose expression was found to be about 44-fold higher in MSCs than in fibroblasts. In addition, another four miRNAs, miR-520f, miR-181a-2, miR-340, and miR-431, show >3-fold higher levels in MSCs than in fibroblasts. In agreement, MEST, the host gene for the inronic mirR-335, was also found to be up-regulated in MSCs.

miRNA Expression Signatures of Human Embryonic Stem Cells Embryonic stem (ES) cells share several unique features, including unlimited self-renewal and the ability to differentiate into any of the three embryonal lineages – ectoderm, endoderm, and mesoderm. For the cell fate decision to be made in response to internal and/or niche-specific signals, a complex set of dynamic

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1 Detection, Profiling, and Quantification of miRNA Expression

feedback loops and cross-regulation of pathways is required. Recently, miRNA expression in stem cells has been shown to differ significantly from other cell types tested to date (Houbaviy et al. 2003; Suh et al. 2004; Tang et al. 2006). The role of miRNA-mediated regulation of stem cell division (Hatfield et al. 2005), as well as adipocyte (Esau et al. 2004), cardiac (Zhao et al. 2005), neural (Kuwabara et al. 2004; Wu and Belasco 2005) and hematopoietic lineage differentiation (Chen et al. 2004, 2005), is well known. More recently, a unique set of miRNAs has been shown to be associated with mouse ES cells and embryoid body (EB) formation (Houbaviy et al. 2003; Tang et al. 2006a, b; Murchison et al. 2005). In the study performed by Suh et al. (2004), 14 miRNAs were found to be expressed in a human ES cell-specific manner; miR-302b*, miR-302b, miR-302c*, miR-302c, miR-302a*, miR-302d, miR-367, miR-200c, miR-368, miR-154*, miR371, miR-372, miR-373*, and miR-373 (Suh et al. 2004). This study suggests that the expression patterns of miRNAs cloned from ES cells can be classified into four groups: (1) miRNAs that are expressed in ES cells as well as in EC cells; miR302b*, miR-302b, miR-302c*, miR-302c, miR-302a*, hsc-3, miR-302d, and miR367. These miRNAs may have conserved roles in mammalian pluripotent stem cells. (2) miRNAs that are expressed specifically in ES cells but not in other cells including EC cells; miR-200c, miR-368, miR-154*, miR-371, miR-372, miR-373*, and miR-373. These miRNAs may have functions specific to ES cells. (3) miRNAs that are rare in ES cells but abundant in HeLa and STO cells; let-7a, miR-301 (hsc11), miR-374, miR-21, miR-29b, and miR-29. These stage-specific miRNAs may play roles in the regulation of development and differentiation, like let-7 in C. elegans. (4) The last class consists of miR-16, miR-17-5p, miR-19b, miR-26a, miR-92, miR-103, miR-130a, and miR-222. These are expressed in most tested cell lines, so they may contribute to basic cellular functions. Lakshmipathy et al. (2007a, b) identified the differences in miRNA expression between undifferentiated ES cells and their corresponding differentiated cells that underwent differentiation in vitro over a period of 2 weeks, confirming the identity of a signature miRNA profile in pluripotent cells, comprising a small subset of differentially expressed miRNAs in ES cells. ES cells express high levels of miR200c, miR-371, miR-372, miR-302a, miR-320d, miR-373, miR-302c, miR-21, miR-222, miR-296, miR-494, miR-367, miR-miR154, miR-29a, miR-143, miR29c, and let7a, relative to their corresponding differentiated cells. By comparison, miR17M, miR-92, and miR-93 are more abundantly expressed in differentiated cells than in ES cells.

miRNA Signatures During Human Erythroid Cell Differentiation Although the stages of erythroid cell differentiation are well-characterized, the molecular mechanisms that orchestrate the coordinated changes from erythroid lineage commitment to terminal maturation remain poorly understood. In a study documented by Zhan et al. (2007), the authors explored the expression profile of miRNAs in erythroid cells at different stages of differentiation using miRNA

1.2 Human Disease-Related Expression Profiles of miRNAs

17

microarray analysis. Real-time RT-PCR was used to confirm the results of miRNA microarray. MEL cells were used, which are derived from Friend virus transformed mouse spleen erythroid precursors that are blocked at about the pronormoblast stage of differentiation. Their studies show that, of 295 miRNAs assayed, more than 100 are expressed in erythroid cells with varied abundances. Of the miRNAs on the array, miR-298 was the most abundant miRNA with signals of about 50,000 in uninduced MEL cells. miR-320 was the second most abundant miRNA in uninduced MEL cells. miR-29b, miR-140*, miR-193, miR-382, and miR-434-5p, which were undetectable in untreated MEL cells, became detectable following induction of erythroid differentiation In contrast, the levels of both miR-298 and miR-320, the two most abundant miRNAs in untreated MEL cells, decreased upon induction of erythroid maturation. The level of miR-451 is the most significantly increased (more than sevenfold), whereas the levels of several miRNAs, such as miR-29a, miR-26a, miR-22, miR-144, miR-15b, miR-292-5p, and miR-30a-5p, increased more than twofold upon induction of erythroid differentiation. Functional studies using gain of function and loss of function approaches showed that miR-451 is associated with erythroid maturation. The findings indicate that dynamic changes in miRNA expression occurred during erythroid differentiation, with an overall increase in the levels of miRNAs upon terminal differentiation of erythroid cells.

1.2.2

miRNA Transcriptome and Human Physiology

1.2.2.1

Glucose-Regulated miRNAs from Pancreatic b Cells

Tang et al. (2009b) carried out a screen in the pancreatic b-cell line MIN6 to identify miRNAs with altered abundance in response to changes in glucose concentrations. This screen resulted in identification of 61 glucose-regulated miRNAs from a total of 108 miRNAs detectable in MIN6 cells. Fifty of the identified miRNAs, including miR-124a, miR-107, and miR-30d, are up-regulated in the presence of high glucose. Only a few of the miRNAs, including miR-296, miR484, miR-612, miR-638, and miR-690, are significantly down-regulated by high glucose treatment. Overexpression of miR-30d increases insulin gene expression, while inhibition of miR-30d abolished glucose-stimulated insulin gene transcription. Overexpression or inhibition of miR-30d has no effect on insulin secretion. The findings suggest that the putative target genes of miR-30d may be negative regulators of insulin gene expression. Several miRNAs including miR-375 and miR-124a, which are highly expressed. in pancreatic b cells cells (Baroukh et al. 2007; Kloosterman et al. 2007; Poy et al. 2007), have been implicated to negatively regulate insulin exocytosis (Poy et al. 2004; Krek et al. 2005; Lovis et al. 2008a, b). In addition, both miR-375 and miR124a appear to have important roles in pancreas development (Baroukh et al. 2007; Kloosterman et al. 2007; Poy et al. 2007).

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1.2.2.2

1 Detection, Profiling, and Quantification of miRNA Expression

miRNAs in Human Omental and Subcutaneous Adipose Tissue

miRNAs have been shown to regulate metabolic processes, which are associated with type 2 diabetes including insulin signaling and glucose homeostasis (Poy et al. 2004; Gauthier and Wollheim 2006). To gain further insight into the association between miRNA expression in human adipose tissue and parameters of obesity, fat distribution, and glucose metabolism, Klo¨ting et al. (2009) performed a global miRNA gene expression array in paired omental and abdominal subcutaneous adipose tissue samples from 15 overweight or obese individuals with either normal glucose tolerance or type 2 diabetes. Expression of 155 miRNAs was carried out using the TaqManHMicroRNA Assays Human Panel Early Access Kit (Applied Biosystems, Darmstadt, Germany). These authors identified expression of 106 (68%) miRNAs in human omental and subcutaneous adipose tissue, but did not observe any miRNAs exclusively expressed in either fat depot, suggesting common developmental origin of both fat depots. They further identified significant correlations between the expressions of miRNA-17-5p, miR-132, miR-99a, miR-134, miR-181a, miR-145, and miR-197 and both adipose tissue morphology and key metabolic parameters, including visceral fat area, HbA1c, fasting plasma glucose, circulating leptin, adiponectin, and interleukin-6. They suggested that miRNA expression differences may contribute to intrinsic differences between omental and subcutaneous adipose tissue; human adipose tissue miRNA expression correlates with adipocyte phenotype, parameters of obesity, and glucose metabolism.

1.2.2.3

miRNAs in Circadian Rhythmicity

The daily cycling of light and temperature, generated by the earth’s rotation, is one of the most important driving forces in the evolution of the circadian clock, allowing organisms to anticipate and adapt to their daily (and seasonally) changing environment. Recently, a few studies have suggested that miRNAs may be important regulators of circadian rhythmicity, providing a new dimension (posttranscriptional) of our understanding of biological clocks. In one study with mice, miR-132 and miR-219-1 show daily oscillation in the suprachiasmatic nucleus (SCN), but not in other regions, peaking during the subjective day, which was abolished in mCry1/mCry2 double mutant, strongly indicating a clock control of the expression of these miRNAs in vivo. In another recent study on the mouse, 78 miRNAs were found expressed in adult mouse retina, 21 of which are potentially retina-specific (Xu et al. 2007b). This study identified a polycistronic, sensory organ-specific paralogous miRNA cluster that includes miR-96, miR-182, and miR-183 on mouse chromosome 6qA3 with conservation of synteny to human chromosome 7q32.2. In situ hybridization showed that members of this cluster are expressed in photoreceptors, retinal bipolar and amacrine cells. To identify miRNAs potentially involved in circadian rhythm regulation of the retina, these authors performed miRNA expression profiling with retinal RNA harvested at noon (Zeitgeber time 5) and midnight (Zeitgeber time 17) and identified a subgroup of 12 miRNAs,

1.2 Human Disease-Related Expression Profiles of miRNAs

19

including members of the miR-183/96/182 cluster with diurnal variation in expression pattern. The daily cycling of expression of a number of miRNAs including miR-96, miR-124a, miR-103, miR-182, miR-106b, miR-422a, and miR-422b was found in the retina.

1.2.2.4

miRNAs in Human Pregnancy and Parturition

Montenegro et al. (2009) performed to determine gestational age-dependent changes in miRNA expression in the chorioamniotic membranes and to assess the significance of miRNAs in human pregnancy and parturition. The expression profile of 455 miRNAs was compared between patients at term without labor, in labor, and preterm labor using microarrays. A total of 39 miRNAs are differentially expressed between term and preterm cases, of which 31 (79.5%) are downregulated at term. A comparison between the preterm labor and term labor groups revealed differential expression of ten miRNAs, all of which are down-regulated at term. A comparison between the preterm labor and term without labor cases also showed decreased expression of 28 miRNAs and increased expression of ten miRNAs at term. The down-regulation of nine miRNAs at term (miR-25, miR338, miR-101, miR-449, miR-154, miR-135a, miR-142-3p, miR-202∗, and miR136) was shared by the two groups compared, preterm labor versus term labor and preterm labor versus term without labor. Overall, the majority (79.5%) of all differentially expressed miRNAs have decreased expression at term. Decreased expression of miR-338, miR-449, miR-136, and miR-199a∗ at term was confirmed by qRT-PCR.

1.2.2.5

Gender difference of miRNAs in liver

It has been shown that hepatic transcript profiles are different in men and women. An interesting study tested the role of miRNAs in the gender-differences of gene expression (Cheung et al. 2009). Using microarrays, miRNA screening was performed to identify sex-dependent miRNAs in rat liver. Out of 324 unique probes on the array, 254 were expressed in the liver and eight (3% of 254;) of those were found to be different between the sexes, with miR-21, miR-148a, miR-451, and miR-526c being male-predominant and miR-29b, miR-122a, miR-193, and miR205 being female-predominant. Among the eight putative sex-different miRNAs, only one female-predominant miRNA (miR-29b) was confirmed using quantitative real-time PCR. Furthermore, 1 week of continuous growth hormone treatment in male rats reduced the levels of miR-451 and miR-29b, whereas mild starvation (12 h) raised the levels of miR-451, miR-122a, and miR-29b in both sexes. The biggest effects were obtained on miR-29b with growth hormone treatment. It appears that hepatic miRNA levels depend on the hormonal and nutritional status of the animal and that miR-29b is a female-predominant and growth hormoneregulated miRNA in rat liver.

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1.2.3

1 Detection, Profiling, and Quantification of miRNA Expression

miRNA Transcriptome in Diseased States

The first evidence of the involvement of miRNA in a human disease was obtained in 2002, when for the first time a deregulation of two miRNAs (miR-15a and miR-161) was related to CLL (Calin et al. 2002). Thus, if a disease occurs specifically in a given tissue, the miRNAs specifically expressed in that tissue will have a great potential to be related to that disease. The dissection of these relationships will be valuable for studying the functions of these miRNAs and their mechanisms in diseases and can be used to discovering novel disease-associated miRNAs. For example, 5 of the 6 papers revealed a down-regulation of let-7 in cancers, which is consistent with previous report, while all papers reported an up-regulation of let-7 in Alzheimer’s disease (see our HMDD database). 1. miRNAs tend to show similar or different dysfunctional evidences for the similar or different disease clusters, respectively. 2. A negative correlation between the tissue specificity of a miRNA and the number of diseases it associated. 3. An association between miRNA conservation and disease. 4. miRNAs associated with the same disease tend to emerge as predefined miRNA groups. 5. Disease-associated miRNAs show various dysfunctions, such as mutation, up-regulation, deletion, and down-regulation. 6. The miRNAs that have a higher degree of conservation tend to be associated with diseases with a higher probability significantly. miRNA conservation is associated with human disease susceptibility. 7. Revealed that miRNAs in 57% of the diseases have at least one family member in that disease associated miRNAs, which is significantly higher than the random. This finding suggested that the miRNA family members might have similar functions and play roles in similar biological processes and therefore their dysfunction would lead to similar phenotype. 8. Bartel and his colleagues reported that neighboring miRNAs show significant coexpression by a microarray profiling analysis (Farth et al. 2005). We found that miRNAs in 46% of the diseases have at least one neighboring member, which is significantly higher than the random. For example, all the six miRNAs implicated in hematopoietic malignancies are located in the miR-17 cluster. This result indicated that neighboring miRNAs might be regulated by common regulators at similar conditions and function together, and then their dysfunctions would result in the same disease.

1.2.3.1

Cancers

The initial study establishing a role for miRNAs in cancer came with the observation that two miRNA genes, miR-15a and miR-16a, are deleted in the majority of B

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cell chronic lymphocytic leukemia patients (Calin et al. 2002). Since then, roles for miRNAs in various cancers have been reported, primarily on the basis of differential miRNA profiling between cancer and normal tissue or from genetic screens with miRNA expression libraries. An important step toward the understanding of miRNA transcriptomes as biomarkers of human cancers was reached by the study reported by Volinia et al. (2006). In this study, the authors conducted a large-scale miRNA transcriptome analysis on 540 samples of six solid tumors including lung, breast, stomach, prostate, colon, and pancreatic tumors, and were able to identify a solid cancer miRNA signature composed by a large portion of overexpressed miRNAs. The up-regulated miRNAs in three or more types of solid cancers include miR-17-5p/miR-20a/miR-92-2, miR-21, miR-24, miR-25, miR-29, miR-30c, miR32, miR-106a, miR-107, miR-128b, miR-146, miR-155, miR-181-1b, miR-191, miR-199a-1, miR-214, and miR-221. The predicted targets for the differentially expressed miRNAs are significantly enriched for protein-coding tumor suppressors and oncogenes. Nonetheless, apart from the common panel aberrantly altered miRNAs, each type of tumor entities has its own miRNA signature, which can be used as a biomarker for distinguishing the types and progonosis of cancers.

Solid Cancers Probably the most classical study linking miRNAs and human cancers is the one reported by Volinia et al. (2006), describing a large-scale detailed analysis of the miRNA profiles in 540 samples from six solid tumors. The clustering of miRNA expression profiles derived from 228 miRNAs in 363 solid cancer and 177 normal samples. Comparison of all tumors against all normal tissues identified 26 overexpressed and 17 underexpressed miRNAs, out of 137 miRNAs expressed in at least 90% of the samples and shows a very good separation between the different tissues. These results indicated that, in solid cancers, the spectrum of expressed miRNAs is very different from that of normal cells (43 of 137 miRNAs, 31%). Among these miRNAs are some with well characterized cancer association, such as miR-17-5p, miR-20a, miR-21, miR-92, miR-106a, and miR-155. Strikingly, miR21, miR-191, and miR-17-5p are significantly overexpressed in all the tumor types examined or in five of six. The predicted targets for the differentially expressed miRNAs are significantly enriched for protein-coding tumor suppressors and oncogenes.

Breast Cancer The human “breast”-specific signature is characterized by the expression profile of 23 miRNA (Heneghan et al. 2009). In this study, the breast has been reported to be the tissue with the lower number of detected miRNA. Expression of 222 premiRNA was studied by real-time PCR in 32 commonly used cell lines, including

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1 Detection, Profiling, and Quantification of miRNA Expression

five breast cancer ones (T47D, SKBR3, MDA361, MCF7, and MDA231) (Jiang et al. 2005). This study revealed that let-7f-1 is increased by seven-fold in the epithelial-derived breast, lung, and colorectal cancer cells (Silveri et al. 2006). The miRNA microarray was used to evaluate miRNA expression profiles in ten normal and 76 neoplastic breast tissues (Iorio et al. 2005). Twenty-nine miRNAs were found to be differentially regulated, of which a set of 15 could be used to predict the nature of the cell sample analyzed with 100% accuracy (i.e., tumor or normal breast tissue). Expression of some miRNA could be correlated with specific breast cancer biopathologic features, such as estrogen and progesterone receptor expression, tumor stage, vascular invasion, or proliferation index. Among the differentially expressed miRNAs, miR-10b, miR-125b, miR145, miR-21, and miR-155 emerged as the most consistently deregulated in breast cancer. Three of them, miR-10b, miR-125b, and miR-145, were down-regulated and the remaining two, miR-21 and miR-155, were up-regulated, suggesting that they may potentially act as tumor suppressor genes or oncogenes, respectively. miR-145 and miR-21, whose expression could differentiate cancer versus normal tissues, were also differentially expressed in cancers with different proliferation indexes or different tumor stages. In particular, miR-145 is progressively down-regulated from normal breast to cancer with high proliferation index. Similarly, but in opposite direction, miR-21 is progressively up-regulated from normal breast to cancers with high tumor stage. It was observed that breast cancer primary tumors have a decreased expression level of miR-125b compared to normal breast tissue, suggesting that downregulation of miR-125b impairs the differentiation capability of cancer cells.

Lung Cancer Yanaihara et al. (2006) found that expression levels of the five miRNAs (hsa-mir155, hsa-mir-17-3p, hsa-mir-let-7a-2, hsa-mir-145, and hsa-mir-21) were statistically altered in lung cancers, and these also had a prognostic effect on patient survival. An expression meta-analysis of predicted gene targets of three lung-enriched miRNAs, miR-34b/miR-34c/miR-449, identifies a diagnostic signature for lung cancer (Liang 2008). The study by Markou et al. (2008) suggests that overexpression of mature miR21 is an independent negative prognostic factor for overall survival in non–small cell lung cancer patients.

Liver Cancer The miRNA expression profiles in a large set of 52 human primary liver tumors consisting of premalignant dysplastic liver nodules and hepatocellular carcinomas were examined by qRT-PCR (Varnholt et al. 2008). Eighty miRNAs were examined

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in a subset of tumors, which yielded 10 up-regulated and 19 down-regulated miRNAs compared to normal liver. Subsequently, five miRNAs (miR-122, miR100, miR-10a, miR-198, and miR-145) were selected on the basis of the initial results and further examined in an extended tumor sample set of 43 hepatocellular carcinomas and nine dysplastic nodules. miR-122, miR-100, and miR-10a were overexpressed whereas miR-198 and miR-145 were up to five-fold down-regulated in hepatic tumors compared to normal liver parenchyma.

Colon Cancer Dı´az et al. (2008) assessed in 110 colon cancer patients the levels of miR-17-5p, miR-106a, miR-126, E2F1, and EGFL7 by quantitative real-time RT-PCR. Altered expression of miR-17-5p, miR-106a, and EGFL7 was associated with pathological tumor features of poor prognosis. Down-regulation of miR-106a predicted shortened disease-free survival and overall survival. miR-17-5p correlated with disease-free survival only at early stages. Inverse correlations were found between miR-17-5p and miR-106a levels and their target expression, E2F1. No correlation was found between miR-126 expression and its host gene levels, EGFL7. miR-106a deregulation was therefore considered as a marker of disease-free survival and overall survival independent of tumor stage. The lack of association between expression of miR-126 and its host gene EGFL7 suggests their regulation by independent stimuli. Inverse correlation between miR-17-5p and miR-106a and E2F1 levels supports E2F1 as a target mRNA for the two miRNAs. In a separate study, Schetter et al. (2008) reported that 37 miRNAs were differentially expressed in colon tumors from the test cohort. Selected for validation were miR-20a, miR-21, miR-106a, miR-181b, and miR-203, and all five were enriched in tumors from the validation cohort. Higher miR-21 expression was present in adenomas and in tumors with more advanced TNM staging. In situ hybridization demonstrated miR-21 to be expressed at high levels in colonic carcinoma cells. The 5-year cancer-specific survival rate was 57.5% for the Maryland cohort and was 49.5% for the Hong Kong cohort. High miR-21 expression was associated with poor survival, independent of clinical covariates, including TNM staging, and was associated with a poor therapeutic outcome. Among the miRNAs identified, some have a well-characterized association with colon cancer progression, eg miR-10b, miR-21, miR-30a, miR-30e, miR125b, miR-141, miR-200b, miR-200c, and miR-205 (Baffa et al. 2009). The expression levels of miR-143 and miR-145 were decreased in human colon tumors (Akao et al. 2007; Schepeler et al. 2008; Takagi et al. 2009). A decrease of miR101 levels could represent one of the leading causes of COX-2 overexpression in colon cancer cells (Strillacci et al. 2009). Guo et al. reported a ubiquitous loss of miR-126 expression in colon cancer lines when compared to normal human colon epithelia that may provide a selective growth advantage during colon carcinogenesis (Guo et al. 2008a).

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1 Detection, Profiling, and Quantification of miRNA Expression

Colorectal Cancer Role of miRNAs in human colorectal cancer has been evidenced by several studies (Tang and Fang 2009). Ng et al. (2009) tested miRNAs using real-time PCR-based miRNA profiling on plasma and tissues from colorectal cancer. Of the panel of 95 miRNAs analyzed, five miRNAs were up-regulated both in plasma and tissue samples (miR-17-3p, miR-135b, miR-222, miR-92, and miR-95). The plasma levels of these markers were significantly reduced after surgery in ten colorectal cancer patients. Further validation with an independent set of plasma samples indicated that miR-92 differentiates colorectal cancer from gastric cancer, IBD, and normal subjects. Another study presented the expression of 156 mature miRNAs with several bioinformatics algorithms in colorectal tumors and adjacent non-neoplastic tissues from patients and colorectal cancer cell lines (Bandre´s et al. 2006). A group of 13 miRNAs were significantly altered in their expression in this tumor. The most significantly deregulated miRNA being miR-31, miR-96, miR-133b, miR-135b, miR-145, and miR-183. In addition, the expression level of miR-31 was correlated with the stage of colorecta tumor.

Gastric Cancer Human gastric cancer is among the major causes of cancer mortality worldwide. Using miRNA microarray assay, a group reported the miRNA expression profile in gastric cancer as compared with non-tumor tissues. The study revealed that the most highly expressed miRNAs in non-tumorous tissues are miR-768-3p, miR-1395p, miR-378, miR-31, miR-195, miR-497, and miR-133b. The most highly expressed miRNAs in gastric cancer tissues include miR-20b, miR-20a, miR-17, miR-106a, miR-18a, miR-21, miR-106b, miR-18b, miR-421, miR-340*, miR-19a, and miR-658. Unfortunately, verification of this expression pattern was not done with more accurate methods. The expression levels of three miRNAs (miR-34b, miR-34c, and miR-128a) were significantly up-regulated and those of three miRNAs (miR-128b, miR-129, and miR-148) were down-regulated in undifferentiated gastric cancer tissue when compared with those of the paired normal tissues (Katada et al. 2009). The probability of survival was significantly lower in patients with high expression levels of miR-20b or miR-150. There was a correlation between miR-27a and lymph node metastasis.

Esophageal Cancer Esophageal cancer is the sixth leading cause of death from cancer and one of the least studied cancers worldwide. With cryopreserved esophageal cancer tissues using advanced microRNA microarray techniques, Guo et al. (2008a, b) identified

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seven miRNAs that could distinguish malignant esophageal cancer lesions from adjacent normal tissues. Among the seven miRNAs, three miRNAs (miR-25, miR424, and miR-151) showed up-regulation and four miRNAs (miR-100, miR-99a, miR-29c, and mmu-miR-140*) showed reduction in cancer versus normal tissue. Some of these miRNAs could be correlated with the different clinicopathologic classifications. High expression of miR-103 and miR-107 correlates with poor survival. Five miRNAs (miR-335, miR-181d, miR-25, miR-7, and miR-495) correlate with gross pathologic classification (fungating versus medullary) and two miRNAs (miR-25 and miR-130b) correlate with differentiation classification (high versus middle versus low).

Thyroid Papillary Carcinomas A significant increase in miRNAs miR-221, miR-222, and miR-181b was detected in thyroid papillary carcinomas in comparison with normal thyroid tissue (Pallante et al. 2006). These results were further confirmed by Northern blot and quantitative RT-PCR analyzes. Moreover, RT-PCR revealed miR-221, miR-222, and miR-181b overexpression in fine needle aspiration biopsies corresponding to thyroid nodules, which were eventually diagnosed as papillary carcinomas after surgery. Finally, miR-221, miR-222, and miR-181b overexpression was also demonstrated in transformed rat thyroid cell lines and in mouse models of thyroid carcinogenesis. Functional studies, performed by blocking miR-221 function and by overexpressing miR-221 in human thyroid papillary carcinomas-derived cell lines, suggest a critical role of miR-221 overexpression in thyroid carcinogenesis.

Pancreatic Cancer Differential expression of 95 miRNAs with their potential functions related to cancer biology, cell development, and apoptosis was analyzed by qRT-PCR for pancreatic cancer tissue samples or cancer cell lines in comparison with those in relatively normal pancreatic tissues or normal human pancreatic ductal epithelial (HPDE) cells (Zhang et al. 2009). Human pancreatic tissue with chronic pancreatitis also was included for analysis. Analysis was performed on ten pancreatic cancer cell lines and 17 pairs of pancreatic cancer/normal tissues. Eight miRNAs were significantly up-regulated in most pancreatic cancer tissues and cell lines, including miR-196a, miR-190, miR-186, miR-221, miR-222, miR-200b, miR-15b, and miR95. The incidence of up-regulation of these eight genes between normal control subjects and tumor cells or tissues ranged from 70 to 100%. The magnitude of increase of these miRNAs in pancreatic cancer samples ranged from 3- to 2,018fold of normal control subjects. Roldo et al. (2006) showed that the expression of has-miR-103 and has-miR-107 and lack of expression of has-miR-155 could discriminate pancreatic tumors from normal pancreatic tissues.

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1 Detection, Profiling, and Quantification of miRNA Expression

Hematological Cancers Acute myeloid leukemia (AML). AML is the most common acute leukemia in adults; it is characterized by clonal expansion of hematopoietic stem cells blocked at different stages of erythroid, granulocytic, monocytic, or megakaryocytic differentiation. It was shown that miRNA expression profiles are linked to the karyotype (Dixon-McIver et al. 2008) and expression of a specific miRNA (miR-181a) correlated with AML morphological subtype (Debernardi et al. 2007). miRNA signatures were associated with cytogenetic abnormalities in AML and the high expression of miR-191 and miR-199a correlated with patients having poor prognosis (Garzon et al. 2008). Chronic myeloid leukemia (CML). CML is a multi-step chronic BM disorder involving progression from chronic phase to an accelerated phase characterized by a translocation involving chromosomes 9 and 22, generating the Philadelphia chromosome. In CML CD34þ cells from patients from chronic phase, the miR17–92 polycistron, also called oncomir-1 comprising seven miRNA (miR-17-5p, miR-17-3p, miR-18a, miR-19a, miR-19b-1, miR-20a, and miR-92a-1) was up-regulated compared to blast phase (Venturini et al. 2007). Acute lymphocytic leukemia (ALL). ALL arises from either T or B lymphocyte precursors; however, B-ALL is the most common type. ALL is the predominant cancer in childhood and has a favorable prognosis compared to AML. Using the TaqMan MicroRNA Assays Human Panel (Applied Biosystems), Zanette et al. (2007) analyzed miRNA expression profiles in leukemia samples and CD19þ samples from healthy individuals.The five most highly expressed miRNAs were miR-128b, miR-204, miR-218, miR-331, and miR-181b-1 in ALL, and miR-331, miR-29a, miR-195, miR-34a, and miR-29c in CLL (Zanette et al. 2007), allowing for distinction between ALL and CLL. The miR-17-92 cluster was also found to be up-regulated in ALL. Another study demonstrated that several miRNAs were differentially expressed between AML and ALL, where miR-128a, miR-128b, miR-223, and let-7b were the most significant and discriminatory. The authors found that a signature of only two of these four miRNA was sufficient to discriminate these diseases. A subset of AML patients showed up-regulation of miR-155, which may repress genes implicated in hematopoietic development and disease (O’Connell et al. 2008). Chronic lymphocytic leukemia (CLL). CLL is characterized by elevated number of clonal lymphocytes B in circulation, usually arrested in G0/G1 phase. Calin et al. (2004a) reported that a unique 13-miRNA expression signature (hsa-miR-15a, hsamiR-195, hsa-miR-221, miR-23b, miR-155, miR-223, miR-29a-2, miR-24-1, miR29b-2, miR-146, miR-16-1, miR-16-2, and miR-29c) was a prognostic indicator of CLL. miR-15a and miR-16 are in a cluster located in 13q14.3, a chromosome region frequently deleted in CLL patients, which could explain the loss or downregulation of these miRNAs (Calin et al. 2002). These two miRNAs have potential for CLL prognosis: patients with good prognosis showed down-regulation of miR15a and miR-16, whereas bad prognosis was associated with down-regulation of miR-29 (Calin et al. 2002, 2004a). The miR-143 and miR-145 could also be used as

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disease markers, in combination with miR-15a and miR-16, as they were significantly down-regulated in B-cell diseases, including CLL (Akao et al. 2007). By using a cloning-based method, miR-21, miR-150, and miR-155 were shown to be up-regulated in CLL (Fulci et al. 2007), while miR-181a, let-7a and miR-30d were down-regulated (Marton et al. 2008). In addition, Marton et al. (2008) identified five new miRNA (miR-1200, let-7i*, miR-1201, miR-1202, and miR-1203) associated to CLL. Zanette et al. (2007) described up-regulation of miR-331, miR-29a, miR-195, miR-34a, and miR-29c in CLL.

Hodgkin Lymphoma Hodgkin lymphoma (HL) is derived from preapoptotic germinal center B cells, although a general loss of B cell phenotype is noted. HL is one of the most frequently occurring lymphomas, with an annual incidence rate of three to four new cases per 100,000 persons in the Western world. Using quantitative reverse transcription–polymerase chain reaction and miRNA microarray, we determined the miRNA profile of HL and compared this with the profile of a panel of B-cell non–Hodgkin lymphomas. The two methods showed a strong correlation for the detection of miRNA expression levels. The HL-specific miRNA included miR-1792 cluster members, miR-16, miR-21, miR-24, and miR-155. Using a large panel of cell lines, we found differential expression between HL and other B-cell lymphoma-derived cell lines for 27 miRNAs. A significant down-regulation in HL compared to non-Hodgkin lymphoma was observed only for miR-150.

Ovarian Cancer Epithelial ovarian cancer is the sixth most common cancer in women worldwide. In comparison to normal ovary, miRNAs are aberrantly expressed in human ovarian cancer. The most significantly overexpressed miRNAs were miR-200a, miR-141, miR-200c, and miR-200b, whereas miR-199a, miR-140, miR-145, and miR-125b1 were among the most down-modulated miRNAs (Iorio et al. 2007). The overall miRNA expression could clearly separate normal versus cancer tissues. The expression of these miRNAs can also be correlated with specific ovarian cancer biopathologic features, such as histotype, lymphovascular and organ invasion, and involvement of ovarian surface.

Prostate Cancer Expression profile of 40 prostatectomy specimens from stage T2a/b, early relapse, and non-relapse cancer patients was examined (Tong et al. 2009). Paired analysis was carried out with microdissected, malignant and non-involved areas of each specimen, using high-throughput liquid-phase hybridization (mirMASA) reactions

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and 114 miRNA probes. Five miRNAs (miR-23b, -100, -145, -221, and -222) were significantly down-regulated in malignant tissues. Ectopic expression of these miRNAs significantly reduced LNCaP cancer cell growth, suggesting growth modulatory roles for these miRNAs. Patient subset analysis showed that those with post-surgery elevation of prostate-specific antigen (chemical relapse) displayed a distinct expression profile of 16 miRNAs, as compared with patients with non-relapse disease. A trend of increased expression (>40%) of miR-135b and miR-194 was confirmed by qRT-PCR in 11 patients from each clinical subset.

Kidney Cancer The expression profiles of 245 miRNAs in kidney primary tumors were analyzed using a microarray containing 245 human and mouse miRNA genes (Gottardo et al. 2007). The specimens include a total of 27 kidney specimens (20 carcinomas, four benign renal tumors, and three normal parenchyma). A set of four human miRNAs (has-miR-28, has-miR-185, has-miR-27, and has-let-7f-2) were found significantly up-regulated in renal cell carcinoma compared to normal kidney. Of the kidney cancers studied, there was no differential miRNA expression across various stages, whereas with increasing tumor-nodes-metastasis staging in bladder cancer, miR26b showed a moderate decreasing trend.

Bladder Cancer Urothelial carcinoma is the most common form of cancer in the bladder and can be divided into two groups: the low-grade tumors which are always papillary and usually superficial, and the high-grade tumors which can be either papillary or nonpapillary and often invasive. Superficial tumors account for 75–80% of bladder neoplasms, while the remaining 20–25% are invasive or metastatic. Neely et al. (2009) performed microarray analysis and identified several miRNAs that were differentially expressed between the noninvasive and invasive bladder carcinoma cell lines including miR-21, miR-31, miR-200a, miR-200c, miR-205, miR-373*, miR-487b, miR-498, and miR-503. Cell lines characterized as invasive showed a miR-21:miR-205 ratio at least ten-fold higher than the quantitative ratio obtained from non-invasive cell lines. miR-21 and miR-205 expression levels were further determined in 53 bladder tumors (28 superficial and 25 invasive). The results suggest a miR-21:miR-205 expression ratio as a biomarker for distinguishing between invasive and noninvasive bladder tumors with high sensitivity and specificity. miRNA expression was analyzed in 27 bladder specimens (25 urothelial carcinomas and two normal mucosa) (Gottardo et al. 2007). Human miRNAs hasmiR-223, has-miR-26b, has-miR-221, has-miR-103-1, has-miR-185, has-miR-23b, has-miR-203, has-miR-17-5p, has-miR-23a, and has-miR-205 were significantly up-regulated in bladder cancers compared to normal bladder mucosa.

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Clear Cell Renal Cell Carcinoma A genome-wide expression profiling of miRNAs using microarray analysis and qRT-PCR was performed in clear cell renal cell carcinoma (ccRCC), with matched malignant and non-malignant tissue samples from two independent sets of 12 and 72 ccRCCs. The microarray-based experiments identified 13 overexpressed and 20 down-regulated miRNAs in malignant samples. Expression in ccRCC tissue samples compared to non-malignant samples measured by RT-PCR was increased on average by 2.7- to 23-fold for the miR-16, miR-452*, miR-224, miR-155, and miR-210, but decreased by 4.8- to 138-fold for miR-200b, miR-363, miR-429, miR-200c, miR-514, and miR-141. No significant associations between these differentially expressed miRNAs and the clinico-pathological factors tumor stage, grade, and survival rate were found. Nevertheless, malignant and non-malignant tissue could clearly be differentiated by their miRNA profiles. A combination of miR-141 and miR-155 resulted in a 97% overall correct classification of samples.

Endometrioid Adenocarcinoma The miRNA expression profile was studied in ten pairs of endometrioid adenocarcinoma and adjacent nontumorous endometrium using human miRNA microarray (Wu et al. 2009a, b). Seventeen miRNAs exhibited higher expression and six miRNAs exhibited lower expression in endometrioid adenocarcinoma samples than those in the nontumorous samples in the microarray. Of those, the miR-205, miR-449, and miR-429 were greatly enriched; in contrast, the miR-204, miR-99b, and miR-193b were greatly down-regulated in adenocarcinoma tissues. The expressions of these six miRNAs were validated using real time reverse transcription-PCR.

Head and Neck Cancer Head and neck cancer is the term given to a variety of malignant tumors that develop in the oral cavity, larynx, pharynx, and salivary glands and are predominantly squamous cell carcinomas. Molecular signatures using gene expression analysis have been identified that describe these tumors by location. However, head and neck squamous cell carcinoma (HNSCC) of the oral cavity is noted for its heterogeneity and has defied substantive molecular classification. In contrast to gene expression analysis of tumors, a relatively small number of miRNAs can be used to classify tumors and thus less heterogeneity should exist in miRNA expression profiles of various HNSCCs. Ramdas et al. (2009) studied miRNA expression profiles in HNSCC and adjacent normal tissue. They found that several miRNAs were determined to be differentially regulated in the HNSCC samples when compared with their normal tissue counterparts: miR-7, miR-15b, miR-21, miR-25, miR-34b, miR-93, miR-155, miR-181a, miR-181c, miR-182, miR-185,

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and let-7f increased by >2-folds, and miR-125a and miR-125b decreased by >40%. Quantitative RT-PCR was used to validate the expression of miR-21, miR-155, miR-103, miR-107, miR-93, miR-23b, miR-125b, and let7i miRNAs in both tumor and adjacent normal tissue samples.

Retinoblastoma Differential expression miRNAs in human retinoblastoma tissues was analyzed by miRNA microarray in conjunction with Northern blot analysis and in situ hybridization. A group of miRNAs were identified as highly expressed in retinoblastoma, including hsa-miR-494, hsa-let-7e, hsa-miR-513-1, hsa-miR-513-2, hsa-miR518c*, hsa-miR-129-1, hsa-miR-129-2, hsa-miR-198, hsa-miR-492, hsa-miR-498, hsa-miR-320, hsa-miR-503, and hsa-miR-373*.

Metastasis The spread and growth of cells from a primary tumor site (known as metastasis) is the most common cause of death for cancer patients and may occur through organ damage caused by growing lesions, paraneoplastic syndromes, or treatment complications. At the cellular level, the early stages of metastasis are characterized by the loss of contact with neighboring cells and an increase in invasive capacity. For a metastatic lesion to arise, tumor cells must disseminate by intravasating into the blood or lymphatic system. This requires the breaking of local cellcell contacts and invasion into the surrounding stroma and may be enhanced by neo-angiogenesis into the primary tumor site which is a pre-requisite for continued tumor growth. Several miRNAs have been linked to merastasis. miRNA profiling of human breast cancer MDA-MB-231 cells that were selected for being highly metastatic to the bone or lung, in comparison with the parental unselected cell population has identified three miRNAs (miR-335, miR-126, and miR-206) that have lower expression in both metastatic cell lines and in the metastases that developed after the injection of primary human malignant cells into mice (Tavazoie et al. 2008). Importantly, breast cancer patients with tumors expressing decreased levels of miR-335, miR-206, and miR-126 had a shorter time to metastatic relapse. Inhibition of miR-335 in non-metastatic cells was sufficient to increase metastasis to the lung. Overexpression of miR-335 also decreased the migration and invasion of cells in vitro. Another study identified miRNAs of interest to cancer on the basis of their differential expression between normal mammary tissue and primary breast carcinoma using microarray profiling data. Among these, Ma et al. identified miR-10b as being highly expressed only in metastatic cells (Ma et al. 2007). Inhibition of miR10b decreased invasion in matrigel, while miR-10b overexpression is sufficient to promote cell motility and invasion in otherwise non-invasive cell lines and can

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drive the metastasis of these normally non-metastatic cell lines when they are injected into mouse mammary fat pads. The overexpression of miR-373 and miR-520c increased in vitro invasion through matrigel and increased in vivo metastasis to both bone and lung when MCF7 cells overexpressing these miRNAs were injected into the tail veins of mice. Conversely, inhibition of miR-373 was sufficient to inhibit in vitro migration of otherwise invasive MDA-MB-435 and HCT-15 cells. miR-21 is perhaps the most frequently reported miRNA up-regulated in tumors, with several studies correlating miR-21 overexpression with increased metastasis (Roldo et al. 2006; Slaby et al. 2007). Inhibition of miR-21 in MDA-MB-231 cells was found to decrease invasion in vitro and decrease lung metastasis of cells injected into the tail vein of mice (Zhu et al. 2008). Inhibition of miR-21 also decreased intravasation across the chorioallantoic membrane and reduced lung metastases in chicken embryos (Asangani et al. 2008). In human breast and ovarian tumors, a strong correlation exists between miR200 and E-cadherin expression (Gregory et al. 2008; Park et al. 2008). Consistent with the pro-invasive properties associated with EMT and the inhibition of EMT by miR-200, inhibition of miR-200 promotes cell migration (Gregory et al. 2008) while miR-200 expression reduces migration (Park and Tang 2009). miR-205 has also been implicated in the maintenance of mouse mammary epithelial progenitor cells (Ibarra et al. 2007) and therefore may have a role in cancer-associated stem cells with which EMT has been recently linked (Mani et al. 2008). Altered miR-205 expression has also been reported in various cancers (Feber et al. 2008; Gottardo et al. 2007; Sempere et al. 2007; Iorio et al. 2007; Volinia et al. 2006).

1.2.3.2

Cardiovascular Diseases

Cardiac Hypertrophy and Heart Failure In response to stress (such as hemodynamic alterations associated with myocardial infarction, hypertension, aortic stenosis, valvular dysfunction, etc.), the adult heart undergoes remodeling process and hypertrophic growth to adapt to altered workloads and to compensate for the impaired cardiac function. Hypertrophic growth manifests enlargement of cardiomyocyte size and enhancement of protein synthesis through the activation of intracellular signaling pathways and transcriptional mediators in cardiac myocytes. The process is characterized by a reprogramming of cardiac gene expression and the activation of “fetal” cardiac genes (McKinsey and Olson 2005). Recent studies revealed an important role for specific miRNAs in the control of hypertrophic growth and chamber remodeling of the heart and point to miRNAs as potential therapeutic targets in heart disease. The first common finding is that an array of miRNAs is significantly altered in their expression, some up- and some down-regulated. The second common finding

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1 Detection, Profiling, and Quantification of miRNA Expression

is that single miRNAs can critically determine the progression of cardiac hypertrophy. For example, Olson’s group reported >12 miRNAs that are up- or downregulated in cardiac tissue from mice in response to transverse aortic constriction (TAC) or expression of activated calcineurin, stimuli that induce pathological cardiac remodeling (van Rooij et al. 2006). Many of these miRNAs were found similarly regulated in failing human hearts. Forced overexpression of stress-inducible miRNAs induced hypertrophy in cultured cardiomyocytes. Particularly, overexpression of miR-195 alone, which is up-regulated during cardiac hypertrophy, is sufficient to induce pathological cardiac growth and heart failure in transgenic mice. The same group later found that miR-208, encoded by an intron of the alpha myosin heavy chain (aMHC) gene, is required for cardiomyocyte hypertrophy, fibrosis, and expression of aMHC in response to stress and hypothyroidism (van Rooij et al. 2007). The study showed that miR-208 mutant mice failed to undergo stress-induced cardiac remodeling, hypertrophic growth, and bMHC up-regulation, whereas transgenic expression of miR-208 was sufficient to induce bMHC. Abdellatif’s group reported an array of miRNAs that are differentially and temporally regulated during cardiac hypertrophy (Sayed et al. 2007). They found that miR-1 was singularly down-regulated as early as day 1, persisting through day 7, after TAC-induced hypertrophy in a mouse model. A study from Condorelli’s group focuses on the role of miR-133 and miR-1 in cardiac hypertrophy with three murine models: TAC mice, transgenic mice with selective cardiac overexpression of a constitutively active mutant of the Akt kinase, and human tissues from patients with cardiac hypertrophy (Care` et al. 2007). They showed that cardiac hypertrophy in all three models results in reduced expression levels of both miR-133 and miR-1 in the left ventricle. In vitro overexpression of miR-133 or miR-1 inhibits cardiac hypertrophy. In contrast, suppression of miR133 induces hypertrophy, which is more pronounced than that after stimulation with conventional inducers of hypertrophy. In vivo inhibition of miR-133 by a single infusion of an anti-miRNA antisense oligonucleotide (AMO) against miR-133 causes marked and sustained cardiac hypertrophy. Identified 19 deregulated miRNAs in hypertrophic mouse hearts after aortic banding. Knockdown of miR-21 expression via AMO-mediated depletion has a significant negative effect on cardiomyocyte hypertrophy induced by TAC in mice or by angiotensin II or phenylephrine in cultured neonatal cardiomyocytes. Consistently, another independent group identified 17 miRNAs up-regulated and three miRNAs down-regulated in TAC mice, and seven up-regulated and four downregulated in phenylephrine-induced hypertrophy of neonatal cardiomyocytes. They further showed that inhibition of endogenous miR-21 or miR-18b that are most robustly up-regulated augments hypertrophic growth, while introduction of either of these two miRNAs into cardiomyocytes represses cardiomyocyte hypertrophy (Tatsuguchi et al. 2007). A study directed to the human heart identified 67 significantly up-regulated miRNAs and 43 significantly down-regulated miRNAs in failing left ventricles

1.2 Human Disease-Related Expression Profiles of miRNAs

33

versus normal hearts (Thum et al. 2007). Interestingly, 86.6% of induced miRNAs and 83.7% of repressed miRNAs are regulated in the same direction in fetal and failing heart tissues compared with healthy heart tissues, consistent with the activation of “fetal” cardiac genes in heart failure. Collectively, with respect to hypertrophy, it is evident that in addition to the muscle-specific miRNAs miR-1, miR-133, and miR-208, other miRNAs, including miR-195, miR-21, miR-18b, also play an important role. It appears that multiple miRNAs are involved in cardiac hypertrophy and each of them can independently determine the pathological process. The most consistent changes reported by these studies using microarray are up-regulation of miR-21 (6 of 6 studies), miR-23a (4 of 6), miR-125b (5 of 6), and miR-214 (4 of 6), and down-regulation of miR-150 (5 of 6 studies) and miR-30 (5 of 6) (Wang et al. 2008; Yang et al. 2008a; Barringhaus and Zamore 2009).

Myocardial Ischemia/Reperfusion Injuries It is estimated that >50% of the death due to cardiovascular disease can be attributed to ischemic heart disease leading to myocardial infarction, a syndrome characterized by insufficient blood supply to the myocardium. Despite advances in treatment of myocardial infarction through removal of the occlusion, morbidity and mortality remain substantial, with about 5–6% of patients having a subsequent cardiovascular event within 30 days. The abrupt reperfusion of ischemic myocardium can itself cause tissue damage, a phenomenon termed myocardial reperfusion injury. We first described up-regulation of miR-1 in a rat model of acute myocardial ischemia and in ventricular samples of patients with coronary artery disease (Yang et al. 2007). This up-regulation of miR-1 results in significant cardiac arrhythmogenisis. Shan et al. (2009) confirmed our observation in a similar model but with prolonged time of infarction (1–4 weeks of coronary artery occlusion) and further established the role of miR-1 in apoptotic cell death. The study reported by Roy et al. (2009) revealed that in myocardial tissue following 2 and 7 days of ischemia/ reperfusion subjected to miRNA expression profiling and quantification, using a Bioarray system that screens for human-, mice-, rat-, and Ambi-miRNAs, 13 miRNAs were up-regulated on day 2 post ischemia/reperfusion, nine miRNAs were up-regulated on day 7, and 6 miRs were down-regulated on day 7. The up-regulated miRNAs include miR-21, miR-214, miR379, miR-146b, miR142-5p, miR-15b, miR-17-5p, miR-20b, and miR-106a and the down-regulated miRNAs include miR-150, miR-204, and miR-499. It is unclear yet whether these affected miRNAs are involved in the pathological conditions, what influences they produce, damaging or protective, and how they are acting under this setting. Interesting to note is that some of the miRNAs demonstrated the opposite directions of changes in their expression between ischemic myocardium and hypertrophic hearts. For example, miR-1, let-7, miR-181b, miR-29a, and miR30a/e, which are up-regulated in ischemic myocardium, are down-regulated in hypertrophy. Similarly, miR-208, miR-214, miR-320, and miR-351, which are

34

1 Detection, Profiling, and Quantification of miRNA Expression

down-regulated in ischemic myocardium, are up-regulated in hypertrophy. This fact further reinforces the notion that different pathological conditions have different expression profiles. Earlier, we discovered that miR-1 promotes apoptotic cell death induced by oxidative stress (Xu et al. 2007a, b). This observation was later reproduced by two groups. Yu’s laboratory found that glucose induces apoptosis of cardiomyocytes via miR-1 that targets insulin-like growth factor 1 (IGF-1). Subsequently, the same group further observed that miR-1 and miR-206 expression were significantly increased, and IGF-1 protein was markedly reduced without obvious change of its mRNA level after myocardial infarction induction. Position 175-196 of rat IGF-1 30 UTR was identified to be required for efficient down-regulation by miR-1/miR206. In the serum withdrawal and hypoxic conditions, caspase-3 activity and mitochondrial potential were significantly increased in H9C2-miR-1 cells compared with the control group, respectively. These results indicate that miR-1 and miR-206 are involved in apoptotic cell death in myocardial infarction by posttranscriptional repression of IGF-1 (Shan et al. 2009). More recently, Tang et al. (2009a, b) reported that miR-1 is closely related with ischemia/reperfusion injury in a rat model. In vitro, the level of miR-1 was dramatically increased in response to oxidative stress. Overexpression of miR-1 facilitated H2O2-induced apoptosis in cardiomyocytes. Inhibition of miR-1 by antisense inhibitory oligonucleotides caused marked resistance to H2O2. Another miRNA miR-320 was also found to be involved in the regulation of cardiac ischemia/reperfusion (I/R) injury (Ren et al. 2009). The authors found that only miR-320 expression was significantly decreased in the hearts on I/R in vivo and ex vivo. Gain-of-function and loss-of-function approaches were employed in cultured adult rat cardiomyocytes to investigate the functional roles of miR-320. Overexpression of miR-320 enhanced cardiomyocyte death and apoptosis, whereas knockdown was cytoprotective, on simulated I/R. Furthermore, transgenic mice with cardiac-specific overexpression of miR-320 revealed an increased extent of apoptosis and infarction size in the hearts on I/R in vivo and ex vivo relative to the wild-type controls. Conversely, in vivo treatment with antagomir-320 reduced infarction size relative to the administration of mutant antagomir-320 and saline controls. miR-320 produced antithetical regulation of Hsp20 (Ren et al. 2009).

Preconditioning Protection to Ischemic Injuries Mice subjected to cytoprotective heat-shock (HS) showed a significant increase of miR-1, miR-21, and miR-24 in the heart. miRNAs isolated from HS mice and injected into non-HS mice significantly reduced infarct size after ischemia/reperfusion (I/R) injury, which was associated with the inhibition of pro-apoptotic genes and increase in anti-apoptotic genes. Chemically synthesized miR-21 also reduced infarct size, whereas a miR-21 inhibitor abolished this effect. Overall, this study provided evidence for the potential role of endogenously synthesized miRNAs in cardioprotection following I/R injury (Yin et al. 2008).

1.2 Human Disease-Related Expression Profiles of miRNAs

35

Yin et al. (2009) examined their hypothesis that miRNAs induced after ischemic preconditioning (IPC) in the heart may create a preconditioned phenotype through up-regulating proteins including endothelial nitric oxide synthase (eNOS)/inducible nitric oxide synthase (iNOS) and heat shock protein (HSP)70, which are implicated in the late-phase protection of IPC. miRNAs were extracted from hearts of ICR mice following IPC. The purified miRNAs were injected in vivo into the left ventricular wall of mice, and, 48 h later, the hearts were subjected to regional ischemia/reperfusion injury by left anterior descending artery ligation for 30 min followed by reperfusion for 24 h. IPC caused no changes in miR-23b and miR-483 whereas miR-1, miR-21,and miR-24 were significantly increased. The IPC-miRNA treatment caused an increase in eNOS mRNA and protein, whereas iNOS was not changed. HSF-1 (heat shock transcription factor 1) and HSP70 were also increased with IPC-miRNA treatment versus control. Moreover, injection of IPC-miRNA protected the hearts against ischemia/reperfusion injury, as shown by a reduction of infarct size as compared with saline or non-IPC miRNA-treated control.

Vascular Angiogenesis Reactive oxygen species (ROS), such as superoxide and hydrogen peroxide, are involved in the pathogenesis of many vascular diseases by modulating expression of a large number of genes related to vascular cell differentiation, proliferation, migration, and apoptosis. In this respect, increased ROS are associated with a variety of vascular disorders such as atherosclerosis, hypertension, restenosis after angioplasty or bypass, diabetic vascular complications, transplantation arteriopathy, and vascular aneurysm. ROS-mediated gene expression regulation has recently been extensively studied at epigenetic and transcriptional levels. Exposure of vascular cells to ROS modulates oxidation-sensitive signaling pathways and transcription factors that could be an important mechanism responsible for ROSmediated expression changes of multiple genes. In the study reported by Lin et al. (2009), 143 miRNAs out of the 238 miRNAs in the microarray were found expressed in rat vascular smooth muscle cell (RVSMCs). After treatment with H2O2 for 6 h, 57 miRNAs were highly expressed and deregulated. These include up-regulation of miR-21, miR-15b, miR-10b, miR-18, miR-20a/b, miR-30b/c/d, miR-195, and let-7b/d/f/I and down-regulation of miR-29b, miR-143, miR-145, miR-328, miR214, etc.

1.2.3.3

Neuronal Diseases

Several studies have implicated miRNAs in diseases of the CNS. For example, a mutation in the target site of miR-189 in the human SLITRK1 gene has been shown to be associated with Tourette’s syndrome (Abelson et al. 2005), while another study has reported altered miRNA profiles in the prefrontal cortex of patients with schizophrenia and schizoaffective disorder (Perkins et al. 2007).

36

1 Detection, Profiling, and Quantification of miRNA Expression

Saba et al. (2008) used microarrays and RT-PCR to profile miRNA expression changes in the brains of mice infected with mouse-adapted scrapie. Fifteen miRNAs were found de-regulated during the disease processes; miR-342-3p, miR-320, let-7b, miR-328, miR-128, miR-139-5p, and miR-146a were over 2.5-fold up-regulated and miR-338-3p and miR-337-3p were over 2.5 fold down-regulated. Computational analysis predicted numerous potential gene targets of these miRNAs, including 119 genes previously determined to be also de-regulated in mouse scrapie. In particular, a correlation between miRNA expression and putative gene targets involved in intracellular protein-degradation pathways and signaling pathways related to cell death, synapse function, and neurogenesis was identified.

1.2.3.4

Human Immunodeficiency Virus Type 1 (HIV-1) Seropositive Individuals

Houzet et al. (2008) profiled the miRNA expression in peripheral blood mononuclear cells (PBMCs) from 36 HIV-1 seropositive individuals and 12 normal controls. The HIV-1 patients were grouped into four classes: Class I with high CD4þ T cell count and low viral load, Class II with high CD4þ T cell count and high viral load, Class III with low CD4þ T cell count and low viral load, and Class IV with low CD4þ T cell count and high viral load. The expression of 327 well-characterized human cellular miRNAs was analyzed using miRNA microarrays. The data revealed the following two points. (1) miRNA expression is deregulated in HIV infected patients and HIV-1 infection generally resulted in the down regulation of most human miRNAs in vivo. Fifty-nine miRNAs were down regulated while three were up regulated when compared to normal PBMCs. Some polycistronic miRNA clusters such as miR-451 and miR-144; and miR-23a, miR-27a, and miR-24 were down regulated simultaneously. (2) The down-regulation of 14 mRNAs (including miR-19a, miR-20b, miR-30a/c/e, miR-101, miR-155, and miR-146b) was specific to class IV, but was absent from class I, II, or III; the changes in four other miRNAs (miR-143, miR-199a, miR30e-3p, miR-335) were unique to class I, but not observed in class II, III, or IV. Eight other miRNAs were changed in both class I and IV patients, but not in class II or III patients; while a further eight miRNAs (let-7a, miR-1, miR-106b, miR-20a, miR-25, miR-29a, miR-34b, and miR-520b) were changed in class I, II, and IV patients, but were absent from class III patients. Lastly, 12 miRNA changes were present in all four classes of patients. These patterns suggest class-specific “signatures” that plausibly correlate stage-specific miRNA alterations with the in vivo course of HIV-1 infection.

1.2.3.5

Human Non-obstructive Azoospermia

Infertility is a worldwide reproductive health problem which affects 10–15% of couples. Half of the cases are due to male factors, and about 60–75% of male infertility cases are idiopathic, as the molecular mechanisms underlying the defects

1.3 miRNAs as Biomarkers for Human Disease

37

remain unknown. A significant proportion of idiopathic male infertility is accompanied by severe oligozoospermia or azoospermia. Altered miRNA expression in patients with non-obstructive azoospermia has been documented by Lian et al. (2009). In this study, 154 miRNAs were found differentially down-regulated and 19 up-regulated in the testes of non-obstructive azoospermia patients, with initial microarray screening followed by qRT-PCR verification. Around 7.8% (13 out of 154) down-regulated miRNAs belong to the testicular miRNAs: miR-19a, miR20b, miR-29c, miR-30a*, miR-30d*, miR-34b*, miR-92a, miR-181a, miR-449a, miR-652, let-7f, let-7f-2* and let-7i* (Ro et al. 2007). Several down-regulated miRNA clusters in patients with non-obstructive azoospermia were identified, such as the oncogenic potential of the miR-17-92 cluster and miR-371,2,3 cluster.

1.2.3.6

Other Diseases

Hepatitis Using real-time polymerase chain reaction, Ura et al. (2009) measured the expression of 188 miRNAs in liver tissues obtained from 12 patients with hepatitis B virus (HBV)-related hepatocellular carcinoma (HCC) and 14 patients with hepatitis C virus (HCV)-related HCC, including background liver tissues and normal liver tissues obtained from nine patients. Global gene expression in the same tissues was analyzed via complementary DNA microarray to examine whether the differentially expressed miRNAs could regulate their target genes. Their data revealed two types of miRNA, one associated with HBV and HCV infections and the other with the stage of liver disease. Out of the 31 miRNAs associated with disease state (HCC vs. chronic hepatitis), 15 (miR-17-3p, miR30a-5p, miR-30e, miR-92, miR-99a, miR-122a, miR-125b, miR-130a, miR-139, miR-187, miR-199a, miR-199a*, miR-200a, miR-200b, and miR-326) are downregulated in HCC, which promote cancer-associated pathways such as cell cycle, adhesion, proteolysis, transcription, and translation; six miRNAs are up-regulated in HCC (miR-21, miR-98, miR-183, miR-221, miR-222, and miR-301), which repress the anti-tumor immune response.

1.3

miRNAs as Biomarkers for Human Disease

The elucidation of miRNA transcriptomes between diseased and normal tissues or between different disease types, stages and grades, gives the chance to identify the miRNAs most probably involved in human disease and to establish new diagnostic and prognostic markers. 1. Even with much less degree of freedom than mRNAs, miRNA expression profiles reflect the developmental lineage and differentiation state of cells and

38

2.

3.

4.

5.

6. 7.

1 Detection, Profiling, and Quantification of miRNA Expression

successfully classify poorly differentiated tumors that could not have definitive diagnosis by histopathology, while the classification based upon the mRNA profiles was highly inaccurate. A very promising diagnostic strategy could arise from miRNAs if they are found in serum and can be detected by RT-PCR. Recent studies clearly indicate that miRNAs have unusually high stability in formalin-fixed and paraffin-embedded tissues and can remain intact in plasma and serum as well. Indeed, while expression patterns of miRNAs in tissue specimens have been well regarded as better biomarkers of human cancer, being characteristic of tumor type, tumor grade, and developmental origin, most recently, circulating miRNAs have also been revealed to be the stable blood-based markers for cancer detection. This is presumably because miRNAs are shorter than mRNAs, and therefore more resistant to ribonuclease degradation. Different diseases have distinct miRNA transcriptomes. The cancer cluster of miRNAs is clearly separated from the cardiovascular disease cluster of miRNAs. Studies indicate that all cancers are connected together by miRNA profiles, suggesting that various cancers may share similar associations at the miRNA level, in which some strong onco-miRNAs or miRNA tumor suppressors may play key roles. In a remarkable study, Lu et al. (2005) showed that expression data of only 217 miRNAs performed better at identifying cancer types than analysis of 16,000 mRNAs. They concluded that miRNAs might help detecting cancer better than other strategies presently available because miRNAs are only several hundreds, compared to tens of thousands of mRNAs and proteins. This can also partly be attributed to the fact that miRNA expression tends to be very strictly defined in time and space (Xi et al. 2006). This result revealed a potential correlation between miRNA tissue specificity and disease, which may be of value in predicting specific disease-related miRNAs by combining the miRNA tissue specificity values. Thus, if a disease occurs specifically in a given tissue, the miRNAs specifically expressed in that tissue will have a great potential to be related to that disease. Disease-associated miRNAs show various dysfunctions, such as mutation, upregulation, deletion, and down-regulation. Finally, miRNA analysis requires no expensive and time-consuming detection strategies using antibodies or mass spectrometry.

1.4

Methods for Analyzing miRNAs Expression

Owing to the uniqueness of miRNAs distinct from protein-coding mRNAs, there are differences in the approaches to detect and quantify miRNAs and mRNAs: (1) The extremely small size of miRNAs renders most conventional biological amplification tools ineffective because of the inability of even smaller primers/promoters (8- to10-nt) to bind on such small miRNA templates. For example, the regular

1.4 Methods for Analyzing miRNAs Expression

39

RT-PCR can only be used to quantify miRNA precursors rather than the mature miRNAs. (2) The close similarities among family members of miRNAs have presented challenges for developing miRNA-specific detection assays. (3) Small RNAs are less efficiently precipitated in ethanol and for this reason during the isolation by standard Trizol protocol of the RNA, resuspension in ethanol should be avoided. (4) miRNAs seem to be more stable than longer RNAs, for example in degraded samples it is still possible to obtain readable miRNA expression data. Moreover, miRNAs have a higher stability compared to mRNAs in samples obtained from formalin-fixed paraffin-embedded tissues or in serum.

1.4.1

Ideal Methods for miRNA Detection

The ideal miRNA profiling method should fulfill several requirements: 1. Sensitive enough to determine miRNA profiles even with small amounts of starting material 2. Specific enough to reproducibly detect a 1-nt difference between miRNAs 3. Able to provide quantitative analysis of miRNA levels 4. Capable of processing multiple samples in parallel 5. Easy to perform and not require equipment or reagents not readily available in a conventional molecular biology laboratory (Takada and Mano 2007)

1.4.2

Classification of Methods for miRNA Detection

To date, several decades of methods have been developed for miRNA detection. Currently available methods can be classified into following categories: 1. Based on the mechanism of miRNA capturing: (a) Hybridization-based techniques (e.g., Northern blots, in situ hybridization, RT-PCR, and microarrays). (b) Amplification-based techniques (e.g., real-time quantitative PCR; gold nanoparticle-initiated silver enhancement). (c) Cloning-based techniques (e.g., miRAGE). 2. Based on throughput of miRNA profiling: (a) Low throughput methods (e.g., Northern blots, in situ hybridization). (b) Medium throughput methods (e.g., multiplex RT-PCR, miRAGE). (c) High throughput methods (e.g., microarrays). 3. Based on quantification: (a) Quantitative (e.g., real-time quantitative PCR). (b) Semi-quantitative (e.g., Northern blots). (c) Non-quantitative (e.g., Northern blots, in situ hybridization).

40

1 Detection, Profiling, and Quantification of miRNA Expression

4. Based on miRNA capture probes: (a) One-probe assays (e.g., Northern blots, in situ hybridization, and microarrays). (b) Two-probe assays or sandwich-type assays (e.g., gold nanoparticle-based assays and enzyme-amplified assays). 5. Based on read-out format: (a) Optical signal detection (including a variety of biochemical and chemical ligation-based techniques and PCR-based assays that use colorimetry, fluorescence, and bioluminescence). (b) Electrical signal (e.g., polyaniline nanowire technique; electrocatalytic nanoparticle tags technique). 6. Based on miRNA labeling: (a) Fluorescence labeling (e.g., Taqman real-time RT-PCR). (b) Luminescence labeling (e.g., electrocatalytic moiety labeling technique). (c) Non-labeling (e.g., electrocatalytic moiety labeling technique). These above methods have all achieved a certain level of success and have been successfully applied to generate miRNA transcriptomes for our understanding of the biological importance of miRNAs in various pathophysiological settings as described in the earlier sections of this chapter, even though none of these methods is perfect and has inherent limitations. Some methods rely on expensive equipment and an advanced read-out system, which might limit their application.

1.4.3

Brief Introduction to the Currently Available miRNA Detection Methods

Numerous techniques for detecting miRNAs have been developed including a variety of microarray-based (e.g., Krichevsky et al. 2003; Barad et al. 2004; Calin et al. 2004a; Liu et al. 2004a, b; Nelson et al. 2004; Shingara et al. 2005) and PCRbased approaches (Schmittgen et al. 2004; Chen et al. 2005; Jiang et al. 2005), padlock and rolling circle amplification (Jonstrup et al. 2006), an Invader assay (Allawi et al. 2004), an ELISA-based assay (Mora and Getts 2006), bead-based assays (Lu et al. 2005), single molecule detection (Neely et al. 2006), a splinted ligation strategy (Maroney et al. 2007), SAGE-based miRAGE (Cummins et al. 2006), RNA-primed array-based Klenow enzyme (RAKE) assay (Nelson et al. 2004, 2006), gold nanoparticle-based assays (Yang et al. 2008b), gold nanowire method (Fan et al. 2007), and signal amplifying ribozymes (Hartig et al. 2004) (see Table 1.2 for comparison of different approaches for miRNA detection). On the basis of published studies to date, the most widely used strategies for miRNA detection and profiling are microarray-based and PCR-based assays (Chen et al. 2005; Thomson et al. 2007). To date, the most straightforward and widely used assay for small RNA detection has been traditional Northern blotting. When the first miRNAs were described, Northern blotting was used to detect these small RNAs. To date, Northern blot remains the gold standard of miRNA

In situ hybridization (ISH)

Northern blot (NB)

Non-quantitative detection of miRNA expression

Validating miRNA signature highlighted by microarray

l

l

Good specificity

The “gold standard” of miRNA detection

l

l

Less reagent-, time-, and laborconsuming l Semi-quantitative

l

Rapid, computationalized analysis l Less expensive per miRNA

l

l

l

Reagent- and time-consuming

Low sensitivity, requiring large amounts of starting RNA

Low specificity (can be improved)

l

l

Low sensitivity

l

Limitation l Non-quantitative

l

Cellular and subcellular localization of miRNAs

l

With resolution at the single cell level or even at subcellular levels when using nonradioactive probes

l

Non-quantitative

May need to handle radioactive materials l Non-quantitative l Slow analysis speed l Unlikely to be used as a routine method for diagnostic purposes l Direct detection of miRNA in l Straightforward visualization of l Slow analysis speed living cells or fixed, embedded signals for miRNA under tissues detection

Differential expression of miRNAs

l

Table 1.2 Comparison of various approaches for miRNA expression detection Application Advantage l High-throughput miRNA l High-throughput miRNA miRNA profiling profiling Microarray

l

(continued )

Specificity and sensitivity can be improved by LNA modification of probes

Specificity and sensitivity can be improved by LNA modification of probes l DIG-labeling of probes can avoid radioactive materials

l

Remarks l Specificity and sensitivity can be improved by LNA modification of probes

1.4 Methods for Analyzing miRNAs Expression 41

Poly(A)-Tailed Universal Reverse Transcription

miR-Q RT-PCR

Stem-loop Realtime RT-PCR (qRT-PCR)

Validating miRNA signature highlighted by microarray

Can be used extensively for clinical diagnosis l Direct detection of miRNA in total RNA extracted from cell lines or tissues

l

Direct detection of miRNA in total RNA extracted from cell lines or tissues l Validating miRNA signature highlighted by microarray

l

Use for quantification of miRNAs between RNA samples of different statuses, locations, ages, genders, etc. l Can be used extensively for clinical diagnosis

l

l

Table 1.2 (continued) Application l Dynamic changes of miRNA expression

Quantitative

l

l

Advanced discriminative power and sensitivity

Providing a linearity of up to eight orders of magnitude detecting as low as 0.2 fM miRNA molecules l Easy to perform

l

Can be multiplex PCR for medium-throughput miRNA profiling l Advanced discriminative power and sensitivity

l

Proven specificity, which is ensured by using TaqMan to distinguish 1-nt difference

l

Advantage l Can be performed with morphologically preserved tissues sections or cell preparations l Highest sensitivity: requiring minute amounts of starting RNA Slow analysis speed

High cost per miRNA

l

l

l

Specificity may be inherently limited by the use of only one miRNA-specific primer and the use of a universal primer

Low-throughput

Only working with SYBR Green that can limit its specificity

Relying on commercial companies to provide TaqMan probes l Low-throughput

l

l

l

Limitation

l

Specificity and sensitivity can be further improved by LNA modification of probes

Remarks

42 1 Detection, Profiling, and Quantification of miRNA Expression

miRNA cloning

miRNA Amplification Profiling (mRAP)

Direct detection of miRNA in total RNA extracted from cell lines or tissues

l

Can lead to discovery of new miRNAs l Use for differential expression profilings of miRNAs between RNA samples of different

l

Can lead to discovery of new miRNAs l May be used for high-throughput profiling

l

l

Can be used extensively for clinical diagnosis

l

l

l

Advantage of identifying new miRNAs

Can be multiplex PCR for highthroughput miRNA profiling

Can be multiplex PCR for medium-throughput miRNA profiling l Combining cloning for new miRNA discovery, highthroughput profiling, and quantification of miRNA levels into one l High sensitivity and specificty

Proven specificity, which is ensured by using TaqMan l Quantitative

Highest sensitivity: requiring minute amounts of starting RNA

l

l

Relatively simple and convenient

l

Validating miRNA signature highlighted by microarray l Medium-throughput miRNA profiling

l

Validating miRNA signature highlighted by microarray l Can be used for clinical diagnosis Multiplexing RT- l Direct detection of miRNA in total RNA extracted from cell PCR lines or tissues

l

Slow analysis speed

High cost per miRNA

l

Complicated procedures with the need for enrichment of short RNAs by fractionation and of ligation of adaptors

May not suitable for clinical use as a diagnostic tool l Labor intensive

l

l

l

Complicated multiple steps

Relying on commercial companies to provide TaqMan probes

l

l

Slow analysis speed

High cost per miRNA

Low-throughput

l

l

l

l

(continued )

Specificity and sensitivity can be further improved by LNA modification of probes

1.4 Methods for Analyzing miRNAs Expression 43

NanoparticleAmplified SPR Imaging

Electrocatalytic Nanoparticle Tags

Validating miRNA signature highlighted by microarray

Medium-throughput miRNA profiling

l

l

l

Use for quantification and profiling of miRNAs between RNA samples of different statuses, locations, ages, genders, etc.

Direct detection of miRNA in total RNA extracted from cell lines or tissues l Validating miRNA signature highlighted by microarray

l

Direct detection of miRNA in total RNA extracted from cell lines or tissues

l

l

Limitation

Requirement of a large amount (1 mg) total RNA as a starting material l Unlikely be used extensively for clinical diagnosis l An opportunity for the lowl Limited application due to the density electrochemical array requirement for multiple in miRNA expression profiling electrochemical detection instruments l This method allows for as low as l Unlikely to be used extensively 5 ng total RNA for a successful for clinical diagnosis miRNA detection l Capable of identifying miRNAs with <2-fold difference in expression levels under two conditions l Quantitative l High sensitivity comparable to l Requirement of sophisticated, PCR methods (10 fM detection expensive instruments (SPR limit) imager) for signal readout l Combining a surface enzyme l Hardly be used extensively for reaction with nanoparticleclinical diagnosis amplified SPR imaging l An opportunity for mediumthroughput array in miRNA expression profiling

Table 1.2 (continued) Application Advantage statuses, locations, ages, genders, etc. l May be used for high-throughput profiling Remarks

44 1 Detection, Profiling, and Quantification of miRNA Expression

l

Gold Nanoparticle Probes

Padlock-Probes and RollingCircle Amplification

Splinted Ligation

Direct detection of miRNA in total RNA extracted from cell lines or tissues l Validating miRNA signature highlighted by microarray

Can be used extensively for clinical diagnosis

Can be used extensively for clinical diagnosis

Direct detection of miRNA in total RNA extracted from cell lines or tissues

l

l

Direct detection of miRNA in total RNA extracted from cell lines or tissues l Validating miRNA signature highlighted by microarray

l

l

Direct detection of miRNA in total RNA extracted from cell lines or tissues l Validating miRNA signature highlighted by microarray

l

Conducting Polymer Nanowires Quantitative Relatively simple Simple, sensitive, and specific

Colorimetric method detect limit of 10 fM

l

l

l

Does not require specialized, sophisticated equipment or any amplification step l Simple to perform

l

Medium sensitive with ~50 times more sensitive than Northern blotting l Quantitative

l

Does not require expensive equipment and an advanced read-out system l No need to use enzymes l High-specificity for miRNA detection l Simple to perform

l

l

Ultrasensitive (detection limit of 5 fM)

l

The obligation of radioactive labeling

Difficulty of determining silver enhancement reaction time to obtain reproducible results

Need to handle toxic chemical

l

The obligation of radioactive labeling

Relatively low sensitivity compared to qRT-PCR methods l Low-throughput

l

l

l

l

(continued )

1.4 Methods for Analyzing miRNAs Expression 45

Single Molecule Method

Invader Assay

Can be used extensively for clinical diagnosis

Can be used extensively for clinical diagnosis

Direct detection of miRNA in total RNA extracted from cell lines or tissues l Validating miRNA signature highlighted by microarray

l

l

Direct detection of miRNA in total RNA extracted from cell lines or tissues l Validating miRNA signature highlighted by microarray

l

l

Table 1.2 (continued) Application l Validating miRNA signature highlighted by microarray

Does not require specialized, sophisticated equipment or any amplification step l Quantitative, able to precisely measure 1.2-fold changes in RNA expression l Specific able to discriminate between miRNAs that differ by a single base, and to distinguishes between miRNAs and their precursors l Sensitive able to detect 1–10 RNA molecules per cell l Simple, rapid (requiring only 2– 3 h incubation) l Does not use radioactivity l Readily to be performed directly in cell lysates l Medium sensitivity with a detection limit of 100 fM of total RNA l Quantitative

l

Advantage l Medium sensitive with a detection limit of a few nanograms of total RNA l Quantitative

l

l

Requires special equipment

Low-throughput

Limitation l Relatively low sensitivity compared to qRT-PCR methods l Low-throughput

Remarks

46 1 Detection, Profiling, and Quantification of miRNA Expression

Direct detection of miRNA in total RNA extracted from cell lines or tissues

Direct detection of miRNA in total RNA extracted from cell lines or tissues

l

l

Validating miRNA signature highlighted by microarray

l

Validating miRNA signature highlighted by microarray l Direct detection of miRNA in total RNA extracted from cell lines or tissues

l

l

High-throughput miRNA profiling Bead-Based Flow l Direct detection of miRNA in total RNA extracted from cell Cytometric lines or tissues assay l Medium-throughput miRNA profiling

RAKE assay

SurfaceEnhanced Raman Spectroscopy

Enzymatic Method

l

Quantitative

Quantitative

Complicated procedures

l

l

Requires 3 ng labeled low molecular weight RNA sample l Requires sophisticated read-out system

l

l

Complicated analysis skill, and convoluted data interpretation and verification

Complicated analysis skill, and convoluted data interpretation and verification l Low throughput l Does not involve the generation l Relatively complicated of a cDNA library or procedures amplification of the RNA sample l Reasonable sensitivity and l Hardly be used extensively for specificity clinical diagnosis l Sensitive, and accurate l Requires sophisticated read-out identification of miRNAs system

l

Simple and yields reproducible data (No preparative enrichment, ligation, or target/ signal amplification steps are required) l Can be medium-throughput with 384-well reactions l High sample throughput for studying the expression of the same miRNA in many different samples l Can be medium-throughput with 384-well reactions l Sensitive and accurate identification of miRNAs

l

(continued )

1.4 Methods for Analyzing miRNAs Expression 47

Electrocatalytic Moiety Labeling Technique

Ribozyme Method

Molecular Beacon

l

Direct detection of miRNA in total RNA extracted from cell lines or tissues

Direct detection of miRNA in total RNA extracted from cell lines or tissues l Validating miRNA signature highlighted by microarray l Can be used extensively for clinical diagnosis l Direct detection of miRNA in total RNA extracted from cell lines or tissues l Validating miRNA signature highlighted by microarray

l

Medium-throughput miRNA profiling l Validating miRNA signature highlighted by microarray

l

Table 1.2 (continued) Application l Validating miRNA signature highlighted by microarray Bioluminescence- l Direct detection of miRNA in total RNA extracted from cell Based lines or tissues Detection

High specificity

No requirement for special equipment l Rapid, nonradioactive

l

l

Discriminate single-nucleotide differences in miRNAs l Low-cost with no requirement for sophisticated equipment l Relatively simple procedures

l

l

Complicated procedures

Difficulty to develop suitable ribozymes

Low sensitivity

l

l

Low sensitivity

l

Advantage Limitation l May be used extensively for clinical samples l Rapid, requiring in a microplate l The procedures are relative format a total assay time of complicated 1.5 h without the need for sample PCR amplification l Super sensitive with a detection limit of 1 fM. l Can be done in a 96-well microtiter plate for medium throughput analysis l May be suitable for application in clinical diagnostics and drug discovery l Extremely simple to perform l Low-throughput

Remarks

48 1 Detection, Profiling, and Quantification of miRNA Expression

l

Direct detection of miRNA in total RNA extracted from cell lines or tissues

l

Simple and easy to perform

Straightforward visualization of signals for miRNA under detection

l

l

Ultrasensitive with miRNA being detected amperometrically at subpicomolar levels with high specificity l May be expanded to low-density array of 50–100 electrodes l Non-invasive for patients

l

l

l

Hardly to be used as a routine method for diagnostic purposes

l

miRNA expression profiling with appropriate methods

l

Proven specificity, which is ensured by using TaqMan to distinguish 1-nt difference

l

Slow analysis speed

The current availability of only a single ELF substrate limits this approach to imaging only a single RNA per cell sample l With resolution at the single cell l Complicated procedures level or even at subcellular levels when using nonradioactive probes l Hardly be used as a routine method for diagnostic purposes l High cost per miRNA Single Cell Stem- l Direct detection of miRNA in a l High sensitivity: requiring single cell level minute amounts of starting Looped RealRNA Time PCR

miRNA expression profiling with appropriate methods l May be used as a routine method for diagnostic purposes l Direct visualization of miRNA Quantitative localization in a single cell LNA-ELFlevel FISH Method

Serum and Plasma miRNA Detection

Validating miRNA signature highlighted by microarray

l

l

(continued )

Specificity and sensitivity can be further improved by LNA modification of probes

1.4 Methods for Analyzing miRNAs Expression 49

Whole-mount In situ hybridization

miRNA FunctionReporter Expression Assay

Cellular and subcellular localization of miRNAs

l

l

With resolution at the single cell level or even at subcellular levels when using nonradioactive probes

Straightforward visualization of signals for miRNA under detection l Monitor developmental changes l Can be performed with of miRNAs and the related morphologically preserved target mRNAs at the whole embryos animal level

Monitor developmental changes of miRNAs and the related target mRNA at the single cell level l Hardly be used as a routine method for diagnostic purposes l Cellular and subcellular localization of miRNAs

l

l

Can be multiplex PCR for medium-throughput miRNA profiling l Straightforward visualization of signals for miRNA under detection

l

Table 1.2 (continued) Application Advantage l May be used as a routine method l Quantitative for diagnostic purposes

l

l

Slow analysis speed

Complicated procedures for constructing dual-fluorescence GFP-reporter/mRFP-sensor (DFRS) plasmids

Limitation Remarks l Relying on commercial companies to provide TaqMan probes l Low-throughput

50 1 Detection, Profiling, and Quantification of miRNA Expression

1.4 Methods for Analyzing miRNAs Expression

51

expression profiling. However, there are several technical limitations that prevent researchers from using Northern blot as a routine miRNA expression profiling tool. Northern Blotting using radioactive probes is very sensitive, but it is very timeconsuming. Northern blotting is not practical in large clinical studies to detect the expression of hundreds of miRNAs and it also requires large amounts (5–25 mg) of total RNA from each sample. A modified version of Northern blot using locked nucleic acid-modified oligonucleotides was developed by Va´lo´czi et al. (2004). The sensitivity was improved by ten-fold as compared to conventional DNA probes. As an improvement to Northern blot, the use of nylon macroarrays for miRNA analysis has also been reported; however, Northern blot and cloning techniques suffer from poor sensitivity and involve laborious procedures. As the number of identified miRNA increases, microarrays appear to be an ideal method for profiling in a highly efficient parallel fashion for a large number of miRNAs in a single run. Besides the limitations mentioned above, all of these methods have a common weakness of low throughput and slow analysis speed. Fortunately, miRNA microarrays can overcome these drawbacks by offering rapid, parallel, and high throughput analysis. Owing to their adaptability and high throughput, microarrays may prove to be the preferred platform for genome-wide miRNA expression analysis. Many different miRNA microarray platforms have already been successfully applied for miRNA studies. However, microarrays have encountered difficulties in reliably amplifying miRNAs without bias. Microarray probe design is severely limited due to the short length of the mature miRNA; there is very little room to fine-tune the hybridization conditions for all miRNAs, which significantly discounts the sensitivity and specificity of microarrays (Liu et al. 2004a, b; Krichevsky et al. 2003). In some designs, the optimization of the sequence specificity of the probe–target interactions relies on the empirical selection of all probes in the microarray to have matched probe-target melting temperature (Tm); thus, not all Tm-balanced probes will be capable of single base discrimination (Castoldi et al. 2007). The use of locked nucleic acid probes allowed optimization of the hybridization conditions suitable for all miRNAs, producing an improved microarray platform that offered single base mismatch (SBM) discrimination capability with a detection limit of 500 fM (Neely et al. 2006). To further enhance the sensitivity of miRNA assay, several quantitative realtime PCR (qRT-PCR) approaches for the simultaneous amplification and quantification of miRNAs have been developed recently (Chen et al. 2005; Raymond et al. 2005). In each of these approaches, the miRNAs were first lengthened to generate extended sequences suitable for subsequent PCR amplifications. It combined the exceptional amplification power of PCR with quantitative detection of the amplified products in real time during each reaction cycle. The stem-loop RT-PCR analysis offers the highest sensitivity (Chen et al. 2005; Mestdagh et al. 2008), and can even profile miRNA from a single cell, though this method does require prior knowledge of the miRNA sequences for analysis. The PCR-based technique is able to detect low copy number with high sensitivity and specificity on both the precursor (Shi and Chiang 2005; Raymond et al. 2005) and the mature form of miRNAs (Jiang et al. 2005). It can be used extensively for clinical samples with minute amounts of

52

1 Detection, Profiling, and Quantification of miRNA Expression

available RNA. Despite the many advantages, a number of challenges need to be addressed in qRT-PCR assays. One major constraint is their limited capacity for multiplexing (Chen et al. 2005; Raymond et al. 2005). Running too many replicates as single reactions for a panel of miRNAs is not cost-effective and not feasible for small or rare samples. In addition, it is limited by high cost. In addition to Northern blot, microarray and RT-PCR as current standard methods, emerging techniques have exhibited superior flexibility and adaptability in miRNA detection and quantification as well as matched or increased sensitivity in comparison to the current standards (Hunt et al. 2009). These new methods have addressed many of the problems associated with miRNA detection through the employment of enzyme-based signal amplification, enhanced hybridization conditions using PNA capture probes, highly sensitive and flexible forms of spectroscopy, and extremely responsive electrocatalytic nanosystems. Earlier attempts to develop ultrasensitive assays for miRNAs were mostly based on optical detections, including a variety of biochemical and chemical ligation-based techniques (Miska et al. 2004; Thomson et al. 2004; Nelson et al. 2004; Babak et al. 2004; Gao and Yang 2006) and PCR-based assays (Krichevsky et al. 2003; Liu et al. 2004a, b; Neely et al. 2006; Chen et al. 2005; Raymond et al. 2005). For example, Miska and co-workers proposed an array-based miRNA expression profiling procedure, in which miRNAs are ligated to 30 and 50 adaptor oligonucleotides followed by RTPCR (Thomson et al. 2004). A T4 RNA ligase-based (Thomson et al. 2004), a RNA-primed, array-based Klenow enzyme assays (Nelson et al. 2004) were also used to couple the 30 end of miRNA to a fluorophore-tagged nucleotide. Babak et al. proposed a cisplatin-based chemical ligation procedure for miRNAs (Babak et al. 2004). Another chemical ligation procedure at the 30 end was developed by Gao and Yang (2006). However, poor reliability and differential ligation efficiencies may compromise the quality of the data, and the sensitivity of these assays is not always satisfactory. According to Fan et al. (2007), electrical methods, on the other hand, are more advantageous than optical methods because electrical detections often directly (label-free or non-labeling) transduce nucleic acid hybridization events into useful electrical signals such as capacitance and conductance (Macanovic et al. 2004). Because of the inherent superiorities of electrical transduction methods, such as excellent compatibility with advanced semiconductor technology, miniaturization, and low cost, nucleic acid biosensors based on electrical detection are able to provide high performance at low-cost with a simple miniaturized readout, and thus are exempt from the problems encountered in the optical detection systems. The detection can be tailored to be extremely sensitive with high multiplexing capability. In addition, combining the unique electronic properties of nanoscale materials, electrical detection systems offer excellent prospects for designing nucleic acid detection devices. A good example is the use of metal nanoparticles, such as gold nanoparticles, as labels for sensitive electronic transduction of different biomolecular recognition events, following the milestone discovery of several interesting physiochemical properties of oligonucleotide-functionalized gold nanoparticles by Mirkin and co-workers (Mirkin et al. 1996), in which gold

References

53

nanoparticles were used to bridge a gap between two microband electrodes (Park et al. 2002). A simple detection of a conductance change resulted in a detection limit of 500 fM (Park et al. 2002). Later, the same group demonstrated the superiority of the oligonucleotide-functionalized gold nanoparticles in a number of bioassays (Elghanian et al. 1997; Taton et al. 2000, 2001; Cao et al. 2002; Nam et al. 2003, 2004). Worth mentioning here is the bio-barcode technology. It offers PCR-like sensitivity for both DNA and proteins (Nam et al. 2003, 2004; Stoeva et al. 2006). At the cost of sensitivity, Diessel et al. proposed a modified version of the gold nanoparticle method by introducing continuous online monitoring of the autometallographic enhancement process, eliminating the need for multistep enhancement and all of the washing, drying, and measurement cycles in between (Diessel et al. 2004). Nonetheless, the eventual acceptance of electrical detection techniques will depend on how these techniques compare with the current gold standards, that is, PCR and ELISA, in terms of simplicity, sensitivity, specificity, reliability, and portability.

References Abelson JF, Kwan KY, O’Roak BJ, Baek DY, Stillman AA, Morgan TM, Mathews CA, Pauls DL, Rasin MR, Gunel M, Davis NR, Ercan-Sencicek AG, Guez DH, Spertus JA, Leckman JF, Dure LS 4th, Kurlan R, Singer HS, Gilbert DL, Farhi A, Louvi A, Lifton RP, Sestan N, State MW (2005) Sequence variants in SLITRK1 are associated with Tourette’s syndrome. Science 310:317–320 Akao Y, Nakagawa Y, Naoe T (2007) MicroRNA-143 and -145 in colon cancer. DNA Cell Biol 26:311–320 Allawi HT, Dahlberg JE, Olson S, Lund E, Olson M, Ma W-P, Takova T, Neri BP, Lyamichev VI (2004) Quantitation of microRNAs using a modified Invader assay. RNA 10:1153–1161 Alvarez-Garcia I, Miska EA (2005) MicroRNA functions in animal development and human disease. Development 132:4653–4662 Ambros V (2004) The functions of animal microRNAs. Nature 431:350–355 Asangani IA, Rasheed SA, Nikolova DA, Leupold JH, Colburn NH, Post S, Allgayer H (2008) MicroRNA-21 (miR-21) post-transcriptionally downregulates tumor suppressor Pdcd4 and stimulates invasion, intravasation and metastasis in colorectal cancer. Oncogene 27:2128–2136 Bae S, Ahn JH, Park CW, Son HK, Kim KS, Lim NK, Jeon CJ, Kim H (2009) Gene and microRNA expression signatures of human mesenchymal stromal cells in comparison to fibroblasts. Cell Tissue Res 335:565–573 Bak M, Silahtaroglu A, Møller M, Christensen M, Rath MF, Skryabin B, Tommerup N, Kauppinen S (2008) MicroRNA expression in the adult mouse central nervous system. RNA 14:432–444 Babak T, Zhang W, Morris Q, Blencowe BJ, Hughes TR (2004) Probing microRNAs with microarrays: tissue specificity and functional inference. RNA 10:1813–1819 Bandre´s E, Cubedo E, Agirre X, Malumbres R, Za´rate R, Ramirez N, Abajo A, Navarro A, Moreno I, Monzo´ M, Garcı´a-Foncillas J (2006) Identification by Real-time PCR of 13 mature microRNAs differentially expressed in colorectal cancer and non-tumoral tissues. Mol Cancer 5:29 Baffa R, Fassan M, Volinia S, O’Hara B, Liu CG, Palazzo JP, Gardiman M, Rugge M, Gomella LG, Croce CM, Rosenberg A (2009) MicroRNA expression profiling of human metastatic cancers identifies cancer gene targets. J Pathol 2009 Jun 1. [Epub ahead of print]

54

1 Detection, Profiling, and Quantification of miRNA Expression

Barad O, Meiri E, Avniel A, Aharonov R, Barzilai A, Bentwich I, Einav U, Gilad S, Hurban P, Karov Y, Lobenhofer EK, Sharon E, Shiboleth YM, Shtutman M, Bentwich Z, Einat P (2004) MicroRNA expression detected by oligonucleotide microarrays: system establishment and expression profiling in human tissues. Genome Res 14:2486–2494 Baroukh N, Ravier MA, Loder MK, Hill EV, Bounacer A, Scharfmann R, Rutter GA, Van Obberghen E (2007) MicroRNA-124a regulates Foxa2 expression and intracellular signaling in pancreatic beta-cell lines. J Biol Chem 282:19575–19588 Barringhaus KG, Zamore PD (2009) MicroRNAs: regulating a change of heart. Circulation 119:2217–2224 Bartel DP (2004) MicroRNAs: genomics, biogenesis, mechanism, and function. Cell 116:281–297 Bauersachs J, Thum T (2007) MicroRNAs in the broken heart. Eur J Clin Invest 37:829–833 Berezikov E, Chung WJ,Willis J, Cuppen E, Lai EC (2007) Mammalian mirtron genes. Mol Cell 28:328–336 Bohnsack MT, Czaplinski K, Gorlich D (2004) Exportin 5 is a RanGTP-dependent dsRNAbinding protein that mediates nuclear export of pre-miRNAs. RNA 10:185–191 Bracken CP, Gregory PA, Khew-Goodall Y, Goodall GJ (2009) The role of microRNAs in metastasis and epithelial-mesenchymal transition. Cell Mol Life Sci 66:1682–1699 Brennecke J, Stark A, Russell RB, Cohen SM (2005) Principles of microRNA-target recognition. PLoS Biol 3:e85 Calin GA, Dumitru CD, Shimizu M, Bichi R, Zupo S, Noch E, Aldler H, Rattan S, Keating M, Rai K, Rassenti L, Kipps T, Negrini M, Bullrich F, Croce CM (2002) Frequent deletions and down-regulation of micro- RNA genes miR15 and miR16 at 13q14 in chronic lymphocytic leukemia. Proc Natl Acad Sci USA 99:15524–15529 Calin GA, Liu CG, Sevignani C, Ferracin M, Felli N, Dumitru CD, Shimizu M, Cimmino A, Zupo S, Dono M, Dell’Aquila ML, Alder H, Rassenti L, Kipps TJ, Bullrich F, Negrini M, Croce CM (2004a) MicroRNA profiling reveals distinct signatures in B cell chronic lymphocytic leukemias. Proc Natl Acad Sci USA 101:11755–11760 Calin GA, Sevignani C, Dumitru CD, Hyslop T, Noch E, Yendamuri S, Shimizu M, Rattan S, Bullrich F, Negrini M, Croce CM (2004b) Human microRNA genes are frequently located at fragile sites and genomic regions involved in cancers. Proc Natl Acad Sci USA 101: 2999–3004 Cao YC, Jin R, Mirkin CA (2002) Nanoparticles with Raman spectroscopic fingerprints for DNA and RNA detection. Science 297:1536–1540 Care` A, Catalucci D, Felicetti F, Bonci D, Addario A, Gallo P, Bang ML, Segnalini P, Gu Y, Dalton ND, Elia L, Latronico MV, Høydal M, Autore C, Russo MA, Dorn GW, Ellingsen O, Ruiz-Lozano P, Peterson KL, Croce CM, Peschle C, Condorelli G (2007) MicroRNA-133 controls cardiac hypertrophy. Nat Med 13:613–618 Carmell MA, Xuan Z, Zhang MQ, Hannon GJ (2002) The Argonaute family: Tentacles that reach into RNAi, developmental control, stem cellmaintenance, and tumorigenesis. Genes Dev 16:2733–2742 Castoldi M, Benes V, Hentze MW, Muckenthaler MU (2007) miChip: A microarray platform for expression profiling of microRNAs based on locked nucleic acid (LNA) oligonucleotide capture probes. Methods 43:146–152 Chen C, Ridzon DA, Broomer AJ, Zhou Z, Lee DH, Nguyen JT, Barbisin M, Xu NL, Mahuvakar VR, Andersen MR, Lao KQ, Livak KJ, Guegler KJ (2005) Real-time quantification of microRNAs by stem-loop RT-PCR. Nucleic Acids Res 33:e179 Chen C-Z, Li L, Lodish HF, Bartel DP (2004) MicroRNAs modulate hematopoietic lineage differentiation. Science 303: 83–86 Chendrimada TP, Gregory RI, Kumaraswamy E, Norman J, Cooch N, Nishikura K, Shiekhattar R (2005) TRBP recruits the Dicer complex to Ago2 for microRNA processing and gene silencing. Nature 436:740–744 Chang TC, Wentzel EA, Kent OA, Ramachandran K, Mullendore M, Lee KH, Feldmann G, Yamakuchi M, Ferlito M, Lowenstein CJ, Arking DE, Beer MA, Maitra A, Mendell JT (2007)

References

55

Transactivation of miR-34a by p53 broadly influences gene expression and promotes apoptosis. Mol Cell 26:745–752 Cheung L, Gustavsson C, Norstedt G, Tollet-Egnell P (2009) Sex-different and growth hormoneregulated expression of microRNA in rat liver. BMC Mol Biol 10:13 Cullen BR (2004) Transcription and processing of human microRNA precursors. Mol Cell 16:861–865 Cummins JM, He Y, Leary RJ, Pagliarini R, Diaz LA Jr, Sjoblom T, Barad O, Bentwich Z, Szafranska AE, Labourier E, Raymond CK, Roberts BS, Juhl H, Kinzler KW, Vogelstein B, Velculescu VE (2006) The colorectal microRNAome. Proc Natl Acad Sci USA 103: 3687–3692 Davis E, Caiment F, Tordoir X, Cavaille´ J, Ferguson-Smith A, Cockett N, Georges M, Charlier C (2005) RNAi-mediated allelic trans-interaction at the imprinted Rtl1/Peg11 locus. Curr Biol 15:743–749 Debernardi S, Skoulakis S, Molloy G, Chaplin T, Dixon-McIver A, Young BD (2007) MicroRNA miR-181a correlates with morphological sub-class of acute myeloid leukaemia and the expression of its target genes in global genome-wide analysis. Leukemia 21:912–916 Denli AM, Tops BB, Plasterk RH, Ketting RF, Hannon GJ (2004) Processing of primary microRNAs by the Microprocessor complex. Nature 432:231–235 Dı´az R, Silva J, Garcı´a JM, Lorenzo Y, Garcı´a V, Pen˜a C, Rodrı´guez R, Mun˜oz C, Garcı´a F, Bonilla F, Domı´nguez G (2008) Deregulated expression of miR-106a predicts survival in human colon cancer patients. Genes Chromosomes Cancer 47:794–802 Diessel E, Grothe K, Siebert HM, Warner BD, Burmeister J (2004) Online resistance monitoring during autometallographic enhancement of colloidal Au labels for DNA analysis. Biosens Bioelectron 19:1229–1235 Dixon-McIver A, East P, Mein CA, Cazier JB, Molloy G, Chaplin T, Andrew Lister T, Young BD, Debernardi S (2008) Distinctive patterns of microRNA expression associated with karyotype in acute myeloid leukaemia. PLoS ONE 3:e2141 Elghanian R, Storhoff JJ, Mucic RC, Letsinger RL, Mirkin CA (1997) Selective colorimetric detection of polynucleotides based on the distance-dependent optical properties of gold nanoparticles. Science 277:1078–1081 Esau C, Kang X, Peralta E, Hanson E, Marcusson EG, Ravichandran LV, Sun Y, Koo S, Perera RJ, Jain R, Dean NM, Freier SM, Bennett CF, Lollo B, Griffey R (2004) MicroRNA-143 regulates adipocyte differentiation. J Biol Chem 279:52361–52365 Fan Y, Chen X, Trigg AD, Tung CH, Kong J, Gao Z (2007) Detection of MicroRNAs using targetguided formation of conducting polymer nanowires in nanogaps. J Am Chem Soc 129:5437–5443 Farth KK, Grimson A, Jan C, Lewis BP, Johnston WK, Lim LP, Burge CB, Bartel DP (2005) The widespread impact of mammalian microRNAs on mRNA repression and evolution. Science 310:1817–1821 Fazi F, Rosa A, Fatica A, Gelmetti V, De Marchis ML, et al. (2005) A minicircuitry comprised of microRNA-223 and transcription factors NFI-A and C/EBPalpha regulates human granulopoiesis. Cell 123:819–831 Feber A, Xi L, Luketich JD, Pennathur A, Landreneau RJ, Wu M, Swanson SJ, Godfrey TE, Litle VR (2008) MicroRNA expression profiles of esophageal cancer. J Thorac Cardiovasc Surg 135:255–260 Fulci V, Chiaretti S, Goldoni M, Azzalin G, Carucci N, Tavolaro S, Castellano L, Magrelli A, Citarella F, Messina M, Maggio R, Peragine N, Santangelo S, Mauro FR, Landgraf P, Tuschl T, Weir DB, Chien M, Russo JJ, Ju J, Sheridan R, Sander C, Zavolan M, Guarini A, Foa` R, Macino G (2007) Quantitative technologies establish a novel microRNA profile of chronic lymphocytic leukemia. Blood 109:4944–4951 Gao Z, Yang Z (2006) Detection of microRNAs using electrocatalytic nanoparticle tags. Anal Chem 78:1470–1477 Garzon R, Garofalo M, Martelli MP, Briesewitz R, Wang L, Fernandez-Cymering C, Volinia S, Liu CG, Schnittger S, Haferlach T, Liso A, Diverio D, Mancini M, Meloni G, Foa R, Martelli

56

1 Detection, Profiling, and Quantification of miRNA Expression

MF, Mecucci C, Croce CM, Falini B (2008) Distinctive microRNA signature of acute myeloid leukemia bearing cytoplasmic mutated nucleophosmin. Proc Natl Acad Sci USA 105: 3945–3950 Gauthier BR, Wollheim CB (2006) MicroRNAs: ‘ribo-regulators’ of glucose homeostasis. Nat Med 12:36–38 Giraldez AJ, Cinalli RM, Glasner ME, Enright AJ, Thomson JM, Baskerville S, Hammond SM, Bartel DP, Schier AF (2005) MicroRNAs regulate brain morphogenesis in zebrafish. Science 308:833–838 Gottardo F, Liu CG, Ferracin M, Calin GA, Fassan M, Bassi P, Sevignani C, Byrne D, Negrini M, Pagano F, Gomella LG, Croce CM, Baffa R (2007) Micro-RNA profiling in kidney and bladder cancers. Urol Oncol 25:387–392 Gregory PA, Bert AG, Paterson EL, Barry SC, Tsykin A, Farshid G, Vadas MA, Khew-Goodall Y, Goodall GJ (2008) The miR-200 family and miR-205 regulate epithelial to mesenchymal transition by targeting ZEB1 and SIP1. Nat Cell Biol 10:593–601 Gregory RI, Yan KP, Amuthan G, Chendrimada T, Doratotaj B, Cooch N, Shiekhattar R (2004) The Microprocessor complex mediates the genesis of microRNAs. Nature 432:235–240 Grun D, Wang YL, Langenberger D, Gunsalus KC, Rajewsky N (2005) microRNAs target predictions across seven Drosophila species and comparison to mammalian targets. PLoS Comput Biol 1:e13 Guo C, Sah JF, Beard L, Willson JK, Markowitz SD, Guda K (2008) The noncoding RNA, miR126, suppresses the growth of neoplastic cells by targeting phosphatidylinositol 3-kinase signaling and is frequently lost in colon cancers. Genes Chromosomes Cancer 47:939–946 Guo J, Miao Y, Xiao B, Huan R, Jiang Z, Meng D, Wang Y (2009) Differential expression of microRNA species in human gastric cancer versus non-tumorous tissues. J Gastroenterol Hepatol 24:652–657 Guo Y, Chen Z, Zhang L, Zhou F, Shi S, Feng X, Li B, Meng X, Ma X, Luo M, Shao K, Li N, Qiu B, Mitchelson K, Cheng J, He J (2008) Distinctive microRNA profiles relating to patient survival in esophageal squamous cell carcinoma. Cancer Res 68:26–33 Haley B, Zamore PD (2004) Kinetic analysis of the RNAi enzyme complex. Nat Struct Mol Biol 11:599–606 Han J, Lee Y, Yeom KH, Nam JW, Heo I, Rhee JK, Sohn SY, Cho Y, Zhang BT, Kim VN (2006) Molecular basis for the recognition of primary microRNAs by the Drosha-DGCR8 complex. Cell 125:887–901 Harris TA, Yamakuchi M, Ferlito M, Mendell JT, Lowenstein CJ (2008) MicroRNA-126 regulates endothelial expression of vascular cell adhesion molecule 1. Proc Natl Acad Sci USA 105:1516–1521 Hartig JS, Gru¨ne I, Hani Najafi-Shoushtari S, Famulok M (2004) Sequence-Specific Detection of MicroRNAs by Signal-Amplifying Ribozymes. J Am Chem Soc 126: 722–723 Hatfield SD, Shcherbata HR, Fischer KA, Nakahara K, Carthew RW, Ruohola-Baker H (2005) Stem cell division is regulated by the microRNA pathway. Nature 435:974–978 Heneghan HM, Miller N, Lowery AJ, Sweeney KJ, Kerin MJ (2009) MicroRNAs as Novel Biomarkers for Breast Cancer. J Oncol 2009:950201 Hohjoh H, Fukushima T (2007) Expression profile analysis of microRNA (miRNA) in mouse central nervous system using a new miRNA detection system that examines hybridization signals at every step of washing. Gene 391:39–44 Houbaviy HB, Murray MF, Sharp PA (2003) Embryonic stem cell-specific MicroRNAs. Dev Cell 5:351–358 Houzet L, Yeung ML, de Lame V, Desai D, Smith SM, Jeang KT (2008) MicroRNA profile changes in human immunodeficiency virus type 1 (HIV-1) seropositive individuals. Retrovirology 5:118 Humphreys DT, Westman BJ, Martin DI, Preiss T (2005) MicroRNAs control translation initiation by inhibiting eukaryotic initiation factor 4E/cap and poly(A) tail function. Proc Natl Acad Sci USA 102:16961–16966

References

57

Hunt EA, Goulding AM, Deo SK (2009) Direct detection and quantification of microRNAs. Anal Biochem 2009 Jan 18. [Epub ahead of print] Hutva´gner G, Zamore PD (2002) A microRNA in a multiple-turnover RNAi enzyme complex. Science 297:2056–2060 Ibarra I, Erlich Y, Muthuswamy SK, Sachidanandam R, Hannon GJ (2007) A role for microRNAs in maintenance of mouse mammary epithelial progenitor cells. Genes Dev 21:3238–3243 Iorio MV, Ferracin M, Liu CG, Veronese A, Spizzo R, Sabbioni S, Magri E, Pedriali M, Fabbri M, Campiglio M, Me´nard S, Palazzo JP, Rosenberg A, Musiani P, Volinia S, Nenci I, Calin GA, Querzoli P, Negrini M, Croce CM (2005) MicroRNA gene expression deregulation in human breast cancer. Cancer Res 65:7065–7070 Iorio MV, Visone R, Di Leva G, Donati V, Petrocca F, Casalini P, Taccioli C, Volinia S, Liu CG, Alder H, Calin GA, Me´nard S, Croce CM (2007) MicroRNA signatures in human ovarian cancer. Cancer Res 67:8699–8707 Jackson RJ, Standart N (2007) How do microRNAs regulate gene expression? Sci STKE 23:243–249 Ji R, Cheng Y, Yue J, Yang J, Liu X, Chen H, Dean DB, Zhang C (2007) MicroRNA expression signature and antisense-mediated depletion reveal an essential role of MicroRNA in vascular neointimal lesion formation. Circ Res 100:1579–1588 Jiang J, Lee EJ, Gusev Y, Schmittgen TD (2005) Real-time expression profiling of microRNA precursors in human cancer cell lines. Nucleic Acids Res 33:5394–5403 Jonstrup SP, Koch J, Kjems J (2006) A microRNA detection system based on padlock probes and rolling circle amplification. RNA 12:1747–1752 Jopling CL, Yi M, Lancaster AM, Lemon SM, Sarnow P (2005) Modulation of hepatitis C virus RNA abundance by a liver-specific MicroRNA. Science 309:1577–1581 Liang Y, Ridzon D, Wong L, Chen C (2007) Characterization of microRNA expression profiles in normal human tissues. BMC Genomics 8:166 Katada T, Ishiguro H, Kuwabara Y, Kimura M, Mitui A, Mori Y, Ogawa R, Harata K, Fujii Y (2009) microRNA expression profile in undifferentiated gastric cancer. Int J Oncol 34:537–542 Khvorova A, Reynolds A, Jayasena SD (2003) Functional siRNAs and miRNAs exhibit strand bias. Cell 115:209–216 Kim VN (2005) MicroRNA biogenesis: coordinated cropping and dicing. Nat Rev Mol Cell Biol 6:376–385 Kloosterman WP, Lagendijk AK, Ketting RF, Moulton JD, Plasterk RH (2007) Targeted inhibition of miRNA maturation with morpholinos reveals a role for miR-375 in pancreatic islet development. PLoS Biol 5:e203 Kloosterman WP, Plasterk RHA (2006) The diverse functions of microRNAs in animal development and disease. Dev Cell 11:441–450 Klo¨ting N, Berthold S, Kovacs P, Scho¨n MR, Fasshauer M, Ruschke K, Stumvoll M, Blu¨her M (2009) MicroRNA expression in human omental and subcutaneous adipose tissue. PLoS One 4:e4699 Krek A, Gru¨n D, Poy MN, Wolf R, Rosenberg L, Epstein EJ, MacMenamin P, da Piedade I, Gunsalus KC, Stoffel M, Rajewsky N (2005) Combinatorial microRNA target predictions. Nat Genet 37:495–500 Krichevsky AM, King KS, Donahue CP, Khrapko K, Kosik KS (2003) A microRNA array reveals extensive regulation of microRNAs during brain development. RNA 9:1274–1281 Kuehbacher A, Urbich C, Zeiher AM, Dimmeler S (2007) Role of Dicer and Drosha for endothelial microRNA expression and angiogenesis. Circ Res 101:59–68 Kuwabara T, Hsieh J, Nakashima K, Taira K, Gage FH (2004) A small modulatory dsRNA specifies the fate of adult neural stem cells. Cell 116:779–793 Lagos-Quintana M, Rauhut R, Yalcin A, Meyer J, Lendeckel W, Tuschl T (2002) Identification of tissue-specific microRNAs from mouse. Curr Biol 12:735–739 Lakshmipathy U, Love B, Adams C, Thyagarajan B, Chesnut JD (2007a) Micro RNA profiling: an easy and rapid method to screen and characterize stem cell populations. Methods Mol Biol 407:97–114

58

1 Detection, Profiling, and Quantification of miRNA Expression

Lakshmipathy U, Love B, Goff LA, Jo¨rnsten R, Graichen R, Hart RP, Chesnut JD (2007b) MicroRNA expression pattern of undifferentiated and differentiated human embryonic stem cells. Stem Cells Dev 16:1003–1016 Landgraf P, Rusu M, Sheridan R, Sewer A, Iovino N, Aravin A, Pfeffer S, Rice A, Kamphorst AO, Landthaler M, Lin C, Socci ND, Hermida L, Fulci V, Chiaretti S, Foa` R, Schliwka J, Fuchs U, Novosel A, Mu¨ller RU, Schermer B, Bissels U, Inman J, Phan Q, Chien M, Weir DB, Choksi R, De Vita G, Frezzetti D, Trompeter HI, Hornung V, Teng G, Hartmann G, Palkovits M, Di Lauro R, Wernet P, Macino G, Rogler CE, Nagle JW, Ju J, Papavasiliou FN, Benzing T, Lichter P, Tam W, Brownstein MJ, Bosio A, Borkhardt A, Russo JJ, Sander C, Zavolan M, Tuschl T (2007) A mammalian microRNA expression atlas based on small RNA library sequencing. Cell 129:1401–1414 Landthaler M, Yalcin A, Tuschl T (2004) The human DiGeorge syndrome critical region gene 8 and Its D. melanogaster homolog are required for miRNA biogenesis. Curr Biol 14:2162–2167 Lawrie CH (2007) MicroRNAs and haematology: small molecules, big function. Br J Haematol 137:503–512 Lee TI, Rinaldi NJ, Robert F, Odom DT, Bar-Joseph Z, Gerber GK, Hannett NM, Harbison CT, Thompson CM, Simon I, Zeitlinger J, Jennings EG, Murray HL, Gordon DB, Ren B, Wyrick JJ, Tagne JB, Volkert TL, Fraenkel E, Gifford DK, Young RA (2002a) Transcriptional regulatory networks in Saccharomyces cerevisiae. Science 298:799–804 Lee Y, Jeon K, Lee JT, Kim S, Kim VN (2002b) MicroRNA maturation: stepwise processing and subcellular localization. EMBO J 21:4663–4670 Lee Y, Hur I, Park SY, Kim YK, Suh MR, Kim VN (2006) The role of PACT in theRNA silencing pathway. EMBO J 25:522–532 Lewis BP, Shih IH, Jones-Rhoades MW, Bartel DP, Burge CB (2003) Prediction of mammalian microRNA targets. Cell 115:787–798 Lewis BP, Burge CB, Bartel DP (2005) Conserved seed pairing, often flanked by adenosines, indicates that thousands of human genes are microRNA targets. Cell 120:15–20 Lim LP, Lau NC, Garrett-Engele P, Grimson A, Schelter JM, Castle J, Bartel DP, Linsley PS, Johnson JM (2005) Microarray analysis shows that some microRNAs downregulate large numbers of target mRNAs. Nature 433:769–773 Lian J, Zhang X, Tian H, Liang N, Wang Y, Liang C, Li X, Sun F (2009) Altered microRNA expression in patients with non-obstructive azoospermia. Reprod Biol Endocrinol 7:13 Liang Y, Ridzon D, Wong L, Chen C (2007) Characterization of microRNA expression profiles in normal human tissues. BMC Genomics 8:166 Liang Y (2008) An expression meta-analysis of predicted microRNA targets identifies a diagnostic signature for lung cancer. BMC Med Genomics 1:61 Lin Y, Liu X, Cheng Y, Yang J, Huo Y, Zhang C (2009) Involvements of microRNAs in hydrogen peroxide-mediated gene regulation and cellular injury response in vascular smooth muscle cells. J Biol Chem 284:7903–7913 Liu C-G, Calin GA, Meloon B, Gamliel N, Sevignani C, Ferracin M, Dumitru CD, Shimizu M, Zupo S, Dono M, Alder H, Bullrich F, Negrini M, Croce CM (2004) An oligonucleotide microchip for genome-wide microRNA profiling in human and mouse tissues. Proc Natl Acad Sci USA 101:9740–9744 Liu Y, Mochizuki K, Gorovsky MA (2004) Histone H3 lysine 9 methylation is required for DNA elimination in developing macronuclei in Tetrahymena. Proc Natl Acad Sci USA 101:1679–1684 Lovis P, Gattesco S, Regazzi R (2008a) Regulation of the expression of components of the exocytotic machinery of insulin-secreting cells by microRNAs. Biol Chem 389:305–312 Lovis P, Roggli E, Laybutt DR, Gattesco S, Yang JY, Widmann C, Abderrahmani A, Regazzi R (2008b) Alterations in microRNA expression contribute to fatty acid-induced pancreatic beta-cell dysfunction. Diabetes 57:2728–2736 Lu J, Getz G, Miska EA, Alvarez-Saavedra E, Lamb J, Peck D, Sweet-Cordero A, Ebert BL, Mak RH, Ferrando AA, Downing JR, Jacks T, Horvitz HR, Golub TR (2005) MicroRNA expression profiles classify human cancers. Nature 435:745–746

References

59

Lu Y, Thomson JM, Wong HY, Hammond SM, Hogan BL (2007) Transgenic over-expression of the microRNA miR-17-92 cluster promotes proliferation and inhibits differentiation of lung epithelial progenitor cells. Dev Biol 310:442–453 Lund E, Guttinger S, Calado A, Dahlberg JE, Kutay U (2004) Nuclear export of microRNA precursors. Science 303:95–98 Luo X, Lin H, Pan Z, Xiao J, Zhang Y, Lu Y, Yang B, Wang Z (2008) Overexpression of Sp1 and downregulation of miR-1/miR-133 activates re-expression of pacemaker channel genes HCN2 and HCN4 in hypertrophic heart. J Biol Chem 283:20045–20052 Ma L, Teruya-Feldstein J, Weinberg RA (2007) Tumour invasion and metastasis initiated by microRNA-10b in breast cancer. Nature 449:682–688 Ma JB, Yuan YR, Meister G, Pei Y, Tuschl T, Patel DJ (2005) Structural basis for 50-endspecific recognition of guide RNA by the A. fulgidus Piwi protein. Nature 434:666–670 Macanovic A, Marquette C, Polychronakos C, Lawrence MF (2004) Impedance-based detection of DNA sequences using a silicon transducer with PNA as the probe layer. Nucleic Acids Res 32:e20 Mani SA, Guo W, Liao MJ, Eaton EN, Ayyanan A, Zhou AY, Brooks M, Reinhard F, Zhang CC, Shipitsin M, Campbell LL, Polyak K, Brisken C, Yang J, Weinberg RA (2008) The epithelialmesenchymal transition generates cells with properties of stem cells. Cell 133:704–715 Markou A, Tsaroucha EG, Kaklamanis L, Fotinou M, Georgoulias V, Lianidou ES (2008) Prognostic value of mature microRNA-21 and microRNA-205 overexpression in non-small cell lung cancer by quantitative real-time RT-PCR. Clin Chem 54:1696–1704 Maroney PA, Chamnongpol S, Souret F, Nilsen TW (2007) A rapid, quantitative assay for direct detection of microRNAs and other small RNAs using splinted ligation. RNA 13:930–936 Marton S, Garcia MR, Robello C, Persson H, Trajtenberg F, Pritsch O, Rovira C, Naya H, Dighiero G, Cayota A (2008) Small RNAs analysis in CLL reveals a deregulation of miRNA expression and novel miRNA candidates of putative relevance in CLL pathogenesis. Leukemia 22:330–338 McKinsey TA, Olson EN (2005) Toward transcriptional therapies for the failing heart: chemical screens to modulate genes. J Clin Invest 115:538–546 Meister G, Tuschl T (2004) Mechanisms of gene silencing by double-stranded RNA. Nature 431:343–349 Mestdagh P, Feys T, Bernard N, Guenther S, Chen C, Speleman F, Vandesompele J (2008) High-throughput stem-loop RT-qPCR miRNA expression profiling using minute amounts of input RNA. Nucleic Acids Res 36:e143 Miranda KC, Huynh T, Tay Y, Ang YS, Tam WL, Thomson AM, Lim B, Rigoutsos I (2006) A pattern-based method for the identification of MicroRNA binding sites and their corresponding heteroduplexes. Cell 126:1203–1217 Mirkin CA, Letsinger RL, Mucic RC, Storhoff JJ (1996) A DNA-based method for rationally assembling nanoparticles into macroscopic materials. Nature 382:607–609 Miska EA, Alvarez-Saavedra E, Townsend M, Yoshii A, Sestan N, Rakic P, Constantine-Paton M, Horvitz HR (2004) Microarray analysis of microRNA expression in the developing mammalian brain. Genome Biol 5:R68 Montenegro D, Romero R, Kim SS, Tarca AL, Draghici S, Kusanovic JP, Kim JS, Lee DC, Erez O, Gotsch F, Hassan SS, Kim CJ (2009) Expression patterns of microRNAs in the chorioamniotic membranes: a role for microRNAs in human pregnancy and parturition. J Pathol 217:113–121 Mora JR, Getts RC (2006) Enzymatic microRNA detection in microtiter plates with DNA dendrimers. Biotechniques 41:420–424 Murchison EP, Partridge JF, Tam OH, Cheloufi S, Hannon GJ (2005) Characterization of Dicerdeficient murine embryonic stem cells. Proc Natl Acad Sci USA 102:12135–12140 Nam JM, Stoeva SI, Mirkin CA (2004) Bio-bar-code-based DNA detection with PCR-like sensitivity. J Am Chem Soc 126:5932–5933 Nam JM, Thaxton CS, Mirkin CA (2003) Nanoparticle-based bio-bar codes for the ultrasensitive detection of proteins. Science 301:1884–1886

60

1 Detection, Profiling, and Quantification of miRNA Expression

Neely LA, Patel S, Garver J, Gallo M, Hackett M, McLaughlin S, Nadel M, Harris J, Gullans S, Rooke J (2006) A single-molecule method for the quantitation of microRNA gene expression. Nat Methods 3:41–46 Neely LA, Rieger-Christ KM, Neto BS, Eroshkin A, Garver J, Patel S, Phung NA, McLaughlin S, Libertino JA, Whitney D, Summerhayes IC (2009) A microRNA expression ratio defining the invasive phenotype in bladder tumors. Urol Oncol [Epub ahead of print] Nelson PT, Baldwin DA, Kloosterman W.P, Kauppinen S, Plasterk RH, Mourelatos Z (2006) RAKE and LNA-ISH reveal microRNA expression and localization in archival human brain. RNA 12:187–191 Nelson PT, Baldwin DA, Scearce LM, Oberholtzer JC, Tobias JW, Mourelatos Z (2004) Microarraybased, high-throughput gene expression profiling of microRNAs. Nat Methods 1:155–161 Nilsen TW (2007) Mechanisms of microRNA-mediated gene regulation in animal cells. Trends Genet 23:243–249 Ng EK, Chong WW, Jin H, Lam EK, Shin VY, Yu J, Poon TC, Ng SS, Sung JJ (2009) Differential expression of microRNAs in plasma of colorectal cancer patients: A potential marker for colorectal cancer screening. Gut . 2009 Feb 9. [Epub ahead of print] O’Connell RM, Rao DS, Chaudhuri AA, Boldin MP, Taganov KD, Nicoll J, Paquette RL, Baltimore D (2008) Sustained expression of microRNA-155 in hematopoietic stem cells causes a myeloproliferative disorder. J Exp Med 205:585–594 Okamura K, Hagen JW, Duan H, Tyler DM, Lai EC (2007) The mirtron pathway generates microRNA-class regulatory RNAs in Drosophila. Cell 130:89–100 Okamura K, Ishizuka A, Siomi H, Siomi MC (2004) Distinct roles for Argonaute proteins in small RNA-directed RNA cleavage pathways. Genes Dev 18:1655–1666 Pallante P, Visone R, Ferracin M, Ferraro A, Berlingieri MT, Troncone G, Chiappetta G, Liu CG, Santoro M, Negrini M, Croce CM, Fusco A (2006) MicroRNA deregulation in human thyroid papillary carcinomas. Endocr Relat Cancer 13:497–508 Park SM, Gaur AB, Lengyel E, Peter ME (2008) The miR-200 family determines the epithelial phenotype of cancer cells by targeting the E-cadherin repressors ZEB1 and ZEB2. Genes Dev 22:894–907 Park CS, Tang SJ (2009) Regulation of microRNA Expression by Induction of Bidirectional Synaptic Plasticity. J Mol Neurosci 38:50–56 Park SJ, Taton TA, Mirkin CA (2002) Array-based electrical detection of DNA with nanoparticle probes. Science 295:1503–1506 Parker JS, Roe SM, Barford D (2005) Structural insights into mRNA recognition from a PIWI domain-siRNA guide complex. Nature 434:663–666 Perkins DO, Jeffries CD, Jarskog LF, Thomson JM, Woods K, Newman MA, Parker JS, Jin J, Hammond SM (2007) microRNA expression in the prefrontal cortex of individuals with schizophrenia and schizoaffective disorder. Genome Biol 8:R27 Pillai RS, Bhattacharyya SN, Filipowicz W (2007) Repression of protein synthesis by miRNAs: how many mechanisms? Trends Cell Biol 17:18–126 Poy MN, Eliasson L, Krutzfeldt J, Kuwajima S, Ma X, Macdonald PE, Pfeffer S, Tuschl T, Rajewsky N, Rorsman P, Stoffel M (2004) A pancreatic islet-specific microRNA regulates insulin secretion. Nature 432:226–230 Poy MN, Spranger M, Stoffel M (2007) microRNAs and the regulation of glucose and lipid metabolism. Diabetes Obes Metab Suppl 2:67–73 Rajewsky N, Socci ND (2004) Computational identification of microRNA targets. Dev Biol 267:529–535 Ramdas L, Giri U, Ashorn CL, Coombes KR, El-Naggar A, Ang KK, Story MD (2009) miRNA expression profiles in head and neck squamous cell carcinoma and adjacent normal tissue. Head Neck 31:642–654 Raymond CK, Roberts BS, Garrett-Engele P, Lim LP, Johnson JM (2005) Simple, quantitative primer-extension PCR assay for direct monitoring of microRNAs and short-interfering RNAs. RNA 11:1737–1744

References

61

Ren XP, Wu J, Wang X, Sartor MA, Qian J, Jones K, Nicolaou P, Pritchard TJ, Fan GC (2009) MicroRNA-320 is involved in the regulation of cardiac ischemia/reperfusion injury by targeting heat-shock protein 20. Circulation 119:2357–2366 Ro S, Park C, Sanders KM, McCarrey JR, Yan W (2007) Cloning and expression profiling of testis-expressed microRNAs. Dev Biol 311:592–602 Roldo C, Missiaglia E, Hagan JP, Falconi M, Capelli P, Bersani S, Calin GA, Volinia S, Liu CG, Scarpa A, Croce CM (2006) MicroRNA expression abnormalities in pancreatic endocrine and acinar tumors are associated with distinctive pathologic features and clinical behavior. J Clin Oncol 24:4677–4684 Roy S, Khanna S, Hussain SR, Biswas S, Azad A, Rink C, Gnyawali S, Shilo S, Nuovo GJ, Sen CK (2009) MicroRNA expression in response to murine myocardial infarction: miR-21 regulates fibroblast metalloprotease-2 via phosphatase and tensin homologue. Cardiovasc Res 82:21–29 Ruby JG, Jan C, Bartel DP (2007) Intronic microRNA precursors that bypass Drosha processing. Nature 448:83–86 Saba R, Booth SA (2006) Target labelling for the detection and profiling of microRNAs expressed in CNS tissue using microarrays. BMC Biotechnol 6:47 Saba R, Goodman CD, Huzarewich RL, Robertson C, Booth SA (2008) A miRNA signature of prion induced neurodegeneration. PLoS ONE 3:e3652 Sayed D, Hong C, Chen IY, Lypowy J, Abdellatif M (2007) MicroRNAs play an essential role in the development of cardiac hypertrophy. Circ Res 100:416–424 Schepeler T, Reinert JT, Ostenfeld MS, Christensen LL, Silahtaroglu AN, Dyrskjøt L, Wiuf C, Sørensen FJ, Kruhøffer M, Laurberg S, Kauppinen S, Ørntoft TF, Andersen CL (2008) Diagnostic and prognostic microRNAs in stage II colon cancer. Cancer Res 68:6416–6424 Schetter AJ, Leung SY, Sohn JJ, Zanetti KA, Bowman ED, Yanaihara N, Yuen ST, Chan TL, Kwong DL, Au GK, Liu CG, Calin GA, Croce CM, Harris CC (2008) MicroRNA expression profiles associated with prognosis and therapeutic outcome in colon adenocarcinoma. JAMA 299:425–436 Schmittgen TD, Jiang J, Liu Q, Yang L (2004) A high-throughput method to monitor the expression of microRNA precursors. Nucleic Acids Res 32:e43 Schwarz DS, Hutvagner G, Du T, Xu Z, Aronin N, Zamore PD (2003) Asymmetry in the assembly of the RNAi enzyme complex. Cell 115:199–208 Sempere LF, Christensen M, Silahtaroglu A, Bak M, Heath CV, Schwartz G, Wells W, Kauppinen S, Cole CN (2007) Altered MicroRNA expression confined to specific epithelial cell subpopulations in breast cancer. Cancer Res 67:11612–11620 Sempere LF, Freemantle S, Pitha-Rowe I, Moss E, Dmitrovsky E, Ambros V (2004) Expression profiling of mammalian microRNAs uncovers a subset of brain-expressed microRNAs with possible roles in murine and human neuronal differentiation, Genome Biol 5:R13 Shan ZX, Lin QX, Fu YH, Deng CY, Zhou ZL, Zhu JN, Liu XY, Zhang YY, Li Y, Lin SG, Yu XY (2009) Upregulated expression of miR-1/miR-206 in a rat model of myocardial infarction. Biochem Biophys Res Commun 381:597–601 Shi R, Chiang VL (2005) Facile means for quantifying microRNA expression by real-time PCR. Biotechniques 39:519–525 Shingara J, Keiger K, Shelton J, Laosinchai-Wolf W, Powers P, Conrad R, Brown D, Labourier E (2005) An optimized isolation and labeling platform for accurate microRNA expression profiling. RNA 11:1461–1470 Silveri L, Tilly G, Vilotte JL, Le Provost F (2006) MicroRNA involvement in mammary gland development and breast cancer. Reprod Nutr Dev 46:549–556 Slaby O, Svoboda M, Fabian P, Smerdova T, Knoflickova D, Bednarikova M, Nenutil R, Vyzula R (2007) Altered expression of miR-21, miR-31, miR-143 and miR-145 is related to clinicopathologic features of colorectal cancer. Oncology 72:397–402 Song JJ, Smith SK, Hannon GJ, Joshua-Tor L (2004) Crystal structure of Argonaute and its implications for RISC slicer activity. Science 305:1434–1437

62

1 Detection, Profiling, and Quantification of miRNA Expression

Stoeva SI, Lee JS, Smith JE, Rosen ST, Mirkin CA (2006) Multiplexed detection of protein cancer markers with biobarcoded nanoparticle probes. J Am Chem Soc 128:8378–8379 Strillacci A, Griffoni C, Sansone P, Paterini P, Piazzi G, Lazzarini G, Spisni E, Pantaleo MA, Biasco G, Tomasi V (2009) MiR-101 downregulation is involved in cyclooxygenase-2 overexpression in human colon cancer cells. Exp Cell Res 315:1439–1447 Suh MR, Lee Y, Kim JY, Kim SK, Moon SH, Lee JY, Cha KY, Chung HM, Yoon HS, Moon SY, Kim VN, Kim KS (2004) Human embryonic stem cells express a unique set of microRNAs. Dev Biol 270:488–498 Takada S, Mano H (2007) Profiling of microRNA expression by mRAP. Nat Protoc 2:3136–3145 Takagi T, Iio A, Nakagawa Y, Naoe T, Tanigawa N, Akao Y (2009) Decreased expression of microRNA-143 and -145 in human gastric cancers. Oncology 77:12–21 Tang F, Hajkova P, Barton SC, Lao K, Surani MA (2006a) MicroRNA expression profiling of single whole embryonic stem cells. Nucleic Acids Res 34:e9 Tang F, Hajkova P, Barton SC, O’Carroll D, Lee C, Lao K, Surani MA (2006b) 220-plex microRNA expression profile of a single cell. Nat Protoc 1:1154–1159 Tang JT, Fang JY (2009) MicroRNA regulatory network in human colorectal cancer. Mini Rev Med Chem 9:921–926 Tang Y, Zheng J, Sun Y, Wu Z, Liu Z, Huang G (2009a) MicroRNA-1 regulates cardiomyocyte apoptosis by targeting Bcl-2. Int Heart J 50:377–387 Tang X, Muniappan L, Tang G, Ozcan S (2009b) Identification of glucose-regulated miRNAs from pancreatic b cells reveals a role for miR-30d in insulin transcription. RNA. 15:287–293 Taton TA, Lu G, Mirkin CA (2001) Two-color labeling of oligonucleotide arrays via size-selective scattering of nanoparticle probes. J Am Chem Soc 123:5164–5165 Taton TA, Mirkin CA, Letsinger RL (2000) Scanometric DNA array detection with nanoparticle probes. Science 289:1757–1760 Tatsuguchi M, Seok HY, Callis TE, Thomson JM, Chen JF, Newman M, Rojas M, Hammond SM, Wang DZ (2007) Expression of microRNAs is dynamically regulated during cardiomyocyte hypertrophy. J Mol Cell Cardiol 42:1137–1141 Tavazoie SF, Alarco´n C, Oskarsson T, Padua D, Wang Q, Bos PD, Gerald WL, Massague´ J (2008) Endogenous human microRNAs that suppress breast cancer metastasis. Nature 451:147–152 Tay Y, Zhang J, Thomson AM, Lim B, Rigoutsos I (2008) MicroRNAs to Nanog, Oct4 and Sox2 coding regions modulate embryonic stem cell differentiation. Nature 455:1124–1128 Thomson JM, Parker J, Perou CM, Hammond SM (2004) A custom microarray platform for analysis of microRNA gene expression. Nat Methods 1:47–53 Thomson JM, Parker J, Perou CM, Hammond SM, John JR, Gregory JH (2007) Microarray analysis of miRNA gene expression. Methods in Enzymology, Academic Press pp107–122 Thum T, Galuppo P, Wolf C, Fiedler J, Kneitz S, van Laake LW, Doevendans PA, Mummery CL, Borlak J, Haverich A, Gross C, Engelhardt S, Ertl G, Bauersachs J (2007) MicroRNAs in the human heart: a clue to fetal gene reprogramming in heart failure. Circulation 116:258–267 Tong AW, Fulgham P, Jay C, Chen P, Khalil I, Liu S, Senzer N, Eklund AC, Han J, Nemunaitis J (2009) MicroRNA profile analysis of human prostate cancers. Cancer Gene Ther 16:206–216 Ura S, Honda M, Yamashita T, Ueda T, Takatori H, Nishino R, Sunakozaka H, Sakai Y, Horimoto K, Kaneko S (2009) Differential microRNA expression between hepatitis B and hepatitis C leading disease progression to hepatocellular carcinoma. Hepatology 49:1098–1112 Va´lo´czi A, Hornyik C, Varga N, Burgya´n J, Kauppinen S, Havelda Z (2004) Sensitive and specific detection of microRNAs by northern blot analysis using LNA-modified oligonucleotide probes. Nucleic Acids Res 32:e175 van Rooij E, Sutherland LB, Liu N, Williams AH, McAnally J, Gerard RD, Richardson JA, Olson EN (2006) A signature pattern of stress-responsive microRNAs that can evoke cardiac hypertrophy and heart failure. Proc Natl Acad Sci USA 103: 18255–18260 van Rooij E, Sutherland LB, Qi X, Richardson JA, Hill J, Olson EN (2007) Control of stressdependent cardiac growth and gene expression by a microRNA. Science 316:575–579

References

63

Varnholt H, Drebber U, Schulze F, Wedemeyer I, Schirmacher P, Dienes HP, Odenthal M (2008) MicroRNA gene expression profile of hepatitis C virus-associated hepatocellular carcinoma. Hepatology 47:1223–1232 Vasudevan S, Tong Y, Steitz JA (2007) Switching from repression to activation: microRNAs can up-regulate translation. Science 318:1931–1934 Venturini L, Battmer K, Castoldi M, Schultheis B, Hochhaus A, Muckenthaler MU, Ganser A, Eder M, Scherr M (2007) Expression of the miR-17-92 polycistron in chronic myeloid leukemia (CML) CD34+ cells. Blood 109:4399–4405 Volinia S, Calin GA, Liu CG, Ambs S, Cimmino A, Petrocca F, Visone R, Iorio M, Roldo C, Ferracin M, Prueitt RL, Yanaihara N, Lanza G, Scarpa A, Vecchione A, Negrini M, Harris CC, Croce CM (2006) A microRNA expression signature of human solid tumors defines cancer gene targets. Proc Natl Acad Sci USA 103:2257–2261 Wang B, Love TM, Call ME, Doench JG, Novina CD (2006) Recapitulation of short RNAdirected translational gene silencing in vitro. Mol Cell 22:553–560 Wang Y, Blelloch R (2009) Cell cycle regulation by MicroRNAs in embryonic stem cells. Cancer Res 69:4093–4096 Wang Z, Luo X, Luy, Yang B (2008) miRNAS at the heart of the matter. J Mol Med 86:772–783 Wu H, Neilson JR, Kumar P, Manocha M, Shankar P, Sharp PA, Manjunath N (2007) miRNA profiling of naı¨ve, effector and memory CD8 T cells. PLoS ONE 2:e1020 Wu L, Belasco JG (2005) Micro-RNA regulation of the mammalian lin-28 gene during neuronal differentiation of embryonal carcinoma cells. Mol Cell Biol 25:9198–9208 Wu W, Lin Z, Zhuang Z, Liang X (2009) Expression profile of mammalian microRNAs in endometrioid adenocarcinoma. Eur J Cancer Prev 18:50–55 Xi Y, Formentini A, Chien M, Weir DB, Russo JJ, Ju J, Kornmann M, Ju J (2006) Prognostic Values of microRNAs in Colorectal Cancer. Biomark Insights 2:113–121 Xiao C, Calado DP, Galler G, Thai TH, Patterson HC, Wang J, Rajewsky N, Bender TP, Rajewsky K (2007) MiR-150 controls B cell differentiation by targeting the transcription factor c-Myb. Cell 131:146–159 Xu C, Lu Y, Lin H, Xiao, J, Wang H, Luo X, Li B, Yang B, Wang Z (2007) The muscle-specific microRNAs miR-1 and miR-133 produce opposing effects on apoptosis via targeting HSP60/ HSP70 and caspase-9 in cardiomyocytes. J Cell Sci 120:3045–3052 Xu S, Witmer PD, Lumayag S, Kovacs B, Valle D (2007) MicroRNA (miRNA) transcriptome of mouse retina and identification of a sensory organ-specific miRNA cluster. J Biol Chem 282:25053–25066 Yanaihara N, Caplen N, Bowman E, Seike M, Kumamoto K, Yi M, Stephens RM, Okamoto A, Yokota J, Tanaka T, Calin GA, Liu CG, Croce CM, Harris CC (2006) Unique microRNA molecular profiles in lung cancer diagnosis and prognosis. Cancer Cell 9:189–198 Yang B, Lin H, Xiao J, Lu Y, Luo X, Li B, Zhang Y, Xu C, Bai Y, Wang H, Chen G, Wang Z (2007) The muscle-specific microRNA miR-1 causes cardiac arrhythmias by targeting GJA1 and KCNJ2 genes. Nat Med 13:486–491 Yang B, Lu Y, Wang Z (2008a) Control of cardiac excitability by miRNAs. Cardiovasc Res 79:571–580 Yang B, Lu Y, Wang Z (2009) MicroRNAs and Apoptosis: Implications in Molecular Therapy of Human Disease. Clin Exp Pharmacol Physiol 2009 Jun 29. [Epub ahead of print] Yang N, Kaur S, Volinia S, Greshock J, Lassus H, Hasegawa K, Liang S, Leminen A, Deng S, Smith L, Johnstone CN, Chen XM, Liu CG, Huang Q, Katsaros D, Calin GA, Weber BL, Bu¨tzow R, Croce CM, Coukos G, Zhang L (2008b) MicroRNA microarray identifies Let-7i as a novel biomarker and therapeutic target in human epithelial ovarian cancer. Cancer Res 68: 10307–10314 Yekta S, Shih IH, Bartel DP (2004) MicroRNA-directed cleavage of HOXB8 mRNA. Science 304:594–596 Yi R, Qin Y, Macara IG, Cullen BR (2003) Exportin-5 mediates the nuclear export of premicroRNAs and short hairpin RNAs. Genes Dev 17:3011–3016

64

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Yin C, Salloum FN, Kukreja RC (2009) A novel role of microRNA in late preconditioning: upregulation of endothelial nitric oxide synthase and heat shock protein 70. Circ Res 104:572–575 Yin C, Wang X, Kukreja RC (2008) Endogenous microRNAs induced by heat-shock reduce myocardial infarction following ischemia-reperfusion in mice. FEBS Lett 582:4137–4142 Ying SY, Lin SL (2005) Intronic microRNAs. Biochem Biophys Res Commun 326:515–520 Yu B, Yang Z, Li J, Minakhina S, Yang M, Padgett RW, Steward R, Chen X (2005) Methylation as a crucial step in plant microRNAs biogenesis. Science 307:932–935 Zanette DL, Rivadavia F, Molfetta GA, Barbuzano FG, Proto-Siqueira R, Silva-Jr WA, Falca˜o RP, Zago MA (2007) miRNA expression profiles in chronic lymphocytic and acute lymphocytic leukemia. Braz J Med Biol Res 40:1435–1440 Zhan M, Miller CP, Papayannopoulou T, Stamatoyannopoulos G, Song CZ (2007) MicroRNA expression dynamics during murine and human erythroid differentiation. Exp Hematol 35:1015–1025 Zhang Y, Li M, Wang H, Fisher WE, Lin PH, Yao Q, Chen C (2009) Profiling of 95 microRNAs in pancreatic cancer cell lines and surgical specimens by real-time PCR analysis. World J Surg 2008 Nov 22. [Epub ahead of print] Zhao JJ, Yang J, Lin J, Yao N, Zhu Y, Zheng J, Xu J, Cheng JQ, Lin JY, Ma X (2009) Identification of miRNAs associated with tumorigenesis of retinoblastoma by miRNA microarray analysis. Childs Nerv Syst 25:13–20 Zhao Y, Samal E, Srivastava D (2005) Serum response factor regulates a muscle specific microRNA that targets Hand2 during cardiogenesis. Nature 436:214–220 Zhu S, Wu H, Wu F, Nie D, Sheng S, Mo YY (2008) MicroRNA-21 targets tumor suppressor genes in invasion and metastasis. Cell Res 18:350–359

Part II miRNA Microarray Methods

Chapter 2

Microarray and Its Variants for miRNA Profiling

Abstract Oligo microarray technology is simply founded on the Watson–Crick base-pairing nature of nucleic acids. Synthesized antisenseprobes are spotted and immobilized onto a nylon support platform using a handheld spotting device (RNA 9:1274–1281, 2003). Microarray-based techniques are particularly attractive for miRNA profiling as they are able to screen large numbers of miRNAs simultaneously. The microarray technology was first developed in 1995 (Science 270:467–470, 1995), on the basis of the ability to perform multiple hybridizations in parallel with the oligo probes pre-spotted on a glass or a quartz support platform (Methods Enzymol 410:3–28, 2006; Methods Enzymol 410:73–98, 2006; Methods Enzymol 410:28–57, 2006). This strategy was later adapted and modified for miRNA profiling. Thus far, the majority of the published papers that reported profiling analysis pertained to the use of microarray technology. Several technical variants of miRNA microarrays have been independently developed in the past few years and the major variations include the oligo probe design, the probe immobilization chemistry, the sample labeling, and the microarray chip signal-detection methods. Most miRNA array platforms that have been described to date make use of DNA oligonucleotides as capture probes (Nature 448:83–86, 2007; Cell 130:89–100, 2007; Methods Enzymol 410:3–28, 2006; Methods Enzymol 410:73–98; Methods Enzymol 410:28–57, 2006; Science 270:467–470, 1995; Curr Biol 12:735–739, 2002; Cell 110:513–520, 2002). Several nucleic acid analogs have emerged over the last few years that demonstrate more favorable hybridization characteristics as compared to standard DNA-based probes. Some of the analogs, including peptide nucleic acids (PNA) (Nature 437:1195–1198, 2005) and locked nucleic acids (LNA), have been used on classical mRNA microarrays. Recently, Castoldi et al. (RNA 12:913–920, 2006, Methods 43:146–152, 2007) have described the application of mixed LNA/ DNA-modified capture probes in array-based profiling of miRNAs and have reported superior sensitivity over conventional DNA-based miRNA arrays. However, microarrays have encountered difficulties in reliably amplifying miRNAs without bias. Also, because of the extremely short nature of miRNAs, there is very little room to

Z. Wang and B. Yang, MicroRNA Expression Detection Methods, DOI 10.1007/978-3-642-04928-6_2, # Springer-Verlag Berlin Heidelberg 2010

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fine-tune the hybridization conditions for all miRNAs, which significantly discounts the sensitivity and specificity of microarrays.

2.1

Introduction

Oligo microarray technology is simply founded on the Watson–Crick base pairing nature of nucleic acids. Synthesized antisenseprobes are spotted and immobilized onto a nylon supportplatform using a handheld spotting device (Krichevsky et al. 2003) (Fig. 2.1). Microarray-based techniques are particularly attractive for miRNA profiling as they are able to screen large numbers of miRNAs simultaneously. The microarray technology was first developed in 1995 (Schena et al. 1995), on the basis of the ability to perform multiple hybridizations in parallel with the oligo probes pre-spotted on a glass or a quartz support platform (DalmaWeiszhausz et al. 2006; Kreil et al. 2006; Wolber et al 2006). This strategy has been adapted and modified for miRNA profiling. Thus far, the majority of the published papers that reported profiling analysis pertained to the use of microarray technology. Several technical variants of miRNA microarrays have been independently developed in the past few years and the major variations include the oligo probe desing, the probe immobilization chemistry, the sample labeling, and the microarray chip signal-detection methods. Most miRNA array platforms that have been described to date make use of DNA-oligonucleotides as capture probes (Ruby

Microarray Expression Profiling 1. Isolate RNA samples, synthesize cDNA probe for microarray hybridization Cultured cells or clinical samples

2. Hybridize labeled probe with DNA microarray on a chip

3. Scan the chip and collect raw data

4. Analyze data, i.e. perform hierarchic clustering and correlate with histoclinical data

Fig. 2.1 Schematic depiction of microarray analysis for miRNA expression profiling

2.1 Introduction

69

et al. 2007; Okamura et al. 2007; Dalma-Weiszhausz et al. 2006; Kreil et al. 2006; Wolber et al. 2006; Schena et al. 1995; Lagos-Quintana et al. 2002; Rhoades et al 2002). Application of microarray technology is challenged by several innate properties of miRNAs: (1) the short length of miRNAs offers little sequence for appending detection molecules at optimized hybridization conditions with maximal binding affinity without compromising specificity; (2) a wide range of predicted melting temperature (Tm) versus their (DNA) capture probes. The design of the probe depends solely on the sequence of the miRNA itself, which determines a different temperature of annealing for each probe::miRNA interaction; (3) low abundance of certain miRNAs representing a small fraction (<0.01%) of total cellular RNA; and (4) many different miRNAs belong to families sharing similar sequences differing by as little as a single nucleotide. To tackle these problems, several nucleic acid analogs have emerged over the last few years that demonstrate more favorable hybridization characteristics as compared to standard DNA-based probes. Some of the analogs, including peptide nucleic acids (PNA) (Gershon 2005) and locked nucleic acids (LNA), have been used on classical mRNA microarrays. Recently, Castoldi et al. (2006, 2007) described the application of mixed LNA/DNA-modified capture probes in array-based profiling of miRNAs and reported superior sensitivity over conventional DNA-based miRNA arrays. LNA is a new class of bicyclic high-affinity RNA analogs in which the furanose ring of the ribose sugar is chemically locked in an RNA-mimicking conformation by the introduction of an O20 ,C40 -methylene bridge, resulting in unprecedented hybridization affinity toward complementary DNA and RNA molecules (Fig. 2.2.) (Vester and Wengel 2004). LNA nucleotides can be mixed with DNA or RNA bases in the oligonucleotide whenever desired. Mixed LNA/DNA- or LNA/RNA-modified miRNA capture probes have many advantages: (1) very efficient binding to complementary nucleic acids, (2) high potency as antisense molecules in vitro and in vivo, and (3) commercial availability as oligonucleotides (fully modified, mix-mers with DNA, RNA, other monomers, and various modifications) and as phosphoramidite building blocks. LNA oligonucleotides can be synthesized using conventional automated phosphoramidite chemistry, and LNA monomers are compatible with other monomers, e.g., DNA, RNA, and 20 -O-Me-RNA, and with phosphorothioate (PS) and/or phosphodiester linkages (Wengel 1999; Kumar et al 1998). LNA resembles natural nucleic acids with respect to Watson–Crick base pairing. LNAs as fully modified oligomers or as mix-mers containing, for example, LNA and DNA or LNA and RNA nucleotides induce very high thermal stability of duplexes toward O

–O

Fig. 2.2 Structure of LNA

Base

O

O P =O

O

O O

O –O P = O O

LNA [b-D-ribo configuration]

Base

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2 Microarray and Its Variants for miRNA Profiling

complementary RNA or DNA (Wengel 1999). In addition, LNA:LNA base pairing, i.e., the self-annealing capacity, is so strong that it must be taken into account when designing LNAs (see www.exiqon.com for the LNA design tool). The thermal stabilities obtained for LNA oligonucleotides depend on the length of the sequence and the number of LNA nucleotides (Christensen et al. 2001). Generally, the largest impact upon introduction of LNA nucleotides is observed for short oligonucleotides with one or more centrally positioned LNA monomers (Petersen and Wengel 2003; Wengel 1999; McTigue et al. 2004). Furthermore, single or multiple, but separated, LNA modifications appear to have a larger impact, in relative terms, than contiguous stretches of LNA nucleotides. The thermal stability and improved mismatch discrimination of short LNA-modified oligonucleotides have made them useful for single nucleotide polymorphism genotyping assays, antisense-based gene silencing and gene expression profiling (Vester and Wengel 2004). LNA nucleotides have been used to increase the sensitivity and specificity of expression in DNA microarrays, FISH probes, real-time PCR probes, and other molecular biology techniques based on oligonucleotides. The ability to increase the density of the spots on the array resulted in a higher number of genes that could be analyzed simultaneously (Dalma-Weiszhausz et al. 2006; Gershon 2005). Three different technologies classically exist to detect nucleic acids (DNA or RNA) on an array platform. The first, commonly used for custom arrays, uses glass slides (poly-lysine coated common microscope slides) and is based on the spotting of unmodified oligonucleotides over the slide (Hughes et al 2006). The second uses glass slides too, and it is also based on the deposition of the probes on the slide. The distinction is that the 50 terminus of the probe is cross linked to the matrix on the glass. The number of probes present on these slides can be much higher compared to that on the former. In the last technology, the probes are photochemically synthesized directly on the surface which is made of quartz (Liu et al. 2008). In the last case, the number of probes rises to millions per area the size of a thumbnail (Wolber et al 2006). Usually, but not always, the first two are based on the comparison of two samples for each glass (one used as reference) stained with different colors. The third uses single color hybridization where each slide is hybridized with only one sample.

2.2

Protocol

2.2.1 1. 2. 3. 4. 5.

Materials

Pipette tips, sterile, RNase-free, and aerosol-resistant Pipette aid and disposable pipetts Sterile, nuclease-free conical tubes (15 and 50 mL) Microcentrifuge tubes, sterile, RNase-free, 1.7 mL CodeLink Activated Slides (GE Healthcare, PN, USA)

2.2 Protocol

6. 7. 8. 9. 10.

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Bioarray rack Omnigrid 100 slide holder 96-well plates 384 well microtiter plate 0.2 mm filter

2.2.2

Instruments

1. Gene Machine Omnigrid 100 Arrayer (Genomic Solution, Inc. 4355 Varsity Dr. Suite E, Ann Arbor MI 48108) 2. Tecan TeMo liquid handler (Tecan TEMO liquid handler, TECAN US Inc. Research Triangle Park) 3. Axon Scanner 4200 (Molecular Device Corp. CA, USA) 4. Micropipettes (10, 20, 200, 1,000 ml) 5. Nanodrop UV spectrophotometer 6. Microcentrifuge, at room temperature and at 4 C 7. Water bath (settings 70, 65, 37 C) 8. Speed-Vac concentrator 9. Vortex 10. Shaking incubator (New Brunswick Innova 4080) 11. Desiccator (Fisher) 12. Tecan TeMo liquid handler 13. Omnigrid 100 slide holder 14. Tecan HS4800 hybridization stations 15. Bioarray Position Tool 16. GenePix software 17. Tecan HS 4800 Hybridization Station 18. Axon GenePix 4000B scanner 19. Computer configured for Axon 4000B Scanner 20. Sigma/Qiagen Centrifuge (4–15 C) (Qiagen) 21. Centrifuge Plate Rotor-2 96 (Qiagen) 22. Pipette tips, sterile, RNase-free, and aerosol-resistant 23. Microcentrifuge tubes, sterile, RNase-free, 1.7 ml 24. Micropipettes 25. Powder-free gloves 26. Microcentrifuge 27. Microtiter plate lid, Black (Corning) 28. Bioarray Processors (GE, Healthcare) 29. Bioarray Rack (GE) 30. Small Reagent Reservoir (GE) 31. Large Reagent Reservoir (GE) 32. Bioarray Removal Tool (GE) 33. Bioarray Position Tool (GE)

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2.2.3

Reagents

2.2.3.1

Chemicals and Enzymes

1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 23. 24.

20 SSC (Sigma) 5 M Sodium chloride (Sigma) Bicine (Sigma B8660-1KG) Taurine (Sigma) Fluorescein (Sigma) Anhydrous monobasic sodium phosphate (Sigma) Dibasic sodium phosphate, heptahydrate (Sigma) 0.5 mg/mL 30 NNNNNNNN-(dA)12 T (biotin) (dA)12-Biotin 50 Oligonucleotide primer 5 First-strand buffer 0.1 M Dithiothreitol (DTT) 10 mM dNTP mix Superscript1 II RnaseH+ reverse transcriptase (200 U/mL) (Invitrogen) 10 mM dNTP mix (Invitrogen) 0.5% NEN Blocking Reagent (Perkin Elmer) Molecular Probes Streptavidin-Alexa Fluor1 647 conjugate (staining solution is a 1:500 dilution in TNB) (Molecular Probes) Nuclease-free H2O (Ambion) 1 PBS; pH 7.4 (Invitrogen/LTI) 1 M Tris–HCl, pH 7.6 (Sigma) 5 M NaCl (Sigma) TweenX-20 (Sigma) Formamide (Sigma) 50 Denhardt Solution (Sigma) Sodium-Fluorescein StreptavidAlexa-647

2.2.3.2

Solution Preparation

1. Phosphate buffer pH8.0 (2, 100 mM): [Resuspend 0.69 mg of anhydrous monobasic sodium phosphate and 25.46 g of dibasic sodium phosphate, heptahydrate in 900 ml of distilled H2O and adjust the solution to pH8.0 by adding 100 mL of 10 N NaOH. Add more distilled H2O to make 1,000 mL volume and filter the solution by 0.22 mm filter unit. Store the 100 mM (2) phosphate buffer at 4 C until use] 2. Fluorescein solution (2 mM): [First prepare a 200 mM solution by dissolving 18.845 mg of fluorescein in 250 mL of H2O. Dilute 5 mL of 200 mM solution with 495 ml of H2O (2 mM Fluorescein solution.] 3. Bicine and taurine array blocking solution (100 mM): [Dissolve 48.9 g of bicine and 37.5 g of taurine in 2,400 mL of H2O. Adjust to pH9.0 by adding 40 mL of 10 N NaOH. Add more H2O to a volume of 3,000 mL.]

2.2 Protocol

73

4. 4.04 SSC/0.1% SDS array washing solution: [Make 4.04 SSC from 20x SSC solution by dilution first. Mix together 2,970 mL of 4.04 SSC and 30 mL of 10% SDS.] 5. Stock solutions for post-hybridization array processing TNT buffer (20 L): 0.1 M Tris–HCl (pH7.6) 0.15 M NaCl 0.05% Tween-20. (a) Rinse a 25 L Carboy with 150 mL of isopropanol. (b) Rinse the carboy twice with 3 L of deionized H2O and completely drain the carboy. (c) Add 2 L 1 M Tris–HCl, 600 mL 5 M NaCl, 10 mL Tween-20, and 17.39 L deionized H2O. (d) Mix well by swirling. (e) Filter TNT through a 0.2 mm filter. This solution can be stored up to 2 weeks at room temperature. 6. 0.75 TNT buffer: Add 25 ml of deionized water to 75 mL of TNT buffer (from above) per 100 ml of buffer required. 7. TNB buffer (0.5 L): 0.1 M Tris–HCl (pH7.6) 0.15 M NaCl 0.5% NEN blocking reagent (Perkin Elmer FP1020) (a) Take 435 ml nuclease-free H2O; (b) add 50 mL of 1 M Tris–HCl (pH7.6); (c) add 15 mL of 5 M NaCl. (d) Slowly add 2.5 g of NEN Blocking reagent in 0.5 g increments until all 2.5 g of blocking reagent is dissolved, while warming in a water bath at 60 C. (e) Filter TNB buffer through a 0.88 mm filter. (f) Aliquot the TNB buffer to 50 ml tubes and store at 20 C. This solution can be stored for up to 12 weeks at 20 C. Thaw immediately before use.

2.2.4

Procedures

2.2.4.1

miRNA Microarray Fabrication

Design of miRNA Capture Oligonucleotide Probes 1. Collect the miRNA precursor sequence from the Sanger Database (http://microrna. sanger.ac.uk/cgi-bin/sequences/browse.pl) (Fig. 2.3). 2. Design two 40-mer miRNA capture oligonucleotide probes that are exactly antisense to the target miRNAs, one for the mature miRNA and the other for precursor miRNA, from the sense strand of both arms of the hairpin structure of the pre-miRNA. LNA-modification of probes is highly recommended to enhance the sensitivity and specificity of hybridization between the capture probes and target miRNAs (Castoldi et al. 2006, 2007). 3. Modify the oligo probes at the 50 end with Amine-C6 linker and order from Integrated DNA technology (IDT; Coralville, IA, USA) at 50 or 100 mM stock concentration in H2O.

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2 Microarray and Its Variants for miRNA Profiling Select a miRNA of interest

Design of miRNA capture oligonucleotide probes

Oligo probe library for miRNA array fabrication

miRNA array fabrication

Total RNA extraction

miRNA array post-printing processes

Synthesis of biotin-labeled first-strand cDNA targets

miRNA microarray hybridization

Scan chip to collect and analyze data

Fig. 2.3 Flowchart of microarray procedures for miRNA expression detection

Oligo Probe Library for miRNA Array Fabrication 1. Synthesize the oligos in a 96 well plate format from the vendor, and then assemble in house 4 96-well plates into 384 well microtiter plate format by Tecan TeMo liquid handler in order to work with the array fabrication system. 2. Prepare the working oligo library at a concentration of 20 mM in 50 mM sodium phosphate buffer (pH8.0) with 2 mM of sodium-fluorescein.

miRNA Array Fabrication 1. The Codelink-activated slides are loaded onto an Omnigrid 100 slide holder. 2. The 384 well oligo library plates are loaded sequentially onto an OmniGrid 100 arrayer and the miRNA arrays are printed with a designed printing protocol (Omnigrid user manual website).

2.2 Protocol

75

miRNA Array Post-Printing Processes 1. Quality control scanning. Scan the printed array slides by Axon Scanner 4200 at 488 nm excitation length in order to detect fluorescein, and save QC image files. 2. Oligo probe coupling. At 70% humidity overnight, the dispensed aminemodified oligo probes on the slides couple with NHS ester group of coated polymer gel matrix and are covalently immobilized on the surface of Codelinkactivated slides. 3. miRNA array blocking. The NHS ester groups surrounding the spots of oligo probe dispensed need to be blocked, by incubating in a shaking incubator, in a solution of 50 C prewarmed 100 mM bicine and 100 mM taurine, at pH9.0 and temperature of 50 C for 60 min. 4. Then rinse the blocked slides with deionized H2O and wash further in 50 C prewarmed 4 SSC/0.1% SDS buffer for 30 min with 50 rpm agitation. 5. Rinse the slides with H2O and spin dry them. 6. The slides are now ready to hybridize the labeled miRNA cDNA targets and should be stored in a desiccator until use.

2.2.4.2

Total RNA Extraction

1. Use TRIzol and TRI Reagent to purify total RNA samples of appropriate quality for small RNA detection. All steps of RNA extraction should be performed according to the manufacturer’s instructions. 2. Treat the RNA sample with DNase I to ensure that the samples will be practically free of genomic DNA and of satisfactory good quality. 3. Do not use minicolumn-based purification methods for RNA extraction as these may result in loss of the small RNA fraction. 4. It is important to assess the quality of the sample by analyzing 1–5 mL of the extracted RNA sample by electrophoresis in 1x TBE buffer using 1.2% agarose gels containing nuclease-free ethidium bromide (0.5–1 mg/mL final concentration).

2.2.4.3

Synthesis of Biotin-labeled First-strand cDNA Targets

1. 5 mg of each total RNA sample (optimal concentration should be determined for each source of total RNA). 2 mL 0.5 mg/ml primer X ll Nuclease-free H2O to 12 mL final volume 12 mL Total volume Add 5 mg of total RNA to 10 mL RNAse free H2O 2 mL of oligo primer (50 biotinAAA-AAA-AAA-AAA-(T-biotin)-AAA-AAA-AAA-AAA-NNN-NNN-NN 30 ) (0.5 mg/mL) (where N stands for random octamer).

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2 Microarray and Its Variants for miRNA Profiling

2. Incubate 10 min in a 70 C water bath and then immediately place the tube on ice. 3. Centrifuge for 5 s to collect the sample at the bottom of the tube and immediately place the tube on ice. 4. With the tube remaining on ice, add the following reagents to the 12 mL of total RNA/control mRNA/primer mix. 5. 4 mL 5 first-strand buffer. 2 mL 0.1 M DTT 1 mL 10 mM dNTP mix 1 mL Superscript1 II RNaseH1 reverse transcriptase (200 U/mL) 20 mL Final volume 6. Incubate 90 min in a 37 C water bath. 7. Centrifuge for 5 s to collect the sample at the bottom of the tube. 8. RNA template degradation. After incubating for 90 min for the first strand synthesis, (a) add 3.5 mL of 0.5 M NaOH/50 mM EDTA into 20 mL reaction mix and (b) incubate at 65 C for 15 min to denature the DNA/RNA hybrids and degrade the RNA template. (c) Then neutralize the reaction with 5 mL of 1 M Tris–HCI (pH 7.6). Each labeled target should be in a volume of 28.5 mL. 9. The sample preparation is now done and the samples should be stored at 80 C until use.

2.2.4.4

miRNA Microarray Hybridization and Data Collection

1. Prime all channels of the Tecan HS4800 hybridization stations and load hybridization chambers to the hybridization station 2. Load pre-printed miRNA array face-up to the hybridization station. Close hybridization chambers on the hybridization station 3. Prime the chip in the hybridization chamber at 23 C with 6 SSPE with 0.05% Tween 20 for 1 min 4. Inject 75 ml prehybridization mix of 6 SSPE/2 Dehardt/30% formamide and prehybridize the chip at 25 C for 30 min 5. Inject the hybridization mix of labeled biotin–cDNA n 6 SSPE/2 Dehardt/ 30% Formamide and hybridize at 25 C for 18 h 6. Wash in 0.75 TNT buffer at 23 C for 5 min 7. Wash in 0.75 TNT buffer at 37 C for 10 min 8. Water rinse on the hybridization station at 23 C for 30 s 9. Unload the chips from the machine for post-hybridization washing 10. Open the hybridization chamber of the hybridization station and remove the slide chips as quickly as possible and then place them into a slot of the Bioarray Rack, which was placed in the large Reagent Reservoir containing 37 C prewarmed 0.75 TNT 11. Move the slide into place using the Bioarray Position Tool, tooth-side down. Wash them in 37 C prewarmed 0.75 TNT with agitation in shaking incubator at 37 C for 40 min with 50 rpm agitation

2.3 Application and Limitation

77

12. Block the chip in TNB blocking buffer at room temperature for 30 min 13. Stain the chips with streptavidAlexa-647 1:500 in TNB buffer at room temperature for 30 min 14. Post-stain wash in 1 TNT at room temperature for 40 min in total with three buffer changes 15. Rinse the chips with distilled water briefly and spin dry them at 1,000 rpm for 1 min 16. Scan the chip by Axon Scanner at a power setting of Power 100 and PMT 800. The image data may be extracted by GenePix software (Fig. 2.3)

2.2.4.5

Experimental Design Guidelines to Prevent RNase Contamination

The following precautions are recommended to prevent RNase contamination when working with any technologies involving RNA handling. 1. Wear gloves at all times while handling reagents, materials, and equipment to prevent RNase contamination from hands. Change gloves after touching nonRNase-free surfaces. 2. Avoid using equipment and work areas that have been exposed to RNases. Clean the equipment and work surfaces with ethanol or commercially available Rnase decontamination solutions. 3. Clean the interior and exterior of micropipette shafts with ethanol or commercially available RNase decontamination solutions and use barrier tips. 4. Use RNase-free plastic ware and Rnasefree buffers and reagents. Guidelines for assay setup. Follow the guidelines (above) to prevent RNase contamination. Also, use the following guidelines for successful assay setup. 5. Thaw reagents on ice, mix thoroughly before use, and immediately return unused materials to –20 C. 6. When preparing working reagents, measure components accurately, mix thoroughly, spin briefly, and keep on ice. 7. Assemble reactions on ice or at indicated temperature throughout the procedure.

2.3

Application and Limitation

As the number of identified miRNA increases, microarrays appear to be an ideal method for profiling in a highly efficient parallel fashion for a large number of miRNAs in a single run. The use of locked nucleic acid probes allowed optimization of the hybridization conditions suitable for all miRNAs, producing an improved microarray platform that offered single base mismatch (SBM) discrimination capability with a detection limit of 500 fM. However, microarrays have encountered difficulties in reliably amplifying miRNAs without bias. Also, because of the extremely short nature of miRNAs, there is

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2 Microarray and Its Variants for miRNA Profiling

very little room to fine-tune the hybridization conditions for all miRNAs, which significantly discounts the sensitivity and specificity of microarrays. Another drawback of microarray technology is the lack of ability to provide quantitative data; the intensity of signals gives no indication of miRNA expression level in the sample and relative intensities of signals do not rank the level of miRNA expression either. Finally, a large quantity of RNA is required for testing; about 30 mg of total RNA is commonly used to hybridize an array (Krichevsky et al 2003).

References Castoldi M, Benes V, Hentze MW, Muckenthaler MU (2007) miChip: A microarray platform for expression profiling of microRNAs based on locked nucleic acid (LNA) oligonucleotide capture probes. Methods 43:146–152 Castoldi M, Schmidt S, Benes V, Noerholm M, Kulozik AE, Hentze MW, Muckenthaler MU (2006) A sensitive array for microRNA expression profiling (miChip) based on locked nucleic acids (LNA). RNA 12:913–920 Christensen U, Jacobsen N, Rajwanshi VK, Wengel J, Koch T (2001) Stopped-flow kinetics of locked nucleic acid (LNA)-oligonucleotide duplex formation: studies of LNA-DNA and DNADNA interactions. Biochem J 354:481–484 Dalma-Weiszhausz DD, Warrington J, Tanimoto EY, Miyada CG (2006) The affymetrix GeneChip platform: an overview. Methods Enzymol 410:3–28 Gershon D (2005) DNA microarrays: more than gene expression. Nature 437:1195–1198 Hughes TR, Hiley SL, Saltzman AL, Babak T, Blencowe BJ (2006) Microarray analysis of RNA processing and modification. Methods Enzymol 410:300–316 Kreil DP, Russell RR, Russell S (2006) Microarray oligonucleotide probes. Methods Enzymol 410:73–98 Krichevsky AM, King KS, Donahue CP, Khrapko K, Kosik KS (2003) A microRNA array reveals extensive regulation of microRNAs during brain development. RNA 9:1274–1281 Kumar R, Singh SK, Koshkin AA, Rajwanshi VK, Meldgaard M, Wengel J (1998) The first analogues of LNA (locked nucleic acids): phosphorothioate-LNA and 2’-thio-LNA. Bioorg Med Chem Lett 8:2219–2222 Lagos-Quintana M, Rauhut R, Yalcin A, Meyer J, Lendeckel W, Tuschl T (2002) Identification of tissue-specific microRNAs from mouse. Curr Biol 12:735–739 Liu CG, Spizzo R, Calin GA, Croce CM (2008) Expression profiling ofmicroRNA using oligo DNA arrays. Methods 44:22–30 McTigue PM, Peterson RJ, Kahn JD (2004) Sequencedependent thermodynamic parameters for locked nucleic acid (LNA)-DNA duplex formation, Biochemistry 43:5388–5405 Okamura K, Hagen JW, Duan H, Tyler DM, Lai EC (2007) The mirtron pathway generates microRNA-class regulatory RNAs in Drosophila. Cell 130:89–100 Petersen M, Wengel J (2003) LNA: a versatile tool for therapeutics and genomics. Trends Biotechnol 21:74–81 Rhoades MW, Reinhart BJ, Lim LP, Burge CB, Bartel B, Bartel DP (2002) Prediction of plant microRNA targets. Cell 110:513–520 Ruby JG, Jan C, Bartel DP (2007) Intronic microRNA precursors that bypass Drosha processing. Nature 448:83–86 Schena M, Shalon D, Davis RW, Brown PO (1995) Quantitative monitoring of gene expression patterns with a complementary DNA microarray. Science 270:467–70

References

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Vester B, Wengel J (2004) LNA (locked nucleic acid): high-affinity targeting of complementary RNA and DNA. Biochemistry 43:13233–13241 Wengel J (1999) Synthesis of 3’-C- and 4’-C-branched oligonucleotides and the development of locked nucleic acid (LNA). Acc Chem Res 32:301–310 Wolber PK, Collins PJ, Lucas AB, De Witte A, Shannon KW (2006) The Agilent in situsynthesized microarray platform. Methods Enzymol 410:28–57

Part III Northern Blotting Methods

Chapter 3

Northern Blotting and Its Variants for Detecting Expression and Analyzing Tissue Distribution of miRNAs

Abstract Northern blot analysis of small RNA species, such as miRNAs, is a widely used technique to assess accumulation levels of miRNAs of interest. This technique, combined with polyacrylamide gel electrophoresis, allows examination of expression properties of target miRNAs, determination of their sizes, and validation of predicted miRNAs. Northern blotting involves the use of electrophoresis to separate RNA samples by size and detection with a hybridization probe complementary to part of or the entire target sequence. When the first miRNAs were described, Northern blotting was used to detect these small RNAs. To date, the Northern blot remains the gold standard of miRNA expression profiling. However, there are several technical limitations that prevent researchers from using the Northern blot as a routine miRNA expression profiling tool. Northern Blotting using radioactive probes is very sensitive, but very time-consuming. Northern blotting is not practical in large clinical studies to detect the expression of hundreds of miRNAs and it also requires large amounts (5–25 mg) of total RNA from each sample. A modified version of the Northern blot using locked nucleicacid-modified oligonucleotides was developed by Valoczi (Nucleic Acids Res 32:e175, 2004). The sensitivity was improved tenfold as compared to conventional DNA probes. Ramkissoona et al. show that an RNA oligonucleotide labeled with a single DIG molecule can successfully be used to identify target miRNAs with equivalent sensitivity to isotope-labeled probes (Mol Cel Probes 20:1–4, 2006). Nonisotopic detection methods are generally faster, more convenient, and reduce radiation exposure to researchers. The ability to use nonisotopic methods and yet obtain sensitive and reliable results offers an advantage to investigators who prefer to avoid isotopes.

3.1

Introduction

The Northern blot is a technique used in molecular biology research to study gene expression by detection of RNA (or isolated mRNA) in a sample (Alberts Z. Wang and B. Yang, MicroRNA Expression Detection Methods, DOI 10.1007/978-3-642-04928-6_3, # Springer-Verlag Berlin Heidelberg 2010

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3 Northern Blotting and Its Variants

et al. 2008; Kevil et al. 1997). With Northern blotting it is possible to observe cellular control over structure and function by determining the particular gene expression levels during differentiation, morphogenesis, as well as abnormal or diseased conditions (Schlamp et al. 2008). The Northern blot analysis of small RNA species, such as miRNAs, is a widely used technique to assess accumulation levels of miRNAs of interest. This technique, combined with polyacrylamide gel electrophoresis, allows examination of expression properties of target miRNAs, determination of their sizes, and validation of predicted miRNAs (Fig. 3.1). Northern blotting involves the use of electrophoresis to separate RNA samples by size and detection with a hybridization probe complementary to part of or the entire target sequence. The term “Northern blot” actually refers specifically to the capillary transfer of RNA from the electrophoresis gel to the blotting membrane; however the entire process is commonly referred to as Northern blotting (Trayhurn 1996). The northern blot technique was developed in 1977 by James Alwine, David Kemp, and George Stark at Stanford University (Alwine et al. 1977). Northern blotting takes its name from its similarity to the first blotting technique, the Southern blot, named after biologist Edwin Southern (Alberts et al. 2008). The major difference is that RNA, rather than DNA, is analyzed in the northern blot (Bor 2006). A general blotting procedure (Trayhurn 1996) starts with the extraction of total RNA from a homogenized tissue sample. The mRNA can then be isolated through the use of oligo (dT) cellulose chromatography to maintain only those

Northern Blot 1. Collect cells or tissue and extract RNA with TRI Reagent (MRC)

5. Wash the membrane and expose with the film

2. Separate RNA on agarose gel

4. Hybridize denatured probe with the membrane

3a. Transfer separated RNA onto membrane

3b. Synthesize labeled cDNA probe Denature purified DNA fragment

Add Klenow DNA Polymerase, random primers and labeled nucleotides Klenow Fragment Klenow Fragment

Fig. 3.1 Schematic depiction of the Northern blot analysis in general

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RNAs with a poly (A) tail (Durand and Zukin 1993; Mori et al. 1991). RNA samples are then separated by gel electrophoresis. Since the gels are fragile and the probes are unable to enter the matrix, the RNA samples, now separated by size, are transferred to a nylon membrane through a capillary or vacuum blotting system. A nylon membrane with a positive charge is the most effective for use in Northern blotting since the negatively charged nucleic acids have a high affinity for them. The transfer buffer used for the blotting usually contains formamide because it lowers the annealing temperature of the probe-RNA interaction preventing RNA degradation by high temperatures (Yang et al. 1993). Once the RNA has been transferred to the membrane it is immobilized through covalent linkage to the membrane by UV light or heat. After a probe has been labeled, it is hybridized to the RNA on the membrane. Experimental conditions that can affect the efficiency and specificity of hybridization include ionic strength, viscosity, duplex length, mismatched base pairs, and base composition (Streit et al. 2008) The membrane is washed to ensure that the probe has bound specifically and to avoid background signals from arising. The hybrid signals are then detected by X-ray film and can be quantified by densitometry. When the first miRNAs were described, Northern blotting was used to detect these small RNAs. To date, the Northern blot remains the gold standard of miRNA expression profiling. However, there are several technical limitations that prevent researchers from using the Northern blot as a routine miRNA expression profiling tool. Northern blotting using radioactive probes is very sensitive, but is very timeconsuming. Northern blotting is not practical in large clinical studies to detect the expression of hundreds of miRNAs; it also requires large amounts (5–25 mg) of total RNA from each sample. A modified version of the Northern blot using locked nucleicacid-modified oligonucleotides was developed by Va´lo´czi et al. (2004).The sensitivity was improved tenfold as compared to conventional DNA probes. Ramkissoona et al. show that an RNA oligonucleotide labeled with a single DIG molecule can successfully be used to identify target miRNAs with equivalent sensitivity to isotope labeled probes (Ramkissoona et al. 2006). Nonisotopic detection methods are generally faster, more convenient, and reduce radiation exposure to researchers. The ability to use nonisotopic methods and yet obtain sensitive and reliable results offers an advantage to investigators who prefer to avoid isotopes.

3.2 3.2.1

Protocol Basic Protocols

The following protocols for Northern blot can be found in Protocol Online. Your Lab’s Reference Book (Contributed by Zuyuan Qian) [http://www.protocol-online. org/prot/Molecular_Biology/RNA/Northern_Blotting/] (see Fig. 3.2).

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3 Northern Blotting and Its Variants Total RNA extraction

Agarose/formaldehyde gel electrophoresis

Transfer of RNA from gel to membrane

Prepare membrane for hybridization

Design and synthesize probes

miRNA/Probe Hybridization

Removal of non-specific, background signals

Data analysis

Fig. 3.2 Flowchart of Northern blotting analysis for miRNA expression detection. (According to Zuyuan Qian) [http://www.protocol-online.org/prot/Molecular_Biology/RNA/Northern_Blotting/]

3.2.1.1 1. 2. 3. 4. 5. 6. 7.

RNase-free dish Whatman 1 mm paper Whatman 3 mm paper Nylon membrane (MSI) Glass pipets ProbeQuantTMG-50 micro columns (Amersham Pharmacia Biotech) Kodak XAR film

3.2.1.2 1. 2. 3. 4. 5. 6.

Materials

Instruments

Vortex Centrifuge Microcentrifuge Eppendorf model 5415C UV transilluminator X-ray intensifying screens

3.2 Protocol

3.2.1.3 1. 2. 3. 4.

87

Reagents

E running buffer Formaldehyde 20x SSPE Redivue [32P] dCTP

3.2.1.4

Procedures

Agarose/Formaldehyde Gel Electrophoresis (1) Prepare gel: (a) Dissolve 0.75 g agarose in 36 mL water and cool to 60 C in a water bath. (b) Place in a fume hood and add 5 mL of 10 E running buffer and 9 mL formaldehyde. (c) Pour the gel and allow it to set. (d) Remove the comb, place the gel in the gel tank, and add sufficient 1 E running buffer to cover to a depth of ~1 mm. (2) Prepare sample: (a) Adjust the volume of each RNA sample to 6 ml with water, then add 2.5 6 mL freshly prepared sample denaturation mix. (b) Mix by vortexing, microcentrifuge briefly to collect liquid, and incubate 15 min at 55 C. (c) Cool on ice for 2 min, then add 2 mL loading dye mix. (3) Run gel: Run the gel in 1 E running buffer at 100 V for 10 min, then at 65 V for 90 min. Transfer of RNA from Gel to Membrane 1. Prepare gel for transfer: Place the gel in an RNase-free dish and rinse with changes sufficient deionized water to cover the gel for 4 20 min. 2. Transfer RNA from gel to membrane: (a) Fill the glass dish with enough 20 SSPE. (b) Cut two pieces of Whatman 1 mm paper, place it on the glass plate, and wet it with 20 SSPE. (c) Place the gel on the filter paper and squeeze out air bubble by rolling a glass pipet. (d) Cut four strips of plastic wrap and place over the edges of the gel. (e) Cut a piece of nylon membrane just large enough to cover the gel and wetted in water. Place the wetted membrane on the surface of the gel. Try to avoid getting air bubbles under the membrane. (f) Flood the surface of the membrane with 20 SSPE. Cut five sheets of Whatman 3 mm paper to the same size as membrane and place on top of the membrane. (g) Put paper towels on top of the Whatman 3 mm paper to a height of ~6 cm, and add a weight to hold everything in place. (h) Leave overnight.

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Prepare Membrane for Hybridization 1. Remove paper towels and filter papers and recover the membrane and flattened gel. 2. Mark in pencil the position of the wells on the membrane and ensure that the up– down and back–front orientation are recognizable. 3. Rinse the membrane in 5 SSPE, then place it on a sheet of Whatman 3 mm paper and allow to dry. 4. Place RNA-side-down on a UV transilluminator (254 nm wavelength), and irradiate for an appropriate length of time.

Hybridization Analysis 1. Prepare DNA or RNA probe (>108dpm/mg): Probes are labeled with Ridiprimer DNA labeling system (Amersham LIFE SCIENCE), according to the manufacturer’s protocols: (a) Dilute the DNA to be labeled to a concentration of 2.5–25 ng in 45 mL of sterile water. (b) Denature the DNA sample by heating to 95–100 C for 5 min in a boiling water bath. (c) Centrifuge briefly to bring the contents to the bottom of the tube, and place on ice for 10 min. (d) Add the denatured DNA to the labeling mix and reconstitute the mix by gently flicking the tube until the blue color is evenly distributed. (e) Add 5 mL of Redivue [32P] dCTP and mix by gently pipetting up and down. Centrifuge briefly to bring the contents to the bottom of the tube and then incubate at 37 C for 30 min. (f) The probe is purified using ProbeQuantTMG-50 micro columns. To prepare G-50 micro columns, resuspend the resin in the column by vortexing, loosen the cap one-fourth turn and snap off the bottom closure. (g) Place the column in a 1.5 mL screw-cap microcentrifuge tube for support, then pre-spin the column for 1 min at 3,000 rpm in an Eppendorf model 5415C. (h) Place the column in a new 1.5 mL tube and slowly apply 50 ml of the sample to the top–center of the resin, being careful not to disturb the resin bed. (i) Spin the column at 3,000 for 2 min. The purified sample is collected at the bottom of the support tube. 2. Hybridization (a) Prehybridization. Wet the membrane in the 5 SSPE and place it RNAside-up in a hybridization tube and add 5 ml prehybridization solution, then place the tube in the hybridization oven and incubate with rotation 6 h at 42 C for DNA probe or 60 C for RNA probe.

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(b) Hybridization. Denature double-stranded probe by heating in a water bath for 10 min at 100 C and then transfer to ice. Pipet the desired volume of probe into the hybridization tube and continue to incubate with rotation overnight at 42 C for DNA probe or 60 C for RNA probe. 3. Autoradiography (a) The membrane is washed twice for 5–10 min with wash-buffer at room temperature, and twice for 15 min at 65 C with prewarmed (65 C) washbuffer. (b) Remove final wash solution and rinse membrane in 5 SSPE at room temperature. Blot excess liquid and cover in UV-transparent plastic wrap. Do not allow membrane to dry out if it is to be reprobed. (c) Blot is exposed at 80 C using Kodak XAR film and X-ray intensifying screens.

3.2.2

Protocols with Improved Sensitivity

3.2.2.1

Materials

1. Denaturing polyacrylamide (19:1) gel

3.2.2.2

Instruments

1. Vertical gel system (protean II Bio-Rad system with gel plates 16 20 cm; Bio-Rad). 2. Semi-dry electroblotter (SciPlas).

3.2.2.3 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12.

Reagents

TRI-reagent (Sigma) Diethylpyrocarbonate (DEPC; Sigma) Deionized formamide (Sigma) Acrylamide/bis (19:1) (Sigma) Urea (Fluka) Tetramethylethylenediamine (Sigma) Ammonium persulfate (Sigma) MOPS (Roche) Ethidium bromide (EtBr; Sigma) Nylon membrane (Hybond NX; Amersham/Pharmacia) 3 MM Whatman chromatography paper (Schleicher & Schuell) Decade RNA markers (Ambion)

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3 Northern Blotting and Its Variants

13. g[32P] ATP (Amersham/Pharmacia) 14. 1-Methylimidazole (Sigma-Aldrich) 15. 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide (EDC: Sigma)

DEPC-Treated Water 1. 2. 3. 4. 5.

Add 1 mL DEPC to 1,000 ml distilled water Shake vigorously to mix Incubate in fume hood, at room temperature (20 C) overnight Autoclave to inactivate residual DEPC and cool before use DEPC-treated water can be stored at room temperature

50 MOPS–NaOH (pH7.0) 1. Prepare 1 M MOPS in distilled water (pH7.0) with NaOH 2. Store at 4 C

dPAG Denaturing polyacrylamide (19:1) gel prepared with 1. 10–15% acrylamide 2. 7 M urea 3. Buffer with 20 mM MOPS–NaOH (pH7.0)

g[32P] ATP-labeled Decade RNA Markers Follow manufacturer’s (Ambion) instructions for labeling.

6 Loading Dye 1. Dissolve 0.25% (wt/vol) bromophenol blue in 30% (vol/vol) glycerol 2. Store at 20 C for later use.

EtBr Working Solution 0.5 mg/ml 1. Prepare fresh, dilute stock (10 mg/mL) with 20 mM MOPS–NaOH (pH7.0) to use at 0.5 mg/mL.

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91

EDC Cross-Linking Solution 1. Add 245 mL of 12.5 M 1-methylimidazole to 9 mL DEPC-treated water 2. Adjust pH to 8.0 with 1 M HCl (usually requires B300 mL). This can be prepared 1–2 h before use and kept at room temperature 3. Immediately before use, add 0.753 g EDC and make the volume up to 24 mL with DEPC-treated water. This gives a working solution of 0.16 M EDC in 0.13 M 1-methylimidazole at pH8.0 and is sufficient to saturate 320 cm2 (20 16 cm2) of 3 MM Whatman paper.

3.2.3

Protocols with Improved Specificity and Sensitivity

As already described in the previous chapter, LNA monomers are bicyclic, highaffinity RNA analogs having a modified ribose moiety (Kauppinen et al. 2006). In LNAs the furanose ring in the sugar–phosphate backbone is chemically locked in an N-type (C30 -endo) conformation by the introduction of a 20 -O,40 -C methylene bridge. This chemical modification induces a conformational change that brings about enhanced base stacking and phosphate backbone pre-organization (Petersen and Wengel 2003). Since LNAs possess similar water solubility as DNAs or RNAs, LNA-modified oligonucleotides can be used easily in many biological experimental applications. Moreover, the introduction of LNA residues results in increased stability against endo- and 30 -exonucleases (Frieden et al. 2003). The superior properties of LNA-modified oligonucleotides make them an ideal tool for detecting small RNA molecules. In Northern blot hybridization assays, LNA-modified oligonucleotide probes detected mature miRNAs by at least a tenfold higher efficiency than traditional DNA probes (Va´lo´czi et al. 2004). Moreover, LNA-modified oligonucleotide probes were highly specific, as demonstrated by the use of different single- and double-mismatched LNA probe molecules. The enhanced efficiency, stability, and discriminatory power of LNA-modified oligonucleotide probes render them especially useful in miRNA Northern blotting (Va´rallyay et al. 2007). The experimental procedures described below are based primarily upon the study reported by blotting Va´rallyay et al. (2007). 3.2.3.1

Materials

1. Vertical gel 2. Quick Spin Column (Roche) 3.2.3.2

Instruments

1. Gel Penguin Electrophoresis System 2. Stratalinker UV Crosslinker (Stratagene)

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3.2.3.3 1. 2. 3. 4.

5. 6. 7.

8. 9. 10. 11. 12. 13. 14. 15. 16.

17. 18.

Reagents

TRIZOL (Invitrogen) or TRI Reagent (Sigma) 10 TBE buffer: [0.9 M Tris, 0.9 M boric acid, 0.02 M EDTA, pH8.0] Penguin electrophoresis system or equivalent 12% acrylamide solution (can be adjusted between 8 and 15%): [24 mL 40% acrylamide/bis (19/1) solution (Ambion), 8 mL 10 TBE, 40 g urea, and water to 80 mL] TEMED (Sigma) Ammonium persulfate (APS): [Prepare 10% solution in water (immediately freeze in aliquots for single use at 20 C) Small RNA loading dye: [10 mL deionized formamide, 200 mL 0.5 M EDTA (pH8.0), add xylene cyanol and bromphenol blue to obtain a faint blue solution. Avoid a high concentration of dye since it may interfere with the correct separation of small RNA species] FDE: [10 mL deionised formamide, 200 mL 0.5 MEDTA (pH8.0), 10 mg xylene cyanol, 10 mg bromphenol blue] Ethidium bromide solution: 10 mg/mL in water Nytran N membrane (Schleicher & Schuell, Germany) or Hybond-N+ membrane (Amersham Pharmacia Biotech) 20 SSC: [3 M NaCl, 0.3 M sodium citrate-2H2O, pH7.0] Synthetic RNAs for RNA size markers LNA-modified oligonucleotides for probes (Exiqon, Denmark) T4 polynucleotide kinase (Fermentas, Lithuania) [g-32P]ATP or equivalent Hybridization solution: [50% deionized formamide, 0.5% SDS, 5· SSPE, 5 Denhardt’s solution (50 stock from Invitrogen), and 20 mg/mL sheared, denatured salmon sperm DNA] RNase A stock solution 10 mg/mL RNase A buffer: [0.5 M NaCl, 10 mM Tris, pH 7.5, 1 mM EDTA, 20 mg/mL RNase A]

3.2.3.4

Procedures

RNA Extraction 1. Use TRIzol and TRI Reagent to purify total RNA samples of appropriate quality for small RNA detection. All steps of RNA extraction should be performed according to the manufacturer’s instructions. 2. Treat the RNA sample with DNase I to ensure that the samples will be practically free of genomic DNA and satisfactory good quality. 3. Do not use minicolumn-based purification methods for RNA extraction since these may result in loss of the small RNA fraction.

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4. It is important to assess the quality of the sample by analyzing 1–5 mL of the extracted RNA sample by electrophoresis in 1x TBE buffer using 1.2% agarose gels containing nuclease-free ethidium bromide (0.5–1 mg/mL final concentration).

Denaturing PAGE 1. Use the Gel Penguin Electrophoresis System, a 20 20 cm glass plate gel sandwich with 1.5 mm spacers. Make sure that the apparatus is cleaned with detergent and rinsed thoroughly with autoclaved, sterile, distilled water. 2. After setting up the gel apparatus using 1.5 mm spacers, prepare 80 mL acrylamide solution by adjusting acrylamide content (about 8–15%) of the gel depending on the required resolution. 3. Warm up the solution carefully by using a microwave oven or water bath to dissolve the urea. 4. Cool the solution to room temperature and bring up to 80 mL with water. Add 480 mL 10% APS and 32 mL TEMED and pour the gel, allowing it to polymerize at least for 1 h. 5. Remove the comb very carefully to maintain intact wells, and assemble the gel sandwich on the apparatus. 6. Rinse wells thoroughly with 1 TBE using a syringe and a needle. No leaks should be observed. The gel must be pre-run at 400 V (40 mA) for 60 min. Make sure that during the pre-run the gel warms up because this guarantees proper denaturing conditions. 7. Before loading the samples on the warm gel, rinse the wells again.

Sample Preparation, Gel Electrophoresis and Capillary Gel Transfer 1. Since the advantage of LNA modified oligonucleotide probes is their enhanced sensitivity, the amount of RNA samples used should be adjusted accordingly. Quantities ranging from 1 to 100 mg total RNA can be loaded into the wells. This might be determined by the expected abundance of the small RNA species of interest. Generally, further purification of the small RNA fraction is not necessary. However, if the target miRNA is poorly expressed, the small RNA fraction can be separated from higher molecular weight RNA species and concentrated. The most reliable way to achieve small RNA purification is to purchase a small RNA purification system (for example, the flashPAGETM Fractionator (Ambion)). 2. Add the adjusted amount of RNA sample in 20 mL water to 20 mL small RNA loading dye. It is possible to reduce the volume of the samples to about 10 mL (5 mL RNA sample and 5 mL small RNA loading dye) depending on the amount of samples and size of the wells. The reduced sample volume may result in sharper bands.

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3 Northern Blotting and Its Variants

3. Denature samples at 65 C for 20 min, chill them on ice for 1 min, and spin in a microcentrifuge for 15 s at high speed. 4. Load samples and markers into wells of the warm gel. Also, 10 mL FDE into each well not used for samples; FDE contains a higher concentration of dye which helps to monitor progress of the electrophoresis. 5. Run the gel at 200 V while the entire sample volumes enter into the gel, and then run at 400 V until the bromophenol blue reaches the bottom of the gel. 6. Dismantle the apparatus and submerge the gel in 500 mL 1 TBE containing 0.5 mg/mL ethidium bromide to stain tRNAs and 5S RNA for loading control. 7. Gently shake for 10 min and rinse the gel in 1 TBE for 5–10 min to wash away excess ethidium bromide. 8. Perform the washing step in 1 TBE even if quantification by ethidium bromide staining is not required. An alternative method to check the uniformity of loading is stripping the membrane and rehybridizing with probes detecting, for example, 5S rRNA. 9. Transfer of RNAs onto the membrane is performed by capillary blotting subsequently to soaking the gel in 20 SSC for 10 min following routine protocols. Alternatively, the transfer can be performed using a semi-dry transfer cell (e.g., Bio-Rad) according to the manufacturer’s instructions. 10. After blotting, rinse the membrane with 2 SSC for 2 min and allow it to dry. Using a pencil, mark the positions of lanes on the RNA sample side of the membrane. 11. Fix RNA onto the membrane by ultraviolet cross-linking (with the RNA sample side facing up); for example, use the Stratalinker UV Crosslinker (Stratagene) according to the manufacturer’s instructions.

Preparation of Radiolabeled, LNA-modified Oligonucleotide Probes for miRNA Detection 1. LNA-modified oligonucleotide (Exiqon, Denmark) complementary to the target miRNA is labeled by combining 10 pmol of LNA-modified oligonucleotide with 1 mL 10 T4 polynucleotide kinase buffer and then bringing the reaction volume to 8.5 mL with water. (Exiqon has a website for probe design; see http://lnatools.com.) 2. Then, 0.5 mL T4 polynucleotide kinase and 1 mL [g-32P]ATP (0.4 MBq) are added in this order, and the reaction is incubated at 37 C for 60 min. 3. 40 mL of 1 TE is added to the reaction and the efficiency of labeling can then be estimated by spot test, which is performed by dispensing 0.5 mL labeled probe onto a marked piece of membrane. 4. The membrane is UV cross-linked after allowing the spot to dry. The membrane is then washed in 0.1 SC, 0.1% SDS at 65 C or 15 min, and exposed either to X-ray film or a phosphorimager cassette to check labeling. The probe can also be checked by removal of unincorporated, free nucleotides using spin columns, followed by checking the quantity of incorporated radiolabeled nucleotides by

3.2 Protocol

95

scintillation counting. However, for hybridization, it is not necessary to remove unincorporated free nucleotide.

Preparation of Radiolabeled RNA Markers 1. For small RNA Northern blotting, it is advisable to run marker RNAs adjacent to experimental RNA samples in order to correctly assess sizes of the target small RNA species. Synthetic RNAs of different lengths (for example; 18, 21 and 25 nucleotides in length) can be radiolabeled for this purpose by using T4 polynucleotide kinase. 2. For radiolabeled markers, remove unincorporated nucleotides by filtration through a Quick Spin Column. 3. Dilute the marker to 100 mL and load 1 mL onto a denaturing polyacrylamide gel and/or measure radioactivity to determine the optimal amount of radiolabeled marker. Alternatively, RNA marker ladders can be purchased and used according to the manufacturer’s instructions (for example DecadeTM Markers; Ambion).

Hybridization of LNA-modified Oligonucleotide Probes 1. Use hybridization solutions containing formamide for small RNA Northern blot analyses. The UV-fixed membrane is prehybridized at the temperature of hybridization (see below) for 30 min. 2. Heat the labeled LNA probe for 1 min at 95 C and cool it on ice before addition to the hybridization solution. It is recommended to replace the prehybridization solution at the time of probe addition. 3. Select the optimal temperature for hybridization - 50–60 C. 4. Normally, 1-h hybridization is sufficient for efficient detection of the target miRNA. While a 2-h hybridization resulted in a slightly stronger signal, further extending the hybridization period did not improve signal intensity. 5. Wash membranes twice in 2 SSC, 0.1% SDS at the temperature of hybridization for 10 min. If using more than one probe in an experiment, wash the membranes in separate vessels since the extremely effective hybridization of LNA-modified probes may result in cross-hybridization in the wash solutions. 6. Membranes may be exposed to film to check signals; however, make sure that membranes remain damp in case additional washes are required. LNA-modified oligonucleotide probes allow the application of stringent wash conditions (Va´lo´czi et al. 2004). The stringency of wash conditions can be adjusted by gradually decreasing the salt concentration of the buffer and/or increasing the temperature. For high stringency washing of membranes hybridized with LNA-modified oligonucleotide probes, use 0.1 SSC, 0.1% SDS at 65 C for 5 min.

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Removal of Nonspecific, Background Signals In some cases, use of LNA-modified oligonucleotide probes may result in appearance of undesired background hybridization signals. Increasing hybridization temperature can be an effective step to reduce background hybridization. Application of highly stringent wash conditions can also be useful to eliminate background noise. If background hybridization signals stubbornly remain, RNase A digestion can provide an alternative method. 1. This is done by incubating the membrane, subsequently to hybridization, in 50–100 mL RNase A buffer at 37 C for 30 min with gentle shaking. This will remove imperfect duplexes formed during hybridization. The RNase Adigested membrane will become non-reusable. 2. Wash the membrane again for 5 min after RNase A treatment. While these techniques may reduce or eliminate background hybridization, they may lower specific signal intensity and thus require prolonged exposure times for detection.

Stripping Northern Membranes Membranes incubated with LNA-modified oligonucleotide probes can be stripped efficiently for reuse. Never allow membranes to dry after hybridization. 1. Place the membrane in a boiling solution of 0.1% SDS, 5 mM EDTA for 5–10 min. Avoid wrinkling the filter and do not let it dry out before complete stripping of the blot. 2. Expose the membrane for signal detection to ensure that the probe has been stripped completely. Abundant miRNAs require longer treatment times in boiling buffer. However, fresh, nonstripped Northern blots are best for detecting miRNAs.

3.2.3.5

Advantages

1. The technique presented here allows sensitive detection of miRNAs by Northern blot which can be extremely beneficial when small amounts of samples are available, expression levels of target miRNAs are low, or subtle discrimination of related miRNAs (differing only by a few nucleotides) is necessary. 2. Commercial availability of LNA-modified probes (Exiqon, http://www.exiqon. com) and their chemical properties make them highly important and effective tools in many aspects of molecular biology. 3. The higher cost of LNA-modified oligonucleotide probes compared to traditional DNA oligonucleotides can be compensated by the reduced time and effort required for experiments, as well as the higher chance for success. As the LNA-based technologies become increasingly popular and widely used, costs associated with this technology will decrease significantly.

3.2 Protocol

3.2.4

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Protocols with Nonisotopic Detection

Northern blot analysis using radioisotopes is often inconvenient due to many constraints associated with the use of radioisotopes and in many instances not feasible at institutions, which strictly limit the use of isotopes. As an alternative, digoxigenin (DIG)-labeling system has several advantages compared to radioactive labeling techniques: high sensitivity, short exposure times, longer shelf life, and increased safety. DNA oligonucleotides, DNA, and RNA can be efficiently labeled with DIG, and have been used in multiple applications, including Northern blot, Southern blot, dot blot, in situ hybridization, arrays, and ELISA, for detecting genomic, viral, or mRNA targets (Chevalier et al. 1997; Parkin et al. 2005; McCabe et al. 1997). RNA probes labeled with DIG are generally produced by two methods: in vitro transcription method using a linearized plasmid DNA or a PCR product as a template which carries a suitable promoter added during amplification and direct chemical addition of DIG to RNA using total RNA and poly(A)+ mRNA. Kajigaya and colleagues set up a nonisotopic Northern analysis method for miRNA detection using 30 -DIG-labeled RNA oligo probes (Ramkissoona et al. 2006). They applied this method to miRNA or total RNA fractions extracted from human leukemic cell lines using either 32P- or DIG-labeled RNA probe for miR181, miR-155 or miR-16. They were able to establish that the use of DIG-labeled RNA probes is equally sensitive compared to 32P-labeled probes in detecting miRNA quantities as low as 50 ng. The protocols described in this section are primarily the same as Ramkissoona’s study (Ramkissoona et al. 2006).

3.2.4.1

Materials

1. 15% TBE-Urea gels (Invitrogen, Carlsbad, CA) 2. BrightStar-Plus positively charged nylon membranes (Ambion) 3. Kodak Biomax MR film

3.2.4.2

Instruments

1. Power supply for Northern blot gel

3.2.4.3 1. 2. 3. 4.

Reagents

mirVana miRNA Isolation Kit (Ambion, Austin, TX) Gel Loading Buffer II (Ambion) Ultrahyb-Ultrasensitive Hybridization Buffer (Ambion) DIG Luminescent Detection Kit (Roche, Penzberg, Germany)

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5. Anti-DIG, alkaline phosphatase conjugated antibody 6. Chemiluminescent substrate CSPD

3.2.4.4

Procedures

Extraction of miRNA and Total RNA 1. Isolate miRNAs and total RNAs from cell lines using the mirVana miRNA Isolation Kit and selectively precipitate enriched miRNA or total RNA fractions using differing amounts of ethanol, followed by column purification according to the manufacturer’s protocol.

Probes 1. Select miRNAs based on published sequences listed in the miRNA Registry (http://www.sanger.ac.uk/Software/Rfam/mirna/). 2. miRNA probe sequences are designed to be exactly antisense to the selected miRNAs. 3. Synthesize the 30 -DIG-labeled RNA oligonucleotides probes using the services provided by Proligo Primer & Probes (Proligo, Boulder, Co). RNA oligonucleotides are synthesized with 30 terminal C7 amine moieties, through which DIG is conjugated in a post-synthesis conjugation reaction. 4. The probes should be synthesized at HPLC grade or perform HPLC purification on your own to yield 30 -DIG-labeled RNA oligo probes.

Northern Blot Analysis 1. Add synthetic miRNAs (1,000, 500, 200, 100 and 50 ng) or total RNAs (5.0, 2.5, and 1.0 mg) into Gel Loading Buffer II and heat at 95 C for 3 min. 2. Load the samples onto denaturing 15% TBE-Urea gels. 3. Transfer onto BrightStar-Plus positively charged nylon membranes. 4. Prehybridize at 65 C for 1 h using Ultrahyb-Ultrasensitive Hybridization Buffer. 5. Incubate the blots with 30 -DIG-labeled RNA probe (100 ng/mL) at room temperature. 6. Perform probe detection using the DIG Luminescent Detection Kit according to manufacturer’s protocol. In brief, incubate blots in blocking solution for 30 min and then in antibody solution (anti-DIG, alkaline phosphatase conjugated antibody) for 30 min, followed by washing twice in washing buffer. After equilibration in detection buffer, incubate the blots with chemiluminescent substrate CSPD and exposed to Kodak Biomax MR Film.

References

3.3

99

Application and Limitation

Traditionally, DIG-labeled RNA probes are several hundred bases in length and are prepared using in vitro transcription or PCR to incorporate DIG-dUTP nucleotides. However, Ramkissoona et al. show that an RNA oligonucleotide labeled with a single DIG molecule can be used successfully to identify target miRNAs with equivalent sensitivity to isotope labeled probes (Ramkissoona et al. 2006). This finding suggests that 30 -DIG-labeled RNA oligonucleotide probe hybridization to miRNA molecules is not significantly affected by the presence of a linked DIG molecule. An added benefit of DIG-labeled probes is that detection is possible by a chemiluminescent reaction, which provides a bright signal in a short time period. In order to detect weak 32 P signals, exposure times may take several days; DIG probes, while capable of luminescing for days, provide results in minutes to hours. Nonisotopic detection methods are generally faster, more convenient, and reduce radiation exposure to researchers. The ability to use nonisotopic methods and yet obtain sensitive and reliable results offers an advantage to investigators who prefer to avoid isotopes. In summary, Northern blotting is considered the “gold standard” of miRNA detection, but it is very time consuming and requires large amounts of RNA samples. It is unlikely to be used as a routine method for diagnostic purposes.

References Alberts B, Johnson A, Lewis J, Raff M, Roberts K, Walter P (2008) Molecular Biology of the Cell, 5th ed. Garland Science, Taylor & Francis Group, NY, pp 538–539 Alwine JC, Kemp DJ, Stark GR (1977) Method for detection of specific RNAs in agarose gels by transfer to diazobenzyloxymethyl-paper and hybridization with DNA probes. Proc Natl Acad Sci USA 74:5350–5354 Bor (2006) Northern Blot analysis of mRNA from mammalian polyribosomes. Nature Protoc doi:10.1038/nprot.2006.216 Chevalier J, Yi J, Michel O, Tang XM (1997) Biotin and Digoxigenin as labels for light and electron microscopy in situ hybridization probes: where to we stand? J Histochem Cytochem 45:481–491 Durand GM, Zukin RS (1993) Developmental Regulation of mRNAs Encoding Rat Brain Kainate/ AMPA Receptors: A Northern Analysis Study. J Neurochem 61:2239–2246 Frieden M, Hansen HF, Koch T (2003) Nuclease stability of LNA oligonucleotides and LNADNA chimeras. Nucleosides Nucleotides Nucleic Acids 22:1041–1043 Kauppinen S, Vester B, Wengel J (2006) Locked nucleic acid: high-affinity targeting of complementary RNA for RNomics. Handb Exp Pharmacol 173:405–422 Kevil CG, Walsh L, Laroux FS, Kalogeris T, Grisham MB, Alexander JS (1997) An Improved, Rapid Northern Protocol. Biochem Biophys Research Comm 238:277–279 McCabe MS, Power JB, de Laat AM, Davey MR (1997) Detection of single copy genes in DNA from transgenic plants by nonradioactive Southern blot analysis. Mol Biotech 7:79–84 Mori H, Takeda-Yoshikawa Y, Hara-Nishimura I, Nishimura M (1991) Pumpkin malate synthase Cloning and sequencing of the cDNA and Northern blot analysis. Eur J Biochem 197:331–336 Streit S, Michalski CW, Erkan M, Kleef J, Friess H (2008) Northern blot analysis for detection of RNA in pancreatic cancer cells and tissues. Nature Protocols 4:37–43

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Parkin RK, Boeckh MJ, Erard V, Huang ML, Myerson D (2005) Specific delineation of BK polyomavirus in kidney tissue with a digoxigenin lableled DNA probe. Molec Cell Probes19:87–92 Petersen M, Wengel J (2003) LNA: a versatile tool for therapeutics and genomics. Trends Biotechnol 21:74–81 Ramkissoona SH, Mainwaring LA, Sloand EM, Young NS, Kajigaya S (2006) Nonisotopic detection of microRNA using digoxigenin labeled RNA probes. Mol Cel Probes 20:1–4 Schlamp K, Weinmann A, Krupp M, Maass T, Galle PR, Teufel A (2008) BlotBase: A northern blot database. Gene 427:47–50 Trayhurn P (1996) Northern Blotting. Pro Nutrition Soc 55:583–589 Va´lo´czi A, Hornyik C, Varga N, Burgya´n J, Kauppinen S, Havelda Z (2004) Sensitive and specific detection of microRNAs by northern blot analysis using LNA-modified oligonucleotide probes. Nucleic Acids Res 32:e175 Va´rallyay E´, Burgya´ J, Havelda Z (2007) Detection of microRNAs by Northern blot analyses using LNA probes. Methods 43:140–145 Yang H, McLeese J, Weisbart M, Dionne J-L, Lemaire I, Aubin RA (1993) Simplified high throughput protocol for Northern hybridization. Nucleic Acids Res 21:3337–3338

Part IV In Situ Hybridization Methods

Chapter 4

In Situ Hybridization and Its Variants for Detecting Expression and Analyzing Cellular Distribution of miRNAs

Abstract In situ hybridization is an important tool for analyzing gene expression and developing hypotheses about gene functions. Normal hybridization requires the isolation of DNA or RNA, separating it on a gel, blotting it onto nitrocellulose, and probing it with a complementary sequence. For hybridization histochemistry, sample cells and tissues are usually treated to fix the target transcripts in place and increase access of the probe. As noted above, the probe is either a labeled complementary DNA or, now most commonly, a complementary RNA. The probe hybridizes to the target sequence at elevated temperature, and then the excess probe is washed away. Solution parameters such as temperature, salt, and/or detergent concentration can be manipulated to remove any non-identical interactions. The probe that was labeled with either radio-, fluorescent- or antigen-labeled bases is then localized and quantitated in the tissue using autoradiography, fluorescence microscopy, or immunohistochemistry, respectively. ISH can also use two or more probes, labeled with radioactivity or the other non-radioactive labels, to simultaneously detect two or more transcripts. Alenius and colleagues have applied LNAmodified miRNA capture oligo probe to ISH to enhance the detection sensitivity and specificity, compared to normal DNA probes (RNA 12:1161–1167, 2006; Nature Protocols 2:1508–1514, 2007). LNA-modified oligonucleotide probes display markedly increased hybridization affinities toward complementary RNAs compared to traditional RNA or DNA based probes (Biochemistry 43:13233– 13241, 2004). This property allows for very stringent hybridization conditions, increasing the specificity and sensitivity of miRNA detection (Nucleic Acids Res 32:e175, 2004; Biochemistry 43:13233–13241, 2004). Most recently, Schmittgen and colleagues from Ohio State University Medical Center (Columbus, OH, USA) (Methods 46:115–126, 2009) have developed a novel method for in situ detection of the mature miRNA, based on the extension of the molecule after its hybridization, to an ultramer template in formalin-fixed, paraffin-embedded tissues. The major advantages of the ultramer extension method over the LNA probe method is that it is specific to the mature, active form of the miRNA, is easier to optimize with a broader window between signal and background, is less sensitive to changes in stringency, and is much less expensive. Z. Wang and B. Yang, MicroRNA Expression Detection Methods, DOI 10.1007/978-3-642-04928-6_4, # Springer-Verlag Berlin Heidelberg 2010

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4.1

Introduction

In situ hybridization, as the name suggests, is a type of hybridization that uses a labeled complementary DNA or RNA sequence to localize a specific DNA or RNA sequence in morphologically preserved tissues sections or cell preparations by hybridizing the complementary strand of a nucleotide probe to the sequence of interest. This is distinct from immunohistochemistry, which localizes proteins in tissue sections. DNA ISH can be used to determine the structure of chromosomes. Fluorescent DNA ISH (FISH) can, for example, be used in medical diagnostics to assess chromosomal integrity (Fig. 4.1). RNA ISH (hybridization histochemistry) is used to measure and localize mRNAs and other transcripts within tissue sections or whole mounts. The basic principles for in situ hybridization are the same, except that one is utilizing the probe to detect specific nucleotide sequences within cells and tissues. The sensitivity of the technique is such that threshold levels of detection are in the region of 10–20 copies of mRNA per cell. Normal hybridization requires the isolation of DNA or RNA, separating it on a gel, blotting it onto nitrocellulose and probing it with a complementary sequence. For hybridization histochemistry, sample cells and tissues are usually treated to fix the target transcripts in place and increase access of the probe. As noted above, the probe is either a labeled complementary DNA or, now most commonly, a complementary RNA (riboprobe). The probe hybridizes to the target sequence at elevated

FISH

Nick Translation

Gene

Fixed Cells (on slide)

DNase (random cuts)

DIG-dUTP (or Biotin-dUTP) dATP + dCTP + dGTP

Denatruation (75oC)

Denatruation (Formamide 42oC)

Hybridization (on slide)

+ Epifluorescent Microscopy

Antibodies anti-Dig (or Avidin) linked with a fluoropfore

Fig. 4.1 Schematic depiction of fluorescent in situ hybridization in general

4.1 Introduction

105

temperature, and then the excess probe is washed away (after prior hydrolysis using RNase in the case of unhybridized, excess RNA probe). Solution parameters such as temperature, salt, and/or detergent concentration can be manipulated to remove any non-identical interactions (i.e., only exact sequence matches will remain bound). Then, the probe that was labeled with either radio-, fluorescent-, or antigen-labeled bases (e.g., digoxigenin) is localized and quantitated in the tissue using either autoradiography, fluorescence microscopy, or immunohistochemistry, respectively. ISH can also use two or more probes, labeled with radioactivity or the other non-radioactive labels, to simultaneously detect two or more transcripts. For several years after the discovery that large numbers of miRNAs were present in animals, a major limitation for understanding miRNA function was the difficulty in determining detailed spatial expression patterns for specific miRNAs. However, two approaches for detection of miRNAs by ISH have been developed recently. Plasterk and colleagues have used an ISH method based on Locked Nucleic Acid (LNA) oligonucleotide probes to detect mature miRNAs in whole mounts of zebrafish and mouse embryos (Kloosterman et al. 2006; Wienholds et al. 2005). The LNA-based ISH methodology has been applied to whole-mount embryos from a variety of vertebrate species, including chicken and medaka (Darnell et al. 2006; Ason et al. 2006), and for the analysis of adult human brain sections (Nelson et al. 2006) as well as mouse tissue sections (Ryan et al. 2006; Wulczyn et al. 2007). Independently, we have developed an miRNA ISH method based upon RNA oligonucleotide probes and have used this method for the detection of mature miRNAs in tissue sections from embryonic and adult mice (Deo et al. 2006), as well as from rat and human brains (this article) and zebrafish (our unpublished observations). We have also detected miRNAs by ISH in cultured cells (Deo et al. 2006). The LNA-based miRNA ISH method derives a high degree of sequence specificity from the base-pairing properties of LNA probes. In contrast, we have used high-stringency wash conditions based on tetramethylammonium chloride (TMAC) in combination with RNase –a A treatment to remove unhybridized probe to generate high sequence specific conditions for miRNA ISH with RNA probes. Both methods appear to generate similar results based on the comparison of published expression patterns. One potential advantage of RNA probe based ISH is that the use of TMAC washes allows probes of different sequence compositions (with potentially different melting temperatures) to be processed under identical conditions (Wood et al. 1985; Jacobs et al. 1988). Although miRNAs are generated by processing from longer precursor and primary transcripts that contain the same miRNA sequence, miRNA ISH using either RNA or LNA probes, appears to primarily detect mature miRNA (Wienholds et al. 2005; Deo et al. 2006). Most likely, the discrimination between mature miRNA and precursor or primary transcript RNAs reflects the accumulation of mature miRNAs to high levels in cells relative to mRNAs (Lim et al. 2003) and the location of the target miRNA sequences within a stem-loop secondary structure in miRNA precursors and primary transcripts, which seems likely to reduce access of probes for hybridization during ISH. It also seems possible that miRNAs basepaired with a target mRNA in RNP complexes are less accessible to hybridization

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106

during ISH, in which case miRNA ISH may preferentially detect mature miRNAs that are not bound to a target mRNA; but this possibility has not been tested. To date, most published studies on miRNA ISH have used probes labeled with nonradioactive haptens (digoxigenin or fluorescein). Bound hapten-labeled probes are detected by histochemical enzymatic reactions after application of alkaline phosphatase (AP)-conjugated anti-hapten antibodies. For RNA oligonucleotide probes, we have used fluorescein as a hapten, since it is convenient to commercially synthesize RNA oligonucleotides with a 50 end fluorescein. Nonradioactive ISH methods are rapid and permit the precise localization of mRNA or miRNA expression to specific cells or even cellular compartments, as well as facilitate whole-mount analyses, but these methods are less amenable to quantitative analysis of miRNA levels. However, ISH with radioactively labeled probes allows accurate semi-quantitative or quantitative comparisons of mRNA expression levels using autoradiographic methods (e.g., X-ray film, photographic emulsions, and phosphorimagers). Alenius and colleagues have applied LNA-modified miRNA capture oligo probe to ISH to enhance the detection sensitivity and specificity, compared to normal DNA probes (Obernosterer et al. 2006, 2007). LNA-modified oligonucleotide probes display markedly increased hybridization affinities towards complementary RNAs, compared to traditional RNA or DNA based probes (Vester and Wengel 2004). This property allows for very stringent hybridization conditions, increasing the specificity and sensitivity of miRNA detection (Va´lo´czi et al. 2004; Vester and Wengel 2004). Most recently, Schmittgen and colleagues from Ohio State University Medical Center (Columbus, OH, USA) (Nuovo et al. 2009) have developed a novel method for in situ detection of mature miRNA based on the extension of the molecule after its hybridization to an ultramer template in formalin-fixed, paraffin-embedded tissues. The major advantages of the ultramer extension method over the LNA probe method include the fact that it is specific to the mature, active form of the miRNA, is easier to optimize with a broader window between signal and background, is less sensitive to changes in stringency, and is much less expensive.

4.2

Protocol

4.2.1

Basic Protocol

4.2.1.1

Materials

1. 2. 3. 4. 5. 6.

Silane-coated glass slides (Fisher Scientific) Coverslips (Fisher Scientific) Protocol mounting media (Fisher Scientific) Secure-Seal hybridization chamber (RPI Corp) Coverslips or Lifterslips (Eric Scientific) Slides with Aqua-Poly/mount (Polysciences, Inc)

4.2 Protocol

4.2.1.2 1. 2. 3. 4. 5. 6.

107

Instruments

Incubator (Fisher Scientific) Hot plate (up to 100 1C) (Enzo Diagnostics) Microscope and digital camera Ventana Medical Systems Benchmark or Discovery-automated Immunohistochemistry and in situ hybridization system Humidified chamber

4.2.1.3

Reagents

Chemicals and Enzymes 1. Acetic anhydride (Sigma). 2. Sheep serum: Normal sheep serum (Jackson ImmunoResearch Laboratories Inc.). 3. Formalin, 10%, neutral-buffered (Fisher Scientific). 4. Diethylprocarbonate (DEPC) water (Fluka). 5. 30 -End tailing oligonucleotide kit (Enzo Diagnostics). 6. Biotin-modified dUTP (Enzo Diagnostics). 7. Streptavidin – alkaline phosphatase detection kit (Enzo Diagnostics; contains washes, alkaline phosphatase conjugate, NBT, BCIP chromogen, counterstain, proteinase K, and silane-coated glass slides). 8. 20 SSC (Ventana Medical Systems). 9. Antidigoxigenin alkaline phosphatase conjugate (Roche Diagnostics). 10. RNase-free DNase (Roche Diagnostics). 11. RNase inhibitor (Roche Diagnostics). 12. Bovine albumin (MP Biomedicals). 13. In situ hybridization buffer (Enzo Diagnostics). 14. Pepsin powder (Dako). 15. Digoxigenin-11-dUTP (Roche). 16. EZ rTth kit (Applied Biosystems). 17. Ultraview universal alkaline phosphatase red detection kit for immunohistochemistry (Ventana Medical Systems). 18. Proteinase K solution (Ventana Medical Systems). 19. Antibodies for immunohistochemistry. Preparation of Solutions 20. DEPC-treatment of solutions: [Where indicated, treat solutions with DEPC to inhibit RNases [27]. Add DEPC to 0.1% final concentration, mix, and leave overnight at room temperature in a fume hood. Autoclave solutions the next day. Note that solutions that contain Tris cannot be treated with DEPC. We set aside a stock of Tris for use only with RNA].

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21. PFA (4% paraformaldehyde in 1 PBS, 50 mL): [Microwave 47.5 mL of DEPC-treated water 30 s. Add 2 g PFA. Mix and heat with stirring. Add 50 mL of 10 N NaOH. Keep at 68 C with mixing until PFA dissolves completely. Add 40 mL of concentrated HCl. Add 2.5 mL 20 PBS (DEPC treated). pH should be ~7.4. Filter the solution using a 0.45 mm filter. This solution can be stored at 4 C for up to 7 days]. 22. 20 PBS (1 L): [Dissolve the following chemicals in 800 mL of water. Adjust the pH to 7.4 with HCl. Adjust the volume to one liter with water. Treat with DEPC [27]]. NaCl KCl Na2HPO4 KH2PO4

160.0 g 4.0 g 28.8 g 4.8 g

23. Proteinase K (PK) buffer: [50 mM Tris (pH7.5) (stock 1 M; cannot be DEPC treated), 5 mM EDTA (stock 0.5 M; DEPC treated)]. 24. Proteinase K (Roche): [Dilute 66 mL of 7.5 mg/mL stock in 50 mL of PK buffer just prior to use, for final concentration of 10 mg/mL]. 25. 1 M triethanolamine, pH8.0 (10 stock): [Add 66.5 ml triethanolamine to 413.5 mL DEPC-treated water then adjust pH with 20 ml HCl]. 26. Hybridization solution (50 mL): 50% formamide 5 SSC 0.3 mg/ML yeast RNA (ICN) 100 mg/mL heparin (Sigma) 1 Denhardt’s solution 0.1% Tween 20 0.1% CHAPS 5 mM EDTA 3 mL/mL DNA random primer

25 mL formamide 12.5 mL of 20 SSC (see below) 0.3 mL of 50 mg/mL stock in DEPC 100 mL of 50 mg/mL stock in DEPC 0.5 mL of 100 stock (see below) 0.5 mL of 10% Tween 20 in DEPC 0.5 mL of 10% CHAPS in DEPC 0.5 mL of 0.5 M EDTA (pH8.0) in DEPC 150 mL of stock (see below)

27. 20 SSC (1 L): [NaCl 175.3 g, Sodium citrate 88.2 g (pH7.0 with HCl). DEPC (0.1%) treat solution overnight in chemical hood. Autoclave solution next morning [27]]. 28. 100 Denhardt’s solution: [2% bovine serum albumin (ICN), 2% polyvinylpyrrolidone (PVP-40), 2% Ficoll 400; Make a slurry in DEPC-treated water and then dilute to final volume. Filter, sterilize, and store in aliquots at 20 C [27,28]]. 29. DNA random primers: [The inclusion of random 12-mer DNA oligonucleotides (customsynthesized by Invitrogen) in the hybridization solution reduces background, presumably by blocking nonspecific binding of probes. Resuspend oligonucleotides in DEPC-treated water at 0.1 nmole/mL and store in aliquots at 20 C]. 30. RNase A solution (10 mg/mL): [200 mL of 10 mg/mL RNase A stock (Roche) in 200 mL of 2 SSC solution (10 mg/mL)].

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109

31. 3 M TMAC solution (200 mL): [5 M TMAC 120 mL, 10% Tween 20 4 mL, 1 M Tris pH 8.0 10 mL; Add DEPC-treated water to 200 mL]. 32. PBT solution: [1 PBS with 0.1% Triton X-100]. 33. Anti-fluorescein antibody: [Anti-fluorescein-AP (Roche). Pre-absorb the antibody [28] before use to reduce background. To produce 1 mL pre-absorbed antibody add ~50 mL volume of mouse embryo powder to 0.5 mL 1 PBT and incubate at 70 C for 30 min. Vortex for 10 min. Cool mixture on ice. Add 5 mL sheep serum þ1 mL antibody. Mix for 1 h at 4 C. Centrifuge at 14,000 rpm for 4 min. Collect supernatant and dilute to 1 mL of 1 PBT/20% sheep serum, for a final antibody concentration of 1:1,000. Store at 4 C]. 34. AP buffer: 100 mM Tris (pH9.5) 50 mM MgCl2 100 mM NaCl 0.1% Tween 20 Add sterile water to final volume. The AP staining reaction should generate a purple precipitate on sections. If precipitate color is brown, check the pH of the AP buffer. 35. AP substrate solution (50 mL): Nitro-blue tetrazolium chloride (NBT, Roche): 37.5 mL of 100 mg/mL stock 5-Bromo-4-chloro-30 -indolyphosphate (BCIP, Roche): 350 mL of 50 mg/mL stock Dilute in 50 mL of AP buffer just prior to use and mix well. Final concentrations are NBT: 75 mg/mL and BCIP: 350 mg/mL. 36. Pepsin solution: [dilute 13 mg of the pepsin into 9.5 mL of DEPC-treated water and 0.5 mL of 0.2 N HCl. The solution can be frozen at 20 C in 1-mL aliquots and thawed immediately before use].

4.2.1.4

Procedures

The ISH protocols described in this chapter are essentially the same as reported by Thompson et al. (2007) (see Fig. 4.2).

Design of miRNA Capture Probes 1. Select miRNAs based on published sequences listed in the miRNA Registry (http://www.sanger.ac.uk/Software/Rfam/mirna/). 2. Design RNA oligonucleotide probes with 20 nt complementarity to a specific target miRNA. (Since the TMAC-based wash conditions are not sensitive to

110

4 In Situ Hybridization and Its Variants for Detecting Expression Select a miRNA of interest

Design of miRNA capture oligonucleotide probes

Preparation of RNA oligonucleotide probe

Gel purification of fluorescein-labeled RNA oligonucleotides

5' end labeling of RNA oligonucleotides with 33P

Preparation of tissue sections or cells for miRNA ISH

Pre-hybridization

Hybridization

Stringency washes

Immunohistochemistry

Fig. 4.2 Flowchart of fluorescent in situ hybridization (FISH) analysis for miRNA expression detection. According to Thompson et al. (2007)

sequence composition, the choice of 20 nt sequence within a miRNA longer than 20 nt to be used for probe generation appears to be arbitrary. Probes with 21 or 22 nt complementary to a target miRNA appears to function similarly (Deo et al. 2006). In case it is desirable to distinguish between closely related miRNAs, probes should be positioned on the target miRNAs so as to maximize the number of mismatches with non-target miRNA(s), so that potential duplexes between probe and non-target miRNA(s) will be as short as possible (to reduce crosshybridization). However, it is unlikely that any miRNA ISH procedure will be able to distinguish between miRNAs that differ by only a single nucleotide at one end (e.g., miR-128a and miR-128b)).

4.2 Protocol

111

Design of Control Probes 1. Several different types of control RNA oligonucleotide probes are possible. To confirm the sequence specificity of ISH, control probes can include mutations that create mismatches with the target miRNA at 1–3 internal positions (Kloosterman et al. 2006; Deo et al. 2006). It is important to avoid mismatch mutations that would allow a probe to hybridize to other members of a miRNA gene family unintentionally. 2. To detect probe trapping or other types of sequence-independent probe binding that can lead to elevated background, a reversed or scrambled sequence RNA probe can be used (after comparison with Genband to check for unanticipated targets). One limitation of reversed or scrambled sequence controls is that it is not easy to verify that they are functional probes, since they have no endogenous target. 3. An alternative is to use a probe that detects a known miRNA which is not expressed in the tissue of interest (or which is expressed in a distinct pattern within the tissue of interest). In this case, the integrity and function of the control probe can be confirmed by ISH using the appropriate tissue. When comparing ISH using control probes to ISH with probes for miRNAs of interest, it is important to use identical processing for all probes.

RNA Oligonucleotide Probe Preparation 1. RNA oligonucleotide probes can be custom synthesized commercially (Invitrogen, Dharmacon). For nonradioactive ISH, RNA oligonucleotides are synthesized with a 50 end fluorescein modification. 2. For radioactively labeled ISH, RNA oligonucleotides are custom synthesized without any 50 end modification (50 hydroxyl) and are enzymatically labeled at the 50 end with 33P using T4 DNA kinase. Note that 50 fluorescein-modified RNA oligonucleotides cannot be enzymatically labeled with T4 DNA kinase.

Gel Purification of Fluorescein-Labeled RNA Oligonucleotides 1. Load 20 mg of fluorescein-labeled RNA oligonucleotide onto an 18% polyacrylamide/TBE gel. 2. After electrophoresis, identify the RNA oligonucleotide band by fluorescence from the incorporated fluorescein (using a standard UV transilluminator). 3. Isolate the highest molecular weight band. Cut out RNA in smallest band possible and transfer to 100 mL of diethyl pyrocarbonate (DEPC)-treated 1 PBS. 4. Crush gel into as small pieces as possible and transfer to a microfuge tube. 5. Incubate overnight at 37 C. 6. Centrifuge at 11,000 g for 15 min and collect supernatant.

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4 In Situ Hybridization and Its Variants for Detecting Expression

7. To precipitate probe: add 1/10 volume DEPC-treated 3 MNaOAc, 2.5 volumes 100% ethanol, and store at 20 C for 1 h. Centrifuge at 11,000 g for 15 min. Remove ethanol and allow pellet to air dry. Pellet will be visibly yellow. Resuspend in 10 mL of RNase-free water and store at 20 C. 8. Integrity of the purified probe can be confirmed by gel elecrophoresis using a small amount of the purified probe on an 18% polyacrylamide/TBE gel (the unpurified probe can be used for size comparison). 50 End Labeling of RNA Oligonucleotides with 33P 1. Incubate 5–10 pmol of RNA oligonucleotide and ~100 mCi of 1,000 Ci/mmol 33 P-cATP with T4 DNA kinase according to the enzyme manufacturer’s recommendations (typically 30–45 min at 37 C). 2. 33P-labeled RNA oligonucleotides are purified from unincorporated 33P-cATP using P6 Bio-Rad Spin columns according to the manufacturer’s directions. Typical incorporation percentages are ~50%. Since incorporation rates are similar across different RNA oligonucleotides, use ~0.5–1.0 106 cpm of labeled RNA oligonucleotide per 40 ml of hybridization buffer.

Preparation of Tissue Sections for miRNA ISH 1. As with conventional mRNA ISH, tissue can be fixed at the time of collection by immersion or perfusion, or freshfrozen tissue sections can be used. Embryos or small pieces of tissue can be fixed by overnight immersion in 4% paraformaldehyde (PFA) in PBS at 4 C. It is better to fix large adult tissues such as brain by perfusion with PFA. 2. After fixation, tissues are rinsed in 1 PBS, dehydrated in 15% sucrose until the tissue sinks, and embedded in OCT (Fisher). 3. Tissue sections (12 mm) are cut using a cryostat and transferred to SuperFrost/ plus slides (Fisher). 4. Slides are stored at 20 C until ISH. 5. Alternately, for fresh-frozen tissue, tissues are collected as quickly as possible and immediately frozen in dry ice-chilled isopentane baths (35 C). Frozen tissues are then stored at 80 C until sectioning. Frozen tissue sections (10–14 mm thick) are thaw mounted onto clean, SuperFrost microscope slides and stored at 80 C until ISH.

Preparation of Cultured Cells for miRNA ISH 1. Cells are plated on poly-L-lysine and laminin-coated microscope slides. This coating helps retain cells during the ISH processing steps. Microscope slides are coated with poly-L-lysine (10 mg/mL) in 1 PBS for 3.5 h, followed by air

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drying in a laminar flow hood. Slides are subsequently rinsed three times with distilled water and stored at 4 C (for up to 2 weeks). 2. Prior to use, slides are coated with mouse laminin (2 mg/mL, Invitrogen) overnight, then washed with 1 PBS twice prior to plating cultured cells onto the slides in appropriate media. 3. At the desired time point, cells are washed with 1 D-PBS (Invitrogen) for 5 min and fixed in PFA for 20 min. 4. Fixed cells are rinsed three times with DEPC-treated 1 PBS and stored in DEPC-treated 1 PBS at 4 C until in situ hybridization (cells can be stored for a few days, but we have not systematically tested whether the signal deteriorates with storage). Process for miRNA ISH as described for tissue sections below, but omit proteinase K and RNase A as indicated. RNase contamination of reagents or slide processing labware prior to hybridization can compromise any RNA in situ hybridization procedure. To ensure that all glassware, reagent bottles, tubes, slide holders, etc., are RNase free, either purchase RNase-free products (sterile disposable tubes) or clean with commercial solutions designed to inactivate RNases (e.g., RNase Away, Ambion). Similarly, maintenance of an RNase-free work environment can be vital to the generation of maximal in situ hybridization results.

miRNA ISH Procedure All steps below should be performed at room temperature unless indicated otherwise. Day 1 1. For sections of immersion or perfusion fixed tissues. Slides are removed from storage at 20 C and air dried at 37 C for 30 min. Slides are then placed into 4% paraformaldehyde in PBS (made fresh) for 20 min in a fume hood. Continue at step (2). For sections of fresh-frozen tissues. Sections are removed from the 80 C freezer and immediately placed into 4% paraformaldehyde in PBS (made fresh) for 20 min at room temperature in a fume hood. High resolution autoradiographic images can be collected using photographic emulsions with potentially longer exposure times (again this must be empirically determined) followed by photographic development in standard photographic development solutions. 2. Wash slides twice in 1 PBS for 10 min each. 3. Treat the slides with 10 mg/mL proteinase K for 6 min at room temperature. This step is omitted when performing ISH on cultured cells. 4. Wash slides in 1 PBS for 10 min. 5. Fix slides in 4% paraformaldehyde in PBS for 15 min in fume hood. 6. Rinse slides in DEPC-treated water. 7. Acetic anhydride treatment: make 0.1 M TEA by diluting 20 mL 1 M Triethanolamine (TEA) (pH8.0) in 180 mL DEPC-treated water. In fume hood, add

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8. 9.

10.

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0.5 mL acetic anhydride to 0.1 M TEA and make certain that acetic anhydride is well dispersed. Immediately submerge slides in this solution for 5 min. Repeat this step with fresh dilution of acetic anhydride in TEA for 5 min. Wash slides in 1 PBS for 10 min. Pre-hybridization: for each slide, 200 mL of hybridization solution is added into a Secure-Seal hybridization chamber with the gasket side up. The slide is then carefully inverted onto the chamber such that the sections face the solution and the gasket forms a seal along the edge of the slide. Avoid bubbles. Alternately, it is possible to add pre-hybridization solution directly onto the slides (covering the sections) and then use conventional coverslips or Lifterslips. Incubate for 2.5 h at 37 C (in incubator or in a covered dish in a water bath). If coverslips/Lifterslips are used, the slides must be incubated in a sealed humidified chamber (cover bottom with filter paper soaked in 50% formamide and place slides on plastic supports above filter paper). Option 1: hybridization with fluorescein-labeled RNA oligonucleotide probes. Pre-warm hybridization solution with probe at 37 C for 10 min before applying to slides. Remove hybridization chamber and discard the pre-hybridization solution. Add 200 ll of hybridization solution containing 1 lg/ml RNA oligonucleotide probe and re-apply Secure-Seal hybridization chamber as described for pre-hybridization above. Continue at step (11). Option 2: hybridization with radioactively labeled RNA oligonucleotide probes. Pre-warm hybridization solution with probe at 37 C for 10 min before applying to slides. Remove gasket/coverslip and discard pre-hybridization solution. Dilute 0.5–1.0 106 cpm of 50 -33P-labeled RNA oligonucleotide probe into 40–50 mL of hybridization buffer (for two rodent brain sections, 40–50 mL of hybridization buffer is typically sufficient). Add solution over sections and apply coverslip/LifterSlips as described above. When radioactive probes are used, employ appropriate precautions for handling radioactive materials at this and subsequent steps. Hybridize overnight at 37 C in incubator or in a covered dish in a water bath (if coverslips/LifterSlips are used, slides must be incubated in a humidified chamber) Note: the precise hybridization temperature is important; higher hybridization temperatures may decrease signal substantially.

Day 2 1. Remove probe. Wash in 2 SSC for 15 min at 37 C. Used fluorescein-labeled probes can be saved and re-used. Store used probes at 20 C. 2. RNase A treatment: incubate slides in 10 mg/mL RNase A in 2 SSC for 30 min at 37 C. The RNase A treatment is not required for in situs on cultured cells. Note: the concentration of RNase A may need to be titrated depending on the batch and/or supplier. 3. Rinse in 2 SSC twice for 15 min each. 4. High-stringency TMAC washes: dilute 3 M TMAC þ 0.2% Tween 20 in 50 mM Tris (pH8.0) to make TMAC wash solution. TMAC is toxic: employ appropriate precautions when handling. Pre-warm the TMAC wash solution to

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6. 7. 8. 9.

10. 11.

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54 C prior to adding slides. Wash slides twice for 5 min each at 54 C, then wash slides once for 10 min at 54 C. These steps are performed in a dish in a shaking water bath. Note: the precise temperature and duration of these washes are critical, as is pre-warming the wash solution. Measure the bath temperature to confirm the temperature. Detection of radioactively labeled probes: after TMAC washes, rinse slides once in 2 SSC for 10 min, then dehydrate the slides through a graded series of 50–100% ethanol. Allow slides to air dry, then expose to X-ray film (BioMax MR Film, Kodak). Exposure times can vary from 18 h to 20 days, depending upon the relative abundance of each miRNA within tissue areas/ regions, and must be determined empirically. Detection of fluorescein-labeled probes: after TMAC washes, perform the additional washing and antibody-based detection steps starting at (7). Rinse slides twice in 1 PBT for 10 min each. Blocking: incubate slides in 1 PBT/20% sheep serum for 1 h to reduce nonspecific antibody binding. Add 200 mL of the pre-absorbed 1:1,000 diluted anti-fluorescein antibody in PBT/20% sheep serum to each slide. Use the Secure-Seal hybridization chambers as described for prehybridization. Incubate for 4 h at room temperature. Wash slides in 1 PBT for 10 min. Wash slides in 1 PBT overnight at 4 C.

Day 3 1. Wash slides in AP buffer twice for 5 min each. 2. Detection: remove last AP buffer wash and add AP substrate solution (make just prior to use). Signal on sections will be purple and may take 3–10 h (or more) to develop at room temperature, depending on the probe. Probes complementary to abundant miRNAs usually produce detectable signal within the first hour. When longer staining times are necessary, it may improve staining to change the AP substrate solution every 3–5 h (in addition, if the AP substrate solution becomes purple or pink it should be changed). Staining can be restarted by returning sections to the AP substrate solution. Staining is performed with the slides laid flat in plastic dishes, so that the formation of the AP precipitate can be monitored under a stereomicroscope. Minimize exposure of sections to bright light while staining; avoid shaking or mixing AP substrate solution while staining as this can inhibit precipitate formation. 3. After staining reaction is complete, wash slides twice in 1 PBS for 10 min each. 4. Fix slides in 4% paraformaldehyde for 10 min (or longer) to completely and permanently halt the AP staining reaction. Rinse in 1 PBS. 5. Slides can be stored without coverslips in 1 PBS at 4 C indefinitely. For analysis of staining, semipermanently mount coverslips on slides with AquaPoly/mount. Coverslips mounted with Aqua-Poly/mount can be removed from sections by soaking in water if necessary. Slides with mounted coverslips can be stored at room temperature.

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4.2.2

LNA-Modified Protocol with Enhanced Sensitivity and Specificity

Similar to the case of the Northern blot analysis of miRNA detection, LNAmodified miRNA capture oligo probe has also been applied to in situ hybridization to enhance detection sensitivity and specificity, compared to normal DNA probes. Alenius and colleagues from the Institute of Molecular Biotechnology of the Austrian Academy of Sciences (IMBA) and the Research Institute of Molecular Pathology (IMP), Vienna, Austria were the first to make such an application (Obernosterer et al. 2006, 2007). Each incorporated LNA monomer increases the melting temperature (Tm) of a DNA/RNA hybrid by 2–10 C. The increased thermal stability of the LNA-DNA/ RNA duplex makes it possible to make probes as short as ~20 nt with high melting temperatures (70 C), providing the basis for high stringency required for in situ hybridizations. Obernosterer et al designed the protocol for digoxigenin (DIG)conjugated LNA probes which can be used with two different alkaline phosphatase color reactions – nitro blue tetrazolium/5-bromo-4-chloro-3-indolyl phosphate (NBT/BCIP) and Fast Red – to enhance the signal strength. The NBT/BCIP colorimetric method generates a blue precipitate that can be detected with light microscopy and is highly sensitive. Fast Red is a less sensitive substrate but is fluorescent, which makes it suitable for double staining in conjunction with detection of proteins, cell typespecific markers, or putative target genes (Obernosterer et al. 2006, 2007). We have successfully used the protocol on sections of various tissues like brain, liver, and whole mouse embryos (Obernosterer et al. 2006).

4.2.2.1 1. 2. 3. 4. 5. 6.

Silane-coated glass slides (Fisher Scientific) Coverslips (Fisher Scientific) Protocol mounting media (Fisher Scientific) Secure-Seal hybridization chamber (RPI Corp) Coverslips or Lifterslips (Eric Scientific) Slides with Aqua-Poly/mount (Polysciences, Inc)

4.2.2.2 1. 2. 3. 4. 5. 6.

Materials

Instruments

Super Frost Slides (Henzel Glaeser, 51800AMNZ) Coverslips (Menzel-Glaeser, 24 60 mm) Hybridization oven (Biometra Compact Line OV04) Water bath (GFL, 1002) Glass beakers (Bake at 180 C to destroy RNases) Slide carousel (Bake at 180 C to destroy RNases)

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7. Slide mailers (Bake at 180 C to destroy RNases) 8. RNase-free chamber (Nunc Bio-Assay Dish, 245 245 25) 9. Hybridization chamber: add 3 MM paper soaked with 5 SSC/50% formamide to a Nunc Bio-Assay dish (245 245 25) to make a hybridization chamber 10. Humidified chamber: add 3MM paper soaked in tap water to a Nunc Bio-Assay Dish (245 245 25) to make a humidified chamber 11. Confocal microscope (Zeiss Axioplan-2) 12. Digital camera (Coolsnap HQ, Photometrics) 13. Imaging Software (Adobe Photoshop 7.0) 4.2.2.3

Reagents

Chemicals & enzymes 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 23. 24. 25. 26.

Sucrose (Fluka) Paraformaldehyde (PFA) (Sigma) Formamide (Riedel) Diethylpyrocarbonate (DEPC) (Fluka) Triethanolamine (Fluka) Conc. HCl (Riedel) Acetic anhydride (Sigma) 50 Denhardt’s (Sigma) Yeast tRNA (Sigma) Salmon sperm DNA (Sigma) Blocking reagent (Roche) Anti-digoxigenin antibody (Roche) Levamisol (Sigma) BCIP (Roche) NBT (Roche) Tissue Tek OTC (Sakura) Fast Red Substrate (Dako) 1 M Tris pH8.2 10% (w/w) CHAPS (Sigma) 10% and 20% (w/w) Tween (Fluka) 5 M NaCl (Merck) 1 M MgCl2 (Fluka) PBS 20 SSC Dry ice Terminal transferase reaction (Roche DIG High-Prime)

Preparation of solutions 1. DEPC-treated PBS (2 L): [Add 2 mL of DEPC to 2 L of PBS and autoclave]. 2. DEPC-treated water (1 L): [Add 1 mL of DEPC to 1 L of distilled water and autoclave].

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3. 4% PFA for fixation: [Add 4 g PFA to 100 mL boiling PBS þ 10 mL 10 N NaOH]. Shake until dissolved, chill on ice and store at 4 C. Use within 3 days of preparation. 4. Acetylation solution: [Prepare 590 mL of DEPC-treated water in a beaker. Add 8 mL of triethanolamine and 1,050 mL conc. HCl (min. 37%). Mix gently with a magnetic stirrer. Add 1.5 mL acetic anhydride; wait until it has fully dissolved]. 5. Hybridization solution 20 mL (final concentration given in parentheses): [10 mL formamide (50%), 5 mL 20 SSC (5), 2 mL 50 Denhardt’s (5), 250 mL 20 mg/mL yeast RNA (200 mg/mL), 1,000 mL 10 mg/mL salmon sperm DNA (500 mg/mL), 0.4 g Roche blocking reagents, and 1.75 mL DEPCtreated water. Can be stored for months at 20 C. Denature salmon sperm DNA at 96 C for 5 min before adding to the hybridization solution]. 6. Denaturating hybridization solution: [Prepare as hybridization solution but add 500 mL of 10% CHAPS, 100 mL of 20% Tween, and 1,150 mL (instead of 1.75 mL) of DEPC-treated water]. 7. 5 SSC/50% formamide: [In a 50 Lalcon tube mix 20 mL formamide, 10 mL 20 SSC, and 10 mL DEPC-treated water]. 8. Solution B1 (0.1 M Tris pH 7.5/0.15 M NaCl): [100 mL 1 M Tris pH7.5 and 30 mL 5 M NaCl. Make up to 1 L with sterile water]. 9. Blocking solution (20 mL): [2 mL FCS and 18 mL B1. For higher stringency, include 100 ml 10% Tween]. 10. Solution B3 (0.1 M Tris pH9.5/0.1 M NaCl/50 mM MgCl2; 1 L): [200 mL 0.5 M Tris pH9.5/0.5 M NaCl, 50 mL 1 M MgCl2, and milliQ water. Use a 0.45 mm filter (Nalgene) to filter the solution, otherwise the MgCl2 will precipitate]. 11. NBT/BCIP developer solution: [3.4 mL 100 mg/mL NBT, 3.5 mL 50 mg/mL BCIP, 2.4 mL 24 mg/mL levamisol, 5 mL of 10% Tween, and 986 mL B3] 12. Fast Red solution: [Add one tablet of Fast Red to 2 mL substrate buffer. Let it stand in the dark for 8 min. Vortex the solution until the tablet is completely dissolved. A 2 mL staining solution will be sufficient for 8–9 slides. Pour the solution into the supplied substrate container and place on a provided filter top and apply the Fast Red to the slides].

4.2.2.4

Procedures

LNA Probe Design 1. For mature miRNAs, design LNA probes complementary to the full ~19–23 nt long miRNA. Identify unpaired regions in the primary or pre-miRNA by using secondary structure prediction programs such as the RNAfold program [20]. According to Obernosterer et al. (2007), probes targeting regions of extensive base-pairing will not hybridize efficiently.) 2. Within those regions, design ~23 nt long probes with a GC content of ~40–60%.

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3. Apply local sequence alignment search tools (e.g., SWIFT, BLAST [21,22]) to scan the transcriptome for potential probe-binding sites. Discard probes with an overall sequence identity of more than 60% to other RNAs as these are likely to bind nonspecifically. 4. Used probes with ~30% LNA content and the position of LNA incorporations can be determined by the supplier (Exiqon). 5. Use DIG epitope and probes can either be ordered from Exiqon to be directly 30 - or 50 -DIG labeled or synthesized without a DIG label (Exiqon, SIGMAPROLIGO) and 30 -DIG nucleotides added afterwards by a terminal transferase reaction. (For the detection of low-expressedmiRNAs, probes with both 30 - and 50 -DIG labels might be used to enhance the detection signal). 6. Design a 30% LNA scrambled version of the probe as a negative control. Note that it is important to design this probe so that the sequence combination used does not bind within the transcriptome. Tissue Preparation 1. Dissect and pre-fix the tissue(s) of interest or embryos for 30 min up to 2 h in 4% PFA at room temperature (25 C). For detailed fixation times. After fixation, place the specimen in PBS with 30% sucrose, to minimize freeze fracturing of the tissue. 2. Incubate overnight at 4 C. Embedding 1. The next day, remove the specimen from the PBS/sucrose solution and place it in a freeze mold. Add tissue Tek OCT to cover the tissue and remove any air bubbles with a pipet tip. Try to arrange the tissue in an appropriate orientation, making use of the marks at the bottom of the freeze mold. Also try to place the tissue in the center and at the bottom of the mold; this is done to facilitate the sectioning step. 2. Place the block on top of dry ice and wait until the tissue Tek OCT becomes white and frozen. The frozen block can be stored at 80 C for at least a month. However, with increased freezing time, the in situ signal strength decreases and also the Tissue Tek OCT dries out, which directly affects sectioning. Sectioning and tissue fixation 1. Take out the freeze molds to be sectioned and place them in the cryostat and leave them there for at least 30 min to equilibrate to the sectioning temperature, which is normally around 15 to 20 C. 2. Section 10 mm thick slices in the cryostat and collect them on Superfrost PLUS slides. Remember to include one extra slide for the scrambled probe (negative

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control). Note that if the sections are thicker than 15 mm, longer proteinase K (PK) treatments might be needed. 3. Let slides dry at room temperature for at least 30 min. 4. Fix the dried slides in 4% PFA for 10 min at room temperature.

Acetylation of the Tissue 1. Wash slides twice for 3 min in 1 PBS in a slide mailer at room temperature. During the washing steps, prepare the acetylation solution. 2. Transfer the slides to a metal slide carousel submerged in 1 PBS in a beaker and wash for 3 min. During this step, add 1.5 ml acetic anhydride to the acetylation solution. 3. Immerse the slides in the beaker of acetylation solution and stir gently for 10 min. 4. After the acetylation step, transfer the slide carousel to a beaker with 1 PBS solution and wash for 5 min at room temperature without agitation.

PK Treatment 1. Transfer the slides from the beaker into a slide mailer containing 75 mL 1 PBS. Add 37.5 mL PK stock solution (10 mg/mL in DEPC-treated water, yielding a final concentration of 5 mg/mL) and perform PK treatment at room temperature for 5 min. 2. Wash the slides three times for 3 min in 1 PBS at room temperature. During the washing steps, prepare the hybridization chamber.

Pre-Hybridization 1. Place the slides horizontally in the hybridization chamber and add 700 mL of hybridization buffer to each slide. Incubate slides at room temperature for 4–8 h.

Hybridization 1. For each slide, prepare 150 mL denaturizing hybridization buffer and add 0.1 mL (1 pM) of the LNA DIG-labeled probe. 2. Denature probes by heating them up to 80 C for 5 min. Thereafter, quickly place them on ice. 3. Pipet the probe mix carefully onto the tissues and apply glass coverslips. Hybridize slides at 50–60 C overnight.

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Stringency Washes 1. Soak the slides in pre-warmed 60 C 5 SSC and carefully remove coverslips. Note that after the hybridization step, RNase-free conditions are no longer required. 2. Move the slides to a 0.2 SSC slide mailer. Incubate slides in 0.2 SSC at 60 C for 1 h. 3. Finally, incubate slides in B1 solution at room temperature for 10 min.

Immunohistochemistry 1. Prepare a humidified chamber by placing a tap water-soaked 3MM paper at the bottom of the chamber. 2. For each X slides, make X times 500 mL blocking solution. 3. Remove surplus B1 solution and place the slides horizontally in the chamber. Add 500 mL of blocking solution to each slide and leave them for 1 h at room temperature. 4. Dilute the anti-DIG-alkaline phosphatase antibody 1:2,000 in blocking solution (500 mL per slide) and pipet carefully onto the sections. If multiple labeling is being performed in order to detect proteins, then the appropriate primary antibodies should also be included. Incubate at 4 C overnight.

Color Reactions 1. This protocol can be used with different alkaline phosphatase substrates such as NBT/BCIP (option A) or the Fast Red Substrate (option B). NBT/BCIP produces a purple blue precipitate and supports long exposure times and a high sensitivity due to low background signals. Fast Red generates a red fluorescent precipitate that can be detected using standard fluorescent microscopy and is suitable for double staining with antibodies, thus providing the possibility to correlate the miRNA expression with proteins of choice. However, one drawback is that Fast Red generates a yellowish background after some hours of exposure and thus cannot be used to detect lowly expressed miRNAs. (A) NBT/BCIP Reaction 1. Place the slides into a slide mailer and wash three times for 5 min in B1 solution at room temperature. 2. Equilibrate slides for 10 min in B3 solution. 3. For each slide, prepare 150 mL of developer solution. 4. Place slides in a humidified chamber and pipet carefully the developer solution on the slides and place a parafilm or glass coverslip on top.

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5. Develop at room temperature in the dark ranging from 10 min up to 4 days, depending on the miRNA expression levels. The reaction can be monitored using a light microscope; terminate when a strong blue staining is observed. 6. Stop the color reaction by washing the slides for 3 10 min in 1 PBT. 7. Mount the slides in glycerol or any water-soluble mounting media. 8. Visualize the staining by using standard light microscopy. (B) Fast Red Reaction 1. Place the slides into a slide mailer and wash for 3–5 min in B1 solution. 2. If the fluorescent detection of a protein is not required, proceed directly to Step (iii). If fluorescent detection of a protein is required, incubate slides for 1 h at room temperature in the dark with an appropriate green fluorescent secondary antibody such as Alexa 488 (Molecular Probes) diluted 1:500 in blocking solution. 3. Place the slides horizontally in the chamber and equilibrate slides with 700 ml 1 M Tris pH 8.2 for 10 min at room temperature. 4. Add five drops of the Fast Red solution to each slide and place a glass coverslip on top. 5. Develop at room temperature in the dark for 30 min up to 5 h, depending on expression levels. A red precipitate might appear but if not, incubate until background or any unspecific staining on the glass is evident. 6. Stop the color reaction by washing 3 10 min in 1 PBT. 7. Mount the slides in VECTA-Shield or any similar water-based fluorescent mounting medium. If you want to visualize the nucleus, include DAPI in the mounting media15. 8. Use a fluorescence microscope to see the Fast Red staining, where it can be visualized with the Cy-3 filter.

4.2.3

Ultramer Extension Protocol with Reduced Stringency and Expense

As described above, the in situ detection of miRNAs has been assisted by the use of LNA–modified nucleotides (Martin et al. 2007; Nuovo 2008). However, one limitation with the use of LNA probes is their low signal strength. LNA in situ hybridizations on cultured cells have previously been performed with directly conjugated fluorescent dyes but could only detect abundant miRNAs (Thomsen et al. 2005). The LNA nucleotides are much more rigid in three-dimensional space, which results in a substantial increase in the Tm of the small LNA-modified oligonucleotide probe hybridized to its target miRNA (Christensen et al. 2001). It has been demonstrated by some that the LNA-modified probe can detect either the miRNA precursors or mature miRNA depending on the specific sequence in the pre-miRNA targeted (Obernosterer et al. 2006, 2007).

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Schmittgen and colleagues from Ohio State University Medical Center (Columbus, OH, USA) (Nuovo et al. 2009) have recently developed a novel method for the in situ detection of the mature miRNA based on the extension of the molecule after its hybridization to an ultramer template in formalin-fixed, paraffin-embedded tissues. This method, like RNA-primed, array-based, Klenow enzyme (RAKE) analysis, is based on the RNA molecule acting as a primer to initiate the reaction (Nelson et al. 2006). The method involves the labeled extension of miRNA hybridized to a ~100-nucleotide–long ultramer template containing the complementary sequence of the miRNA at its 30 terminus. Pretreatment of the tissue involves incubation with protease to expose the genomic DNA to DNase digestion, thereby eliminating the ultramer-independent DNA synthesis process inherent in paraffin-embedded tissue. By direct comparison with real-time reverse transcriptase (RT)–PCR, RT in situ PCR, and standard in situ hybridization using a LNA probe, it was evident that the ultramer extension method detects only the mature miRNA, is easier to optimize, results generally in a stronger signal, and is much less expensive than the LNA probe method currently used.

4.2.3.1 1. 2. 3. 4. 5. 6.

Silane-coated glass slides (Fisher Scientific) Coverslips (Fisher Scientific) Protocol mounting media (Fisher Scientific) Secure-Seal hybridization chamber (RPI Corp) Coverslips or Lifterslips (Eric Scientific) Slides with Aqua-Poly/mount (Polysciences, Inc)

4.2.3.2 1. 2. 3. 4. 5. 6.

Instruments

Incubator (Fisher Scientific) Hot plate (up to 100 1C) (Enzo Diagnostics) Microscope and digital camera Ventana Medical Systems Benchmark or Discovery-automated Immunohistochemistry and in situ hybridization system Humidified chamber

4.2.3.3 1. 2. 3. 4.

Materials

Reagents

g-P32 ATP T4 kinase rTth reverse transcriptase (Applied Biosystems, Foster City, CA, USA) rTth buffer (Applied Biosystems)

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5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16. 17.

Taq DNA polymerase (Promega, Madison, WI, USA) MMLV reverse transcriptase (Invitrogen, Carlsbad, CA, USA) MultiScribe reverse transcriptase (Applied Biosystems) T7 RNA polymerase (Epicenter, Madison, WI, USA) 12% polyacrylamide 6 M urea denaturing gel 10% buffered formalin Nitroblue tetrazolium (NBT; 300 mg/mL) Bromochloroindolyl phosphate (BCIP; 200 mg/mL) Nuclear fast red (Enzo Life Sciences, Farmingdale, NY) RNase-free DNase I (Roche Diagnostics, Indianapolis, IN, USA) Anti-digoxigenin antibody (75 U/100 mL; Roche Diagnostics) Pepsin (2 mg/mL)

4.2.3.4

Procedures

Solution Phase Extension of miRNAs 1. Custom synthesize mature miRNAs of interest by IDT (Coralville, IA, USA). 2. End-label the miRNA oligomers (1 mg each ) with g-P32 ATP and T4 kinase. The labeled miRNA oligomers will be used to prime miRNA ultramers, and the oligos are extended using a variety of enzymes including rTth reverse transcriptase, Taq DNA polymerase, MMLV reverse transcriptase, MultiScribe reverse transcriptase, and T7 RNA polymerase, according to each company’s protocol. 3. Verify the extension products on a 12% polyacrylamide, 6 M urea denaturing gel.

Cell Lines and Tissue Samples 1. Fix tissue preparations or cell samples immediately in 10% buffered formalin for 4–12 h at room temperature; buffered formalin is the optimal fixative for RNA in situ hybridization.

Ultramer Extension Method The ultramer is at least 100 nucleotides in size. At its 50 end, it consists of a series of four 20-nucleotide repeats (GACCCCTTAATGCGTCTAAA), ending at the 30 end with the complementary sequence of the miRNA of interest. A DNA or RNA polymerase will extend the miRNA using the ultramer as the template. As the miRNA is being extended, the newly synthesized DNA incorporates the reporter nucleotide digoxigenin. The repetitive sequence allows the use of a complementary oligonucleotide (TTTAGACGCATTAAGGGGTC) that can be added at the onset

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of the reaction that will inhibit the extension of the miRNA. This serves as an additional negative control to document the specificity of the reaction. The RT in situ PCR and ultramer extension methods below are optimized for formalin-fixed, paraffin-embedded tissue (Nuovo et al. 2009). Heating the paraffin to 65 C during the embedding process induces nicks and short gaps in the genomic DNA, whereas formalin fixation causes DNA- and RNA-protein crosslinks (Nuovo 1997). Any DNA polymerase will utilize these nicks and short gaps to synthesize DNA that will incorporate the reporter nucleotide, either through gap repair or – if the polymerase has 50 –30 exonuclease activity – via nick translation. This will result in a nonspecific nuclear signal as previously shown with in situ PCR in paraffin tissue sections (Lee et al. 2007, 2008; Nuovo 1997, 2008). In order to prevent this, it is necessary to degrade the genomic DNA by DNase pre-treatment to the point where it will no longer support this nonspecific DNA synthesis pathway. Efficient DNase pre-treatment requires the removal by protease digestion of the formalin-induced DNA-protein adducts in the genomic DNA, which may sterically interfere with DNase activity. Optimal protease digestion is defined by the generation of an intense nuclear signal in the extension/labeling reaction of a test tissue section or cell preparation, reflecting loss of the steric interference of polymerase activity by the DNA-protein adducts (Lee et al. 2007, 2008; Nuovo 1997, 2008). If the protease digestion is not sufficient, residual DNA-protein crosslinks will allow the persistence of sufficiently intact DNA not degraded by DNase, resulting in a nonspecific nuclear signal. If protease digestion is too strong, then the morphology of the tissue or cell preparation is lost and thus the results are not interpretable (Nuovo 1997). The key components of the ultramer extension method are as follows: 1. Protease digestion. Cells or deparaffinized tissue sections are digested in protease (2 mg/mL pepsin in RNase-free water) at room temperature. The optimal protease time is defined as the number of minutes of digestion that will elicit an intense nuclear-based signal in all cell types, in a test tissue section or cell preparation with direct incorporation of the reporter nucleotide in an extension/ labeling reaction. This signal is completely eliminated by overnight digestion with RNase-free DNase I (Nuovo 1997; Lee et al. 2007). 2. DNase digestion. After optimal pro tease digestion, ~200 U of RNase-free DNase I is applied to each tissue section or cell preparation and incubated for 15 h at 37 C (Nuovo 1997, 2008). The DNase is removed by successive washes in RNase-free water and 100% ethanol. 3. miRNA extension solution. The ultramer extension solution consists of the following: 10 mL rTth buffer, 1.6 mL of each of the four dNTPS (10 mM), 1.6 mL of 2% BSA, 12.4 mL of 10 mM mangenese acetate, 2 mL of the ultramer (500 pmol), 0.6 mL of 1 mM digoxigenin dUTP, 2 mL of rTth DNA polymerase, and 15 mL of RNase free water. 4. In situ miRNA extension. After DNase digestion, the tissue section or cell preparation is covered with the miRNA extension solution and then with a polypropylene coverslip and mineral oil to prevent evaporation. The extension

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reaction proceeds for 30–60 min at 60 C. The incorporated digoxigenin is detected with an alkaline phosphatase conjugated to an antidigoxigenin antibody (75 U/100 mL) diluted 1:150 with Tris buffer (pH7.0). As with the RT in situ PCR method, the chromogen is nitroblue tetrazolium (NBT; 300 mg/mL) and bromochloroindolyl phosphate (BCIP; 200 mg/mL) and the counterstain is nuclear fast red. An alternative method can be used to confirm the specificity of the ultramerextended miRNA. The ultramer extension can proceed as described above, but minus the reporter nucleotide. The ultramer can be end-tailed with biotin (Christensen et al. 2001) and used as a probe to detect the unlabeled extended miRNA using the method described below, under “In situ hybridization with LNADNA probes.” A successful reaction with direct incorporation of the reporter nucleotide is defined as a strong nuclear signal in the no-DNase control, and its complete elimination in the DNase control in which neither ultramer nor a scrambled ultramer was used. The latter negative control section should be on the same slide as the test section in order to assure uniformity of variables such as protease digestion. As with RT in situ PCR, these two results indicate optimal protease digestion and successful degradation of the genomic DNA to the point that it can no longer support detectable DNA synthesis. If a signal is seen in the DNase negative control (no ultramer), which is nuclear-based, the reaction must be redone with increased protease digestion time. If the tissue morphology has been destroyed and hence, there is no signal, the reaction must be redone with decreased protease digestion time (Christensen et al. 2001).

4.3

Application and Limitation

The major advantage of ISH for detecting miRNA expression is the ability to readily and precisely determine sites of miRNA expression, with resolution at the single cell level, or even at subcellular levels when using nonradioactive probes. The use of radioactively labeled probes should permit better quantitative assessments of changes in miRNA levels. The large numbers of animal miRNAs and their presence in various tissues suggest that miRNA expression analyses will continue to provide insights into numerous biological processes, including development, adult organ function, and pathologies such as cancer. LNA-modified oligonucleotide probes display markedly increased hybridization affinities toward complementary RNAs, compared to traditional RNA or DNA based probes (Vester and Wengel 2004). This property allows for very stringent hybridization conditions, increasing the specificity and sensitivity of miRNA detection (Va´lo´czi et al. 2004; Vester and Wengel 2004). The major advantages of the ultramer extension method over the LNA probe method are that it is specific for the mature, active form of the miRNA, is easier to

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optimize with a broader window between signal and background, is less sensitive to changes in stringency, and is much less expensive.

References Ason B, Darnell DK, Wittbrodt B, Berezikov E, Kloosterman WP, Wittbrodt J, Antin PB, Plasterk RH (2006) Differences in vertebrate microRNA expression. Proc Natl Acad Sci USA 103:14385–14389 Christensen U, Jacobsen N, Rajwanshi V.K, Wengel J, Koch T (2001) Stopped-flow kinetics of locked nucleic acid (LNA)-oligonucleotide duplex formation: studies of LNA-DNA and DNADNA interactions. Biochem J 354:481–484 Darnell DK, Kaur S, Stanislaw S, Konieczka JH, Yatskievych TA, Antin PB (2006) MicroRNA expression during chick embryo development. Dev Dyn 235:3156–3165 Deo M, Yu JY, Chung KH, Tippens M, Turner DL (2006) Detection of mammalian microRNA expression by in situ hybridization with RNA oligonucleotides. Dev Dyn 235:2538–2548 Jacobs KA, Rudersdorf R, Neill SD, Dougherty JP, Brown EL, Fritsch EF (1988) The thermal stability of oligonucleotide duplexes is sequence independent in tetraalkylammonium salt solutions: application to identifying recombinant DNA clones. Nucleic Acids Res 16: 4637–4650 Kloosterman WP, Wienholds E, de Bruijn E, Kauppinen S, Plasterk RH (2006) In situ detection of miRNAs in animal embryos using LNA-modified oligonucleotide probes. Nat Methods 3:27–29 Lee EJ, Back M, Gusev Y, Brackett DJ, Nuovo GJ, Schmittgen TD (2008) Systematic evaluation of microRNA processing patterns in tissues, cell lines, and tumors. RNA 14:35–42 Lee EJ, Gusev Y, Jiang J, Nuovo GJ, Lerner MR, Frankel WL, Morgan DL, Postier RG, et al. (2007) Expression profiling identifies distinct microRNA signature in pancreatic cancer. Int J Cancer 120:1046–1054 Lim LP, Lau NC, Weinstein EG, Abdelhakim A, Yekta S, Rhoades MW, Burge CB, Bartel DP. (2003) The microRNAs of Caenorhabditis elegans. Genes Dev 7:991–1008 Martin MM, Buckenberger JA, Jiang J, Malana GE, Nuovo GJ, Chotani M, Feldman DS, Schmittgen TD, Elton TS (2007) The human angiotensin II type 1 receptor þ 1166 A/C polymorphism attenuates miR-155 binding. J Biol Chem 282:24262–24269 Nelson PT, Baldwin DA, Kloosterman WP, Kauppinen S, Plasterk RH, Mourelatos Z (2006) RAKE and LNA-ISH reveal microRNA expression and localization in archival human brain. RNA 12:187–191 Nuovo GJ (1997) RT in situ PCR. In Nuovo GJ (Ed.) PCR in situ hybridization: Protocols and Applications, 3 rd edition. Lippincott Williams and Wilkins Press, Baltimore, MD. pp 271–333 Nuovo GJ (2008) In situ detection of precursor and mature microRNAs in paraffin embedded, formalin fixed tissues and cell preparations. Methods 44:39–46 Nuovo G, Lee EJ, Lawler S, Godlewski J, Schmittgen T (2009) In situ detection of mature microRNAs by labeled extension on ultramer templates. Methods 46:115–126 Obernosterer G, Leuschner PJ, Alenius M, Martinez J (2006) Post-transcriptional regulation of microRNA expression. RNA 12:1161–1167 Obernosterer G, Martinez J, Alenius M (2007) Locked nucleic acid-based in situ detection of microRNAs in mouse tissue sections. Nature Protocols 2:1508–1514 Ryan DG, Oliveira-Fernandes M, Lavker RM (2006) Mol Vis 12:1175–1184 Thompson RC, Deo M, Turner DL (2007) Analysis of microRNA expression by in situ hybridization with RNA oligonucleotide probes. Methods 43:153–161 Thomsen R, Nielsen PS, Jensen TH (2005) Dramatically improved RNA in situ hybridization signals using LNA-modified probes. RNA 11:1745–1748

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Va´lo´czi A, Hornyik C, Varga N, Burgya´n J, Kauppinen S, Havelda Z (2004) Sensitive and specific detection of microRNAs by northern blot analysis using LNA-modified oligonucleotide probes. Nucleic Acids Res 32:e175 Vester B, Wengel J (2004) LNA (locked nucleic acid): high-affinity targeting of complementary RNA and DNA. Biochemistry 43:13233–13241 Wienholds E, Kloosterman WP, Miska E, Alvarez-Saavedra E, Berezikov E, de Bruijn E, Horvitz HR, Kauppinen S, Plasterk RH (2005) MicroRNA expression in zebrafish embryonic development. Science 309:310–311 Wood WI, Gitschier J, Lasky LA, Lawn RM (1985) Base composition-independent hybridization in tetramethylammonium chloride: a method for oligonucleotide screening of highly complex gene libraries. Proc Natl Acad Sci USA 82:1585–1588 Wulczyn FG, Smirnova L, Rybak A, Brandt C, Kwidzinski E, Ninnemann O, Strehle M, Seiler A, Schumacher S, Nitsch R (2007) Post-transcriptional regulation of the let-7 microRNA during neural cell specification. FASEB J 21:415–426

Part V Real-Time RT-PCR Methods

Chapter 5

End-Point Stem-Loop Real-Time RT-PCR for miRNA Quantification

Abstract Due to its high level of sensitivity, accuracy, and practical ease, qRT-PCR is accepted as a powerful technique in comparative expression analysis in life sciences and medicine. Real-time PCR methods have been designed for miRNA detection (Nucleic Acids Res 32:e43, 2004; Nucleic Acids Res 33:e179, 2005; RNA 11:1737–1744, 2005), some capable of detecting miRNAs present in only picograms of total RNA. The first miRNA real-time PCR approach was based on detection and quantification of miRNA precursors by Schmittgen et al. from the College of Pharmacy, Ohio State University (Columbus, OH USA) (Methods 44:31–38, 2004; Nucleic Acids Res 33:5394–5403, 2006). Chen and colleagues from the Applied Biosystems (Foster City, CA USA) (Nucleic Acids Res 33:e179, 2005) have developed a quantitative stem-loop RT-PCR for the detection of mature miRNAs that is based on TaqMan assays. This assay combines the power of PCR for exquisite sensitivity, real-time monitoring for a large dynamic range and TaqMan assay reporters to increase specificity. The major innovation of the technique is the invention of the stem–loop RT primer that is used to hybridize to a miRNA molecule and then reverse transcribed with a MultiScribe reverse transcriptase. Following the hybridization during RT step, the RT products are quantified using conventional TaqMan PCR. According to Chen et al. (Nucleic Acids Res 33: e179, 2005), stem-loop RT primers are better than conventional ones in terms of RT efficiency and specificity. TaqMan miRNA assays are specific for mature miRNAs and discriminate among related miRNAs that differ by as little as one nucleotide. Furthermore, they are not affected by genomic DNA contamination. Precise quantification is achieved routinely with as little as 25 pg of total RNA for most miRNAs. In fact, the high sensitivity, specificity, and precision of this method allows for direct analysis of a single cell without nucleic acid purification. Like standard TaqMan gene expression assays, TaqMan miRNA assays exhibit a dynamic range of seven orders of magnitude. Quantification of five miRNAs in seven mouse tissues showed variation from less than 10 to more than 30,000 copies per cell. This method enables fast, accurate, and sensitive miRNA expression profiling and can identify and monitor potential biomarkers specific to tissues or diseases. Stem-loop RT-PCR can be used for the quantification of other small RNA Z. Wang and B. Yang, MicroRNA Expression Detection Methods, DOI 10.1007/978-3-642-04928-6_5, # Springer-Verlag Berlin Heidelberg 2010

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molecules such as short interfering RNAs (siRNAs). Furthermore, the concept of stem-loop RT primer design could be applied in small RNA cloning and multiplex assays for better specificity and efficiency.

5.1

Introduction

Current methods for detection and quantification of miRNAs are largely based on cloning, northern blotting, or primer extension. Although microarrays could improve the throughput of miRNA profiling, the method is relatively limited in terms of sensitivity and specificity (Krichevsky et al. 2003; Liu et al. 2004). Low sensitivity becomes a problem for miRNA quantification because it is difficult to amplify these short RNA targets. Furthermore, low specificity may lead to false positive signal from closely related miRNAs, precursors, and genomic sequences. More recently, a modified Invader assay has been reported for the quantification of several miRNAs (Allawi et al. 2004). However, Invader assays have limited specificity and sensitivity, requiring at least 50 ng total RNA, or 1,000 lysed cells, per assay. Real-time PCR is the gold standard for gene expression quantification (Livak and Schmittgen 2001; Heid et al. 1996) (Fig. 5.1). Due to its high level RT - Polymerase Chain Reaction (RT-PCR) 1. Collect cells or tissue and extract RNA with TRI Reagent (MRC)

2. Synthesize cDNA using total or polyA RNA, nucleotides, Reverse Transcriptase and dT19 oligonucleitide as a primer AMV RT

cDNA (template) mRNA

3. Add oligonucleotide primers specific for target mRNA, termstabile DNA polymerase and nucleotide triphosphates and start amplification process – One cycle shown

3a. Denature DNA template (95oC)

Repeat as many cycles as required

3b. Anneal oligonucleotide primers (40 – 65oC)

One cycle

3c. Synthesize second strand with Taq DNA polymerase (72oC) Taq DNA Polymerase Taq DNA Polymerase

Fig. 5.1 Schematic representation of the basic idea of real-time PCR-based miRNA expression profiling method

5.1 Introduction

133

of sensitivity, accuracy, and practical ease, qRT-PCR is accepted as being a powerful technique in comparative expression analysis in life sciences and medicine (Mocellin et al. 2003). However, it has been a long challenge for scientists to design a conventional PCR assay from miRNAs averaging ~22 nt in length. Real-time PCR methods have been designed for miRNA detection (Schmittgen et al. 2004, 2008; Chen et al. 2005; Raymond et al. 2005), some capable of detecting miRNAs present in only picograms of total RNA. The first miRNA real-time PCR approach was based on the detection and quantification of miRNA precursors (Schmittgen et al. 2004; Jiang et al. 2005). However, other studies have shown that the cellular steady-state level of miRNA precursors does not correspond to cellular concentrations of mature miRNAs (Lagos-Quintana et al. 2002; Raymond et al. 2005). Chen and colleagues from the Applied Biosystems (Foster City, CA USA) (Chen et al. 2005) have developed a quantitative stem-loop RT-PCR for the detection of mature miRNAs that is based on TaqMan assays. Their assay displays high sensitivity and dynamic range and allows for discrimination between members of miRNA families such as let-7. This and a few other assays have recently become commercialized and are rather costly. Raymond et al. (2005) have designed a SYBR Green real-time RT-PCR for the detection of mature miRNA molecules using LNAmodified primers. Most of their developed assays are described as being sensitive to femtomolar concentrations. However, the performance and sensitivity of 70% of the analyzed assays depend strictly on the use of LNA-modified primers, which increases the costs of this approach. Other PCR-based technologies have been published that include extending the size of the mature miRNA by primer extension or using poly A polymerase (Ro et al. 2006; Mano and Takada 2007; Takada and Mano 2007). Chen et al. developed a novel scheme to design TaqMan PCR assays that specifically quantify miRNA expression levels with superior performance over existing conventional detection methods. They have designed and validated assays for 222 human miRNAs (Chen et al. 2005). This assay combines the power of PCR for exquisite sensitivity, real-time monitoring for a large dynamic range, and TaqMan assay reporters to increase the specificity. The major innovation of the technique is the invention of the stem–loop RT primer that is used to hybridize to a miRNA molecule and then reverse transcribed with a MultiScribe reverse transcriptase (Fig. 5.2). Following the hybridization during RT step, the RT products are quantified using conventional TaqMan PCR. According to Chen et al. (2005) stemloop RT primers are better than conventional ones in terms of RT efficiency and specificity. TaqMan miRNA assays are specific to mature miRNAs and discriminate among related miRNAs that differ by as little as one nucleotide. Furthermore, they are not affected by genomic DNA contamination. Precise quantification is achieved routinely with as little as 25 pg of total RNA for most miRNAs. In fact, the high sensitivity, specificity, and precision of this method allows for direct analysis of a single cell without nucleic acid purification. Like standard TaqMan gene expression assays, TaqMan miRNA assays exhibit a dynamic range of seven orders of magnitude. Quantification of five miRNAs in seven mouse tissues showed variation from less than 10 to more than 30,000 copies per cell. This method enables fast, accurate, and sensitive miRNA expression profiling and can identify and

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5 End-Point Stem-Loop Real-Time RT-PCR for miRNA Quantification

Fig. 5.2 Schematic description of TaqMan miRNA assays, TaqMan-based real-time quantification of miRNAs includes two steps, stem–loop RT and realtime PCR. Stem–loop RT primers bind at its 50 portion to the 30 portion of miRNA molecules and are reverse transcribed with reverse transcriptase. Then, the RT product is quantified using conventional TaqMan PCR that includes miRNA-specific forward primer, reverse primer, and a dye-labeled TaqMan probes. The purpose of tailed forward primer at 50 is to increase its melting temperature (Tm) depending on the sequence composition of miRNA molecules. Modified from Chen et al. (2005)

monitor potential biomarkers specific to tissues or diseases. Stem-loop RT-PCR can be used for the quantification of other small RNA molecules such as short interfering RNAs (siRNAs). Furthermore, the concept of stem-loop RT primer design could be applied in small RNA cloning and multiplex assays for better specificity and efficiency.

5.2 5.2.1

Protocol Materials

1. TaqMan1 Gene Expression Assays for human or mouse glyceraldehyde-3phosphate dehydrogenase (GAPDH) endogenous controls (Applied Biosystems). 2. Stem–loop RT primers (Applied Biosystems).

5.2 Protocol

5.2.2 1. 2. 3. 4. 5.

135

Instruments

Thermal cycler (for pulsed reverse transcription and end-point PCRs) Real-time thermal cycler (for SYBR Green I and UPL probe assays) Centrifugation (Allegra 6, Beckman Coulter, Fullerton, CA) Applied Biosystems 9,700 Thermocycler or equivalent Applied Biosystems 7900HT Sequence Detection System (Applied Biosystems)

5.2.3

Reagents

1. Dulbecco’s phosphate-buffered saline (PBS) without MgCl2 and CaCl2 (Invitrogen, Carlsbad, CA) 2. Nucleic Acid Purification Lysis Solution (Applied Biosystems) 3. RNase inhibitor solution (Applied Biosystems) 4. mirVanaTM miRNA Isolation Kit (Ambion, Austin, TX) 5. mirVanaTM miRNA Detection Kit (Ambion) 6. Cyclone Storage Phosphor System (PerkinElmer, Boston, MA) 7. RT buffer (Applied Biosystems) 8. MultiScribe reverse transcriptase (Applied Biosystems) 9. RNase inhibitor (Applied Biosystems) 10. TaqMan1 PCR kit (Applied Biosystems) 11. TaqMan1 Universal PCR Master Mix (Applied Biosystems)

5.2.4

Procedures

The protocols described in this section are essentially the same as reported in the study of Chen et al. (2005) (Figs. 5.2 and 5.3). The principle of the TaqMan microRNA Assays is similar to conventional TaqMan RT-PCR ones. A major difference is the use of a stem-loop primer during the RT reaction. There are several advantages to using stem-loop RT primers. First, by annealing a short RT priming sequence to the 30 portion of the miRNA, it has better specificity for discriminating similar miRNAs. Second, its double-stranded stem structure inhibits hybridization of the RT primer to miRNA precursors and other long RNAs. Third, the base stacking of the stem enhances the stability of miRNA and DNA heteroduplexes, improving the RT efficiency for relatively short RT primers (the portion bound to the 30 end of miRNAs). Finally, the stem-loop structure, when unfolded, adds sequence downstream of the miRNA after reverse transcription. The resulting longer RT product presents a template more amenable to real-time TaqMan assay design. The high sensitivity and specificity is largely contributed by the specific forward PCR primer and Taq-Man probe.

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5 End-Point Stem-Loop Real-Time RT-PCR for miRNA Quantification Select a miRNA of interest

Design of stem-loop miRNA capture oligonucleotide probes

Preparation of tissue or cells total RNA samples

Reverse transcription with stem-loop probes

PCR with forward and reverse promiers and Taqman probe

Data analysis

Fig. 5.3 Flowchart of stem-loop RT-PCR analysis for miRNA expression detection. According to Chen et al. (2005)

5.2.4.1

Targets, Primers and Probes

1. Obtain the sequences of miRNAs of interest from the Sanger Center miRNA Registry at http://www.sanger.ac.uk/Software/Rfam/mirna/index.shtml 2. Desgin stem-loop miRNA capture probes for reverse transcription (RT). Stem– loop RT primers bind at its 50 portion to the 30 portion of miRNA molecules (Fig. 5.2)

5.2.4.2

Tissue RNA Samples, Cells, Cell Lysates, and Total RNA Preparation

Tissue RNA 1. Purchase tissue total RNA samples from Ambion or isolate total RNA from tissues of concern using Trizol according to the manufacturer’s protocol. 2. Normalize RNA samples based on the TaqMan1 Gene Expression Assays for human or mouse GAPDH endogenous controls (Applied Biosystems).

Cultured Cell RNA 1. Trypsinize cells in culture and count with a hemocytometer. 2. Pellete approximately 2.8 106 suspended cells by centrifugation at 1,500 rpm for 5 min, wash with 1 mL Dulbecco’s PBS without MgCl2 and CaCl2.

5.2 Protocol

137

3. Resuspend the cell pellets in 140 mL PBS. 4. Process with three different sample preparation methods. (1) In the first method, mix a 50 mL sample (106 cells) with an equal amount of Nucleic Acid Purification Lysis Solution by pipeting up and down ten times, and then spin briefly. Dilute the lysate 1/10 with 1 U/mL RNase inhibitor solution before adding the solution to an RT reaction. (2) In the second method, use a 50 ml sample (106 cells) to purify total RNA using the mirVanaTM miRNA Isolation Kit according to the manufacturer’s protocol. Elute purified total RNA in 100 mL of elution buffer. (3) The third method involves diluting cells 1:2 with 1 PBS, heating at 95 C for 5 min, and immediately chilling on ice before aliquotting directly into RT reactions. miRNA Detection Using mirVanaTM miRNA Detection Kit 1. Perform solution hybridization-based miRNA analysis using the mirVanaTM miRNA Detection Kit according to the manufacturer’s protocol. 2. Synthesize RNA probes by IDT. 3. Detect the radioisotope labeled RNA fragments and quantify with a Cyclone Storage Phosphor System (PerkinElmer, Boston, MA).

5.2.4.3

Reverse Transcriptase Reactions

1. Purchase stem–loop RT primers for the miRNAs of interest for detection (Applied Biosystems). 2. Reverse transcriptase reactions contain: RNA samples, 50 nM stem–loop RT primer, 1 RT buffer, 0.25 mM each of dNTPs, 3.33 U/mL MultiScribe reverse transcriptase, and 0.25 U/mL RNase inhibitor. 3. Incubate the 7.5 mL reactions in an Applied Biosystems 9,700 Thermocycler in a 96- or 384-well plate for 30 min at 16 C, 30 min at 42 C, 5 min at 85 C and then hold at 4 C. 4. Run all reverse transcriptase reactions, including no-template controls and RT minus controls, in duplicate.

5.2.4.4

PCR

1. Run real-time PCR using a standard TaqMan1 PCR kit protocol on an Applied Biosystems 7900HT Sequence Detection System. The 10 mL PCR reaction contains: 0.67 mL RT product, 1 TaqMan1 Universal PCR Master Mix, 0.2 mM TaqMan1 probe, 1.5 mM forward primer, and 0.7 mM reverse primer. 2. Incubate the reactions in a 384-well plate at 95 C for 10 min, followed by 40 cycles of 95 C for 15 s and 60 C for 1 min. 3. All reactions should be run in triplicate.

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5 End-Point Stem-Loop Real-Time RT-PCR for miRNA Quantification

4. The threshold cycle (CT) is defined as the fractional cycle number at which the fluorescence passes the fixed threshold. TaqMan1 CT values are converted into absolute copy numbers using a standard curve from synthetic lin-4 miRNA.

5.2.4.5

Robotics Protocol

A robotics protocol for TaqMan miRNA assays can be set up to increase the throughput. 1. Use Beckman Coulter BioMek Fx with a 96-channel head to dilute the RT primers and TaqMan assays, stamp out oligos for reaction plates, and set up the PCR. 2. Use Perkin-Elmer Multiprobe II HT to dispense RT/Sample mix for RT reactions. 3. For RT reaction set up, stamp out 7.5 mL of 2 RT primers into 96-well plates. Prepare a large mixture of 2 RT/Sample mix (which contains 2 RT buffer, 0.5 mM dNTPs, 6.7 U/mL Reverse Transcriptase, 0.5 U/mL RNAse Inhibitor, and 2 ng/mL total RNA sample). Aliquot 1 ml RT/Sample mix into a 1.5 ml microcentrifuge tube. Allow the robot to distribute 7.5 mL RT/Sample mix into the plate containing 7.5 mL 2 RT primers. Cover the plate with a removable seal. Invert plate to mix the reactions. After a brief spin, incubate the plate on ice for 5 min. Perform RT reaction in an Applied Biosystem’s 9,700 instrument with the following condition: 16 C for 30 min, 42 C for 30 min, 85 C for 5 min, and 4 C on hold. 4. For PCR set up, stamp out 10 mL 5 TaqManTM assays into 96-well plates. Using the Biomek FX, add 11 mL water, 25 mL 2 TM Universal Master Mix, and 4 mL RT product. Mix on the robot by pipeting up and down. Transfer PCR from 96-well plate to four new wells in 384-well plate. Cover the 384-well plate with Applied Biosystems clear optical seal. Spin plate briefly. Perform PCR in Applied Biosystems 7,900 instrument as described previously. 5. If liquid handling robots are not available to the user, it is possible to prepare miRNA in 384-well reaction plates using manually-operated multichannel repeating pipets. The following protocol is for assaying 192 precursor miRNAs in duplicate per 384-well plate. Alternatively, 380 different miRNAs (plus internal control genes) may be profiled per 384-well plate. A master mix containing everything for the PCR except the primers is prepared. Prepare enough master mix for 5 mL reactions (384-well plus 10% extra). The master mix contains 2.5 mL of the 2 SYBR green master mix and 0.5 mL of diluted cDNA (dilute cDNA 1:50 in molecular biology grade water). Add 105 mL of the master mix to each well of a 12-well strip tube. Using a 12-well repeating multichannel pipet (Rainin Model E 12–20), transfer 3 mL of the master mix from the strip tube to each well of the 384-well plate. In separate 12-well strip tubes or 96-well plates, have the primer pairs diluted to 2 mM. Using a 12-well repeating multichannel pipet (Rainin Model E 12–20) add 2 mL of the 2 mM primers to each well of the 384-well plate. Add the optical adhesive cover

References

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and seal. Perform a quick spin (up to 1,500 RPM) on a centrifuge equipped with a 96-well plate adapter. Perform PCR using the real-time instrument as per the manufacturers’ protocol - typically 40 cycles of 15 s 95 C and 1 min 60 C. It is advisable to have a template of the plate’s configuration programmed on the instrument’s computer to assist in the data analysis.

5.3

Application and Limitation

There is an increasing need for sensitive and specific whole miRNA profiling. The ability to effectively profile miRNAs could lead to the discovery of diseases- or tissue-specific miRNA biomarkers, as well as contribute to the understanding of how miRNAs regulate stem cell differentiation. Quantification of miRNA by realtime PCR is a powerful tool to be included in the toolbox of techniques to study miRNA. Real-time PCR may be used to validate the expression of miRNAs discovered during high throughput arrays, as a discovery tool when configured as a low density PCR array, to study the expression of individual miRNAs and as demonstrated here, to identify miRNAs whose expression may be regulated at the level of miRNA processing. Stem-loop RT primers are better than conventional ones in terms of RT efficiency and specificity (Chen et al. 2005). TaqMan miRNA assays are specific for mature miRNAs and discriminate among related miRNAs that differ by as little as one nucleotide. Furthermore, they are not affected by genomic DNA contamination. Precise quantification is achieved routinely with as little as 25 pg of total RNA for most miRNAs. In fact, the high sensitivity, specificity, and precision of this method allows for direct analysis of a single cell without nucleic acid purification. Like standard TaqMan gene expression assays, TaqMan miRNA assays exhibit a dynamic range of seven orders of magnitude. Quantification of five miRNAs in seven mouse tissues showed variation from less than 10 to more than 30,000 copies per cell. This method enables fast, accurate, and sensitive miRNA expression profiling and can identify and monitor potential biomarkers specific to tissues or diseases. Stem-loop RT-PCR can be used for the quantification of other small RNA molecules such as short interfering RNAs (siRNAs). Furthermore, the concept of stem-loop RT primer design could be applied in small RNA cloning and multiplex assays for better specificity and efficiency.

References Allawi HT, Dahlberg JE, Olson S, Lund E, Olson M, Ma WP, Takova T, Neri BP, Lyamichev VI (2004) Quantitation of microRNAs using a modified Invader assay. RNA 10:1153–1161 Chen C, Ridzon DA, Broomer AJ, Zhou Z, Lee DH, Nguyen JT, Barbisin M, Xu NL, Mahuvakar VR, Andersen MR, Lao KQ, Livak KJ, Guegler KJ (2005) Real-time quantification of microRNAs by stem-loop RT-PCR. Nucleic Acids Res 33:e179

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Heid CA, Stevens J, Livak KJ, Williams PM (1996) Real time quantitative PCR. Genome Res 6:986–994 Jiang J, Lee EJ, Gusev Y, Schmittgen TD (2005) Real-time expression profiling of microRNA precursors in human cancer cell lines. Nucleic Acids Res 33:5394–5403 Krichevsky AM, King KS, Donahue CP, Khrapko K, Kosik KS (2003) A microRNA array reveals extensive regulation of microRNAs during brain development. RNA 9:1274–1281 Lagos-Quintana M, Rauhut R, Yalcin A, Meyer J, Lendeckel W, Tuschl T (2002) Identification of tissue-specific microRNAs from mouse. Curr Biol 12:735–739 Liu C-G, Calin GA, Meloon B, Gamliel N, Sevignani C, Ferracin M, Dumitru CD, Shimizu M, Zupo S, Dono M, Alder H, Bullrich F, Negrini M, Croce CM (2004) An oligonucleotide microchip for genome-wide microRNA profiling in human and mouse tissues. Proc Natl Acad Sci USA 101:9740–9744 Livak KJ, Schmittgen TD (2001) Analysis of relative gene expression data using real-time quantitative PCR and the 2[-Delta Delta C(T)] Method. Methods 25: 402–408 Mano H, Takada S (2007) mRAP, a sensitive method for determination of microRNA expression profiles. Methods 43:118–122 Mocellin S, Rossi CR, Pilati P, Nitti D, Marincola FM (2003) Quantitative real-time PCR: a powerful ally in cancer research. Trends Mol Med 9:189–195 Raymond CK, Roberts BS, Garrett-Engele P, Lim LP, Johnson JM (2005) Simple, quantitative primer-extension PCR assay for direct monitoring of microRNAs and short-interfering RNAs. RNA 11:1737–1744 Ro S, Park C, Jin J, Sanders KM, Yan W (2006) A PCR-based method for detection and quantification of small RNAs. Biochem Biophys Res Commun 351:756–763 Schmittgen TD, Lee EJ, Jiang J, Sarkar A, Yang L, Elton TS, Chen C (2008) Real-time PCR quantification of precursor and mature microRNAs. Methods 44:31–38 Schmittgen TD, Jiang J, Liu Q, Yang L (2004) A high-throughput method to monitor the expression of microRNA precursors. Nucleic Acids Res 32:e43 Takada S, Mano H (2007) Profiling of microRNA expression by mRAP. Nat Protoc 2:3136–3145

Chapter 6

miR-Q RT-PCR for miRNA Quantification

Abstract Sharbati-Tehrani et al from the Institute of Veterinary-Biochemistry, Freie Universita¨t Berlin (Berlin, Germany) developed a new cost-effective, highly sensitive, and reliable qRT-PCR for quantification of small RNA molecules such as miRNAs (BMC Mol Biol 9:34, 2008). The method called miR-Q is based on primer extension, followed by a novel quantitative PCR (qPCR) approach that is carried out using three DNA-oligonucleotides. The miR-Q assay exhibits high sensitivity, linearity, and discriminative power without the use of complex fluorochromic probes or LNA-modified oligonucleotides. It shows a high dynamic range of 6–8 orders of magnitude comprising a sensitivity of up to 0.2 fM miRNA, which corresponds to single copies per cell. There is nearly no cross reaction among closely-related miRNA family members, which points to the high specificity of the assays. The method has been applied to quantify the expression of let-7b in different human cell lines as well as miR-145 and miR-21 expression in porcine intestinal samples. The results show that miR-Q is a cost-effective and highly specific approach, which neither requires the use of fluorochromic probes, nor Locked Nucleic Acid (LNA)-modified oligonucleotides. Moreover, it provides a remarkable increase in specificity and simplified detection of small RNAs.

6.1

Introduction

Sharbati-Tehrani et al from the Institute of Veterinary-Biochemistry, Freie Universita¨t Berlin (Berlin, Germany) developed a new cost-effective, highly sensitive, and reliable qRT-PCR for quantification of small RNA molecules such as miRNAs (Sharbati-Tehrani et al. 2008). The method called miR-Q is based on primer extension, followed by a novel quantitative PCR (qPCR) approach that is carried out using three DNA-oligonucleotides. (1) During the first step, miRNA molecules are converted into cDNA and simultaneously elongated by reverse transcription using a miRNA-specific oligonucleotide with 50 overhang (RT6-miR-x, x

Z. Wang and B. Yang, MicroRNA Expression Detection Methods, DOI 10.1007/978-3-642-04928-6_6, # Springer-Verlag Berlin Heidelberg 2010

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indicates a particular miRNAs). (2) Subsequently, cDNA molecules are amplified and quantified by means of a novel qPCR approach based on utilizing three DNAoligonucleotides at different concentrations. Primarily, a DNA oligonucleotide molecule (RT6-miR-x) is used for reverse transcription, which comprises two main sequence portions. The terminal six bases at the 30 end of RT6-miR-x are miRNA-specific and hybridized to the template RNA molecule of interest. This oligonucleotide molecule also comprises a second sequence portion, enabling the binding of a universal primer (MP-fw) to prime the exponential DNA amplification. For subsequent qPCR analysis, another oligonucleotide molecule (short-miRx-rev) is employed comprising a miRNA-specific sequence portion at its 30 end. Therefore, short-miR-x-rev can hybridize to the cDNA molecule (complementary to the target miRNA) generated after the reverse transcriptase reaction. In addition, the 50 overhang provides a terminal binding sequence for another universal primer (MP-rev). Consequently, two 50 overhangs are introduced to convert and elongate the small RNA molecule. The final exponential amplification is accomplished by means of universal primers (MP-fw and MP-rev), which hybridize to terminal binding sites of the overhangs. These universal primers are not assay-specific and are therefore employed independently of various analyzed miRNA molecules. The universal overhang sequences of the reverse primers as well as RT primers have been deduced from mycobacterial genome, finally being optimized for negligible complementarities and dimer formation. These universal overhang sequences show no similarity with any known miRNA. (3) The final detection and quantification of template miRNA molecules are performed by real-time acquisition of the SYBR Green fluorescence, utilizing a calibration curve.

6.2 6.2.1

Protocol Materials

1. Materials for cell culture, RNA isolation, and RT-PCR (e.g. culture dishes, pipettes, pipette tips)

6.2.2

Instruments

1. Agilent 2100 Bioanalyzer and the RNA Nano Chips (Agilent, Waldbronn, Germany) or equivalent 2. Rotor-Gene 3000 real-time Detection System (Corbett Life Science, Sydney, Australia) 3. StepOnePlus™ Real-Time PCR System (Applied Biosystems, Darmstadt, Germany)

6.2 Protocol

6.2.3

143

Reagents

1. mirVana miRNA Isolation Kit (Ambion) 2. RevertAid™ MMuLV Reverse Transcriptase (Fermentas GmbH, St. Leon-Roth, Germany) 3. Immolase DNA Polymerase (Bioline, Luckenwalde, Germany) 4. SensiMix DNA Kit (Quantace Ltd., Berlin, Germany) 5. RevertAid™ M-MuLV Reverse Transcriptase (Fermentas GmbH)

6.2.4

Procedures

The protocols described in this section are essentially the same as reported in the study of Sharbati-Tehrani et al. (2008) (Fig. 6.1).

6.2.4.1

Oligonucleotides and Synthetic miRNA Molecules

1. Obtain miRNA sequences of interest from the miRBase Sequence database Release 9.2 [20]. 2. Design an RT oligonucleotide primer specific for each selected miRNA (RT6miR-x). An RT6-miR-x contains a stretch of binding sequence for forward universal primer and 6 bases complementary to the 30 end sequence of the target miRNA for binding the miRNA (50 -TGTCAGGCAACCGTATTCACCGTGAGTGGT + 6 miRNA-specific complementary nucleotides). 3. Design a PCR oligonucleotide primer specific for each selected miRNA (ShortmiR-x-rev). A Short-miR-x-rev contains a stretch of binding sequence for forward universal primer and ~16 nucleotides complementary to the 50 end sequence of the target miRNA (50 -CGTCAGATGTCCGAGTAGAGGGGGAACGGCG + 16 miRNA-specific nucleotides). 4. Design a Forward universal primer (MP-FW): TGTCAGGCAACCGTATTCACC, which is complementary to the RT6-miR-x. 5. Design a Reverse universal primer (MP-REV): CGTCAGATGTCCGAGTAGAGG, which is complementary to Short-miR-x-rev. 6. Synthetic miRNA molecules are used for the validation of assays. 7. All DNA and RNA-oligonucleotides are synthesized by services provided by quantified companies.

6.2.4.2

Total RNA Isolation from Samples

1. Human total RNA are prepared using the mirVana miRNA Isolation Kit, according to the manufacturer’s protocol.

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6 miR-Q RT-PCR for miRNA Quantification

5’-

-3’ miRNA RT6-miR-x

RT 5’3’-

-3’

5’

5’

SYBR Green qPCR Short-miR-x-rev

5’ -3’

3’-

SYBR Green qPCR 5’ MP-rev

MP-fw

Fig. 6.1 Schematic description of assay design. The miRNA is first converted and simultaneously elongated into a cDNA molecule using an miRNA-specific oligonucleotide with 50 overhang (RT6miR-x) and six complementary bases (red ). Detection and amplification of the relating cDNA are employed, using a novel PCR approach with three different oligonucleotides at different concentrations within the same assay. The cDNA sequence (blue) is first detected and elongated by a specific oligonucleotide with 50 overhang (short-miR-x-rev). Exponential amplification is then performed using two terminal universal primers (MP-fw & MP-rev). Modified from SharbatiTehrani et al. (2008)

2. The RNA quality and quantity is proven using the Agilent 2100 Bioanalyzer and the RNA Nano Chips.

6.2.4.3

Reverse Transcription

1. 250 fmol of the miRNA-specific DNA-oligonucleotide (RT6-miR-x) with 50 overhang and the RevertAid™ MMuLV Reverse Transcriptase are employed to transcribe miRNA into cDNA. 2. A mixture of 50 ng total RNA spiked with synthetic miRNA and RT6-miR-x is first prepared in a 4 mL volume. 3. The mixture is incubated at 70 C for 5 min and chilled on ice. 4. Bring the volume up to 10 mL by adding the RT-Buffer, 1 mM dNTPs, 100 U Reverse Transcriptase and water. 5. Incubate the reaction at 37 C for 5 min followed by 42 C for 60 min. 6. The enzyme is inactivated by heating at 70 C for 10 min.

6.2 Protocol

6.2.4.4

145

Quantitative PCR

1. The optimal annealing temperature for every miRQ assay should be first proven by performing a conventional PCR with an annealing temperature gradient ranging from 53 to 65 C. The reaction is performed with 4 nM short miR-x-rev, 100 nM MP-fw, and 100 nM MP-rev using the Immolase DNA Polymerase. The reaction is carried out according to the qPCR conditions in 25 mL final volume using 2 mL of cDNA, which is obtained from RT reaction with 10 pM of the particular synthetic miRNA. The product size ranges from 82 to 85 bp, depending on the miRNA being detected. 2. Having determined the optimal annealing temperature, quantify the cDNA, using the Rotor-Gene 3000 real-time Detection System as well as the StepOnePlus™ Real-Time PCR System. For this purpose, make triplicate measurements of 2 mL cDNA in 10 mL ﬁnal reaction volume. 3. Perfom SYBR Green qPCR using the SensiMix DNA Kit, 4 nM short-miR-x-rev, 100 nM MP-fw, and 100 nM MP-rev. 4. Run the amplification via the first step at 95 C for 10 min, followed by 40 cycles with 15 s at 95 C, 10 s at the particular annealing temperature, and 20 s at 72 C. 5. Acquire the fluorescence signal at 72 C and convert the Ct values into fM miRNA, using an miRNA-specific calibration curve. 6. Determine miRNA amounts by triplicate measurements for each sample, compared with a calibration curve established by reverse transcription of serially diluted amounts of the particular synthetic miRNA in the presence of 50 ng bacterial total RNA.

6.2.4.5

5S Ribosomal RNA Normalisation

1. Sample-to-sample variation of miRNA expression should be corrected by normalization with 5S rRNA chosen as a housekeeping gene. Firstly, 50 ng of total RNA is reverse-transcribed using the RevertAid™ M-MuLV Reverse Transcriptase, according to the manufacturer’s protocol using random Hexamers. 2. 5S rRNA molecules are quantitated by triplicate measurements of 2 mL cDNA in 10 mL final reaction volume. 3. Perform SYBR Green qPCR using the SensiMix DNA Kit and 0.2 mM of each primer 5S rRNA-fw (gcccgatctcgtctgatct) and 5S rRNA-rev (agcctacagcacccggtatt). 4. Run amplification via the first step at 95 C for 10 min, followed by 40 cycles with 15 s at 95 C, 10 s at 60 C, and 20 s at 72 C, providing an amplicon of 114 bp. 5. Acquire the fluorescence signal at 72 C and convert Ct values into fg 5S rRNA per qPCR reaction using a calibration curve, which is established by serial dilutions of the corresponding PCR product. 6. Perform normalization by calculating the miRNA:5S rRNA expression ratios.

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6 miR-Q RT-PCR for miRNA Quantification

Application and Limitation

miRQ approach is a real-time RC-PCR method of a different sort. It represents a new, alternative method, which exhibits advanced discriminative power and sensitivity. It is based on primer extension, followed by amplification and quantification of the corresponding cDNA, based on the SYBR Green intercalating chemistry. The miR-Q approach provides a linearity of up to eight orders of magnitude detecting as low as 0.2 fM miRNA molecules.

References Sharbati-Tehrani S, Kutz-Lohroff B, Bergbauer R, Scholven J, Einspanier R (2008) miR-Q: a novel quantitative RT-PCR approach for the expression profiling of small RNA molecules such as miRNAs in a complex sample. BMC Mol Biol 9:34

Chapter 7

Poly(A)-Tailed Universal Reverse Transcription

Abstract Zheng’ laboratory from the Beijing Institute of Radiation Medicine (Beijing, People’s Republic of China) and Yan’s laboratory from the Department of Physiology and Cell Biology, the University of Nevada School of Medicine (Reno, NV, USA) independently established the Poly(A)-Tailed Universal Reverse Transcription technique, a simple method to detect the expression of mature miRNAs all in the same year (Mol Biotechnol 32:197–204, 2006; Biochem Biophys Res Commun 351:756–763, 2006). With this method, total RNA is first polyadenylated by poly(A) polymerase, and then cDNA is synthesized by a specific reverse transcriptase (RT) primer and RT, using the poly(A)-tailed total RNA as templates. Next, the cDNA is amplified using a miRNA-specific primer and a universal primer. The expression of several mature miRNAs was assayed by this method. Compared with other RT-PCR methods for miRNA detection, the poly (A)-tailed RT-PCR is a relatively simple, convenient, and accurate method.

7.1

Introduction

Zheng’ laboratory from the Beijing Institute of Radiation Medicine (Beijing, People’s Republic of China) and Yan’s laboratory from the Department of Physiology and Cell Biology, the University of Nevada School of Medicine (Reno, NV, USA) independently established the Poly (A)-Tailed Universal Reverse Transcription technique, a simple method to detect the expression of mature miRNAs all in the same year (Fu et al. 2006; Ro et al. 2006). With this method, total RNA is first polyadenylated by poly(A) polymerase, and then cDNA is synthesized by a specific reverse transcriptase (RT) primer and RT using the poly(A)-tailed total RNA as templates. Next, the cDNA is amplified using a miRNA-specific primer and a universal primer (Fig. 7.1). The expression of several mature miRNAs was assayed by this method. All these data show that the poly(A)-tailed RT-PCR is a simple, convenient, and accurate method to detect the expression of miRNAs. This method

Z. Wang and B. Yang, MicroRNA Expression Detection Methods, DOI 10.1007/978-3-642-04928-6_7, # Springer-Verlag Berlin Heidelberg 2010

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7 Poly(A)‐Tailed Universal Reverse Transcription

5’

miRNA

3’

Polyadenylation miRNA

5’

Polyadenylated miRNA

Poly(A)

3’

Primer Annealing RT Primer

cDNA

Pure cDNA

RT

RNase treatment

qPCR 5’

Universal Primer 3’

miRNA-specific Primer

Fig. 7.1 Schematic illustration of small RNA cDNA (srcDNA) library construction and real-time quantitative PCR (Q-PCR) analysis. Small RNAs were polyadenylated using a poly(A) polymerase. The poly(A)-tailed RNAs were reverse-transcribed using a primer containing oligo dTs flanked by an adaptor sequence to produce srcDNAs. The sample was treated with RNase H to remove the small RNAs from the cDNAs. Conventional PCR or Q-PCR was carried out using a small RNA-specific primer (srSP) and a universal reverse primer, RTQ-UNIr. Modified from Ro et al. (2006)

has several advantages for the detection of miRNAs. First, in comparison with other methods, results can be obtained in a relatively short time. Second, the expression of many miRNAs can be detected simultaneously. Third, in comparison with the Northern blotting analysis, this method requires a small amount of RNA and needs no radioactive isotopes. In addition, compared with other RT-PCR methods for miRNA detection, the poly(A)-tailed RT-PCR is a relatively simple, convenient, and accurate method.

7.2 7.2.1

Protocol Materials

1. 12% polyacrylamide gel 2. pGEM-T vector (Promega)

7.2 Protocol

7.2.2

149

Instruments

1. Thermocycler

7.2.3 1. 2. 3. 4. 5. 6. 7. 8. 9.

Reagents

Trizol (Invitrogen, Carlsbad, CA) mirVanaTM miRNA Isolation kit (Ambion, Austin, TX) Poly(A) polymerase (Takara) SuperScript III (invitrogen) Dithiothreitol [DTT] RNaseout Ethidium bromide (EtBr) Elution buffer: [0.5 M NH4Ac,10 mM Mg(Ac)2,1 mM EDTA] Phenol/chloroform

7.2.4

Procedures

The Poly(A)-Tailed Universal Reverse Transcription protocols described in this chapter are essentially the same as reported by Fu et al. (2006) and Ro et al. (2006) (see Fig. 7.1).

7.2.4.1

Primer Design

1. As one primer of miRNA in the amplification is fixed, the miRNA-specific primers are critical for the specificity of PCR. The following criteria are used for the miRNA primer design. 2. For most miRNAs, the primer sequence is the same as for the miRNA gene. 3. Some miRNAs are homologous to others, therefore their primer sequences are only part of the miRNA gene 50 end to ensure that the 30 ends of the primers are different. 4. The RT primer contains 25 dTs flanked by an adaptor sequence at 50 end: 50 -CCAATTCTAGAGCTCGAGGCAGGCAGGCGACATGGCTGGCTGGCTAGTTAAGCTTGGTACCGAGCTCGGATCCACTAGTCC(T)25 V(A/G/C) N(A/G/C/T)-30 (Ro et al. 2006) or (ATT CTA GAG GCC GAG GCG GCC GAC ATGd(T)30(A, G, or C)(A, G, C, or T)) (Fu et al. 2006). 5. A Universal reverse primer for PCR: 50 -CGAATTCTAGAGCTCGAGGCAGG-30 .

150

7.2.4.2

7 Poly(A)‐Tailed Universal Reverse Transcription

RNA Extraction and Polyadenylation

1. Extract total RNA from the cultured cells using Trizol according to the manufacturer’s protocol, or from tissues of concern using mirVana miRNA Isolation kit, according to the manufacturer’s instructions. 2. Determine the concentration of RNA samples by the absorbance at 260 nm. 3. Polyadenylate RNA by poly(A) polymerase. Of the polyadenylation reaction, 50 mL are set up with 4 mg of total RNA and 5 U of poly(A) polymerase according to the manufacturer’s protocol. Incubate the reaction at 37 C for 30 min. 4. After incubation, recover the poly(A)-tailed RNA by phenol/chloroform extraction and ethanol precipitation.

7.2.4.3

Reverse Transcription (RT)

1. Perform RT using 4 mg total RNA or poly(A)-tailed RNA and 1 mg of RT primer with 200 U of SuperScript III. 2. Incubate a 4-mg aliquot of RNA (10 mL of total volume) with 1 mL of RT primer and 1 mL of dNTP mix (10 mM each) at 65 C for 5 min to remove any RNA secondary structure. 3. Chill the reactions on ice for at least 1 min. 4. Add the remaining reagents (5 buffer, DTT, RNaseout, SuperScript III) as specified in the SuperScript III protocol. 5. Proceed the reaction for 60 min at 50 C. 6. Treat the reaction with RNase H to remove the small RNAs from the cDNAs. 7. Finally, inactivate the RT by a 15-min incubation at 70 C. The minus reverse transcription control should be treated identically as described previously except that the reactions lacked SuperScript III and primer.

7.2.4.4

PCR, and Sequencing

8. Run the amplification of the miRNAs for 25 cycles with a final annealing temperature of 60 C using miRNA-specific primers and the 30 primer: 50 -ATT CTA GAG GCC GAG GCG GCC GAC ATG T-30 . 9. The b-actin is amplified at a final annealing temperature of 55 C using the following pair of primers: 50 - GGC ATC GTG ATG GAC TCC G -30 and 50 GCT GGA AGG TGG ACA GCG A -30 . 10. Analyze the PCR products on 12% polyacrylamide gel with EtBr staining. 11. The gel slices containing DNA with a size of about 80 bp are excised and the DNA are eluted in elution buffer at 37 C and recovered by phenol/chloroform extraction followed by ethanol precipitation. 12. The DNA fragment is directly subcloned into pGEM-T vector. 13. Sequencing, verify the construct.

References

7.3

151

Application and Limitation

This method has several advantages for the detection of miRNAs. First, in comparison with other methods, results can be obtained in a relatively short time. Second, the expression of many miRNAs can be detected simultaneously. Third, in comparison with Northern blotting analysis, this method requires a small amount of RNA and need no radioactive isotopes. In addition, compared with other RT-PCR methods for miRNA detection, the poly (A)-tailed RT-PCR is a relatively simple, convenient, and accurate method. The specifisity of the assay may be inherently limited by the use of only one miRNA-specific primer and the use of a universal primer.

References Fu HJ, Zhu J, Yang M, Zhang ZY, Tie Y, Jiang H, Sun ZX, Zheng XF (2006) A novel method to monitor the expression of microRNAs. Mol Biotechnol 32:197–204 Ro S, Park C, Jin J, Sanders KM, Yan W (2006) A PCR-based method for detection and quantification of small RNAs. Biochem Biophys Res Commun 351:756–763 Wang JW, Cheng JQ (2008) A simple method for profiling miRNA expression. Methods Mol Biol 414:183–190

Chapter 8

Multiplexing RT-PCR for High-Throughput miRNA Profiling

Abstract While the RT-PCR techniques introduced in Chaps. 5–7 are in general considered low throughput methods, they can be modified and upgraded to medium throughput miRNA detection methods. In 2006, Lao and colleagues from the Applied Biosystems, Foster City (CA, USA) investigated the parameters associated with multiplexing RT-PCR to obtain relative abundance profiles of multiple miRNAs in small sample sizes down to the amount of RNA found in a single cell (Biochem Biophys Res Commun 343:85–89, 2006). In the same year, Surani and colleagues from Wellcome Trust/Cancer Research UK Gurdon Institute of Cancer and Developmental Biology, University of Cambridge (Cambridge, UK) (Nucleic Acids Res 34:e9, 2006; Nat Protoc 1:1154–1159, 2006) developed a Single Cell Stem-Looped Real-Time PCR (SC-SL-RT-PCR) protocol for the detection of the expression profile of 220 mature miRNAs in a single embryonic stem cell, cell by stem-looped real-time PCR. More recently, Mestdagh et al. from the Center for Medical Genetics, Ghent University Hospital (Ghent, Belgium) (Nucleic Acids Res 36:e143, 2008) presented the successful evaluation of the Megaplex (450 mature miRNAs) reverse transcription format of the stem-loop primer-based real-time quantitative polymerase chain reaction (RT-qPCR) approach to quantify miRNA expression. In the meantime, Schmittgen et al. (Mol Biol 429:89–98, 2008) from College of Pharmacy, Ohio State University (Columbus, OH, USA) also reported an example of profiling the expression of over 200 miRNA precursors (pre-miRNAs) using high-throughput real-time PCR. The high specificity of RT-qPCR together with a superior sensitivity makes this approach the method of choice for high-throughput miRNA expression profiling (Wark et al. 2008). While PCR will never rival the throughput of microchip arrays, in situations where one is interested in assaying several hundreds of genes, high throughput, real-time PCR is an excellent alternative to microchip arrays. Moreover, multiplexing RT-PCR is quantitative for miRNA expression detection.

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8 Multiplexing RT-CR for High‐Throughput miRNA Profiling

Introduction

In 2006, Lao and colleagues from the Applied Biosystems, Foster City (CA, USA) investigated the parameters associated with multiplexing RT-PCR to obtain relative abundance profiles of multiple miRNAs in small sample sizes down to the amount of RNA found in a single cell (Lao et al. 2006). They showed that judicious sequence partitioning between RT primers and second strand synthesis primers permitted multiplexed RT-PCR amplification of miRNA in very small samples to allow individual real-time PCR quantification. The same group further demonstrated that zip coding the primers and TaqMan probes with sequences specific to each miRNA greatly improves reaction specificity, which permits the profiling of all miRNAs in a single multiplexed RT-PCR reaction (Lao et al. 2006). In the same year, Surani and colleagues from Wellcome Trust/Cancer Research UK Gurdon Institute of Cancer and Developmental Biology, University of Cambridge (Cambridge, UK) (Tang et al. 2006a, b) developed a Single Cell Stem-Looped Real-Time PCR (SC-SL-RT-PCR) protocol for the detection of the expression profile of 220 mature miRNAs in a single embryonic stem cell, cell by stem-looped real-time PCR. More recently, Mestdagh et al. from the Center for Medical Genetics, Ghent University Hospital (Ghent, Belgium) (Mestdagh et al. 2008) presented the successful evaluation of the Megaplex reverse transcription format of the stemloop primer-based real-time quantitative polymerase chain reaction (RT-qPCR) approach to quantify miRNA expression. The Megaplex reaction provides simultaneous reverse transcription of 450 mature miRNAs, ensuring high-throughput detection. Further, the introduction of a complementary DNA pre-amplification step significantly reduces the amount of input RNA needed, even down to singlecell level. Moreover, pre-amplification using 10 ng of input RNA enabled the detection of miRNAs that were undetectable when using Megaplex alone with 400 ng of input RNA. In the meantime, Schmittgen et al. (2008) from College of Pharmacy, Ohio State University (Columbus, OH, USA) also reported an example of profiling the expression of over 200 miRNA precursors (pre-miRNAs) using high-throughput real-time PCR. The high specificity of RT-qPCR together with a superior sensitivity makes this approach the method of choice for high-throughput miRNA expression profiling. While PCR will never rival the throughput of microchip arrays, in situations where one is interested in assaying several hundreds of genes, high throughput, real-time PCR is an excellent alternative to microchip arrays. Moreover, multiplexing RT-PCR is quantitative for miRNA expression detection. Multiplexing RT-PCR for miRNA detection involves the following steps. Step 1 reverse transcribes all the miRNAs in a single reaction and then step 2 PCR amplifies the cDNA products to provide enough samples for step 3. Step 3 is done as simultaneous, individual singleplex TaqMan reactions in 384-well reaction plates to monitor the abundance of each miRNA after the multiplexed RT-PCRs. Sequences for primers and probes will be provided by the authors upon request.

8.2 Protocol

8.2

155

Protocol

8.2.1

Materials

1. Materials for cell culture, RNA isolation, and RT-PCR (e.g., culture dishes, pipettes, pipette tips)

8.2.2

Instruments

1. Routine instruments (such as centrifuges, votexes, ect) 2. AB 7900 HT Sequence Detection System

8.2.3

Reagents

1. 10 cDNA Archiving kit buffer (Applied Biosystems) 2. MMLV reverse transcriptase 3. 2 Universal Master Mix with no UNG (Applied Biosystems)

8.2.4

Procedures

The protocols described in this section are essentially the same as reported in the study of Lao and colleagues (Lao et al. 2007). 8.2.4.1

RNA and DNA

1. Isolate total RNA samples from tissues or cells of concern, using the commercial kits as already described in the previous chapters 2. Purchase synthetic miRNAs from Integrated DNA Technologies Inc, if needed. 3. Synthesize DNA oligonucleotides by Applied Biosystems or Integrated DNA Technologies 8.2.4.2

Reverse Transcription

1. Reverse transcription reactions of 5 mL contain: 0.5 mL of 10 cDNA Archiving kit buffer, 0.335 mL MMLV reverse transcriptase (50 U/mL), 0.25 mL of 100 mM dNTP, 0.065 mL of AB RNase inhibitor 20 U/mL, 0.5 mL of 48- or 190-plex reverse primer, RP (50 nM each), 2 mL of total RNA, and 1.35 mL H2O.

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8 Multiplexing RT-CR for High‐Throughput miRNA Profiling

2. Prepare the reaction mixture by adding 2 mL of RNA sample to 3 mL of freshly prepared stock reaction mixture containing the remaining reaction ingredients for at least ten reactions. 3. Perform the reaction with the following incubation conditions: (20 C/30 s – 42 C/ 30 s – 50 C/1 s) 60 cycles. 4. Inactivate the enzyme by incubation at 85 C for 5 min. 8.2.4.3

Pre-PCR Amplification

1. The pre-real-time PCR mixture of 25 mL contain: 12.5 mL of 2 Universal Master Mix with no UNG, 5 mL of RT sample, 2.5 mL of 48- or 190-plex forward primer at 500 nM each, 1.25 mL of 100 mM universal reverse primer, 1.25 mL of 5 U/mL AmpliTaq Gold, 0.5 mL of 100 mM dNTP, 0.5 mL of 100 mM MgCl2, and 1.5 mL dH2O. 2. The temperature profile for the reaction are: 10 min incubation at 95 C to activate Taq-GOLD, 55 C incubation for 2 min, followed by 14 cycles of 95 C for 1 s and 65 C for 1 min. 8.2.4.4

Real-Time PCR

1. Dilute the pre-amplified RT-PCR mixture to 100 mL by adding 75 mL H2O into 25 mL of pre-amplified sample. The probes for each TaqMan reaction consist of the 30 –18 nucleotides of the RT-RP for each miRNA with the fluorescence dye, FAM at the 50 end and a minor groove binder with non-fluorescence quencher, MGB, on the 30 end. 2. Real-time reaction mixtures contain: 5 mL of 2 Universal Master Mix with no UNG, 2 mL of a 5 mM FP + 1 mM TaqMan probe mixture, 0.1 mL of 100 mM UR, 0.1 mL of 4 diluted pre-amplified RT-PCR sample, and 2.8 mL dH2O. 3. Assemble real-time reaction mixtures by adding 2 mL of individual FP + TaqMan probe to 8 mL of freshly prepared stock solution containing the rest of the real-time PCR reagents. 4. Perform real-time PCR on an AB 7900 HT Sequence Detection System in a 384well plate format, with the temperature regime consisting of a hot start of 95 C for 10 min, followed by 40 cycles of 95 C for 15 s, and 60 C for 1 min. The realtime PCRs for each miRNA were run in duplicate.

8.3

Application and Limitation

While PCR will never rival the throughput of microchip arrays, in situations where one is interested in assaying several hundreds of genes, medium throughput, realtime PCR is an excellent alternative to microchip arrays. Moreover, multiplexing RT-PCR is quantitative for miRNA expression detection.

References

157

References Lao K, Xu NL, Sun YA, Livak KJ, Straus NA (2007) Real time PCR profiling of 330 human micro-RNAs. Biotechnol J 2:33–35 Lao K, Xu NL, Yeung V, Chen C, Livak KJ, Straus NA (2006) Multiplexing RT-PCR for the detection of multiple miRNA species in small samples. Biochem Biophys Res Commun 343:85–89 Mestdagh P, Feys T, Bernard N, Guenther S, Chen C, Speleman F, Vandesompele J (2008) Highthroughput stem-loop RT-qPCR miRNA expression profiling using minute amounts of input RNA. Nucleic Acids Res 36:e143 Schmittgen TD, Lee EJ, Jiang J (2008) High-throughput real-time PCR. Methods Mol Biol 429:89–98 Tang F, Hajkova P, Barton SC, Lao K, Surani MA (2006a) MicroRNA expression profiling of single whole embryonic stem cells. Nucleic Acids Res 34:e9 Tang F, Hajkova P, Barton SC, O’Carroll D, Lee C, Lao K, Surani MA (2006b) 220-plex microRNA expression profile of a single cell. Nat Protoc 1:1154–1159 Wark AW, Lee HJ, Corn RM (2008) Multiplexed detection methods for profiling microRNA expression in biological samples. Angew Chem Int Ed Engl. 2008;47(4):644–652

Chapter 9

miRNA Amplification Profiling (mRAP)

Abstract The PCR methods described in the previous chapters focus on miRNA expression detection, but are not designed for the discovery of new miRNAs. Mano and Takada from the Division of Functional Genomics, Jichi Medical University (Shimotsukeshi, Tochigi, Japan) developed a modified PCR approach, termed miRNA amplification profiling (mRAP) (Nucleic Acids Res 34:e115, 2006; Methods 43:118–122, 2007; Nat Protoc 2:3136–3145, 2007) that relies on the traditional miRNA cloning approach (Curr Biol 12:735–739, 2002) but with the substitution of the SMART (switching mechanism at the 50 -end of RNA templates of reverse transcriptase) (Science 294:862–864, 2001) reaction for the ligation of a synthetic primer to the 50 -end of small RNAs. mRAP combines cloning for new miRNA discovery, high throughput profiling, and quantification of miRNA levels into one. This approach is highly sensitive, readily allowing the isolation of >1 104 independent miRNA-derived cDNAs from 1 104 cells. The mRAP method thus makes it possible to analyze miRNA expression profiles for small quantities of tissue or cells such as fresh clinical specimens. The mRAP method can be performed in a conventional molecular biology laboratory, it readily allows the processing of multiple samples in parallel, it is able to detect a 1- nt difference among miRNAs, and is capable of identifying both new miRNAs and mutations in known miRNAs. mRAP may currently be the best choice for obtaining expression profiles for both known and unknown miRNAs or for identification of sequence alterations in miRNAs with small amounts of starting material. However, the mRAP is a complex, multistep procedure, and may not be practical for clionical use as a diagnostic tool.

9.1

Introduction

The PCR methods described in the previous chapters, focus on miRNA expression detection, but are unable to lead to the discovery of new miRNAs. Mano and Takada developed a modified PCR approach, termed miRNA amplification profiling (mRAP) (Takada et al. 2006; Mano and Takada 2007; Takada and

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Mano 2007) that relies on the traditional miRNA cloning approach (Lagos-Quintana et al. 2002) but with the substitution of the SMART (switching mechanism at the 50 -end of RNA templates of reverse transcriptase) (Lee and Ambros 2001) reaction for the ligation of a synthetic primer to the 50 -end of small RNAs. mRAP combines cloning for new miRNA discovery, highthrouput profiling, and quantification of miRNA levels into one. The mRAP method involves the following steps: (1) The mRAP method starts with the isolation of <200-nt RNA molecules. Concentration of the miRNA fraction is achieved by subjecting these RNA molecules of <200 nt to denaturing PAGE on a 15% gel and then eluting RNAs of defined size from the gel. (2) The size-selected RNAs are then dephosphorylated to prevent their ligation to each other during the next step. (3) The dephosphorylated RNA is ligated with the 30 adaptor (self-ligation of the 30 adaptor cannot occur because its 30 -end is blocked with inverted deoxythymidine) and subjected to RT with the RT primer. (4) The amplification products by PAGE usually contain three major bands of ~70, ~90, and ~120 bp. (5) Cloning of the ~90-bp DNA fragments, which are first subjected to concatamer formation with the use of the restriction endonuclease BanI. The sizeselected concatamers are cloned into a T-vector, and the resulting plasmid library is the miRNA library. (6) Verifying the quality of the library by nucleotide sequencing of the cDNAs and determination of the miRNA expression profile (Figs. 9.1 and 9.2). The mRAP method can be performed in a conventional molecular biology laboratory, it readily allows the processing of multiple samples in parallel, is able to detect a 1-nt difference among miRNAs, and is capable of identifying both new miRNAs and mutations in known miRNAs. The major advantage of mRAP is its high sensitivity. mRAP may currently be the best choice for obtaining expression profiles for both known and unknown miRNAs or for identification of sequence alterations in miRNAs with small amounts of starting material. Analysis of mRAP products with recently developed high throughput sequencing systems such as pyrosequencing (Ronaghi 2001) would be a robust approach to extensively profile miRNAs in a given tissue. However, the mRAP is a complex, multistep procedure, and may not be practical for clionical use as a diagnostic tool.

9.2 9.2.1

Protocol Materials

1. 15% PAGE gel (SequaGel Sequencing System; National Diagnostics, Atlanta, GA) 2. ProbeQuant G50 (GE Healthcare Bio-Sciences, Uppsala, Sweden) 3. D-tube (EMD Biosciences, San Diego, CA)

9.2 Protocol

161 PO4

OH

miRNA

Dephosphorylation OH

PO4

Ligation

Reverse transcription

3’ Adaptor

RT primers

CCC– Annealing

–GGG 5’ Adaptor

–GGG

CCC– Extension

–GGG

–CCC– PCR

PCR primers

BanI Digestion

Ligation

Filling in & (A) Addition

Cloning into T-vector, Sequencing, and miRNA profiling

Fig. 9.1 Schematic representation of the reactions that constitute the miRNA amplification profiling (mRAP) method. miRNAs are dephosphorylated, ligated to the 30 adaptor and subjected to reverse transcription (RT) and the SMART reaction. The resulting cDNAs are annealed with the 50 adaptor and amplified by PCR. The amplification products are digested with BanI and concatamerized. The protruding ends of the products are filled in, and an A nucleotide is added to the 30 -ends of the concatamers for cloning into a T-vector Modified from Takada and Mano (2007)

162

9 miRNA Amplification Profiling (mRAP) Select a miRNA of interest

Preparation of tissue or cells total RNA samples

Design and synthesize: 3' adaptor 5' adaptor RT primer PCR primers

Fractionation of small RNA molecules by PAGE gel

Dephosphorylation of RNA

Ligation of 3' adaptor to dephosphorylated RNA

Reverse transcription (RT) with RT primer

Anealing of cDNA with 5' adaptor

Extension

PCR amplification with PCR primers

BanI digestion of PCR products

Ligation of digested PCR products

T-vector cloinging, sequencing and miRNA profiling

Fig. 9.2 Flowchart of miRAGE analysis for miRNA expression detection. According to Mano and Takada (2007), Takada and Mano (2007)

9.2.2

Instruments

1. Vertical electrophoresis apparatus (Nihon Eido, cat. no. NA1113 or equivalent; System Instruments, cat. no. SE8010 or equivalent)

9.2 Protocol

163

2. Submarine electrophoresis apparatus (Nihon Eido, cat. no. NB1012 or equivalent) 3. ProbeQuant G-50 micro columns (GE Healthcare, cat. no. 27-5335-01) 4. D-Tube (Novagen, cat. no. 71504-3)

9.2.3

Reagents

1. Fresh or cryopreserved tissue or cells (or cell lines) derived from animals or possibly from plants 2. mirVana miRNA isolation kit (Ambion, Austin, TX) 3. Diethylpyrocarbonate-treated water (DEPC water; Sigma-Aldrich) 4. SequaGel sequencing system (National Diagnostics), consisting of SequaGel concentrate, SequaGel diluent and SequaGel buffer 5. SYBR Green II (Cambrex) 6. 2-Butanol (Wako) 7. Ethanol 8. Chloroform 9. Phenol 10. Glycogen (Roche Diagnostics) 11. Calf intestinal alkaline phosphatase (CIAP; New England Biolabs) 12. T4 RNA ligase (New England Biolabs) 13. Acetylated BSA (Invitrogen) 14. ATP (Takara Bio) 15. DMSO (Sigma-Aldrich) 16. PowerScript reverse transcriptase (Clontech) 17. Deoxynucleoside triphosphates (dNTPs; GE Healthcare) 18. AmpliTaq Gold polymerase (Applied Biosystems) 19. Sodium acetate 20. 10 Tris-borate-EDTA (TBE; Sigma-Aldrich) 21. Acrylamide (Bio-Rad) 22. N,N0 -Methylene-bis-acrylamide (bis-acrylamide) 23. Ammonium persulfate (APS; Wako) 24. Tetramethylethylenediamine (TEMED) 25. 25-bp DNA ladder (Invitrogen) 26. Ethidium bromide (EtBr; Sigma-Aldrich) 27. BanI restriction endonuclease (New England Biolabs) 28. NotI restriction endonuclease (Takara Bio) 29. Ligation high (Toyobo) 30. BIOTAQ DNA polymerase (Bioline) 31. Ammonium acetate 32. 1-kb DNA ladder (Invitrogen) 33. pGEM-T Easy vector (Promega) 34. DH5alpha competent cells (Takara Bio)

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35. 30 adaptor: 50 -(Pu)uuAACCGCGAATTCCAG(idT)-30 (lowercase letters indicate RNA, uppercase letters indicate DNA, Pu denotes 50 -phosphorylated Urd, and idT represents 30 -inverted deoxythymidine) (synthesized by Dharmacon) 36. 50 adaptor: 50 -GACCACGCGTATCGGGCACCACGTATGCTATCGATCG TGAGATGGG-30 (synthesized by Operon Biotechnologies) 37. RT primer: 50 -GACTAGCTGGAATTCGCGGTTAAA-30 (synthesized by Operon Biotechnologies) 38. 50 PCR primer: 50 -GCGTATCGGGCACCACGTATGC-3¢ (synthesized by Operon Biotechnologies) 39. 30 PCR primer: 50 -GACTAGCTTGGTGCCGAATTCGCGGTTAAA-30 (synthesized by Operon Biotechnologies) 40. 19-nt RNA oligomer: r(CGUACGCGGAAUACUUCGA)(synthesized by Dharmacon) 41. 24-nt RNA oligomer: r(CGUACGCGGAAUACUUCGAAAUGU) (synthesized by Dharmacon) 42. 33-nt RNA oligomer: r(CCAUCGAUAAAAAAUAUGGAGAGC UUCCCGAAG) (synthesized by Dharmacon) 43. Small RNA markers: Dissolve 10 mg each of 19-, 24- and 33-nt RNA oligomers in 100 mL DEPC water 44. 2 Dye (mirVana) 45. 10 NEB buffer 3 (New England Biolabs, Ipswich, MA) 46. Calf intestinal alkaline phosphatase (New England Biolabs) 47. 10 Ligation Buffer (New England Biolabs) 48. Acetylated bovine serum albumin (Invitrogen, Carlsbad, CA; 1 mg/ml stock) 49. T4 RNA ligase (New England Biolabs) 50. 5 RT buffer (Clontech, Mountain View, CA) 51. PowerScript reverse transcriptase (Clontech) 52. 10 AmpliTaq buffer (Applied Biosystems, Foster City, CA) 53. AmpliTaq Gold DNA polymerase (Applied Biosystems) 54. 10 K buffer (New England Biolabs) 55. BanI (New England Biolabs; 20 U/mL stock) 56. Ligation High solution (Toyobo, Tokyo, Japan) 57. 10 NH4 buffer (Bioline, London, UK) 58. BioTaq DNA polymerase (Bioline) 59. pGEM-Teasy (Promega, Madison, WI)

9.2.4

Procedures

The procedures described herein are entirely from the protocols developed by Mano and Takada (2007). According to their studies, the mRAP method involves the following steps: (1) The mRAP method starts with the isolation of <200-nt RNA molecules. Concentration of the miRNA fraction is achieved by subjecting these RNA molecules of <200 nt to denaturing PAGE on a 15% gel and then eluting

9.2 Protocol

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RNAs of defined size from the gel. The resolution of size fractionation is usually higher with small RNAs of <200 nt as the starting material than with total RNA. (2) The size-selected RNAs are then dephosphorylated to prevent their ligation to each other during the next step. (3) The dephosphorylated RNA is ligated with the 30 adaptor (self-ligation of the 30 adaptor cannot occur because its 30 -end is blocked with inverted deoxythymidine) and subjected to RT with the RT primer. In this reaction, RT often adds a few C nucleotides at the 30 -end of the generated cDNAs as a result of its terminal transferase activity. The poly(C) overhang hybridizes with the 30 -end of the 50 adaptor, and the enzyme extends the cDNA product through to the end of the 50 adaptor according to the SMART reaction. The cDNAs thus obtained are subjected to PCR for amplification of the sequences derived from the size-selected RNAs. (4) Analysis of the amplification products by PAGE usually reveals three major bands of ~70, ~90, and ~120 bp. The ~70-bp band corresponds to the 50 adaptor–30 adaptor dimer, a by-product of the SMART reaction. Given that the RT primer is complementary to the 30 adaptor, they hybridize to each other and the SMART reaction may take place at the 30 -end of the RT primer without the involvement of an miRNA sequence. The ~90-bp band corresponds to the miRNA-derived cDNAs (more precisely, if the miRNA is 22 nt, then the amplicon is 92 bp). Finally, the ~120-bp band corresponds to a 30 adaptor–50 adaptor–50 adaptor trimer, another by-product of the SMART reaction that is synthesized from the first by-product and the 50 adaptor. The SMART reaction may further take place with the ~120-bp product, yielding a by-product of ~170-bp. This latter product is usually not observed in the gel because its generation is less efficient than that of the trimer. The desired band of ~90 bp is thus observed between the two major by-products of ~120 and ~70 bp. (5) The next step of mRAP is cloning of the ~90-bp DNA fragments, which are first subjected to concatamer formation with the use of the restriction endonuclease BanI. Given that nonpalindromic recognition sequences of BanI are present in the 50 and 30 PCR primers, the concatamers are formed by directional ligation in a tandem manner. Cloning (TA) into commercially available T-vectors is facilitated by filling in of the termini of the concatamers and by the addition of an A nucleotide overhang at their 30 ends by Taq DNA polymerase. The concatamers thus generated are then size-fractionated by PAGE on a 10% gel to obtain products of 4500 bp. The size-selected concatamers are cloned into a T-vector, and the resulting plasmid library is the miRNA library. (6) Check the quality of the library by nucleotide sequencing of the cDNAs and determination of the miRNA expression profile (Figs. 9.1 and 9.2).

9.2.4.1

Isolation and Fractionation of Small RNA Molecules

A small-RNA fraction is isolated from cells or tissue, the RNA molecules are separated by denaturing polyacrylamide gel electrophoresis (PAGE), and the region of the gel containing RNA molecules of 19–24 nucleotides (nt) is excised. The portion of the gel containing the sample RNA should not be stained with a dye or exposed to ultraviolet light.

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1. Purify small RNA molecules with the use of a mirVana miRNA isolation kit and the small-RNA fraction option. Other protocols or kits suitable for the enrichment of small RNAs may be used. 2. Prepare the sample mixture: 5 mL of isolated RNA (65 mg) and 5 mL of 2 Dye. 3. Prepare the size-standard mixture: 0.25 mL of size standards (synthesized RNA oligonucleotides of 19, 24, and 33 nt at a concentration of 0.1 mg/mL each), 4.75 mL of water, and 5 mL of 2 Dye. 4. Incubate each mixture at 90 C for 20 s and then place on ice. 5. Subject the sample and size-standard mixtures to PAGE on a 15% gel. 6. Cut out and stain only the marker lane with SYBR Green II or with ethidium bromide for 5–10 min and then photograph the gel aligned with a ruler. 7. Excise the portion of the sample lane containing RNA molecules of 19–24 nt. 8. Chop the isolated gel into small fragments and then transfer them to a microcentrifuge tube. 9. Add 125 mL of water to the tube and maintain it at 4 C overnight with gentle agitation. 10. Separate the gel pieces by brief centrifugation, and transfer 100 mL of the supernatant containing the small RNA molecules to another microcentrifuge tube. 11. Add 250 mL of 2-butanol to the RNA, invert the tube several times, and centrifuge the mixture briefly. 12. Check the volume of the bottom, water phase (should be <10 mL) and discard the upper, 2-butanol phase. 13. Subject the water phase to chloroform extraction followed by ethanol precipitation with 1 mL of glycogen (20 mg/mL stock) as a carrier. 14. Dissolve the purified RNA molecules in 8.75 mL of water.

9.2.4.2

Synthesis of cDNAs, PCR, and Concatamer Formation

The miRNAs isolated by gel electrophoresis are ligated at their 30 ends to a 30 adaptor with the use of RNA ligase. Complementary DNAs corresponding to the miRNAs are then synthesized with the use of reverse transcriptase and an RT primer complementary to the 30 adaptor. Given that some reverse transcriptases possess terminal deoxynucleotidyl transferase activity, the synthesized cDNA strands frequently have a small poly(C) overhang at their 30 ends. After annealing of a long 50 adaptor to such poly(C) overhangs, PCR is used to amplify the miRNA-derived cDNAs. There are usually three main types of PCR product of different sizes (~70, ~90, and ~120 bp). The ~90-bp products are the ones that contain the cDNAs; the smaller and larger products comprise dimers and trimers of the 50 adaptor and 30 adaptor (without cDNA). The desired products are purified, digested with the restriction endonuclease BanI (the target sites of the enzyme are incorporated into the PCR primers), and ligated to generate cDNA concatamers.

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Required Oligonucleotides 30 adaptor: 50 -(Pu)uuAACCGCGAATTCCAG(idT)-30 (lowercase letters indicate RNA, uppercase letters indicate DNA, Pu denotes 50 -phosphorylated uridine, and idT represents 30 -inverted deoxythymidine [Dharmacon, Chicago, IL]) 0 5 adaptor: 50 -GACCACGCGTATCGGGCACCACGTATGCTATCGATCGTGAGATGGG-30 RT primer: 50 -GACTAGCTGGAATTCGCGGTTAAA-30 50 PCR primer: 50 -GCGTATCGGGCACCACGTATGC-30 30 PCR primer: 50 -GACTAGCTTGGTGCCGAATTCGCGGTTAAA-30 Synthesis of cDNAs from miRNAs 1. Mix the small RNAs (8.75 mL) with 1 mL of 10 NEB buffer 3 and 0.25 mL of calf intestinal alkaline phosphatase in a microcentrifuge tube, and incubate the mixture at 50 C for 30 min. 2. Subject the mixture to phenol–chloroform extraction and chloroform extraction followed by ethanol precipitation in the presence of 1 mL of glycogen (20 mg/mL stock) and 0.5 mL of 100 mM 30 adaptor. 3. Isolate the RNA by centrifugation and allow it to dry before dissolving it in 3 mL of water. 4. To the dissolved RNA, add the following: 1 mL of 10 Ligation Buffer, 1 mL of acetylated bovine serum albumin (1 mg/mL stock), 1 mL of 1 mM ATP, and 3 mL of 50% dimethyl sulfoxide. Incubate the mixture at 90 C for 30 s and then place it on ice for 20 s. 5. Add 1 mL of T4 RNA ligase, and incubate the mixture at 37 C for 1 h. Perform phenol–chloroform extraction, chloroform extraction, and ethanol precipitation. Isolate the RNA by centrifugation and dissolve it in 4 mL of water. 6. Add 0.5 mL of 100 mM RT primer and 0.5 mL of 100 mM 50 adaptor. Incubate the mixture at 70 C for 2 min, and then place it on ice. 7. Add the following: 2 mL of 5 RT buffer, 1 mL of 0.1 M dithiothreitol, 1 mL of a mixture of the four deoxynucleoside triphosphates (dNTPs) each at 2.5 mM, and 1 mL of PowerScript reverse transcriptase. Incubate the resulting mixture at 42 C for 1 h. 8. Add 40 mL of water, and incubate the mixture at 72 C for 7 min. Add 160 mL of water. PCR Amplification of miRNA-Derived cDNAs 1. Mix the following: 49 mL of cDNA solution, 1706 mL of water , 245 mL of 10 AmpliTaq buffer, 245 mL of a mixture of the four dNTPs each at 2 mM, 196 mL of a mixture of the 50 PCR primer and 30 PCR primer each at 10 mM, and 9.8 mL of AmpliTaq Gold DNA polymerase. 2. Transfer 50 mL of the reaction mixture to each of 48 PCR tubes (0.5-mL scale).

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3. Perform PCR with an initial incubation at 95 C for 4.5 min; 32 cycles at 95 C for 30 s and at 65 C for 30 s; and a final incubation at 72 C for 5 min. 4. Subject each reaction mixture to sodium acetate–ethanol precipitation as follows: add 1/10th volume of 3 M NaOAc (pH5.5), mix, add two volumes of 100% ethanol, mix, and chill at 70 C for 15 min. Centrifuge the tubes, and dissolve the final precipitates in water.

Concatamer Formation 1. Subject the PCR products and size standards to PAGE on a 10% gel under nondenaturing conditions. 2. Stain the gel with ethidium bromide, and excise the portion of the gel containing DNA molecules of 90–95 bp. 3. Chop the gel portion into small fragments, transfer the fragments to a microcentrifuge tube, and add 200 mL of 0.3 M NaCl. Incubate the mixture at 37 C for P8 h. 4. Briefly centrifuge the tube and harvest the supernatant containing the cDNAs. 5. Subject the cDNA preparation to ethanol precipitation (in the presence of glycogen). Isolate the DNA by centrifugation and dissolve it in 43 mL of water. 6. Add 5 mL of 10 K buffer and 2 mL of BanI (20 U/mL stock). Incubate the mixture at 37 C for 2 h or overnight. 7. Purify the DNA with ProbeQuant G50 and then subject it to phenol-chloroform extraction, chloroform extraction, and ethanol precipitation (in the presence of glycogen). Isolate the DNA by centrifugation and dissolve it in 2 mL of water. 8. Add 2 mL of Ligation High solution, and incubate the resulting mixture at 16 C for 4 h or overnight. 9. Twenty minutes before the end of the ligation incubation, prepare the following mixture (Taq mix): 76 mL of water, 10 mL of 10 NH4 buffer, 3 mL of 50 mM MgCl2, 10 mL of a mixture of the four dNTPs each at 2 mM, and 1 mL of BioTaq DNA polymerase. 10. Incubate the Taq mix at 95 C for 10 min and then maintain it at 72 C. 11. Add the ligation mixture to the Taq mix and incubate at 72 C for an additional 30 min. 12. Subject the mixture to ammonium acetate–ethanol precipitation as follows: add 1/4th volume of 10 M ammonium acetate (i.e., 2 M final), mix, add two volumes of 100% ethanol, mix, and chill at 70 C for 15 min. Centrifuge the tubes, and dissolve the final precipitates in water. 13. Fractionate the DNA molecules by PAGE on a 10% gel under nondenaturing conditions. 14. Excise the portion of the gel containing DNA molecules of 500–2,000 bp. 15. Chop the excised region of the gel into small fragments and transfer them to a D-tube. 16. Subject the fragments to electrophoretic elution at 100 V for 4 h in Tris–borate– EDTA buffer. 17. Harvest the solution and subject it to sodium acetate–ethanol precipitation in the presence of glycogen.

References

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18. Isolate the cDNA concatamers by centrifugation and dissolve them in water. Ligate the concatamers into a TA cloning vector pGEM-Teasy for nucleotide sequencing.

9.2.4.3

Sequencing of mRAP products

1. First assess the quality of the mRAP plasmid library by sequencing the inserts of ~100 randomly isolated plasmids. The plasmid inserts typically contain two to six cDNAs. BLAST searching of the insert cDNAs against the genome sequence of interest has revealed that about one-third of the cDNAs correspond to miRNAs, one-third to ribosomal or transfer RNA, and the remaining one-third to other sequences. 2. Sequencing of mRAP products on such a scale should allow characterization of a complete body map of miRNA profiles for any organism.

9.3

Application and Limitation

mRAP combines cloning for new miRNA discovery, high-throughput profiling, and quantification of miRNA levels into one. The mRAP method can be performed in a conventional molecular biology laboratory, readily allows the processing of multiple samples in parallel, is able to detect a 1-nt difference among miRNAs and is capable of identifying both new miRNAs and mutations in known miRNAs. The major advantage of mRAP is its high sensitivity. mRAP may currently be the best choice for obtaining expression profiles for both known and unknown miRNAs or for identification of sequence alterations in miRNAs with small amounts of starting material. Analysis of mRAP products with recently developed high throughput sequencing systems such as pyrosequencing (Ronaghi 2001) would be a robust approach to extensively profile miRNAs in a given tissue. In addition, modification of mRAP may allow the identification of Piwi-interacting RNAs and other small RNAs which are slightly larger than miRNAs. mRAP and other emerging techniques for highly sensitive miRNA profiling should contribute to the complete understanding of RNA-mediated regulation of protein-coding genes or vice versa. However, the mRAP method is a complex, multistep procedure, and may not be suitable for clinical use as a diagnostic tool.

References Lagos-Quintana M, Rauhut R, Yalcin A, Meyer J, Lendeckel W, Tuschl T (2002) Identification of tissue-specific microRNAs from mouse. Curr Biol 12:735–739 Lee RC, Ambros V (2001) An extensive class of small RNAs in Caenorhabditis elegans. Science 294:862–864

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Mano H, Takada S (2007) mRAP, a sensitive method for determination of microRNA expression profiles. Methods 43:118–122 Ronaghi M (2001) Pyrosequencing sheds light on DNA sequencing. Genome Res 11:3–11 Takada S, Berezikov E, Yamashita Y, Lagos-Quintana M, Kloosterman WP, Enomoto M, Hatanaka H, Fujiwara S, Watanabe H, Soda M, Choi YL, Plasterk RH, Cuppen E, Mano H (2006) Mouse microRNA profiles determined with a new and sensitive cloning method. Nucleic Acids Res 34:e115 Takada S, Mano H (2007) Profiling of microRNA expression by mRAP. Nat Protoc 2:3136–3145

Part VI Cloning Methods

Chapter 10

miRNA Serial Analysis of Gene Expression (miRAGE or SAGE)

Abstract The miRNA serial analysis of gene expression (miRAGE) combines direct miRNA cloning and SAGE (Science 270:484–487, 1995). Similar to traditional cloning approaches, miRAGE starts with the isolation of 18- to 26base RNA molecules to which specifically designed linkers are ligated, and which are reverse-transcribed into cDNA. However, subsequent steps, including amplification of the complex mixture of cDNAs using PCR, tag purification, concatenation, cloning, and sequencing, have been performed by using SAGE methodology optimized for small RNA species (Science 294:853–858, 2001). A combination of miRNA cloning and expression profiling using MiRAGE allows for the discovery of new miRNAs. MiRAGE can be used for Genome-wide miRNA expression analysis. This approach has the advantage of generating large concatemers that can be used to identify as many as 35 tags in a single sequencing reaction. The main limitations of the miRAGE techniques include intensive labor and complicated procedures with the need for enrichment of short RNAs by fractionation. This approach was developed in Velculescu’s laboratory (Proc Natl Acad Sci USA 103:3687–3692, 2006) based on their earlier work the analysis of mRNA expression using a similar technique (Science 270:484–487, 1995). The same strategy has later been applied to detect miRNA precursors (BMC Genomics 7:285, 2006).

10.1

Introduction

The miRNA serial analysis of gene expression (miRAGE) combines direct miRNA cloning and SAGE (Velculescu et al. 1995). Similar to traditional cloning approaches, miRAGE starts with the isolation of 18- to 26-base RNA molecules to which specifically designed linkers are ligated, and which are reverse-transcribed into cDNA. However, subsequent steps, including amplification of the complex mixture of cDNAs using PCR, tag purification, concatenation, cloning, and sequencing, have been performed by using SAGE methodology optimized for small RNA species (Lagos-Quintana et al. 2001). The first half of this protocol resembles the Z. Wang and B. Yang, MicroRNA Expression Detection Methods, DOI 10.1007/978-3-642-04928-6_10, # Springer-Verlag Berlin Heidelberg 2010

173

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10 miRNA Serial Analysis of Gene Expression (miRAGE or SAGE)

GTAC

AAAAAA TTTTTT Nla III Digestion

GTAC GTAC

Cloning, sequencing, bioinformatics analyses, and expression profiling

AAAAAA TTTTTT AAAAAA TTTTTT AAAAAA TTTTTT

Linker Ligation & BsmF I Digestion CATG GTAC

CATG

CATG GTAC

CATG

Ligation CATG GTAC CATG GTAC CATG GTAC

CATG GTAC CATG GTAC CATG GTAC

AAAAAA TTTTTT AAAAAA TTTTTT AAAAAA TTTTTT

AAAAAA TTTTTT AAAAAA TTTTTT AAAAAA TTTTTT

CATG GTAC

Nla III Digestion CATG GTAC

CATG GTAC

Ditag & Amplification CATG GTAC

Ligation

CATG GTAC

Fig. 10.1 Schematic diagram depicting the SAGE procedures

bead-based method in which linkers are ligated to both the 50 and 30 ends of enriched small RNAs for reverse transcription. A PCR reaction is carried out on the resulting cDNA mix, also with the help of biotinlyated primers. Deviating from the beadbased method; the linkers which now contain biotins are cleaved from the PCR products. The mixture containing amplified small RNA sequence and biotinlyated linkers are run through a column of streptavidin-coated beads for purification. Streptavidin acts as a magnet to bind the biotin-tagged linkers. The eluted product, at least in theory, is purified small RNAs. The small RNAs are concatenated, cloned, and sequenced for analysis (Cummins et al. 2006) (Figs. 10.1 and 10.2).

10.2

Protocol

10.2.1 Materials 1. 2. 3. 4.

RNagents kit (Promega) 15% polyacrylamide TBE/Urea Novex gels (Invitrogen) 18-gage needle-pierced centrifuge tube 75% ethanol

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Total RNA extraction

Enrich small RNAs

Dephosphorylation of small RNAs

Design and synthesize miRAGE 3' Linker

Ligation of miRNA with miRAGE 3' Linker

Design and synthesize miRAGE RT primer

Reverse transcription

Tag amplify, isolate, purify, concatenate, clone, and sequence

Bioinformatics analysis I: Grouping and comparing miRAGE tags to known RNAs

Bioinformatics analysis II: Secondary structure analysis and hairpin stability scoring of candidate miRNAs

Bioinformatics analysis III: Determination of hairpin conservation and homology of candidate miRNAs to existing miRNAs

Fig. 10.2 Flowchart of miRAGE analysis for miRNA expression detection. According to Cummins et al. (2006)

5. 6. 7. 8. 9. 10. 11.

SYBR Gold Nucleic Acid Gel Stain (Molecular Probes) Costar Spin-X Centrifuge Tube Filter (VWR Scientific) 100% EtOH Phred sequence analysis software (CodonCode, Dedham, MA) Sage2000 software package Spin-filter units (SpinX, Corning or Mermaid, Qbiogene) Electrocompetent TOP10 bacterial cells (Invitrogen)

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12. Magnetic-bead-based system (mMACS mRNA Isolation Kit for Total RNA, Miltenyi Biotec) 13. NotI-oligo dT primer 14. pZErO-1 plasmid (Invitrogen) 15. 21 G needle

10.2.2 Instruments 1. Rotisserie-style rotator 2. Sage2000 software package

10.2.3 Reagnets 1. miRAGE 30 linker: 50 -phosphate-UCUCGAGGUACAUCGUUdAdGdAdAdGdCdTdTdGdAdAdTdTdCdGdAdGdCdAdGdAdAdAN3-30 2. miRAGE 50 linker: 50 -dTdTdTdGdGdAdTdTdTdGdCdTdGdGdTdGdCdAdGdTdAdCdAdAdCdTdAdGdGdCdTdTdACUCGAGC 3. 18-base RNA standard: 50 -phosphate-ACGUUGCACUCUGAUACC 4. 26-base RNA standard: 50 -phosphate-CCGGUUCAUCACGUCUAAGAAUCAUG. DNA oligonucleotides were obtained from Integrated DNA Technologies (San Jose, CA) 5. miRAGE reverse transcription primer: 50 -TTTCTGCTCGAATTCAAGCTT CT; LongSage PCR primer (forward) 6. 50 -biotin-TTTTTTTTTGGATTTGCTGGTGCAGTACA-30 7. LongSage PCR primer (reverse): 50 -biotin-TTTTTTTTTCTGCTCGAATTCAAGCTTCT-30 8. 0.3 M NaCl 9. Calf intestinal alkaline phosphatase (NEB, Beverly, MA) 10. Phenol/chloroform 11. T4 RNA ligase (NEB) 12. T4 polynucleotide kinase (NEB) 13. SuperScript II RT (Invitrogen) 14. XhoI endonuclease (NEB) 15. SYBRGreen I (Molecular Probes) 16. BamHI/pZErO-1 17. Yeast tRNA 18. Low Salt LB agar (Lennox L) 19. Zeocin (25–50 mg/mL)

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20. LoTE, a low salt Tris-EDTA buffer: [3 mM Tris-HCl pH7.5 and 0.2 mMEDTA pH7.5] 21. Binding buffer: [2 M NaCl, 50 mM EDTA, pH8.0 22. 10 RNaseONE buffer and 25 units RNaseONE (Promega) 23. 2 GC-I buffer (Takara LA-PCR kit) 24. RNaseIN inhibitor (Promega) 25. 1 M NaOAc (pH6.1) 26. 10% SDS 27. 10 mM long-arm biotin hydrazide 28. Wash buffer: [10 mM Tris-HCl pH7.5, 0.2 mM EDTA, 10 mM NaCl, 20% (v/v) glycerol, and 40 mg/mL Yeast tRNA] 29. Alkaline hydrolysis mix: [50 mM NaOH and 5 mM EDTA pH8.0] 30. 1 M Tris-HCl pH7.5

10.2.4 Procedures The protocols described in this section are essentially the same as reported in the study from Velculescu’s laboratory (Cummins et al. 2006) (see Figs. 10.1 and 10.2). The initial protocol is to enrich small RNAs from the total RNA samples, followed by ligating linkers to both the 50 and 30 ends of the enriched small RNAs for reverse transcription. The next step is to carry out a PCR reaction on the resulting cDNA mix, with biotinlyated primers. The linkers, which now contain biotins, are cleaved from the PCR products. The mixture containing amplified small RNA sequence and biotinlyated linkers are run through a column of streptavidin-coated beads for purification. Streptavidin acts as a magnet to bind the biotin-tagged linkers. The eluted product is presumably, purified small RNAs. The small RNAs are finally concatenated, cloned, and sequenced for analysis (Cummins et al. 2006).

10.2.4.1

miRAGE Approach for miRNA Identification

Small RNA Isolation and Linker Ligation 1. Isolate total RNA from cell lines/tissue samples by using the RNagents kit following the manufacturer’s protocol, with the exception that no final 75% ethanol wash to be performed. 2. Fractionate RNA of the 18- to 26-base size range by electrophoresing 1 mg of total RNA alongside 18- and 26-base RNA standards on two 15% polyacrylamide TBE/Urea Novex gels at 180 V for 70 min. The 18- and 26-base RNA standards are used through all subsequent ligation steps to serve as size standards for gel purification.

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3. Visualize RNAs ranging from 18 to 26 bases in length with SYBR Gold Nucleic Acid Gel Stain, excise from the gel, pulverize by spinning at high speed through an 18-gage needle-pierced centrifuge tube, and gel-extract by incubating the gel slices in 0.3 M NaCl at 4 C on a rotisserie-style rotator for 5 h 4. Transfer the contents into a Costar Spin-X Centrifuge Tube Filter, spin into a fresh tube, EtOH-precipitate (by adding three volumes of 100% EtOH), and resuspend in water 5. Dephosphorylate small RNAs with calf intestinal alkaline phosphatase at 50 C for 30 min, phenol/chloroform-extract, re-EtOH precipitate, and ligate to the miRAGE 30 Linker with T4 RNA ligase at 37 C for 1 h 6. After gel purification of 58- to 66-base RNA products and EtOH precipitation (as described above), phosphorylate the samples with T4 polynucleotide kinase at 37 C for 30 min, phenol/chloroform-extracte, EtOH-precipitate, and ligate (as above) to the miRAGE 50 Linker.

Tag Amplification, Isolation, Concatenation, Cloning, and Sequencing 1. After gel purification of RNA products ranging from 98 to 106 bases, reverse transcribe the ligation products by using miRAGE reverse transcription primer and SuperScript II RT for 50 min at 45 C. 2. Next, amplify, isolate, purify, concatenate, clone, and sequence tags with protocols nearly identical to those performed in LongSAGE and Digital Karyotyping (see Sects 10.2.1.6 and 10.2.1.7), except that miRAGE PCR products range in size from 110 to 118 bp, and miRAGE tags (not ditags) are to be released from linkers with XhoI endonuclease. 3. Sequencing analyze the concatemer clones. 4. Sort out the resulting sequence files by using phred sequence analysis software, and extract 18- to 26-bp tags by using the sage2000 software package, which identifies the fragmenting enzyme site between tags, extracts intervening tags, and records them in a database.

10.2.4.2

Bioinformatic Analyses of miRAGE Tags

Grouping and Comparing miRAGE Tags to Known RNAs 1. Assemble all tags sharing a common set of 11 of 12 core internal sequence elements into groups containing all related members. 2. Further analyze the tag with the most counts in each group. Grouping should facilitate analysis by (1) eliminating rare sequencing errors and (2) removing trivial miRNA variants, because miRNAs are known to display both 50 and 30 variation.

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3. Compare the tags to databases of known RNA sequences (miRNAs, mRNAs, rRNAs, etc.), using blast, and remove those tags matching known sequences from further analysis. 4. Then compare the tags obtained by miRAGE with public databases.

Secondary structure analysis and hairpin stability scoring of candidate miRNAs 1. To determine potential miRNA precursor structures, compare each tag to the human genome sequence. For tags with perfect matches, extract a total of 75 bp (60 þ 15 bp) of flanking genomic sequence around each tag. As there are two possible precursors for each tag (i.e., the tag can be located on the 50 or 30 arm of a putative hairpin), extract pairs of theoretical precursors from the human genome at the position of each tag and carry through, the following analysis. 2. Determine the secondary structure and free energy of folding for each pair of precursor structures by using mfold 3.2 (Zuker 2003) and compare it to values obtained for known miRNAs. The values used for thermodynamic evaluation are the free energy of folding of each precursor sequence (DGfolding) and the difference of DGfolding between the two possible precursors (DDGfolding). A miRNA precursor structure should have to have either DGfolding 29 or 29 < DGfolding 22 and DDGfolding > 5. In cases where both precursors fulfilled these criteria, the member of each pair with the lowest DGfolding will be further considered. 3. Precursors that have not been excluded up to this point should be analyzed to determine, whether they conform to generally acceptable miRNA base-pairing standards (base-pairing involving at least 16 of the first 22 nucleotides of the miRNA and the other arm of the hairpin).

Determination of Hairpin Conservation and Homology of Candidate miRNAs to Existing miRNAs 1. Using the University of California at Santa Cruz phastCons database or other equivalents, they classify all candidate miRNAs as either “conserved” or “nonconserved.” This database has scores at each nucleotide, in the human genome that correspond to the degree of conservation of that particular nucleotide in chimpanzee, mouse, rat, dog, chicken, pufferfish, and zebrafish. A hairpin is to be defined as conserved if the average phastCons conservation score over the seven species in any 15-nt sequence in the hairpin stem is at least 0.9. 2. Compare the candidate miRNAs to the miRBase database using the SSEARCH search algorithm, and obtain the expected values for each; Tags with the expected values of <0.05 are considered having the homology to existing miRNAs.

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10.2.4.3

10 miRNA Serial Analysis of Gene Expression (miRAGE or SAGE)

miRNA Microarray Expression Analysis

1. Use miRNA microarray technique to profile the miRNA cloned above

10.2.4.4

Quantitative RT-PCR (qRT-PCR) Expression Analysis

1. Use qRT-PCR to confirm the results from miRNA microarray

10.2.4.5

50 Terminal LongSAGE

First strand cDNA synthesis and full-length (-)DNA selection Total RNA is prepared using Trizol (Invitrogen), as per the manufacturer’s protocols. mRNA (polyA RNA) is then purified from this total RNA using a magneticbead based system. Typical yields are 20–50 mg mRNA from 1 mg total RNA. In the first step, cDNA is synthesized by reverse transcription with a NotI-oligo dT primer. Then, the first strand cDNA/RNA hybrids are subjected to a full-length enrichment procedure by the biotinylation-based cap-trapper approach. 1. The following are mixed: NotI-dT primer (7 mg/mL) 2 mL and PolyA RNA (20 mg) 18 mL 2. The obtained solution is heated to 65 C for 10 min and 42 C for 1 min. 3. Then, spin tube in microfuge and the following substances are preheated and added to the RNA/primer mix: 2 GC-I buffer RNaseIN inhibitor 10 mM dNTP (with methyl-dCTP instead of dCTP) Saturated trehalose in DEPC-H2O 4.9 M sorbitol in DEPC-H2O Superscript II RT (200U/mL)

75 mL 1 mL 4 mL 10 mL 26 mL 15 mL

4. The obtained solution is incubated at 42 C for 40 min, 50 C for 20 min and 55 C for 20 min 5. 2 mL of proteinase K (20 mg/mL) is then added to degrade the RT: the obtained solution is incubated at 45 C for 15 min followed by phenol/chloroform extraction and isopropanol precipitation (do not use glycogen as the subsequent selection depends on diol oxidation) 6. The RNA/(-)DNA heteroduplex is resuspended into 44.5 mL of ddH2O 7. 3 mL of 1.1 M NaOAc pH 4.5 and 2.5 mL of 100 mM NaIO4 are added to oxidize the diol structures of the mRNA: 50 mL of the reaction solution is incubated on ice in the dark for 45 min followed by adding 0.5 mL of 10% SDS, 11 mL of 5 M NaCl and 61 mL of isopropanol to precipitate the RNA/(-)DNA;

10.2 Protocol

181

8. The precipitated RNA/DNA is resuspended in 50 mL of ddH2O 9. 5 mL 1 M NaOAc (pH6.1), 5 mL 10% SDS and 150 mL 10 mM long-arm biotin hydrazide (freshly prepared) are added to biotinylate the RNA: the reaction is incubated at room temperature in the dark, overnight 10. The biotinylated RNA/(-)DNA is precipitated by adding 5 mL 5 M NaCl, 75 mL 1 M RNase-free NaOAc (pH6.1), and 750 mL 100% EtOH or 200 mL of 100% isopropanol: incubate at 80 C for 30 min, and spin 14 krpm at 4 C for 30 min 11. The pellet is washed with 70% EtOH (in DEPC-treated H2O) and 14 krpm spin is carried out at 4 C for 10 min. The pellet is air-dried and resuspended in 400 mL DEPC-H2O 12. RNaseONE digestion to select for full-length first-strand cDNA: 50 mL 10 RNaseONE buffer and 25 units RNaseONE per mg of starting mRNA are added. The obtained solution is incubated at 37 C for 30 min 13. 10 mL of 10 mg/mL Yeast tRNA (Ambion) and 150 mL of 5 M NaCl are added to stop the reaction. The volume at this stage is approximately 600 mL 14. While biotinylating the RNA/(-)DNA heteroduplex, the streptavidin Dynabeads (Dynal) are prepared as follows: 400 mL of M280 streptavidin beads are pipetted into an RNase-free Eppendorf tube, the beads are then placed on a magnet, left for at least 30 s, after which the supernatant is removed. The beads are resuspended in 400 mL 1 binding buffer 15. The tube is placed on a magnet, left for at least 30 s, after which the supernatant is removed. The 1 binding buffer wash is repeated two more times. The beads are resuspended in 400 mL 1 binding buffer with 100 mg of Yeast tRNA, and then incubated at 4 C for 30 min with occasional mixing, to block all nonspecific binding sites 16. The tube is placed on a magnet, left for at least 30 s, and the supernatant is then removed. The beads are washed with 1 binding buffer for three times, and the supernatant removed just before addition of the RNA/(-)DNA heteroduplex 17. The prepared beads and RNA/(-)DNA heteroduplex are mixed (the binding condition is approximately 1 M NaCl) 18. The mixture is rotated at room temperature for 30 min to allow binding. The tube is placed on a magnet stand, left for at least 30 s, and the supernatant then removed (the supernatant was saved as “unbound”) 19. The beads are washed two times with 400 mL of 1 binding buffer. 20. This is followed by a wash with 400 mL of 0.4% (w/v) SDS, plus 50 mg/mL Yeast tRNA, and another wash with 400 mL of 1 wash buffer. 21. Finally, they are washed with 400 mL of 50 mg/mL Yeast tRNA. For all washes the tube is placed on a magnet stand, left for at least 30 s, and the supernatant is removed 22. The first strand cDNA is released by alkaline hydrolysis of RNA. After removing the supernatant, the following is added: 50 mL alkaline hydrolysis mix, and the tube is rotated at 65 C for 10 min 23. The tube is then placed on a magnet stand, and the supernatant (now containing the eluted full-length first-strand cDNA) transferred to another tube containing

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50 mL 1 M Tris-HCl pH7.5 for neutralization. The hydrolysis procedure is repeated two more times. The final volume of the supernatant is approximately 300 mL (containing the full-length first-strand cDNA (FL(-)cDNA)) 24. The cDNA is extracted by phenol/chloroform and precipitated using 1 mL ethanol with glycogen, then resuspended in 10 mL LoTE. The resuspended FL (-)cDNA is then split into 2 aliquots “A” and “B” of 5 mL each. The reason why two different adapters have to be used is that otherwise panhandle structures will form, and inihibit subsequent PCR.

Second Strand cDNA Synthesis 1. The following reagents are added on ice: FL () cDNA aliquot A 1.6 mg Adapter A(N5) 0.4 mg Adapter A(N6) Soln II (Takara ligation kit ver2) Soln I (Takara ligation kit ver2)

5 mL 4 mL 1 mL 10 mL 20 mL

And in a separate tube: FL () cDNA aliquot B 1.6 mg Adapter B(N5) 0.4 mg Adapter B(N6) Soln II (Takara ligation kit ver2) Soln I (Takara ligation kit ver2)

5 mL 4 mL 1 mL 10 mL 20 mL

2. From this step onwards, each aliquot A and B is processed identically and in parallel. The cDNA and linker-adapter mixture is incubated at 16 C overnight, and then the following are added to each of tubes A and B on ice: dH2O 10 ExTaq buffer (Takara) 2.5 mM dNTP (Takara) ExTaq polymerase (Takara)

20 mL 8 mL 8 mL 4 mL

3. Incubate the mixture in a preheated thermocycler at 65 C, for 5 min; at 68 C, for 30 min; and at 72 C, for 10 min, followed by phenol/chloroform extraction and ethanol precipitation with glycogen, and resuspended in 40 mL dH2O.

10.2 Protocol

183

Formation of 50 LongSAGE tags 1. Immbolize the full-length double-stranded cDNA (FL ds cDNA) at the 30 -end so that MmeI digestion can be used to release 50 LongSAGE tags. NotI digestion is used to expose a cohesive site at the 30 -end of the ds cDNA, to which a biotinylated adapter can be ligated. 2. The following are added on ice:

FL ds cDNA NEB Buffer NotI (10 units/mL) (NEB)

40 mL 35 mL 4 mL

3. Incubate the obtained solution at 37 C for 2 h, then at 65 C for 15 min. 4. To remove unwanted digestion products and buffer salts before adapter ligation, Qiaquick-spin PCR purification column (Qiagen) is used as per the manufacturer’s protocols, and the DNA eluted in 40 mL of EB buffer. 5. The following are added in a microfuge tube: Purified, 30 NotI-cohesive ended ds cDNA Adapter C (1 mg/mL) 10 ligation buffer (Invitrogen) T4 DNA ligase (5 units/mL)

40 mL 10 mL 6 mL 4 mL

6. Incubate the reaction at 16 C overnight, then at 65 C for 15 min. (It is important at this stage to remove all unligated Adapter C, as this will otherwise compete for binding to the streptavidin beads. To do this, and simultaneously select for larger sized ds cDNA, cDNA size fractionation columns (Invitrogen) or ChromaSpin 400 columns (Clontech) are used. 7. In the meantime, prepare 400 mL MyOne streptavidin Dynabeads in kilobase binding buffer (Dynal) as per the manufacturer’s protocols. 8. Immobilize the purified, size-fractionated ds cDNA via the 30 -end to the prepared streptavidin Dynabeads by rotating for 3 h at room temperature. 9. The bound cDNA is then equilibrated in 1 NEB Buffer 4 by washing 4 with 500 mL buffer, then resuspended in 100 mL 1 NEB Buffer 4 (with 40 mM SAM). Two mL (8 units) of MmeI was added, and the reaction incubated at 37 C for 1 h with occasional agitation. This MmeI digestion releases 50 LongSAGE tags from A and B into the solution, so the supernatant is recovered. 10. Wash the beads with 100 mL of ddH2O and the supernatant is pooled and phenol/chloroform extracted and ethanol precipitated in the presence of glycogen and ammonium acetate. Resuspend the DNA in 5 mL LoTE. To avoid denaturation, the tags should be kept on ice.

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10 miRNA Serial Analysis of Gene Expression (miRAGE or SAGE)

Formation of 50 LongSAGE ditags and scaling-up using PCR 50 LongSAGE tags from A 50 LongSAGE tags from B 10 T4 DNA ligase buffer T4 DNA ligase (5 units/mL) dH2O

5 5 2 2 6

mL mL mL mL mL

1. Incubate the ligation reaction at 16 C overnight; no heat inactivation. 2. To optimize the conditions needed to retrieve the ligated ditags using PCR, Make serial dilutions of the ligation reaction from 1:10 to 1:80. 3. Set up PCR reactions including no-ligase and no DNA controls, and use biotinylated PCR primers to facilitate subsequent purification of amplicons. Template Primer P1 (10 mM stock) Primer P2 (10 mM stock) 25 mM MgCl2 10 PCR buffer (Invitrogen) 10 mM dNTP Platinum Taq dH2O

1 mL 1 mL 1 mL 1.5 mL 5 mL 1 mL 0.5 mL 39 mL

Thermocycling conditions: Step 1: 94 C Step 2: 94 C Step 3: 55 C Step 4: 72 C

2 min 30 sec 1 min 1 min

Repeat steps 3–4, 30–40 cycles (to be determined empirically) Step 5: 72 C Hold at 16 C

8 min

4. The PCR products are electrophoresed on 2% agarose, and the conditions that produce the most specific result are selected for large-scale amplification of the 50 LongSAGE ditags, usually 4 96 well reactions. 5. After PCR, the 4 96 reactions are pooled, phenol/chloroform extracted and ethanol precipitated, then resuspended in 200 mL LoTE buffer. 6. The required amplicons corresponding to the 120 bp ditags are gel-purified by electrophoresis on a large 15 17 cm 12% TBE-PAGE gel, and the 120 bp ditag band excised after staining in SYBRGreen I. 7. An easy way to elute the excised DNA is to place each gel slice into a 600 mL microfuge tube, the bottom of which has been pierced with a 21 G needle. Each tube is then placed into a larger 1.7 mL tube, and the tubes centrifuged at 14 krpm for 10 min at 4 C. This effectively homogenizes the gel slices, which are collected at the bottom of each 1.7 mL tube. For elution, add 200 mL LoTE buffer to each sample, incubate at 65 C for 2 h. and then separate the eluted DNA from gel residue using spin-filter units.

10.2 Protocol

185

8. Precipitate the collected, purified DNA using ethanol and resuspend in 100 mL LoTE.

Formation of 50 bp Cohesive Ditags and Concatenation Each 120 bp now has the generic structure 50 -Biot-GTAGGCCGATACTCCGTGTCATTACTAGGCTTAGGATCCGAC(G)NNNNNNNNNNNNNNNNNNNXXX XXXX ...... 30 -CATCCGGCTATGAGGCACAGTAATGATCCGAATCCTAGGCTG(C)NNNNNNNNNNNNNNNNNXXXXXXXXX ...... ......XXXXXXXXXX(C)GTCGGATCCATGTGTGATAGATGAGCTAGAGC CTGAGTGA-30 ......XXXXXXXXXX(G)CAGCCTAGGTACACACTATCTACTCGATCTCGGACTCACT-Biot-50 1. To generate cohesive ditags for subsequent concatenation, remove the 50 and 30 ends using BamHI: PCR product (120 bp ditag) 10 BamHI buffer (NEB) 100 BSA BamHI (20 U/mL, NEB) dH2O to

40–70 mg 100 mL 10 mL 10 mL 1 mL

(The choice of value of 40–70 mg of DNA was arbitrary, but should be sufficient to produce several hundred ng of 50 bp cohesive ditag DNA). 2. The 1 mL reaction is divided into 10 100 mL aliquots for more efficient digestion, and incubated at 37 C for 2 h. Heat inactivation is avoided, but phenol/chloroform extraction and ethanol precipitation can be performed. 3. Then, the pellet comprising 50 bp cohesive ditags and the rest of the digestion products is resuspended in 100 mL LoTE buffer. The generic 50 bp ditag structure is: 50 -GATCCGAC(G)NNNNNNNNNNNNNNNNNNNXXXXXXXXXXXXXX XXX(C)GTCG-30 0 3 -GCTG(C)NNNNNNNNNNNNNNNNNXXXXXXXXXXXXXXXXXXX (G)CAGCCTAG-50 4. To remove the unwanted, biotinylated digestion products (the 50 and 30 -ends of the 50 bp cohesive ditags), 250 mL M280 beads are washed, resuspended in 100 mL 1 binding buffer, and added to the 100 mL DNA above. 5. After rotation for 30 min at room temperature, the supernatant is removed, ethanol precipitated, and resuspended in 40 mL LoTE. 6. To ensure complete removal of all biotinylated termini (that would otherwise inhibit subsequent concatenation), the purified 50 bp cohesive ditag DNA above is gel-purified using a large 12% TBE-PAGE gel. The 50 bp band is excised and purified as mentioned previously.

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10 miRNA Serial Analysis of Gene Expression (miRAGE or SAGE)

7. To generate concatemers, the following is added to a 1.7 mL microfuge tube: 50 bp cohesive ditag DNA 10 T4 DNA ligase buffer T4 DNA ligase (5 units/ mL) dH2O to

8. 9. 10. 11. 12.

100–400 ng 2 mL 2 mL 20 mL

The reaction is incubated at 16 C for 1–3 h. Add 4 mL of 6 loading buffer to the reaction and heat the entire sample at 65 C for 15 min. The sample is then loaded in a single well of an 8% TBE-PAGE minigel and run at 200 V for 1 h, or until Bromophenol Blue is about 2 cm from the bottom. Excise the smear of ligation products as two or more fractions, e.g., 500– 750 bp; 750–1,000 bp; >1,000 bp. Perform elution of DNA from the gel pieces as described earlier. Extract the eluate with phenol/chloroform, then ethanol precipitate, and resuspend the DNA pellet in 6 mL LoTE.

Cloning of Concatemers 1. Prepare the cloning vector by digesting 2 mg of pZErO-1 plasmid DNA with 10 units of BamHI for 3 h at 37 C. Phenol/chloroform extract the digested DNA and ethanol precipitate, then resuspend in LoTE at a concentration of approximately 50 ng/mL. 2. The ligation reaction is performed as follows: 3. Incubate at 16 C overnight, with vector self-ligation performed in parallel as a control. Concatemer DNA BamHI/pZErO-1 10 ligase buffer T4 DNA ligase (5 U/mL) dH2O

6 1 1 1 1

mL mL mL mL mL

4. Purify the ligation products before electroporation by phenol/chloroform extraction followed by ethanol precipitation; the pellet is washed three times with 75% ethanol before resuspending in 20 mL LoTE. 5. 2 mL of this DNA is used to transform 50 mL of electrocompetent TOP10 bacterial cells. 6. After recovery in 1 mL LB media, plate 50 mL on a small (100 mm) agar plate (containing Low Salt LB agar plus Zeocin (25–50 mg/mL) and incubate overnight at 37 C. 7. As a background control, bacteria are plated out that have been similarly transformed with the vector self-ligation reaction above. The background is usually between 1–5%.

10.3 Application and Limitation

187

50 LongSAGE Library Quality Check 1. The following day, several (24–48) colonies are picked to check for insert size by PCR. For each reaction, a single colony was picked into a PCR tube containing: Thermocycling conditions: 10 HiFi buffer 10 mM dNTP PMR011 (10 mM) PMR012 (10 mM) Eppendorf TripleMaster polymerase dH2O

2 mL 0.4 mL 1 mL 1 mL 0.2 mL 11.4 mL

2. Visualize the PCR products on a 1% agarose gel (Note: the primer pair PMR011/ Step 1: 95˚C Step 2: 95˚C Step 3: 55˚C Step 4: 72˚C Repeat steps (2–4), 24 Step 5: 72˚C Hold at 16˚C

2 min 30 sec 1 min 3 min 8 min forever

PMR012 gives a band of approximately 300 bp in the absence of any cloned insert. If the quality of the library thus produced appears good, the remaining transformation mixture can be plated out on large agar plates (we use 20 20 cm Q-trays, Genetix) in preparation for DNA sequencing analysis).

10.3

Application and Limitation

Direct miRNA cloning strategies identified many of the initial miRNAs and demonstrated that miRNAs are found in many species. miRAGE offers several advantages in miRNA research: 1. Combination of miRNA cloning and expression profiling using miRAGE allows for the discovery of new miRNAs. For example, sequence analysis of 273,966 small RNA tags from human colorectal cells using the miRAGE methods allowed Cummins et al. (2006) to identify 200 known mature miRNAs, 133 novel miRNA candidates, and 112 previously uncharacterized miRNA* forms. The expression level of the miRNAs detected by miRAGE ranges over four orders of magnitude (from 23,431 observations for miR-21 to 20 miRNAs that were observed only once), suggesting that this approach can detect miRNAs present at varied expression levels. The identified miRNA tags matches 200 of the mature miRNAs present in the public miRBase database, and 52 of these are expressed at significantly different levels between tumor cells and normal

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10 miRNA Serial Analysis of Gene Expression (miRAGE or SAGE)

colonic epithelium. In addition to detecting known or predicted miRNAs, 1,411 of the miRAGE tags represent 100 previously unrecognized miRNA* forms of known miRNAs. miRNA* molecules correspond to the short-lived complementary strand present in initial miRNA duplexes. 2. miRAGE can be used for genome-wide miRNA expression analysis. 3. This approach has the advantage of generating large concatemers that can be used to identify as many as 35 tags in a single sequencing reaction, whereas existing cloning protocols analyze on an average approximately five miRNAs per reaction (Lagos-Quintana et al. 2001). The main limitations of the miRAGE techniques include (1) Labor intensive; (2) Complicated procedures with the need for enrichment of short RNAs by fractionation; (3) Requirement of a large amount (1 mg) total RNA as a starting material, and (4) The throughput of this approach is low, and cloning approaches have appeared to approach saturation (Lagos-Quintana et al. 2001).

References Cummins JM, He Y, Leary RJ, Pagliarini R, Diaz LA Jr, Sjoblom T, Barad O, Bentwich Z, Szafranska AE, Labourier E, Raymond CK, Roberts BS, Juhl H, Kinzler KW, Vogelstein B, Velculescu VE (2006) The colorectal microRNAome. Proc Natl Acad Sci USA 103:3687–3692 Lagos-Quintana M, Rauhut R, Lendeckel W, Tuschl T (2001) Identification of novel genes coding for small expressed RNAs. Science 294:853–858 Velculescu VE, Zhang L, Vogelstein B, Kinzler KW (1995) Serial analysis of gene expression. Science 270:484–487 Zuker M (2003) Mfold web server for nucleic acid folding and hybridization prediction. Nucleic Acids Res 31:3406–3415

Part VII Nanoparticle Methods

Chapter 11

Electrocatalytic Nanoparticle Tags Technique for High-Sensitivity miRNA Expression Analysis

Abstract The Electrocatalytic Nanoparticle Tags (ENT) technique for miRNA detection is based on the utilization of an indium tin oxide electrode and the nanoparticle tags (isoniazid-capped OsO2 nanoparticles). It involves four steps, starting with the immobilization or coating of oligonucleotide capture probes (antisense to miRNAs for testing) on the electrode. Total RNA sample containing miRNAs, prepared from the tissues or cells of interest, is treated with periodate. The periodate-treated miRNAs are then hybridized with the oligonucleotide capture probes on the electrode. Finally, the nanoparticle tags are brought to the electrode through a condensation reaction to chemically amplify the signal. The resulting electrode exhibits electrocatalytic activity toward the oxidation of hydrazine at 0.10 V, and reduces the oxidation overpotential by as much as 900 mV, which can be readily detected. A detection limit of 80 fM in 2.5-mL droplets and a linear current-concentration relation up to 200 pM are obtained following a 60-min hybridization. Successful attempts are made in the miRNA expression analysis of HeLa cells. This technique was initially developed by Gao and Yang from the Institute of Bioengineering and Nanotechnology, Singapore (Anal Chem 78:1470– 1477, 2006).

11.1

Introduction

Though not yet mainstream in the area of biotechnology, nanotechnology has found its way into the miRNA profiling realm. Three nanoparticle methods have been developed, all requiring base pairing of nucleic acids but not sample amplification. The first example is the Electrocatalytic Nanoparticle Tags (ENT) technique (Gao and Yang 2006) based on the utilization of an indium tin oxide electrode and the nanoparticle tags (isoniazid-capped OsO2 nanoparticles). It involves four steps, starting with the immobilization or coating of oligonucleotide capture probes (antisense to miRNAs for testing) to the electrode, periodate treating of RNA, hybridization of the periodate-treated miRNAs with the oligonucleotide capture Z. Wang and B. Yang, MicroRNA Expression Detection Methods, DOI 10.1007/978-3-642-04928-6_11, # Springer-Verlag Berlin Heidelberg 2010

191

192

11 Electrocatalytic Nanoparticle Tags Technique

probes on the electrode, and chemical amplification of the signal using the nanoparticle tags on the electrode. The second nanotechnology-based method utilizes surface plasmon resonance imaging (SPRI) in combination with traditional molecular biology enzymatic reactions (Fang et al. 2006). This multiplexed platform uniquely consists of a single-stranded LNA capture probe per miRNA. A RNA sample is hybridized to this microarray, followed by poly(A) tailing of bound miRNA. Instead of using cy3 or cy5 labeled poly(T)s to the base pair with the synthesized poly(A)s, gold nanoparticles attached to poly(T) oligos are used. The microarray image is then obtained from a scanner that detects gold nanoparticles. The third nanotechnology used for miRNA detection utilizes a biosensor that has the capability to detect and quantitate miRNAs in the femtomolar range. The core of this biosensor consists of a microscopic platform made with interlocking gold and titanium microelectrodes with wells in between. Capture probes are chemically fixed into these wells. As samples are hybridized onto this platform, miRNAs are bound to their probe in specific wells. The anionic nature of the miRNA phosphate backbone then catalyzes the formation of polyaniline nanowires from a solution of cationic aniline particles, forming a complete electrical circuit between the electrodes, and resulting in an immediate digital readout (Fan et al. 2007). Another nanotechnology-based method, the gold nanoparticle probe technique (Yang et al. 2008), utilizes three probes, miRNA-specific biotinylated probe (capture probe), miRNA-specific poly(A) probe, and gold nanoparticle probe, which are hybridized to the complementary target miRNA in a sandwich assay format. The hybridization complex is immobilized onto the surface of a streptavidin-coated microplate. The signal of adsorbed gold nanoparticle is amplified by silver enhancement and recorded with colorimetric absorbance by a microplate reader (Fig. 11.1). The ENT technique is based on a direct chemical ligation procedure that involves a chemical reaction to tag miRNAs with the OsO2 nanoparticles (Gao and Yang 2007). The nanoparticles effectively catalyze the oxidation of hydrazine and greatly enhance the detectability of miRNAs, thereby lowering the detection limit to femtomolar range. It is well-known that many transition metal salts tend to hydrolyze under neutral or alkaline conditions, forming metal hydroxides or oxides. Hydrolysis has been widely used to synthesize transition metal oxide nanoparticles (Murray et al. 2000). In most cases, nanoparticle nucleation and growth occur via a simple precipitation reaction from homogeneous solution, involving reaction of a metal salt solute with hydroxide or water. To achieve the desired size and size distribution, the growth of the nanoparticles is often arrested by a capping agent. This strategy was adopted by Gao and Yang (2006) in the preparation of OsO2 nanoparticles, using isoniazid as the capping agent. OsO2 nanoparticles in the range of 5–50 nm can be prepared by modulating the reaction conditions. The capping, resulting in some loss of the nanoparticles, significantly narrows the particle size distribution by eliminating (dissolving) the smaller ones. The tags chemically amplify analytical signals to hybridized electrodes, as compared to nonhybridized ones. The differences in amperometric currents are used for quantitation purposes. In a similar way, the nanoparticles were evaluated as novel electrocatalytic tags for

11.1 Introduction

193

Select miRNAs of your interests

Design of oligonucleotide capture probe

Electrode Fabrication I: silanize an ITO slide

Electrode Fabrication II: probe immobilization

Isolate RNA from tissues or cells

miRNA/Probe hybridization

K2OsCl6

NaOH

Preparation of OsO2 nanoparticles

Treatment with OsO2 nanoparticles

Amperometric detection of miRNAs

Fig. 11.1 Flowchart of Electrocatalytic Nanoparticle Tags (ENT) technique for miRNA expression detection, according to Gao and Yang (2006)

possible applications in an ultrasensitive miRNA assay (Gao and Yang 2007). It is immediately apparent that the nanoparticles exhibit a dramatic improvement in response to the oxidation hydrazine: The oxidation of hydrazine appeared at 0.1 V, essentially the same potential as that of the nanoparticles themselves. There is a very strong catalytic effect by the nanoparticles, since the current at potentials in the vicinity of the nanoparticles’ redox potential increased dramatically and the overpotential of hydrazine oxidation was reduced by as much as 900 mV, indicating that the nanoparticles are being turned over by the oxidation of hydrazine. miRNA can selectively hybridize with its complementary capture probe on the electrode surface with very little cross-hybridization; the nanoparticle tags can be successfully ligated on the hybridized miRNA molecules; and the nanoparticles effectively catalyze the oxidation of hydrazine, producing a much enhanced analytical signal. Amperometric detection at a significantly lower operating potential can, therefore, minimize the potential interference and reduce the background signal, yielding an improved signal-to-noise ratio and a lower detection limit.

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11 Electrocatalytic Nanoparticle Tags Technique

11.2

Protocol

11.2.1 Materials 1. Indium tin oxide (ITO)-coated glass slides (Delta Technologies Ltd, Stillwater, MN) 2. miRNAs for the study, with 50 -terminal aldehyde-modified oligonucleotide capture probes [custom-made by Invitrogen Corporation (Carlsbad, CA)] 3. Conducting epoxy (Ladd Research, Williston, VT) 4. Copper wire 5. YM-50 Montage spin column (Millipore Corp., Billerica, MA)

11.2.2 Instruments 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11.

CHI 660A electrochemical workstation Low-current module (CH Instruments, Austin, TX) 2.0-mm-diameter ITO working electrode Miniature Ag/AgCl reference electrode (Cypress Systems, Lawrence, KS) Platinum wire counter electrode V-570 UV/vis/NIR spectrophotometer (JASCO Corp., Japan) Finnigan/MAT LCQ mass spectrometer (ThermoFinnigan, San Jose, CA) Elan DRC II ICP-MS spectrometer (PerkinElmer, Wellesley, MA) VG ESCALAB 220I-XL XPS system (Thermo VG Scientific Ltd., UK) Scanning electron microscope (SEM) JSM-7400F electron microscope (Joel Ltd., Tokyo, Japan)

11.2.3 Reagents 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11.

K2OsCl6 (>99%) Isoniazid (99%, as a capping agent) Sodium periodate (99%) Sodium borohydride (>99%) 3-Aminopropyl trimethoxysilane (APTES, 97%) Mono-n-dodecyl phosphate (MDP) [all from Sigma-Aldrich (St Louis, MO)] Sodium acetate buffer (0.2 M, pH 6.0) containing 2.0 mM sodium periodate (to be used as the hybridization buffer) Diethyl pyrocarbonate RNaseZap (Ambion, TX) Phosphate-buffered saline: [PBS, 0.15 M NaCl + 20 mM phosphate, pH 8.0] TE buffer: [10 mM Tris-HCl, 1.0 mM EDTA, and 0.1 M NaCl]

11.2 Protocol

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

195

Water/ethanol (20/80) solvent containing NaOH Hydrazine (30 mM) Aniline (99.5%) 1,4-Phenylenediisothiocyanate (PDITC, 98%) Horseradish peroxidase (HRP) (200 units/mg) Hydrogen peroxide (31%)

11.2.4 Procedures The protocols described in this section are essentially the same as reported in the study by Gao and Yang (2006) (see Fig. 11.1).

11.2.4.1

Preparation of the OsO2 Nanoparticles

1. Prepare a water/ethanol (20/80) mixture solvent 2. Add slowly a solution of K2OsCl6 in 100 ml of the mixture solvent to NaOH dissolved in water/ethanol (50/50). Make the final concentration of NaOH between 50 and 200 mM 3. Mix the precursors for 5 min, then heat the mixture to 40 C for ~30 min to produce the nanoparticles 4. Dissolve isoniazid in the mixture solvent 5. Add isoniazid to the nanoparticles solution till a final concentration of 10 mM is obtained 6. Stir for 30 min, then add 100 ml of ethanol 7. Centrifuge the mixture at 10,000 rpm 8. Wash the nanoparticles with ethanol several times

11.2.4.2

Design of Oligonucleotide Capture Probe (OCP)

1. Select miRNAs of interest and pull out the mature miRNA sequences from the miRNA Registry (microrna.sanger.ac.uk) 2. Design the OCPs exactly antisense to the selected miRNAs 3. Synthesize the OCPs using the services provided by companies like IDT (Integrated DNA Technologies, Coralville, IA, USA)

11.2.4.3

Electrode Fabrication

1. Prior to OCP immobilization, silanize an ITO slide following a published procedure (Hedges et al. 2004).

196

11 Electrocatalytic Nanoparticle Tags Technique

2. Assemble a patterned 2-mm-thick, adhesive, spacing/insulating layer on top of the slide to form a low-density electrode array of 20–30 individual electrodes of 2-mm-diameter. 3. Add 5.0 ml aliquots of aldehyde-modified OCPs (0.5 mM) in an acetate buffer (0.1 M, pH 6.0) to the individual electrodes and incubate for 3 h at room temperature in a moisture-saturated environmental chamber. 4. After incubation, rinse the electrodes successively with 0.1% SDS and water. 5. Incubate the electrodes in 2.5 mg/ml sodium borohydride solution made of PBS: ethanol (3:1) for 5 min to reduce imine and to improve the stability of the capture probe-coated electrode by generating a stable secondary amine linkage (Horton et al. 1997). 6. Soak the electrodes in vigorously stirred hot water (90–95 C) for 2 min, rinse with water, and blow dry with a stream of nitrogen. 7. To further improve the quality and stability of the electrodes and to minimize the nonhybridization-related nanoparticle uptake, immerse the capture probe-coated electrodes in 2.0 mg/ml MDP for 3–5 h. Subsequently, MDP molecules fill in the defects via strong interaction between phosphate and ITO, forming a mixed monolayer with the capture probes (Popovich et al. 2003). 8. Rinse unreacted MDP molecules, and wash the electrodes by immersing in stirred ethanol for 10 min, followed by a thorough rinsing with water. The surface density of the immobilized capture probes should be in the range of 5–8 pM/cm2 (Steel et al. 1998; Gao and Yang 2006).

11.2.4.4

Total RNA Extraction

1. Extract total RNA from test tissue or cells using TRIzol reagent, according to the manufacturer’s recommended protocol. 2. Enrich small RNAs in the total RNA sample using a YM-50 Montage spin column. 3. Determine RNA concentration by UV–vis spectrophotometry.

11.2.4.5

RNA Hybridization and Detection

Perform hybridization and nanoparticle tagging of miRNA and its amperometric detection in three steps. 1. Place the electrodes in an environmental chamber. Add 2-ml of the total RNA solution in the acetate buffer (0.2 M, pH 6.0) on the electrodes, then add 0.5 ml of sodium periodate (10 mM) and mix thoroughly with the total RNA solution. 2. Perform hybridization and derivation of the 30 overhangs of the miRNAs at 25 C in the dark for 60 min.

References

197

3. After a washing thoroughly with sodium sulfite (0.1 mM) in the acetate buffer, add 5-mL aliquots of 0.1 mg/mL nanoparticles in the acetate buffer, and incubate the electrodes at 30 C for 4 h. 4. After another thorough wash with acetate buffer, characterize the electrodes electrochemically. Perform amperometric detection of the miRNAs on the electrode array at 0.10 V in 30 mM hydrazine in PBS. Apply a 10-ml aliquot of the PBS test solution to the individual electrode in the remaining open circuit. Disable the electrode by withdrawing the test solution. In the case of lower miRNA concentrations, apply smoothing after each measurement to remove random noises. Carry out all experiments at room temperature, unless otherwise stated.

11.3

Application and Limitation

It has been demonstrated that the use of direct chemical ligation, in conjunction with a chemical amplification, offers new possibilities for more sensitive miRNA analysis. The association of the nanoparticles to the hybridized miRNA molecules leads to the formation of the electrocatalytic system, generating a measurable current upon adding substrate to the solution. This electrochemical miRNA assay is easily extendable to a low-density array of 50–100 electrodes, offering an excellent opportunity for the low-density electrochemical array in miRNA expression profiling. Such ultrasensitive electrocatalysis strategies may enable the development of a simple, low-cost, and portable electrochemical miRNA profiling system (Gao and Yang 2006). This technique has been tested with let-7b, miR-106, and miR-139 in HeLa cells (Gao and Yang 2006). The results are in good agreement with those obtained by the Northern blot assay for the same sample, and consistent with previously published data for miRNA expression profiling. This method allows for as low as 5 ng total RNA for a successful miRNA detection, corresponding to 150 HeLa cells (Allawi et al. 2004; Lim et al. 2003). The relative errors associated with the assay was found to be <15% in the concentration range of 1–200 pM. Thus, the assay is capable of identifying miRNAs with <2-fold difference in expression levels under two conditions. However, the need for multiple electrochemical detection instruments, that are unavailable in most of the molecular biology laboratories, may limit its application.

References Fan Y, Chen X, Trigg AD, Tung CH, Kong J, Gao Z (2007) Detection of MicroRNAs using target-guided formation of conducting polymer nanowires in nanogaps. J Am Chem Soc 129:5437–5443

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Fang S, Lee HJ, Wark AW, Corn RM (2006) Attomole microarray detection of microRNAs by nanoparticle-amplified SPR imaging measurements of surface polyadenylation reactions. J Am Chem Soc 128:14044–14046 Gao Z, Yang Z (2006) Detection of microRNAs using electrocatalytic nanoparticle tags. Anal Chem 78:1470–1477 Hedges DH, Richardson DJ, Russell DA (2004) Electrochemical control of protein monolayers at indium tin oxide surfaces for the reagentless optical biosensing of nitric oxide. Langmuir 20:1901–1908 Yang WJ, Li XB, Li YY, Zhao LF, He WL, Gao YQ, Wan YJ, Xia W, Chen T, Zheng H, Li M, Xu SQ (2008) Quantification of microRNA by gold nanoparticle probes. Anal Biochem 376:183–188

Chapter 12

Nanoparticle-Amplified SPR Imaging for High-Sensitivity miRNA Profiling

Abstract The nanoparticle-amplified surface plasmon resonance imaging (SPRI) method is a miRNA profiling technique that employs a combination of surface poly (A) enzyme chemistry and nanoparticle-amplified SPRI measurements on a microarray platform. The surface reaction of poly(A) polymerase creates poly(A) tails on miRNAs hybridized onto locked nucleic acid (LNA) microarrays. DNA-modified nanoparticles are then adsorbed onto poly(A) tails and are detected using SPRI. This technique was initially developed by Corn’s laboratory in the Department of Chemistry, University of California (J Am Chem Soc 128:14044–14046, 2006). It is an ultrasensitive method for the detection of multiple miRNAs at attomole levels.

12.1

Introduction

The nanoparticle-amplified surface plasmon resonance imaging (SPRI) method is a miRNA profiling technique for the detection and quantification of multiple miRNAs (Fang et al. 2006). It employs a combination of surface poly(A) enzyme chemistry and nanoparticle-amplified SPRI measurements on a microarray platform. The first step in the miRNA detection process is the hybridization of miRNAs onto a three-component microarray of complementary single-stranded LNA 16-mers. LNAs are commercially available nucleic acid analogs containing one or more nucleotide monomers where the ribose moiety is modified with an additional bridge connecting the 20 -O and 40 -C atoms (Castoldi et al. 2006) to enhance specificity and affinity of hybridization. Hybridization adsorption of miRNA to the LNA probe creates an overhang at least six bases long at the 30 -OH end of the miRNAs; this overhang is required to ensure a high efficiency of the poly-(A) reaction. Following hybridization adsorption, the surface-bound miRNAs are polyadenylated with poly(A) polymerase. Finally, the arrays are exposed to a solution containing DNA probes with T30-coated gold (Au) nanoparticles to allow the 50 -thiol-modified, 30 -OH miRNAs to attach to the gold surface

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12 Nanoparticle-Amplified SPR Imaging for High-Sensitivity miRNA Profiling

via a surface thiol/maleimide reaction to create an miRNA monolayer to further enhance the SPRI signal (Figs. 12.1 and 12.2). The nanoparticle-amplified SPRI technique demonstrates several advantages over other miRNA expression detection techniques. It is ultrasensitive, is able to detect low abundance of miRNA in cells and in a small quantity of RNA samples down to attomole levels. Currently, a 5 attomole detection limit is about 50 times more sensitive than the fluorescence-based microarray detection method mentioned previously (Shingara et al. 2005). Thus, the 10 fM detection limit demonstrated in this study (Fang et al. 2006) represents a remarkable 105 signal enhancement, which corresponds to the detection of a miRNA fractional surface coverage of 106 (Lee et al. 2006). A more striking feature is that the method offers a platform for a high-throughput quantification of miRNAs, which is distinct from other methodologies that either are high throughput or quantitative, but not both.

SPRI Measurement

T30-coated Au NPs OH

miRNA

Poly(A) Polymerase

Au

Poly(A) Tail Au

Au

OH

5’

5’

5’

Hybridization

Polyadenylation

5’

LNA Probe

miRNA/Probe

Au Nanoparticle Amplification

5’

5’

Polyaneylated miRNA Au NP-coated miRNA

Fig. 12.1 Schematic illustration of miRNA detection procedures using the nanoparticle-amplified SPRI technique. Adapted from Fang et al. (2006). (a) LNA probe immobilization to fabricate an array platform. (b) Hybridization adsorption of miRNA onto a complementary LNA probe array. (c) Polyadenylation to add poly(A) tails to the surface-bound miRNAs using poly(A) polymerase. (d) Hybridization adsorption of T30-coated gold nanoparticles to the poly(A) tails. (e) Detection of miRNAs using SPRI measurements. Modified from Fang et al. (2006)

12.2 Protocol

201 Select miRNAs of your interests

Design of oligonucleotide capture probe

LNA microarray fabrication

Isolate RNA from tissues or cells

HAuCl4

Hybridization of miRNAs to probes in microarray

Preparation of Au nanoparticles

Surface polyadenylation reaction to link poly(A) to 3' end of miRNAs

Coat Au nanoparticle with T30 oligo

Gold nanoparticle amplification by anealing poly(A) in miRNAs with T30 in NP

SPRI measurements of miRNAs

Fig. 12.2 Flowchart of the nanoparticle-amplified surface plasmon resonance imaging (SPRI) method for miRNA detection. According to Fang et al. (2006)

12.2

Protocol

12.2.1 Materials 1. 2. 3. 4.

SF-10 glass (Schott Glass) Thin gold films (45 nm) Software package V++ (Digital Optics) Software package NIH Image V. 1.63

12.2.2 Instruments 1. UV–vis spectroscopy 2. Denton DV-502A metal evaporator 3. SPR imager (GWC Technologies)

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12 Nanoparticle-Amplified SPR Imaging for High-Sensitivity miRNA Profiling

12.2.3 Reagents 1. Poly(A) polymerase 2. 11-Mercaptoundecylamine (MUAM; Dojindo) 3. N-Hydroxysuccinimidyl ester of methoxypoly(ethylene glycol) propionic acid MW 2000 (PEG-NHS; Nektar) 4. 9-Fluorenylmethoxycarbonyl-N-hydroxysuccinimide (Fmoc-NHS; Novabiochem) 5. Sulfosuccinimidyl 4-(N-maleimidomethyl)-cyclohexane-1-carboxylate (SSMCC; Pierce) 6. Yeast poly(A) polymerase (USB) 7. Proteinase K (USB) 8. Absolute ethanol or autoclaved Millipore deionized water 9. 30 -Thiol-modified locked nucleic acid (LNA)-modified DNA oligonucleotide probes (Integrated DNA Technologies) 10. ATP 11. mirVana miRNA Isolation Kit (Ambion) 12. 39 mM sodium citrate solution 13. Reaction buffer: [20 mM Tris-HCl, 60 mM KCl, 4 mM MnCl2, 0.5 mM DTT; pH 7.1] 14. Proteinase K solution: [50 mM Tris-HCl, 100 mM KCl, 5 mM CaCl2; pH 7.4] 15. 1 M NaCl/100 mM phosphate buffer (pH 7.4) 16. M NaCl/10 mM phosphate buffer (pH 7.4) 17. M NaCl/10 mM phosphate buffer (pH 7.4) 18. TRIzol reagent (Invitrogen, Carlsbad, CA)

12.2.4 Procedures The protocols described in this section are essentially the same as reported in the study conducted by Corn’s laboratory (Fang et al. 2006) (Figs. 12.1 and 12.2).

12.2.4.1

Preparation of DNA and RNA Oligonucleotides

1. Select miRNAs of interest for the study and obtain the sequences of these miRNAs from the miRNA Registry (microrna.sanger.ac.uk). Purchase synthetic miRNAs from Integrated DNA Technologies (IDT) or Dharmacon RNA Technologies, or other companies. Determine the concentration of the received miRNAs by UV–vis spectroscopy or a NanoDrop device prior to its use. 2. Design the locked nucleic acid probes (LNA probes) using LNA oligonucleotide design software (www.exiqon.com) to maintain a similar melting temperature for all probes. LNA probes should be in the form of DNA oligonucleotides of

12.2 Protocol

3.

4. 5. 6.

203

16 nts in length designed to bind to the 50 ends of miRNAs of interest, and should be exact antisense to their respective target miRNAs. Add the sequence (C)15(CH2)3-S-S to the 30 end of the probes to make 30 -thiol modification. Design five LNA bases for each probe, with two at the 50 end and the other three evenly distributed across the remaining sequence. For example, miR-1 sequence is 50 -UGGAAUGUAAAGAAGUAUGUAU-30 and its antisense sequence is 50 -ATACATACTTCTTTACATTCCA-30 . By removing the 50 end 6 nts and adding the 30 end modification, we have the LNA probe for miR1: 50 -ACTTCTTTACATTCCA(C)15(CH2)3-S-S-30 (LNA bases are underlined). Synthesize the probes from IDT at RNase-free HPLC grade. Use the 50 -thiol-modified DNA sequence [50 -S-S-(CH2)6-(T)30-30 ] for the coating of gold nanoparticles. Use the following two probes for surface dilution experiments that characterize the polyadenylation and nanoparticle amplification steps: RNA probe (R1) ¼ 50 S-S-(CH2)6(C)8CGUGUUAGCCUCAAGUG-30 ; DNA probe (D1) ¼ 50 -ACGTAGTCAGAGCAGAGTTT(CH2)3-S-S-30 (used as a negative control).

12.2.4.2

LNA Microarray Fabrication

1. Deposit thin gold films (45 nm) with a 1-nm underlayer of chromium onto SF-10 glass using a metal evaporator. 2. Modify the gold substrates with a self-assembled monolayer of MUAM by allowing the MUAM surface to react with the hydrophobic protecting group, Fmoc-NHS. 3. UV photopatterning of the Fmoc surface with a quartz mask containing 500 mm square features to create bare gold spots within the hydrophobic Fmoc background. 4. Expose the bare gold spots to a MUAM solution for a minimum of 2 h before spotting a solution of the heterobifunctional crosslinker SSMCC. 5. Manually spot thiol-modified LNA probes onto the SSMCC array elements and allow to react overnight. 6. Remove the Fmoc background with a mild basic solution and allow to react with PEG-NHS, which resists non-specific adsorption of biomolecules.

12.2.4.3

Preparation of Oligonucleotide-Modified Gold Nanoparticles

Preparation of Gold Nanoparticles 1. Add 20 mL of 39 mM sodium citrate solution to a 200 mL boiling solution of 1 mM HAuCl4 and stir vigorously under reflux conditions for 15 min until no further color change in solution can be observed. 2. Filter the colloidal solution through a 0.2-mm pore membrane, store at 4 C, and autoclave prior to use. Gold nanoparticles prepared using this method should have a lmax of 518 nm with an average diameter of ~12–13 nm.

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12 Nanoparticle-Amplified SPR Imaging for High-Sensitivity miRNA Profiling

Preparation of DNA-Coated Gold Nanoparticle 3. Add 50 mL of 100 mM 50 -thiol-modified T30 oligonucleotide to 800 mL of gold nanoparticle solution and keep the solution at 37 C for at least 4 h. 4. Add 230 mL of H2O and 120 mL of 1 M NaCl/100 mM phosphate (pH 7.4). 5. After aging overnight at 37 C, centrifuge the solution at 12,000 rpm for 40 min, remove the supernatant and resuspend the gold pellet in 0.1 M NaCl/10 mM phosphate buffer (pH 7.4). Repeat this process at least three times. 6. After the final centrifugation, re-suspend the gold pellet in 0.3 M NaCl/10 mM phosphate buffer (pH 7.4) to achieve an optical density of 2, corresponding to a final concentration of ~10 nM. This value can be obtained from UV–vis spectroscopy using an extinction coefficient of ~2 108/M/cm. 12.2.4.4

miRNA Detection

miRNA Hybridization Adsorption to LNA Arrays 1. Isolate total mRNA from the tissue of your choice using the mirVana miRNA Isolation Kit, according to the manufacturer’s instructions. 2. Mix an aliquot containing ~250 ng of total RNA into 500 mL of 0.3 M NaCl/ 10 mM phosphate buffer (pH 7.4). 3. Add the mixture to the LNA array surface. 4. For each single experiment, allow the hybridization adsorption to proceed for ~4 h at a flow rate of 1 mL/min in a circulating 100 mL cell with a total volume of 500 mL sample solution. This reaction time is sufficient to reach a steady-state surface coverage of miRNA from femtomolar solutions given a miRNA hybridization adsorption rate constant of ~104/M/s (Lee et al. 2006). Surface Polyadenylation Reaction 5. After miRNA hybridization adsorption, add a mixture of yeast poly(A) polymerase (2.4 units/mL) and 5 mM ATP in the reaction buffer to the array surface and incubate for 1 h. 6. After the reaction, wash the surface with 100 nM proteinase K solution for 30 min to remove the nonspecifically adsorbed enzyme. To optimize the conditions for the surface enzyme reaction, use a high concentration (500 nM) target miRNA solution to follow the surface polyadenylation reaction with real-time SPRI measurements. The changes in percent reflectivity (r%R) measured from the polyadenylation of miRNA adsorbed onto the LNA microarray should be a function of time. The polymerization reaction data should fit into an exponential equation of the form R1(1 - exp(-t/t)), where R1 represents the steady-state value and t is the time constant for this surface polymerization reaction.

12.3 Application and Limitation

205

Gold Nanoparticle Amplification 7. Expose the array to 300 mL of freshly prepared T30-DNA coated gold nanoparticles for 10 min.

12.2.4.5

SPRI Measurements

1. A SPR imager using near-infrared excitation from an incoherent white light source should be used for all SPRI experiments. Use collimated p-polarized light to illuminate a flow cell/gold chip/prism assembly at a fixed incident angle near the SPR angle. The reflected light is directed through a band-pass filter centered at l ¼ 830 nm and collected with a CCD camera. 2. Collect data using the software package V++ and further analyze the data using the software package NIH Image V.1.63. 3. SPR images obtained before and after the amplification of gold nanoparticle should be subtracted to obtain a difference image. For quantification of miRNAs, a standard calibration curve must be established for each miRNA using synthetic miRNAs equivalent to the test miRNAs. The SPR signal should respond linearly for the synthetic miRNAs over a concentration range of 10–500 fM. For higher miRNA concentrations, the nanoparticle concentration can be reduced to bring the SPRI signal back into the linear response range. Above a miRNA concentration of 100 pM, no nanoparticle amplification is required.

12.2.4.6

Characterization of Nanoparticle Amplification

This step is required only if the reactivity of poly(A) polymerase at chemically modified gold surfaces and the subsequent hybridization adsorption of DNA-coated nanoparticles needs to be characterized. The characterization can be done by measuring the polyadenylation of single-stranded RNA (ssRNA) directly attached to a gold surface with SPRI measurements (Chen et al. 2005).

12.3

Application and Limitation

The combined use of a surface enzyme reaction and DNA-coated nanoparticle enhancement with SPRI offers many advantages for miRNA detection. 1. Multiple miRNAs can be detected in a microarray format for high-throughput profiling, using a novel approach that combines a surface enzyme reaction with nanoparticle-amplified SPR imaging (SPRI) (Fang et al. 2006).

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12 Nanoparticle-Amplified SPR Imaging for High-Sensitivity miRNA Profiling

2. The enzymatically amplified SPRI methodology described here can be used to quantitatively measure miRNAs with excellent sensitivity in total RNA samples down to femtomolar concentrations. The minimum amount of miRNA detected in the study reported by Fang et al. (2006) is 5 attomoles (a 10 fM solution with a total volume of 500 mL). Currently, a 5 attomole detection limit is about 50 times more sensitive than the fluorescence-based microarray detection method mentioned previously (Shingara et al. 2005). Thus, the 10 fM detection limit demonstrated in this study (Fang et al. 2006) represents a remarkable 105 signal enhancement, which corresponds to the detection of a miRNA fractional surface coverage of 106 (Lee et al. 2006). Similar combination has also been recently reported for ultrasensitive single nucleotide polymorphism genotyping (Li et al. 2006). 3. The use of a surface-based enzyme amplification strategy to form the poly(A) tail greatly simplifies the removal of reactants and reagents from the miRNA, and allows for the binding of multiple DNA-coated nanoparticles to a single miRNA adsorption site (Fang et al. 2006). The nanoparticle-amplified SPRI method has been validated for three miRNAs, miR-16, miR-23b, and miR-122b from mouse liver tissue. The amount of miRNAs in the total RNA sample was estimated to be 20 fM, 50 fM, and 2 pM for miR-16, miR-23b, and miR-122b, respectively (Fang et al. 2006). The main technical limitation of this technique is the requirement of sophisticated, expensive instruments (SPR imager) for signal readout.

References Castoldi M, Schmidt S, Benes V, Noerholm M, Kulozik AE, Hentze MW, Muckenthaler MU (2006) A sensitive array for microRNA expression profiling (miChip) based on locked nucleic acids (LNA). RNA 12:913–920 Chen C, Ridzon DA, Broomer AJ, Zhou Z, Lee DH, Nguyen JT, Barbisin M, Xu NL, Mahuvakar VR, Andersen MR, Lao KQ, Livak KJ, Guegler KJ (2005) Real-time quantification of microRNAs by stem-loop RT-PCR. Nucleic Acids Res 33:e179 Fang S, Lee HJ, Wark AW, Corn RM (2006) Attomole microarray detection of microRNAs by nanoparticle-amplified SPR imaging measurements of surface polyadenylation reactions. J Am Chem Soc 128:14044–14046 Lee HJ, Wark AW, Corn RM (2006) Creating advanced multifunctional biosensors with surface enzymatic transformations. Langmuir 22:5241–5250 Li Y, Wark AW, Lee HJ, Corn RM (2006) Single-nucleotide polymorphism genotyping by nanoparticle-enhanced surface plasmon resonance imaging measurements of surface ligation reactions. Anal Chem 78:3158–3164 Shingara J, Keiger K, Shelton J, Laosinchai-Wolf W, Powers P, Conrad R, Brown D, Labourier E (2005) An optimized isolation and labeling platform for accurate microRNA expression profiling. RNA 11:1461–1470

Chapter 13

Conducting Polymer Nanowires Technique for High-Sensitivity miRNA Expression Analysis

Abstract The Conducting Polymer Nanowires technique is an ultrasensitive, nonlabeling approach for direct and RT–PCR-free miRNA expression profiling. A nanogapped microelectrode-based biosensor array is fabricated for ultrasensitive electrical detection of miRNAs. After peptide nucleic acid (PNA) capture probes are immobilized in the nanogaps of a pair of interdigitated microelectrodes and hybridization is performed with their complementary target miRNA, the deposition of conducting polymer nanowires, polyaniline nanowires, is carried out by an enzymatically catalyzed method, where the electrostatic interaction between anionic phosphate groups in miRNA and cationic aniline molecules is exploited to guide the formation of the polyaniline nanowires onto the hybridized target miRNA. The conductance of the deposited polyaniline nanowires correlates directly with the amount of hybridized miRNA. Under optimized conditions, the target miRNA can be quantified in a range from 10 fM to 20 pM with a detection limit of 5 fM. The biosensor array can be applied to the direct detection of miRNA in total RNA extracted from cell lines or tissues. This technique was initially developed by Gao’s group from the Institute of Microelectronics, Singapore (J Am Chem Soc 129:5437–5443, 2007).

13.1

Introduction

It has been documented that peroxidases, such as horseradish peroxidase (HRP), effectively catalyze the polymerization of aniline in the presence of H2O2 under very mild conditions (Ryu et al. 2000; Liu et al. 1999), which opened up new possibilities to use polyaniline in a biological system. The use of polyelectrolyte as templates has markedly improved the processibility and electrical properties of polyaniline through aligning aniline molecules along the polyelectrolyte templates (Trakhtenberg et al. 2005). This approach was subsequently applied to the fabrication of polyaniline nanowires using DNA as templates (Ma et al. 2004). The above studies paved the way for the development of an electrical procedure for the

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13 Conducting Polymer Nanowires Technique

detection of miRNA utilizing polyaniline nanowires as a signal generator for the transduction of miRNA hybridization events. According to Fan et al. (2007), the hybridization of the target miRNA to PNA capture probes results in a negatively charged surface originating from the phosphate groups on the miRNA backbone. When the hybridized biosensor array is incubated in the mixture of aniline/HRP/ H2O2 at pH 4.0, the protonated aniline molecules (pKa = 4.6) (Lide 1993) are concentrated and aligned around the hybridized miRNA strands through electrostatic interaction between the protonated aniline molecules and phosphates in the miRNA. This high proton concentration around the miRNA provides a local environment of high acidity that permits polymerization of aniline in a much less acidic medium than that in conventional electrochemical and chemical approaches, and facilitates a predominantly head-to-tail coupling and deters parasitic branching during polymerization, offering the much desired structure for high conductance. The polyaniline/miRNA complex in which polyaniline wraps around the miRNA template (polyaniline nanowires) is utilized for miRNA sensing. For the complementary miRNA sample, the obvious increase in conductance of the nanogaps with polyaniline nanowires deposited cannot be immediately obtained because the conductance of the as-prepared polyaniline nanowires is very low (Liu et al. 1999). In the study reported by Fan et al. (2007), a sizable increase in conductance was found for the complementary miRNA, whereas only a slight increase was observed for the control sample when compared to a blank biosensor array (PNA functionalized biosensor arrays without undergoing miRNA hybridization, polyaniline nanowire deposition, and doping). This clearly demonstrates that the formation of polyaniline nanowires in the nanogaps is guided by the hybridized miRNA molecules, and the resulting polyaniline nanowire network bridges the gaps, producing a measurable conductance change. The result of the control sample implies that the nonhybridization-related signal of this biosensor array is extremely low, which facilitates the detection of miRNA at ultralow concentrations. This may be attributed to the use of PNA other than conventional anionic oligonucleotides as the capture probes because, on the one hand, PNA has higher affinity toward target miRNA (Fortina et al. 2005), which can suppress the occurrence of nonspecific capture probe-target binding, and, on the other hand, its neutral N-(2-aminoethyl)glycine backbone can prevent the undesired adsorption of aniline monomer, which reduces the background noise. The conductance between the nanogapped electrodes is primarily dependent on the amount (density) of the polyaniline nanowires formed along the target miRNA strands in the gaps. The more the target miRNA molecules hybridized, the more the polyaniline nanowires deposited along the miRNA strands, thus the higher is the conductance. Because the ratio of the polyaniline nanowire to target miRNA molecule is fixed at 1:1, the amount of PNA capture probes immobilized in the gaps and hybridization efficiency determine the amount of target miRNA bound to the biosensor and thereby the amount of polyaniline nanowires, implying that the target miRNA molecules hybridized in the nanogaps are directly correlated to the conductance, and thus a simple and straightforward linear relationship between the conductance and miRNA concentration can be expected (Fan et al. 2007).

13.2 Protocol

209

The Conducting Polymer Nanowires miRNA detection procedures consist of five main steps. The first step is the fabrication of a nanogapped microelectrodebased biosensor array. Then, peptide nucleic acid (PNA) capture probes are immobilized in the nanogaps of a pair of interdigitated microelectrodes. Hybridization of PNA probes is subsequently performed with their complementary target miRNA. The deposition of conducting polymer nanowires, polyaniline nanowires, is carried out by an enzymatically catalyzed method, where the electrostatic interaction between anionic phosphate groups in miRNA and cationic aniline molecules is exploited to guide the formation of the polyaniline nanowires onto the hybridized target miRNA. Finally, the conductance of the deposited polyaniline nanowires is measured, which correlates directly to the amount of the hybridized miRNA (Figs. 13.1 and 13.2). Under optimized conditions, the target miRNA can be quantified in a range from 10 fM to 20 pM, with a detection limit of 5 fM.

13.2

Protocol

13.2.1 Materials 1. Indium tin oxide (ITO)-coated glass slides (Delta Technologies Ltd, Stillwater, MN) 2. miRNAs for the study, with 50 -terminal aldehyde-modified oligonucleotide capture probes [custom-made by Invitrogen Corporation (Carlsbad, CA] 3. Conducting epoxy (Ladd Research, Williston, VT) 4. Copper wire 5. YM-50 Montage spin column (Millipore Corp., Billerica, MA)

13.2.2 Instruments 1. Alessi REL-6100 probe station (Cascade Microtech) 2. Advantest R8340A ultrahigh resistance meter (Advantest Corp., Tokyo, Japan)

13.2.3 Reagents 1. 2. 3. 4. 5.

Amino-terminated PNA capture probes (Eurogentec, Herstal, Belgium) 3-Aminopropyl triethoxysilane (APTES, 99%; Sigma-Aldrich) Aniline (99.5%; Sigma-Aldrich) 1,4-Phenylenediisothiocyanate (PDITC, 98%; Sigma-Aldrich) HRP (200 units/mg; Boehringer Mannheim GmbH, Germany)

210

13 Conducting Polymer Nanowires Technique

Biosensor Fabrication

miR-1 3'-ACCTTACATTTCTTCATACATA-5' miR-133 3'-AAACCAGGGGAAGTTGGTCGAC-5' miR-328 3'-GACCGGGAGAGACGGGAAGGCA-5'

Nanogapped biosensor array

RNA capture probes

+

+

Anniline H2O2 HRP

GACCGGGAGAGACGGGAAGGCA-5' CUGGCCCUCUCUGCCCUUCCGU-5'

AAACCAGGGGAAGTTGGTCGAC-5' UUUGGUCCCCUUCAACCAGCUG-5'

GACCGGGAGAGACGGGAAGGCA-5' CUGGCCCUCUCUGCCCUUCCGU-5'

AAACCAGGGGAAGTTGGTCGAC-5' UUUGGUCCCCUUCAACCAGCUG-5'

ACCTTACATTTCTTCATACATA-5' UGGAAUGTAAAGAAGUATGTAT-5'

Deposition of Polyaniline Nanowire

ACCTTACATTTCTTCATACATA-5' UGGAAUGTAAAGAAGUATGTAT-5'

GACCGGGAGAGACGGGAAGGCA-5' CUGGCCCUCUCUGCCCUUCCGU-5'

AAACCAGGGGAAGTTGGTCGAC-5' UUUGGUCCCCUUCAACCAGCUG-5'

ACCTTACATTTCTTCATACATA-5' UGGAAUGTAAAGAAGUATGTAT-5'

Hybridization

GACCGGGAGAGACGGGAAGGCA-5'

miRNA under test

AAACCAGGGGAAGTTGGTCGAC-5'

miR-1 3'-UGGAAUGUAAAGAAGUAUGUAU-5' miR-133 3'-UUUGGUCCCCUUCAACCAGCUG-5' miR-328 3'-CUGGCCCUCUCUGCCCUUCCGU-5'

ACCTTACATTTCTTCATACATA-5'

PNA Probe Immobilization

Electrical Detection

Fig. 13.1 Schematic illustration of the conducting polymer nanowires method for miRNA detection using miR-1, miR-133, and miR-328 as examples. The procedure primarily consists of five steps: Fabrication of a nanogapped microelectrode-based biosensor array. Immobilization of PNA capture probes to nanogaps of a pair of interdigitated microelectrodes. Hybridization of PNA probes with their complementary target miRNA. Deposition of conducting polymer nanowires, polyaniline nanowires catalyzed by HRP in the presence of cationic aniline molecules and H2O2. Electrical detection of the signals using a resistance meter. PNA, peptide nucleic acid; HRP, horseradish peroxidase; H2O2, hydrogen peroxide. Modified from Fan et al. (2007)

13.2 Protocol

211 Select miRNAs of your interests

Design of peptide nucleic acid (PNA) miRNA capture probes

Nanogapped biosensor array fabrication

Immobilize probes to the array

Isolate RNA from tissues or cells

Hybridization of miRNAs to probes in biosensor microarray

Deposition of conducting polymer nanowires to the biosensor array

Conductance measurements to detect miRNAs

Fig. 13.2 Flowchart of the conducting polymer nanowires miRNA detection procedures for miRNA detection, according to Fan et al. (2007)

6. 7. 8. 9. 10. 11. 12. 13. 14.

Hydrogen peroxide (31%; Santoku BASF, Japan or equivalent) Diethyl pyrocarbonate RNaseZap (Ambion, TX) TE buffer: [a 10 mM Tris-HCl, 1.0 mM EDTA, and 0.1 M NaCl] TRIzol reagent (Invitrogen, Carlsbad, CA) 0.1% trifluoroacetic acid solution 50 mM sodium carbonate buffer (pH 9.0) Methanol Dimethylformamide solution containing ethanolamine and diisopropylethylamine 15. 0.1 M NaAc buffer (pH 4.0) 16. HCl

13.2.4 Procedures The protocols described in this section are essentially the same as reported in the study by Gao’s group (Fan et al. 2007) (see Figs. 13.1 and 13.2).

212

13.2.4.1

13 Conducting Polymer Nanowires Technique

Nanogapped Biosensor Array Fabrication

The biosensor array used by Fan et al. (2007) consists of 100 pairs of interlocking comb-like microelectrodes (gold 15 nm, titanium 10 nm) with 150–200 fingers, each 700 nm wide and 200 mm long, and with a 300 nm gap. It is fabricated as a 10 10 array on a silicon chip with 500 nm coating of SiO2 by standard photolithography, as described below. (1) Thoroughly clean the array with chloroform and acetone to remove any possible organic contaminants, followed by rinsing with 1.0 M NaOH and a thorough wash with H2O (Liu and Bazan 2005). (2) Bake the array in an oven at 120 C for 20 min. (3) Soak the array in absolute ethanol containing 2% APTES and 1% H2O (v/v) for silanization. (4) Wash with absolute ethanol and allow it to dry under mild nitrogen flow before aging at 120 C for 20 min. (5) Use the bifunctional coupling agent PDITC to immobilize the PNA capture probes on the now amino-terminated array. (6) Add 50 mg PDITC into a mixture solvent of dimethylformamide and pyridine. (7) Allow the array to react with the solution for 2 h, followed by washing with dimethylformamide and dichloromethane, and subsequent drying under nitrogen flow.

13.2.4.2

Design and Immobilization of Peptide Nucleic Acid Capture Probes

1. Select miRNAs of your interest and pull out the mature miRNA sequences from the miRNA Registry (microrna.sanger.ac.uk). 2. Design the PNA probes exactly antisense to the selected miRNAs. 3. Synthesize the PNA probes using the services provided by companies like IDT (Integrated DNA Technologies, Coralville, IA, USA). 4. Dissolve the PNA capture probes in 0.1% trifluoroacetic acid solution and dilute to a concentration of 1 mM with 50 mM sodium carbonate buffer (pH 9.0). 5. Spot a 100 mL aliquot of the PNA capture probe solution onto the array, and carry out the reaction in a humid chamber at 37 C for 5 h. 6. Remove unreacted PNA capture probes by a thorough wash with water and methanol sequentially. 7. Use a dimethylformamide solution containing ethanolamine and diisopropylethylamine to passivate the biosensor array surface.

13.2.4.3

Total RNA Extraction

1. Extract total RNA from test tissue or cell using TRIzol reagent, according to the manufacturer’s recommended protocol.

13.2 Protocol

213

2. Enrich small RNAs in the total RNA sample using a YM-50 Montage spin column. 3. Determine the RNA concentration by UV–vis spectrophotometry. To minimize the effect of RNases on the stability of miRNAs, all solutions should be treated with diethyl pyrocarbonate, and surfaces must be decontaminated with RNaseZap. 13.2.4.4

Hybridization of PNA Probes with miRNAs

1. Perform hybridization in TE buffer at room temperature for 60 min using synthetic miRNAs or total RNA samples. 2. After hybridization, rinse thoroughly the biosensor array with the hybridization buffer to remove unhybridized miRNA. 13.2.4.5

Deposition of Conducting Polymer Nanowires

1. For deposition of polyaniline nanowires after hybridization, add a 2 mL aliquot of 0.2 mg/mL HRP and stoichiometric amount of H2O2 to a solution of 2 mM aniline in 0.1 M NaAc buffer (pH 4.0). 2. Directly apply a 200 mL aliquot of the mixture to the biosensor array and keep for 30–40 min. 3. Thoroughly wash the biosensor array with blank NaAc buffer solution and water to remove any residual enzyme and aniline monomer, followed by drying under nitrogen flow. The target-guided formation of polyaniline nanowires is highly dependent on the electrostatic interaction between cationic aniline monomers and anionic phosphate groups in miRNA. An acidic buffer solution at pH 4.0 should be selected for polyaniline deposition because it is the most optimal acidity to provide adequate protonation of aniline, and thus strong enough electrostatic interaction between aniline and the phosphate groups. It also maintains sufficient activity of HRP (Ma et al. 2004). Besides, this relatively low pH is found to favor the generation of continuous polyaniline nanowires other than insulating polyaniline nanoparticles (Akkara et al. 1991). To maximize the biosensor performance, the influence of aniline, HRP, and the deposition time on the conductance of the resulting polyaniline should also be evaluated (Fan et al. 2007). 13.2.4.6

Electrical Detection

1. Take a brief (10–20 s) doping with HCl vapor. 2. Perform the conductance measurements under ambient conditions with a probe station and an ultrahigh resistance meter. 3. All measurements were conducted within the first 30 min after HCl doping.

214

13.2.4.7

13 Conducting Polymer Nanowires Technique

Construction of Calibration Curve

To construct the calibration curve for quantification of miRNAs, 20 measurements described above need to be performed for each concentration to obtain the average of the conductance.

13.3

Application and Limitation

The Conducting Polymer Nanowires technique is an ultrasensitive, nonlabeling approach for direct and PCR-free miRNA expression profiling. This method directly utilized chemical ligation and amplification for signal read-out and thus avoids using labeling probes, which greatly simplifies the detection procedure. In principle, a much lower detection limit could be realized when working with longer target nucleic acids because the bridging of the nanogaps by the polyaniline nanowires can be realized with fewer long nucleic acid molecules. Multiplex detection can be performed by introducing different capture probes onto the biosensor array. Such an in situ amplification strategy may enable the development of a simple, low-cost, and portable electrical array for miRNA expression profiling, opening the door to routine gene expression profiling and molecular diagnostics. The applicability of the biosensor in miRNA detection of real world samples has been examined by analyzing let-7b in total RNA extracted from HeLa cells and lung cancer cells (Fan et al. 2007). The results obtained are consistent with published data of miRNA expression profiling (Nelson et al. 2004; Allawi et al. 2004; Barad et al. 2004). Considering the detection limit of 5 fM for the biosensor array, it is possible that a dozen cells are able to provide an adequate amount of total RNA for miRNA detection (Lim et al. 2003). The relative standard derivation of the biosensor array was found to be <15%, which provides satisfactory accuracy to distinguish slight miRNA expression differences. Indeed, the detection of the single nucleotide mutations is possible at the biosensor array with a single nucleotide mutation selectivity factor of 20:1, much higher than that of the optical microarray and most other previously reported methods (Rosi and Mirkin 2005; Xie et al. 2004), readily allowing discrimination between the perfectly matched and mismatched miRNAs. Comparing the gold nanoparticle labeling and silver enhance methods for detection of nucleic acids (Yang et al. 2008), the sensitivity of the present assay is two orders of magnitude higher, indicating that the polyaniline nanowires bridge the nanogap much more effectively than gold nanoparticles, greatly enhancing the sensitivity of the biosensor array and thereby lowering the detection limit to femtomolar levels. In practice, this sensitivity of the assay meets the requirements for direct miRNA expression profiling. The electrical miRNA biosensor array is fabricated using the target-guided deposition method. Phosphate groups on the backbone of the hybridized miRNA serve directly as the chemical ligation centers, thus eliminating the second hybridization with the gold nanoparticle detection probe conjugates and multiple silver

References

215

enhancing and washing for signal amplification, enabling a more simplified electrical detection with minimal background and with significantly enhanced sensitivity (Fan et al. 2007).

References Allawi HT, Dahlberg JE, Olson S, Lund E, Olson M, Ma WP, Takova T, Neri BP, Lyamichev VI (2004) Quantitation of microRNAs using a modified Invader assay. RNA 10:1153–1161 Barad O, Meiri E, Avniel A, Aharonov R, Barzilai A, Bentwich I, Einav U, Gilad S, Hurban P, Karov Y, Lobenhofer EK, Sharon E, Shiboleth YM, Shtutman M, Bentwich Z, Einat P (2004) MicroRNA expression detected by oligonucleotide microarrays: system establishment and expression profiling in human tissues. Genome Res 14:2486–2494 Fan Y, Chen X, Trigg AD, Tung CH, Kong J, Gao Z (2007) Detection of MicroRNAs using target-guided formation of conducting polymer nanowires in nanogaps. J Am Chem Soc 129:5437–5443 Fortina P, Kricka LJ, Surrey S, Grodzinski P (2005) Nanobiotechnology: the promise and reality of new approaches to molecular recognition. Trends Biotechnol 23:168–173 Lide DR (1993) Handbook of Chemistry and Physics, 68th ed.; CRC Press: Boca, Raton, FL; pp D159–D161 Lim LP, Lau NC, Weinstein EG, Abdelhakim A, Yekta S, Rhoades MW, Burge CB, Bartel DP (2003) The microRNAs of Caenorhabditis elegans. Genes Dev 7:991–1008 Liu B, Bazan GC (2005) Methods for strand-specific DNA detection with cationic conjugated polymers suitable for incorporation into DNA chips and microarrays. Proc Natl Acad Sci USA 102:589–593 Liu Z, Yang H, Zhou P (1999) Studies on the metabolism pathway of aniline degradation by Comamonas acidovorans AN3. Wei Sheng Wu Xue Bao 39:448–453 Ma Y, Zhang J, Zhang G, He H (2004) Polyaniline nanowires on Si surfaces fabricated with DNA templates. J Am Chem Soc 126:7097–7101 Nelson PT, Baldwin DA, Scearce LM, Oberholtzer JC, Tobias JW, Mourelatos Z (2004) Microarray-based, high-throughput gene expression profiling of microRNAs. Nat Methods 1:155–161 Rosi NL, Mirkin CA (2005) Nanostructures in biodiagnostics. Chem Rev 105:1547–1562 Ryu H, Lee JH, Kim KS, Jeong SM, Kim PH, Chung HT (2000) Regulation of neutrophil adhesion by pituitary growth hormone accompanies tyrosine phosphorylation of Jak2, p125FAK, and paxillin. Immunol 165:2116–2123 Trakhtenberg S, Hangun-Balkir Y, Warner JC, Bruno FF, Kumar J, Nagarajan R, Samuelson LA (2005) Photo-cross-linked immobilization of polyelectrolytes for enzymatic construction of conductive nanocomposites. J Am Chem Soc 127:9100–9104 Xie H, Yu YH, Xie F, Lao YZ, Gao Z (2004) A nucleic acid biosensor for gene expression analysis in nanograms of mRNA. Anal Chem 76:1611–1617 Yang WJ, Li XB, Li YY, Zhao LF, He WL, Gao YQ, Wan YJ, Xia W, Chen T, Zheng H, Li M, Xu SQ (2008) Quantification of microRNA by gold nanoparticle probes. Anal Biochem 376:183–188

Chapter 14

Gold Nanoparticle Probe Method for miRNA Quantification

Abstract The gold nanoparticle probe (GNP-probe) technique is a unique and elegant method for sensitive and specific detection and quantification of miRNAs using gold nanoparticle and silver enhancement. The GNP-probe technique allows a lower detection limit of 10 fM with a linear dynamic range from 10 pM to 10 fM or 2 ng of total RNA, and a high specificity to discriminate one single oligonucleotide mismatch of the target miRNA. The method does not require expensive equipment and an advanced read-out system, and can be performed in any standard laboratory. This technique was invented by Xu’s group in 2008 (Anal Biochem 376:183–188, 2008) from MOE Key Laboratory of Environment and Health, School of Public Health, Tongji Medical College, Huazhong University of Science and Technology, China, based on their earlier work on polynucleotides detection using the same strategy (Zhao et al. 2006).

14.1

Introduction

The gold nanoparticle probe (GNP-probe) technique is a simple, sensitive, and specific colorimetric method for the detection and quantification of miRNA using gold nanoparticle (GNP) and silver enhancement (Yang et al. 2008). In brief, the assay utilizes three probes, miRNA-specific biotinylated probe (capture probe), miRNA-specific poly(A) probe, and GNP-probe, which are hybridized to the complementary target miRNA in a sandwich assay format. The hybridization complex is immobilized onto the surface of a streptavidin-coated microplate. After a sufficient rinse of the microplate, the signal of absorbed GNP is amplified by silver enhancement and recorded with colorimetric absorbance by a microplate reader (See Figs. 14.1 and 14.2).

Z. Wang and B. Yang, MicroRNA Expression Detection Methods, DOI 10.1007/978-3-642-04928-6_14, # Springer-Verlag Berlin Heidelberg 2010

217

218

14 Gold Nanoparticle Probe Method for miRNA Quantification

5’–UGGAAUGUAAAGAAGUAUGGAG–3’

–(CH2)6–ACCTTACATTT–5’

miR-1

MSB-probe

Biotin

(25°C 10min)

5’–UGGAAUGUAAAGAAGUAUGGAG–3’ –(CH2)6 –ACCTTACATTT CTTCATACCTC(AAAAAAAAAA)3–5’

MPA-probe (25°C 10min) 5’–UGGAAUGUAAAGAAGUAUGGAG –(CH2)6 –ACCTTACATTTCTTCATACCTC(AAAAAAAAAA)3–5’

GN

5’–(TTTTTTTTTT)3–(CH2)6 –SH

Universal T-probe (25°C 10min)

5’–(TTTTTTTTTT)–(CH2)6 –SH

GNP-probe 5’–UGGAAUGUAAAGAAGUAUGGAG(TTTTTTTTTT)3–(CH2)6–SH –(CH2)6–ACCTTACATTTCTTCATACCTC(AAAAAAAAAA)3–5’

SH

) 6–

2

CH

( )–

TT

5’

3

SES

TT

T TT

TT

(T AG

GG

U UA

AG

A AG

(25°C 30min)

AA

GG

–U

5’

C

H2

Streptavidin

–(

(OD at 630 nm)

H

S ) 6– H2

CA

TA

CT

AC ) 6–

A

AT

TC

T TC TT

GU

U AA

A

AA

AA

A C(

T CC

(25°C 3min)

)–

AA AA

(T AG

Microplate

GG

AU

GU

A GA AA

UA

G AU

A GG

–U

5’

H2

C

Streptavidin

–(

T CA

TA

CT

AC ) 6–

’

5 )–3

AA

AA

A

TC

CT TT

A AA

AA

C(A

CT

C TA

)3

TT

TT

TT

T TT

C –(

Microplate

Fig. 14.1 Schematic illustration of the gold nanoparticle probe method for miRNA detection and quantification. (1) Hybridization miRNA-specific biotin probe (MSB probe) to the 50 end half sequence of target miRNA (e.g., miR-1) in a total RNA sample. (2) Hybridization of miRNAspecific poly(A) probe (MPA probe) to the 30 end half sequence of the target miRNA to form a complex with a free poly(A) tail. (3)(4) Hybridization of T30 gold nanoparticle probe (GNP probe) to the free poly(A) tail of the complex. (5) Immobilization of the hybridization complex onto the surface of a streptavidin-coated microplate. (6) Preparation of silver enhancement. (7) Read optical density at 630 nm in a microplate reader to obtain relative quantity of the target miRNA in the RNA sample, according to Yang et al. (2008)

14.1 Introduction

219 Select miRNAs of your interests

Synthesis of miRNA-specific DNA biotin probe: 5'-Bio-(CH2)6-XXXXXXXXXXX-3' (MSB-probe) Synthesis of miRNA-specific poly(A) probe: 5'-XXXXXXXXXX-(A)30-3' (MPA-probe)

Isolate RNA from tissues or cells

Hybridization of miRNAs with the probes HAuCl4

Synthesis of a probe: 5'-(T)30-(CH2)6- S-S -3'

Preparation of Au nanoparticles

Preparation of gold nanoparticle probes (GNP-probe)

Hybridization of GNP-probe to the poly(A)

Preparation of silver enhancing solution

Measure the degree of surface silver gray scale with colorimetric absorbance by a microplate reader

Fig. 14.2 Flowchart of the gold nanoparticle probe technique for miRNA detection, according to Yang et al. (2008)

The GNP-probe technique allows a lower detection limit of 10 fM, with a linear dynamic range from 10 pM to 10 fM or 2 ng of total RNA and a high specificity to discriminate one single oligonucleotide mismatch of the target miRNA. This method does not require expensive equipment and an advanced read-out system, and can be performed in any standard laboratory.

220

14.2

14 Gold Nanoparticle Probe Method for miRNA Quantification

Protocol

14.2.1 Materials 1. Enzyme immunoassay plates (microplates; Corning Star, Genetimes Technology, Inc) 2. GNP (15 nm, Sino-American Biotechnology Co., SABC) 3. Modified oligonucleotides (with thiol group or biotin, Invitrogen). All oligonucleotides are dissolved in TE buffer. For example, For miR-1: MSB probe: MST probe: MT-MST probe: For miR-328: MSB probe: MST probe: MT-MST probe:

50 -uggaauguaaagaaguauggag-30 50 -Bio-A10CTCCATACTTC-30 50 -TTTACTTACC-(CH2)6-SH-30 50 -TTTAgTTACC-(CH2)6-SH-3’ 50 -cuggcccucucugcccuuccgu-30 50 -Bio-A10ACGGAAGGGCA-30 50 -GAGAGGGCCAG-(CH2)6-SH-30 50 -GAGAcGGCCAG-(CH2)6-SH-30

14.2.2 Instruments 1. Microplate reader 2. Spectrophotometer

14.2.3 Reagents 1. 2. 3. 4. 5. 6. 7. 8. 9.

Streptavidin AgNO3 (0.25 g in 1.0 ml bidistilled water) Citric acid Trisodium citrate (Na3C6H5O72H2O) Hydroquinone (0.085 g in 1.5 ml bidistilled water) TRIzol TE buffer: 10 mM Tris–HCl, 1 mM EDTA, pH 7.4 PBN buffer: [0.3 M NaNO3 and 10 mM Na2HPO4/NaH2PO4 buffer, pH 7.0] 1 mM HAuCl4 (hydrogen tetrachloroaurate): [dissolve 0.1 g HAuCl4 (Aldrich G4022) in 500 mL distilled water. This stock solution of gold(III) ions can be prepared in advance and stored in a brown bottle.] 10. NaCl 11. Citrate buffer: [0.255 g citric acid, 0.235 g trisodium citrate dehydrate in 1.0 ml bidistilled water, pH 3.8] 12. Lysis buffer: [20 mM Tris–HCl (pH8.5) 0.5% NP-40, 20 mg/mL tRNA (in the absence of added divalent cations)

14.2 Protocol

221

14.2.4 Procedures The protocols described in this section are essentially the same as reported in the study by Xu’s group in 2008 (Yang et al. 2008) (See Figs. 14.1 and 14.2).

14.2.4.1

Preparation of Gold Nanoparticle

The formation of GNPs can be observed by a change in color since small nanoparticles of gold are red in color (Rao et al. 2000). A layer of absorbed citrate anions on the surface of the nanoparticles keep the nanoparticles separated. The presence of this colloidal suspension can be detected by the reflection of a laser beam from the particles. Switching to a smaller anion allows the particles to approach more closely and another color change is observed. (1) Add 20 mL of 1.0 mM HAuCl4 to a 50 mL Erlenmeyer flask on a stirring hot plate. Add a magnetic stir bar and bring the solution to a boil. (2) To the boiling solution, add 2 mL of a 1% solution of trisodium citrate dihydrate, Na3C6H5O72H2O. (3) Stir vigorously under reflux conditions for 15 min until no further color change in solution is observed. The gold sol gradually forms as the citrate reduces the gold(III). (4) Filter the colloidal solution through a 0.2 mm pore membrane, store it at 4 C and autoclave it prior to its use. GNPs prepared using this method have a lmax of 518 nm, with an average diameter of approximately 12–13 nm. (One nanometer is approximately one ten-thousandth (1/10,000) of the width of a human hair.) (5) The presence of a colloidal suspension can be detected by the reflection of a laser beam from the particles. Because a laser pointer emits polarized light, the pointer can be oriented such that the beam appears to disappear. The beam from the laser is visible in one view, while it is invisible in the view perpendicular to the first view. Five to ten drops of 1 M NaCl solution are added to the tube on the right. Note the color change of the solution as the nanoparticles get closer together.

14.2.4.2

Preparation of Gold Nanoparticle Probe

1. Synthesize 50 -(T)30-(CH2)6-S-S-30 (by Invitrogen) 2. Centrifuge 0.5 ml GNP solution at 14,000 rpm for 10 min, removing the supernatant aliquot carefully 3. Add 50 ml of 300 pM thiol-modified oligonucleotides to the tube to re-disperse the GNP 4. Incubate the solution at 4 C for 12 h, followed by addition of another 50 ml of TE buffer, and then store at 4 C for 24 h

222

14 Gold Nanoparticle Probe Method for miRNA Quantification

5. Transfer the supernatant to a new tube and centrifuge at 14,000 rpm for 10 min 6. Remove the supernatant carefully and wash the pellet with 0.15 M NaCl, 10 mM phosphate (pH 7.4) 7. Centrifuge at 14,000 rpm for 10 min and remove the supernatant 8. Wash the pellet twice to remove unconjugated oligonucleotides from the pellet 9. Finally, re-disperse the pellet in 50 ml of buffer containing 0.15 M NaCl, 10 mM phosphate (pH 8.5) and store at 4 C for use. For a more detailed description of the preparation of GNP-probes, please consult the protocols developed by Mirkin et al. (1996) and Nam et al. (2004).

14.2.4.3

Preparation of miRNA-Specific Biotin Probe (MSB-Probe)

1. Synthesize a miR-1-specific DNA oligomer in HPLC grade (by companies): 50 -Bio-(CH2)6-CTCCATACTTC-30 2. Dissolve MSB-probe in TE buffer at 100 nM 3. Aliquot MSB-probe and store at 80 C for later use

14.2.4.4

Preparation of miRNA-Specific Poly(A) Probe (MPA-probe)

1. Synthesize another miR-1-specific DNA oligomer in HPLC grade (by companies): 50 -TTTACTTACC-(A)30-30 2. Dissolve MSB-probe in TE buffer at 100 nM 3. Aliquot MSB-probe and store at 80 C for later use

14.2.4.5

Preparation of Total RNA Sample

(1) Prepare total RNA samples from cultured cells or tissues using TRIzol reagent according to the manufacturer’s recommended protocol (2) Use a spectrophotometer to estimate the concentration of extracted RNA samples from the absorption at 260 nm (assuming that 1 A260 ¼ 40 mg/mL) (3) Store the samples at 80 C for use (4) Dilute the RNA samples to a final concentration of 20 mg/mL before use

14.2.4.6 1. 2. 3. 4. 5.

Preparation of Cell Lysis

Suspend approximately 1 106 cells Pellet the cells by centrifugation at 1,000 g for 3 min Wash once with 1 ml PBS (no MgCl2, no CaCl2; Invitrogen) Spin down at 1,000g for 3 min Suspend the cell pellet in 100 ml of 10 mM MOPS (pH 7.5), 100 mM KCl

14.2 Protocol

223

6. Take 2 ml cell aliquot and dilute with 98 ml of lysis buffer containing 20 mM Tris–HCl (pH 8.5), 0.5% NP-40, 20 mg/mL tRNA (in the absence of added divalent cations) 7. Heat the sample for 15 min at 80 C 8. Centrifuge at 1,000 g for 3 min to remove debris

14.2.4.7

Hybridization

1. Mix the following: l l l l

2.20 mg/ml total RNA sample 3.1 ml of 100 nM MSB-probe 4.1 ml of 100 nM MPA-probe 5.20 ml of TE buffer (pH 8.5)

2. Keep the mixture at 25 C for 10 min 3. Add 3 ml of GNP-probe to the mixture and incubate for 10 min at 25 C 4. Transfer the mixture to the streptavidin-coated microplate and incubate at 25 C for 30 min 5. Wash the microplate with 1 PBS solution for 3 min 6. Wash twice with 2 PBN (0.3 M NaNO3 and 10 mM Na2HPO4/NaH2PO4 buffer, pH 7.0) for 3 min each time It is suggested that the hybridization temperature and the rinse temperature should be optimized for each specific miRNA (Yang et al. 2008). 14.2.4.8

Preparation of Silver Enhancing Solution (SES)

1. Mix the following components at a ratio of 75:25:3: l l

l

Hydroquinone (0.085 g in 1.5 ml bidistilled water) Citrate buffer (0.255 g citric acid, 0.235 g trisodium citrate dehydrate in 1.0 ml bidistilled water, pH 3.8) Silver nitrate (AgNO3, 0.25 g in 1.0 ml bidistilled water)

2. Vortex the mixture vigorously until the solution turns transparent 14.2.4.9

Quantification of miRNA by Gold Nanoparticle Probes

1. Add 100 ml silver enhancing solution (SES) into each microplate well simultaneously 2. Keep the reaction at room temperature in dark for around 150 s 3. Terminate the reaction by rinsing the wells with bidistilled water 4. Measure the degree of surface silver gray scale with colorimetric absorbance by a microplate reader

224

14 Gold Nanoparticle Probe Method for miRNA Quantification

14.2.4.10

Quantification of miRNA by Stem-Loop RT-PCR

cDNA of miR-122a was synthesized using a TaqMan MicroRNA Reverse Transcription Kit (Part: 4366596; Applied Biosystem Inc.) and subsequently quantified on an Applied Biosystems 7900HT Sequence Detection System using a TaqMan miRNA Assay Kit (Part: 4373151; Applied Biosystem Inc.) according to the manufacturer’s protocol.

14.3

Application and Limitation

The GNP technique is a unique assay for miRNA detection and quantification based on GNP-probe with silver enhancement, which has been proven to be a sensitive, specific, and simple method. The lower detection limit is 10 fM miRNA or 2 ng of total RNA. Moreover, GNP-probes possess several properties that make them ideal for diagnostic applications: 1. Allows increased sensitivity by several orders of magnitude; light scattered from one nanoparticle is equivalent to the light emitted from 5 105 fluorophores. Tests that employ GNP functionalized with antibodies, for example, are 2–3 orders of magnitude more sensitive than ELISA-based methods. 2. Enables high specificity for miRNA detection and is able to distinguish one single nucleotide mismatch. The single-base pair specificity is achieved by assay reaction kinetics where GNP-probes, comprised of target-specific oligonucleotides, permit hybridization to target miRNA over a very narrow temperature range [43]. 3. The method offers an easy, quantitative detection of miRNAs. 4. Reduces background noise (i.e., signal-to-signal) due to minimal nonspecific binding of the GNP-probes, which in turn, create an enhanced assay signal. 5. The nanoparticle probes are extremely stable, have a long shelf-life, and are nontoxic. 6. The method does not require an expensive equipment or an advanced read-out system, and can be performed in any standard laboratory. The GNP technique has been applied to characterize the distribution of miR122a and miR-128 in mouse brain and liver tissue, and to quantify the synthetic miR-122a (Yang et al. 2008).

References Mirkin CA, Letsinger RL, Mucic RC, Storhoff JJ (1996) A DNA-based method for rationally assembling nanoparticles into macroscopic materials. Nature 382:607–609 Nam JM, Stoeva SI, Mirkin CA (2004) Bio-bar-code-based DNA detection with PCR-like sensitivity. J Am Chem Soc 126:5932–5933

References

225

Rao RCN, Kulkarni GU, Thomasa PJ, Edwards PP (2000) Metal nanoparticles and their assemblies. Chem Soc Rev 29:27–35 Yang WJ, Li XB, Li YY, Zhao LF, He WL, Gao YQ, Wan YJ, Xia W, Chen T, Zheng H, Li M, Xu SQ (2008) Quantification of microRNA by gold nanoparticle probes. Anal Biochem 376:183–188 Zhao L, Li, Hei, Li X, Li Y, Xu S (2006) A colorimetric polynucleotides detection method based on nanoparticle probes with silver staining enhancement. Chin J Biochem Mol Biol 22:919–923

Part VIII

Other Methods

Chapter 15

Splinted Ligation Method

Abstract The Splinted-ligation technology was initially developed by Moore and Query (Methods Enzymol 317:109–123, 2000; Center for RNA Molecular Biology and Department of Biochemistry, Case Western Reserve University, School of Medicine, Ohio, USA) for joining two or more RNA fragments by including bridging splint DNA templates to create RNA:RNA/DNA complexes which are nicked specifically at the desired site of ligation substrates of the T4 DNA ligase. This strategy was later adapted by Maroney et al. (Nat Struct Mol Biol 13:1102– 1107, 2006, RNA 13:930–936, 2007 and Nat Protoc 3:279–287, 2008) for detecting miRNA expression. The Splinted-ligation technology is a protocol for the direct labeling and quantitative detection of small RNAs, based on nucleic acid hybridization technology for characterizing small RNAs of known sequence, such as mature miRNAs. Compared with Northern blotting, the Splinted Ligation technique possesses the advantage of simplicity: it takes advantage of liquid hybridization kinetics and avoids the transfer, prehybridization, and washing steps required for Northern blotting. It also allows easy processing of multiple samples. The Splinted Ligation technology is ~50 times more sensitive than Northern blotting using DNA probes (RNA 13:930–936, 2007). The technology is convenient and straightforward, and does not require specialized, sophisticated equipment or any amplification step. The Splinted-ligation technology has the same applications as Northern blot and many other methods such as expression analysis and validation of microarray studies. It has been successfully used to detect and quantify different classes of small RNAs in unfractionated RNA samples from nanogram to microgram amounts of total RNA without an amplification step.

Z. Wang and B. Yang, MicroRNA Expression Detection Methods, DOI 10.1007/978-3-642-04928-6_15, # Springer-Verlag Berlin Heidelberg 2010

229

230

15.1

15 Splinted Ligation Method

Introduction

The splinted-ligation technique is a protocol for the direct labeling and quantitative detection of small RNAs (Moore and Query 2000), based on nucleic acid hybridization technology for characterizing small RNAs of known sequence, such as mature miRNAs (Maroney et al. 2007, 2008). The technology requires two properly designed oligonucleotide fragments: bridge oligonucleotide and ligation oligonucleotide (Fig. 15.1). Another requirement is that the small RNAs must have a 30 -hydroxyl group to permit ligation between the small RNA and the 50 -phosphate group of the ligation oligonucleotide; miRNAs meet this requirement. The procedures start with a nucleic acid hybridization assay with a bridge oligonucleotide based on perfect Watson–Crick complementarity to a target small RNA and a 50 -end-radiolabeled ligation oligonucleotide. For quantitative detection of miRNAs, simultaneous basepairing of both a miRNA and a ligation oligonucleotide to a bridge oligonucleotide yields a double-stranded structure with a nick on one strand, which can be subsequently ligated by T4 DNA ligase, thus labeling the target miRNA. Then, the label present on the unligated oligonucleotide can be removed by incubation with phosphatase after the ligation step, while the labeled phosphate in the ligation oligonucleotide is kept intact as it has become resistant to phosphatase activity. Following the splinted-ligation reaction, labeled miRNAs carrying a nucleotide extension equivalent to the length of the ligation oligonucleotide and any residual-labeled ligation oligonucleotides can then be separated by denaturing gel electrophoresis and visualized by autoradiography or phosphorimaging (Figs. 15.1 and 15.2).

15.2

Protocol

15.2.1 Materials 1. Filtered tips (aerosol-resistant pipette tips, RNase free) 2. A 13 15 cm 0.75 mm gel (for a tall-gel apparatus) 3. A 36 43 cm 0.8 mm gel (for a sequencing gel system)

15.2.2 Instruments 1. Thermalcycler PCR machine 2. Vertical gel-electrophoresis apparatus for SDS-PAGE or DNA-sequencing applications 3. Electrophoresis power supply for SDS-PAGE or DNA-sequencing applications

15.2 Protocol

231 Ligation Oligonucleotide (L-Oligo)

CGCTTATGACATTC OH

PO4

T4 Poly-nt Kinase

32

P

L-Oligo labelling

ADP

miR-1

UGGAAUGUAAAGAAGUAUGUAU OH

PO4

+

PCGCTTATGACATTC OH Labelled L-Oligo

32

Bridge Oligonucleotide (B-Oligo)

3'-ACCTTACATTTCTTCATACATTGCGAATACTGTAAG-5’ miR-1 lapturing 32 CGCTTATGACATTC OH UGGAAUGUAAAGAAGUAUGUAU P OH |||||||||||||||||||||| |||||||||||||| 3'-ACCTTACATTTCTTCATACATTGCGAATACTGTAAG-5’ PO4

miR-1—L-Oligo ligation

T4 Ligase

Ligated miR-1

Ligated L-Oligo

UGGAAUGUAAAGAAGUAUGUAU PCGCTTATGACATTC OH |||||||||||||||||||||||||||||||||||| 3'-ACCTTACATTTCTTCATACATTGCGAATACTGTAAG-5’ PO4

Unligated L-Oligo

32

32

32

PCGCTTATGACATTC OH

32 CGCTTATGACATTC OH P PCGCTTATGACATTC OH

Post-ligation phosphatase preatment Denaturing Gel for letecting miR-1 by autoradiography / phosphorimaging PO4

CGCTTATGACATTC OH

UGGAAUGUAAAGAAGUAUGUAU P CGCTTATGACATTC 32

CGCTTATGACATTC OH CGCTTATGACATTC OH

OH

Fig. 15.1 Schematic depicting the principle and procedures of the splinted-ligation technology modified from Maroney et al. (2008). T4 Poly-nt kinase: T4 polynucleotide kinase; T4 ligase: T4 DNA ligase

Total RNA extraction

Design and synthesize bridge oligonucleotide

Synthesize of radiolabelled ligation oligonucleotide

miRNA capture by the probes

Ligation of miRNA and probes

Phosphatase treatment

Urea-polyacrylamide gel electrophoresis

Image analysis

Fig. 15.2 Flowchart of the splinted-ligation technique for miRNA detection. According to Maroney et al. (2008)

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15 Splinted Ligation Method

4. Autoradiography film cassette and Kodak BioMax MR film (Kodak) or cassette and storage phosphor screen 5. Film developer for film autoradiography 6. Phosphorimager instrument storage phosphor screen phosphorimaging

15.2.3 Reagents 1. Ligation oligonucleotide (50 -CGCTTATGACATTC/dC-30 ) 2. Bridge oligonucleotide 3. Synthetic RNA positive control (synthetic RNA oligonucleotide corresponding to the sequence of a known small RNA) 4. Bridge oligonucleotide for synthetic RNA positive control 5. Low molecular weight markers, 10–100 nt 6. [g-32P]-ATP (6,000 Ci/mM, 150 mCi/mL) (Perkin Elmer) 7. OptiKinase (10 U/mL) with 10 OptiKinase reaction buffer (0.5 M Tris–HCI; pH7.5, 100 mM MgCl2 and 50 mM DTT) 8. 10 Capture buffer (100 mM Tris–HCl (pH7.5) and 750 mM KCl) 9. PrepEase sequencing dye clean-up kit (USB). Other gel matrix spin columns for nucleic acid clean-up and purification can be used 10. Ligate-IT rapid ligation kit (USB). Other commercially available T4 DNA Ligase kits can be used, but the ligation efficiency may be affected by the difference in buffer compositions (If a rapid ligation kit is not used, the following buffer can be used (final concentrations indicated), 10 mM MgCl2, 1 mM DTT, 1 mM ATP, 7.5% (wt/vol) PEG 6000, 66 mM Tris; pH7.5. In this case, the concentration of T4 DNA ligase needed for optimal ligation efficiency must be determined empirically) 11. RNase Inhibitor (Human Placenta) 12. Shrimp alkaline phosphatase (1 U/ml) 13. 2 Formamide loading dye (95% formamide, 20 mM EDTA, 0.025% bromophenol blue and 0.025% xylene cyanol) 14. RapidGel 40% liquid acrylamide stock solution (19:1) 15. 30% Acrylamide/bis solution (29:1) 16. Urea 17. Glycerol-tolerant gel buffer, 20 solution (1.78 M Tris, 0.57 M taurine and 0.01 M EDTA) 18. TBE buffer, 5 solution (Tris–Borate–EDTA buffer: 0.445 M Tris, 0.445 M boric acid and 0.01 M EDTA) 19. Ammonium persulfate 20. TEMED 21. TE buffer, 1 solution (10 mM Tris–HCl (pH7.5) and 1 mM EDTA) 22. TRIzol reagent (Invitrogen) 23. Phenol:chloroform 24. Glycogen

15.2 Protocol

233

25. PureLink miRNA isolation kit (Invitrogen) 26. Urea–polyacrylamide gel

15.2.4 Procedures The protocols described in this section are essentially the same as reported in the study by Maroney et al. (2007, 2008) (See Figs. 15.1 and 15.2).

15.2.4.1

Total RNA Sample Preparation

Prepare total RNA using guanidine isothiocyanate such as TRIzol reagent and phenol:chloroform according to standard total RNA isolation protocols with the exception that an inert carrier such as glycogen or linear polyacrylamide is added to each sample. Maroney et al. (2007, 2008) recommend adding 20 mg glycogen per 1 mL during alcohol precipitation to increase the recovery of small RNAs. Samples can also be prepared by commercially available column-based methods for small RNA isolation. Dilute RNA sample with TE buffer or RNase-free water. The purified RNA should be kept at 80 C. Avoid leaving the RNA at room temperature (25 C) or 4 C and avoid multiple freeze–thaw cycles after isolation.

15.2.4.2

Urea–Polyacrylamide Gel Preparation

A 13 15 cm 0.75 mm gel (for a tall-gel apparatus) requires 15 mL gel solution and a 36 43 cm 0.8 mm gel (for a sequencing gel system) requires 120 mL gel solution. A mini-protein gel system requires 20 mL gel.

12% Tall-Gel 1. Add the following: Urea (7 M) 40% Acrylamide/bis solution (19:1) 5 Tris–borate–EDTA (TBE) buffer

2. 3. 4. 5.

6.3 g 4.5 mL 3 mL

Adjust to the final volume with nuclease-free water Stir and warm solution at 40–50 C to dissolve urea Cool the mixture to room temperature Add the following reagents immediately before pouring the gel: TEMED 10% ammonium persulfate in nuclease-free water

7.5 mL 75 mL

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15 Splinted Ligation Method

6. Allow to polymerize at room temperature for at least 30 min to 1 h 7. Run in 1 TBE running buffer (diluted with deionized water) Total volume:

15 mL

12% Sequencing Gel 1. Add the following: Urea (7 M) 40% Acrylamide/bis solution (19:1) 5 Tris–borate–EDTA (TBE) buffer

2. 3. 4. 5.

50.4 g 36.0 mL 24.0 mL

Adjust to the final volume with nuclease-free water Stir and warm solution at 40–50 C to dissolve urea Cool the mixture to room temperature Add the following reagents immediately before pouring the gel: TEMED 10% ammonium persulfate in nuclease-free water

60 mL 600 mL

6. Allow to polymerize at room temperature for at least 30 min to 1 h 7. Run in 1 TBE running buffer (diluted with deionized water) Total volume:

120 mL

16% Mini Gel 1. Add the following: Urea (7 M) 30% Acrylamide/bis solution (29:1) 20 glycerol-tolerant gel (GTG) buffer

8.4 g 11 mL 1 mL

2. Adjust to the final volume with nuclease-free water 3. Stir and warm solution at 40–50 C to dissolve urea TEMED 10% ammonium persulfate in nuclease-free water

4. 5. 6. 7.

10 mL 100 mL

Cool the mixture to room temperature Add the following reagents immediately before pouring the gel Allow to polymerize at room temperature for at least 30 min to 1 h Run in 1 TBE running buffer (diluted with deionized water) Total volume:

20 mL

15.2 Protocol

15.2.4.3

235

Design and Preparation of Bridge Oligonucleotide

The bridge oligonucleotide is a DNA oligonucleotide complementary to both the ligation oligonucleotide and a specific small RNA at its 50 - and 30 -ends, respectively (Fig. 15.1). Thus, the bridge oligonucleotide sequence is designed to contain (1) a ~22-nt sequence at the 30 -end complementary to the miRNA of interest and (2) a 14nt sequence complementary to the ligation oligonucleotide) at the 50 -end. This bridge oligonucleotide allows a single labeling reaction of the ligation oligonucleotide for detecting any miRNA of interest. In general, adding unligatable modifications to the ends of the bridge oligonucleotide is not always necessary; however, in some cases it is desirable to block the 30 -end or both the 50 - and the 30 -ends of the bridge oligonucleotide by incorporating modification(s) such as C3 spacer, amino-modifier, inverted dT or dideoxy-C. This ensures that unwanted side ligation reactions do not take place. Maroney et al. (2007, 2008) recommend using an unmodified bridge oligonucleotide as the first option for the assay. The bridge oligonucleotide requires a standard desalting purification after synthesis. Further purification by HPLC or denaturing PAGE is usually unnecessary. Resuspend the bridge oligonucleotide with TE buffer or RNase-free water to 100 mM and store at 20 C. Dilute the stock solution to 100 nM (0.1 pM/ml) with10 capture buffer and use 1 mL in a 10-mL assay reaction. The bridge oligonucleotides may be prediluted to 1 mM with RNase-free water before preparing the 100 nM stock solution in 10 capture buffer. The concentration of the bridge oligonucleotide could affect the output signal independent of the amount of RNA in the test sample. For most applications, the concentration of the bridge oligonucleotide should be carefully measured to yield 0.1 pmol per reaction. It is important that the concentrations of bridge and ligation oligonucleotides are equivalent. Therefore, if the RNA to be quantitated is highly abundant, total RNA should be diluted before the assay. If the RNA to be quantitated is rare, more total RNA should be used or total RNA should be prefractionated to enrich for small RNAs.

15.2.4.4

Preparation of Ligation Oligonucleotide

1. A ligation oligonucleotide is designed to contain 14 nts (50 -CGCTTATGACATTC-30 ), which is unique and will not basepair any miRNA or proteincoding genes (Fig. 14.1). Previous studies reported splints that extend 8–10 nucleotides (nt) on either side of the junction as sufficient for quantitative ligation, and that ligation efficiencies fall off dramatically with shorter templates (Moore and Query 2000). Synthesize the ligation oligonucleotide using services provided by commercial companies. 2. 50 -End label the ligation oligonucleotide with [g-32P]-ATP and remove the unincorporated isotope. Resuspend the ligation oligonucleotide with TE buffer or RNase-free water to 100 mM and store at 20 C. Dilute the stock solution to 10 mM with TE buffer or RNase-free water for the 50 -end-labeling reaction.

236

15 Splinted Ligation Method

3. Thaw frozen reagents on ice, mix thoroughly followed by a spin at a speed of 16,000 g in a microcentrifuge for 10 s at room temperature, and then place on ice. 4. Prepare [32P]-labeled ligation oligonucleotide by combining the following components at room temperature (Replace the ligation oligonucleotide with marker such as low molecular weight marker (100 ng per fragment and 1 mg per labeling reaction) when preparing radiolabeled markers): 10 mM Ligation oligonucleotide RNase-free water 10 OptiKinase reaction buffer [g-32P]-ATP (6,000 Ci/mM, 150 mCi/ml) OptiKinase Total volume

2 mL 12 mL 2 mL 2 mL 2 mL 20 mL

5. Mix thoroughly followed by a spin at maximum speed in a microcentrifuge for 10 s at room temperature. Incubate for 30–60 min at 37 C. 6. While the reactions are incubating, prepare the PrepEase sequencing dye cleanup kit for removing the unincorporated [-32P]-ATP. Centrifuge the column at 750 g for 30 s at room temperature to collect the dry resin at the bottom of the column. 7. Hydrate the resin by adding 600 mL RNase-free water and vortex. Remove air bubbles by vortexing or tapping the column. Incubate for at least 30 min at room temperature (the column can be hydrated overnight at 4 C). 8. Resuspend the settled resin by inverting the column several times. Ensure that no air bubbles are visible. Remove the bottom plug and place in a 2-ml collection tube. 9. Centrifuge at 750 g for 2 min at room temperature to remove the remaining water. Discard the flow-through. 10. After incubating for 30–60 min, dilute the labeling reactions to 100 mL by adding 80 mL RNase-free water. 11. Place the column from (8) in a clean 1.5-ml microcentrifuge tube. Without disturbing the gel bed, carefully apply the diluted sample (from (9) directly onto the top of the gel bed). 12. After loading the sample, centrifuge the column at 750 g for 4 min at room temperature. Discard the used column in a radioactive waste container. The concentration of the labeled detection oligonucleotide should be 100 nM (0.1 pM/ml). Store at 20 C if not required immediately. Keep on ice when in use.

15.2.4.5

miRNA Capture, Ligation, and Phosphatase Treatment

1. Thaw frozen reagents (for Step 12) on ice, mix thoroughly followed by a spin at maximum speed in a microcentrifuge for 10 s at room temperature, and then place on ice.

15.2 Protocol

237

2. Assemble the capture reaction on ice by making a master mix of 1 mL of bridge oligonucleotide in 10 capture buffer plus 1 mL radiolabeled ligation oligonucleotide per sample. Add 2 mL of this master mix to each test sample, which had been diluted to 8 mL with RNase-free water: RNA sample Synthetic RNA positive control Adjust to 8 mL with RNase-free water 0.1 pmol/mL Bridge oligonucleotide in 10 capture buffer Radiolabeled ligation oligonucleotide Total volume

0 mL 1 mL 1 mL 1 mL 10 mL

3. Mix thoroughly followed by a spin at a speed of 16,000g in a microcentrifuge for 10 s at room temperature. 4. Incubate the mixture at 94 C for 1 min, 65 C for 2 min and 37 C for 10 min in a thermalcycler PCR machine. 5. Make a ligase mastermix by combining 3 mL of 5 Ligate-IT buffer, 1 mL Ligate-IT enzyme, and 1 mL RNase-free water, per sample. Add 5 mL of the ligase mastermix to each sample. 6. Mix thoroughly followed by a spin at a speed of 16,000g in a microcentrifuge for 10 s at room temperature. Incubate for 1 h at 30 C in a thermalcycler PCR machine. 7. Add 1 mL shrimp alkaline phosphatase to each reaction. 8. Mix thoroughly followed by a spin at a speed of 16,000g in a microcentrifuge for 10 s at room temperature. Incubate for 15 min at 37 C.

15.2.4.6

Electrophoretic Analysis

1. Prepare a 12 or 15% urea–polyacrylamide gel with 1 running buffer 2. Prerun and warm the gel for 20–30 min 3. Transfer an aliquot (up to 15 mL) of the reaction (from Step xx) into a new tube. Add an equal volume of formamide loading dye 4. Transfer an aliquot (up to 15 mL) of the recommended [32P]-labeled low molecular weight marker into a new tube. Add an equal volume of formamide loading dye 5. Mix the tubes from Steps 20 and 21 thoroughly followed by a brief spin in a microcentrifuge and incubate for 3 min at 95 C to denature the samples. Cool on ice immediately 6. Flush wells of the gel thoroughly to remove acrylamide debris, urea, and air bubbles 7. Load 2–15 mL of samples (from Step 20) onto the gel and include the [32P]labeled markers (from Step 21) on each gel 8. Run the gel at 20–30 mA for a 13 15 cm gel, or at 60 mA for a 36 43 cm gel and stop when the bromophenol blue dye front has migrated to the bottom, or middle of the gel for these gel sizes, respectively

238

15 Splinted Ligation Method

9. At the end of the electrophoretic separation, RNA detection can be completed using either a phosphorimager (option A) or an X-ray film (option B) (a) Detection of RNA using phosphorimaging l

l

Transfer the gel onto a sheet of paper, dry in a gel dryer, wrap with Saran wrap, and expose to a phosphorimager screen Process the phosphorimager screen according to the manufacturer’s instructions

(b) Detection of RNA using X-ray film l

l

Transfer the gel onto a sheet of nondiffusible support material, such as processed film, wrap with Saran wrap and expose to X-ray film Expose the gel to X-ray film with an intensifying screen. Store for 2 h to overnight at 80 C. The gel can be re-exposed several times if required

The entire procedure can be completed in a single day.

15.2.4.7

Control Experiments

1. Prepare a “positive control” to assess assay components and procedure by substituting the RNA sample with a synthetic RNA positive control. The positive control is a premix of a 20–30-nt synthetic RNA oligonucleotide and a bridge oligonucleotide for capturing the synthetic RNA. Resuspend the synthetic RNA positive control oligonucleotide with TE buffer or RNase-free water to 100 mM and store at 80 C. Dilute the stock solution to 0.2–20 nM (0.2–20 fM/ mL) with TE buffer or RNase-free water for standard and positive control reactions. 2. Prepare a “no RNA negative control” to assess sample background signal by substituting the RNA sample with RNase-free water. 3. For use as an “internal/loading control,” it is desirable to detect small RNAs known to be constitutively expressed in the test samples by substituting the target-specific bridge oligonucleotide with a control-specific bridge oligonucleotide.

15.3

Application and Limitation

Compared with Northern blotting, the Splinted-ligation technique possesses the advantage of simplicity: it takes advantage of liquid hybridization kinetics and avoids the transfer, prehybridization, and washing steps required for Northern blotting. It also allows easy processing of multiple samples. The Splinted-ligation technology is ~50 times more sensitive than Northern blotting using DNA probes

References

239

(Maroney et al. 2007). The technology is convenient and straightforward, and does not require specialized, sophisticated equipment or any amplification step. The Splinted-ligation technology has the same applications as Northern blot and many other methods such as expression analysis and validation of microarray studies. It has been successfully used to detect and quantify different classes of small RNAs in unfractionated RNA samples from nanogram to microgram amounts of total RNA without an amplification step and was shown to be consistent with previously determined tissue-specific miRNA expression patterns in animals and plants, the expression of viral miRNAs in infected fibroblasts, the testis-specific expression of piRNAs and the expression of low abundance ta-siRNAs and other small RNAs in Arabidopsis (Hu¨ttenhofer and Vogel 2006). This method was also used to determine miR-21 distribution on polyribosomes before and after hypertonic stress (Maroney et al. 2006), and other miRNAs as well, including, miR-1, miR-9, miR16, miR-20a, miR-21, miR-26a, miR-124a, and miR-375 (Maroney et al. 2007). The major disadvantages of the Splinted-ligation technology are the obligation of radioactive labeling, relatively low sensitivity compared to qRT-PCR methods, and non-high-throughput.

References Maroney PA, Chamnongpol S, Souret F, Nilsen TW (2007) A rapid, quantitative assay for direct detection of microRNAs and other small RNAs using splinted ligation. RNA 13:930–936 Maroney PA, Chamnongpol S, Souret F, Nilsen TW (2008) Direct detection of small RNAs using splinted ligation. Nat Protoc 3:279–287 Maroney PA, Yu Y, Fisher J, Nilsen TW (2006) Evidence that microRNAs are associated with translating messenger RNAs in human cells. Nat Struct Mol Biol 13:1102–1107 Moore MJ, Query CC (2000) Joining of RNAs by splinted ligation. Methods Enzymol 317:109–123 Hu¨ttenhofer A, Vogel J (2006) Experimental approaches to identify non-coding RNAs. Nucleic Acids Res 34:635–646

Chapter 16

Padlock-Probes and Rolling-Circle Amplification

Abstract The padlock-probes and rolling-circle amplification technology is a simple, sensitive, and reliable miRNA expression detection protocol. A padlockprobe is a linear DNA probe where the two terminal aims are designed to be exactly antisense to the 5’-end and 3’-end sequences of a target miRNA of interest. Rolling circle amplification allows linear amplification of the target sequence in a highly quantitative manner without relying on thermocyclers or other advanced read-out systems. The technology was initially developed by Nilsson et al. (Science 265:2085–2088, 1994); Department of Molecular Biology, University of Aarhus, Denmark for detecting localized DNA, and was later modified for detection and distinction of RNA by the same group (Hum Mutat 19:410–415, 2000; Hum Mutat 19:410–415, 2002). Recently, Jonstrup et al. (RNA 12:1747–1752, 2006) further developed the methods into a unique technology for detecting, amplifying, and quantifying miRNA expression. The technology is a simple and reliable miRNA detection protocol and can be performed without specialized equipment and is capable of measuring the content of specific miRNAs in a few nanograms of total RNA. The techniques have been validated for six miRNAs, namely miR-16, miR17-5p, miR-20a, miR-21, miR-27a, and miR-92.

16.1

Introduction

The padlock-probes and rolling-circle amplification technology is a simple, sensitive, and reliable miRNA detection protocol. A padlock-probe is a linear DNA probe where the two terminal aims are designed to be exactly antisense to the 5’end and 3’-end sequences of a target miRNA of interest (Jonstrup et al. 2006). The padlock-probe annealing to the target miRNA is circularized, leaving a nick between the head and tail of the probe (Fig. 16.1). Upon addition of DNA ligase, ligation will occur between the termini of the padlock-probe on a perfectly matching miRNA template (Nilsson et al. 1994, 2000; Jonstrup et al. 2006). Following

Z. Wang and B. Yang, MicroRNA Expression Detection Methods, DOI 10.1007/978-3-642-04928-6_16, # Springer-Verlag Berlin Heidelberg 2010

241

242

16 Padlock-Probes and Rolling-Circle Amplification

miR-1

a

5’-UGGAAUGUAAAGAAGUAUGUAU-3’

miR-1 Padlock Probe

5’-TTTACATTCCATTTATTTCCTCAATGCTGCTGCTGTACTACTAGTGATTTACTTGGATGTCTATACATACTTC-3’

b miRNA

Rolling Circle

Padlock Probe Ligation (DNA ligase)

Extension (DNA polymerase)

Amplification (DNA polymerase)

Fig. 16.1 Schematic drawing showing the principle of padlock probe recognition of miRNAs and the rolling circle amplification. (a) Design of padlock probe. (b) Procedures of padlock-probe rolling-circle amplification. (i) Padlock probes are designed to specifically recognize a particular miRNA. (ii) The padlock probes annealing to the miRNA are circularized upon addition of DNA ligase. (iii) After ligation, the annealed miRNA serves as a primer for extension by a phi29 DNA polymerase. (iv) The phi29 DNA polymerase facilitates rolling circle amplification, thereby producing a DNA product containing multiple copies of the miRNA sequence. Modified from Nilsson et al. (2000)

ligation, the miRNA that is used as a template can subsequently be used as primer for rolling circle amplification, thereby linearly amplifying the target sequence in a process that has been shown to be highly quantitative (Nilsson et al. 2002). Within such a theme, the padlock-probe is used as a probe to hybridize to target the miRNA of interest and then as a template for rolling-circle amplification; miRNA to be detected is initially used as a template for ligation of padlock-probe and subsequently as a primer for rolling-cycle amplification. The ligation is highly specific for the perfectly matched duplex between the padlock-probe and the target miRNA, thereby accurately distinguishing matched and mismatched substrates. It has previously been shown that padlock-probes are very sensitive to mismatches when placed at the ligated junction (Nilsson et al. 2000). The method, therefore, has the power to distinguish between closely related miRNAs, especially if the miRNAs differ at a position near the middle region of the miRNA. Jonstrup et al. (2006) demonstrated that the padlock-probe is able to discriminate between miR-17–5p (5’-CAAAGUGCUUACAGUGCAGGUAG-3’)

16.3 Materials

243

and miR-20a (5’-UAAAGUGCUUAUAGUGCAGGUAG-3’) that differ only in their 50 and 30 terminus and by a single C to U change in the middle of the mature miRNAs. Indeed, both miR-17-5p padlock-probe (pad-miR-17–5p) and pad-miR20a have the ability to discriminate between miR-17–5p and miR-20a. Moreover, the amplification strategy ensures yet another level of specificity. Since the miRNA is used as a primer, only duplexes that are perfectly matched at the 30 -end of the miRNA will be a substrate for phi29 DNA polymerase. This should also prevent any signal from genomic DNA or pri-miRNA contaminations. Whereas other miRNA expression detection technologies require advanced laboratory equipment and chemically modified oligonucleotides. The padlock-probes and rolling-circle amplification technology requires inexpensive DNA oligonucleotides and only equipment no more advanced than for Northern blotting.

16.2

Protocol

Nilsson et al. (2000, 2002)

16.3

Materials

1. Padlock-probes (miRNA-specific; chemically synthesized with a phosphate at the 50 -end) 2. 8% polyacrylamide gel (100 mM Tris Borate, pH7.5, 1 mM EDTA) 3. 2 SSC-soaked hybond N þ filter (Amersham) 4. P32 50 -end-labeled DNA-probe antisense to the miRNA 5. Materials for Northern blot (see Chap. 3)

16.3.1 Instruments 1. A slot-blot apparatus 2. UV-X-linking instrument 3. Instruments for Northern blot (see Chap. 3)

16.3.2 Reagents 1. Trizol reagent (Invitrogen) 2. A mirVana miRNA isolation kit (Ambion) 3. mirVana miRNA probe construction kit (Ambion)

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16 Padlock-Probes and Rolling-Circle Amplification

Ligation buffer (10 mM Tris–HCl pH 7.5, 10 mM MgCl2, 10 mM ATP) T4-DNA ligase (New England Biolabs) 90% formamide P32 50 -end-labeled 25-bp DNA marker (Invitrogen) Rolling circle amplification buffer (50 mM Tris–HCl pH 7.5, 10 mM MgCl2, 10 mM (NH4)2SO4, 4 mM dithiothreitol, 200 mg/mL BSA, 10 U of phi29 DNA polymerase (New England Biolabs), and 0.2 mM dNTPs) 9. Reagents for Northern blot (see Chap. 3)

4. 5. 6. 7. 8.

16.3.3 Procedures The protocols described in this section are essentially the same as reported in the studies by Nilsson et al. (2000, 2002).

16.3.3.1

Design of Padlock-Probes

A padlock-probe is an oligodeoxynucleotide fragment containing three motifs: antisense to 5’-half of a miRNA sequence þ connector þ antisense to 3’-half of the miRNA sequence. Taking miR-1 as an example, miR-1 sequence is 5’UGGAAUGUAAAGAAGUAUGUAU-3’ and the padlock-probe for miR-1 will be 5’-TTTACATTCCATTTATTTCCTCAATGCTGCTGCTGTACTACTAGTGATTTAC TTGGATGTCTATACATACTTC-3’ (Fig. 15.1). For negative control, a padlock-probe with a mutation in each of the two arms needs to be constructed.

16.3.3.2

RNA Isolation

1. Extract total RNA with Trizol reagent 2. Enrich for small RNAs using a mirVana miRNA isolation kit, according to the manufacturer’s protocols or purified on an 8% polyacrylamide gel selectively for RNAs in the range of 15–30 nt (20-nt pool)

16.3.3.3

In vitro transcription of miRNAs

1. In vitro transcribe RNA using a mirVana miRNA probe construction kit, according to the manufacturer’s protocols.

16.3 Materials

16.3.3.4

245

Ligation Assay

1. Mix the following: Varying amounts of in vitro transcribed miRNAs 2.5 fM of P32 50 end-labeled of the corresponding padlock-probe 4.5 mL ligation buffer 2. Heat the samples at 65 C for 3 min, cool slowly to room temperature (RT) over 10 min 3. Add 200 units of T4-DNA ligase in a total reaction volume of 5 mL 4. Incubate the reactions at 37 C for 2 h 5. Add two volumes of RNA load buffer containing 90% formamide to stop the reactions 6. Analyze the ligation products on an 8% polyacrylamide gel alongside a P32 50 -endlabeled 25-bp DNA marker and 2.5 fM of untreated P32 50 -end-labeled pad-miRNA

16.3.3.5

Ligation Followed by Rolling Circle Assay

The ligation procedures are similar to the ones described above with the following modifications. 1. Use in vitro transcribed RNA, miRNA-enriched RNA, or gel-purified 15–30-nt fractions from total RNA depending on the experiment. If one wants to measure exactly mature miRNA, we recommend that a gel purification of miRNA-sized RNAs is performed first, especially if the mature miRNA is situated in the 30 arm of the pre-miRNA. 2. Add padlock-probes containing a non-radioactive 50 phosphate to the sample (instead of the radioactively labeled probes). 3. After the ligation, amplify the samples in a volume of 100 mL rolling circle amplification buffer. 4. Incubate the reactions for 8 h at 30 C followed by 10 min at 65 C to inactivate the polymerase. (A nearly linear relationship is expected between incubation time and signal, indicating that a strong gain in signal can be obtained after 8 h without compromising on linearity). 5. Transfer the samples to a 2 SSC-soaked hybond N þ filter using a slot-blot apparatus. 6. UV-X-link the DNA to the filter with 1,200 uJ. 7. Perform hybridization reactions as described with the Northern blots. Use a P32 50 -end-labeled DNA-probe antisense to the miRNA being detected to visualize the rolling circle amplification. 8. For internal control, an artificial miRNA is added at a fixed concentration immediately after cell lysis and subsequently measured using an artificial matching padlock-probe.

246

16 Padlock-Probes and Rolling-Circle Amplification

Fig. 16.2 Flowchart of the Padlock-Probes and RollingCircle Amplification technology for miRNA expression detection, according to Nilsson et al. (2000)

Select miRNAs of your interests

Design of Padlock miRNA capture probe

Isolate RNA from tissues or cells

Hybridization of miRNA and Padlock probe

Ligation of Padlock probe

Rolling circle amplification

Norteh blotting analysis

16.3.3.6

Northern Blots

Follow the protocols described in Chap. 3. Load RNA samples (20 mg total RNA or 1 mg miRNA-enriched RNA) on a 15% polyacrylamide gel. After running the gels, blot RNA onto a Hybond N+ membrane using a semidry blotter. Hybridize DNA probes complementary to the cloned miRNAs (Fig. 16.2).

16.4

Application and Limitation

16.4.1 Applications The techniques have been validated for six miRNAs including miR-16, miR-17-5p, miR-20a, miR-21, miR-27a, and miR-92, with total RNA derived from three different cell types including HEK293 human embryonic kidney cell, Chang liver cell, N2a murine neuroblastoma cell, and miRNA-enriched RNA. 1. A sensitive padlock-based method to detect and quantify miRNA expression. 2. Providing a simple low-tech method for analysis of miRNAs quantitatively in a few nanograms of total RNA. 3. The method can discriminate between closely related miRNAs and measure miRNA expression in a few nanograms of total RNA.

References

247

4. The inventors of padlock-probes and rolling-cycle amplification technology proposed that the technique could possibly be developed into an in situ hybridization assay to visualize miRNA localization in cells. No additional primer needs to be added since the miRNA itself can function as a primer. Such a method has the potential of being more sensitive than the conventional methods already developed for in situ miRNA detection (Wienholds et al. 2005; Kloosterman et al. 2006). Indeed, padlock-probes have been used for in situ hybridizations (Lizardi et al. 1998; Landegren et al. 2004;and Larsson et al. 2004). 5. Importantly, the rolling circle product contains repeated miRNA sense sequences, which can be detected using miRNA-specific oligonucleotide arrays. Hence, our method holds the possibility of being developed into an array assay combining several different padlock-probes in the same reaction.

16.4.2 Limitations 1. The method is probably not useful for quantification of plant miRNA, since this RNA contains a 20 -OMe at the 3-end, from which Phi29 will likely not extend. 2. It is also limited by the use of radioactive isotope.

References Jonstrup SP, Koch J, Kjems J (2006) A microRNA detection system based on padlock probes and rolling circle amplification. RNA 12:1747–1752 Kloosterman WP, Wienholds E, de Bruijn E, Kauppinen S, Plasterk RH (2006) In situ detection of miRNAs in animal embryos using LNA-modified oligonucleotide probes. Nat Methods 3:27–29 Landegren U, Nilsson M, Gullberg M, Soderberg O, Jarvius M, Larsson C, Jarvius J (2004) Prospects for in situ analyses of individual and complexes of DNA, RNA, and protein molecules with padlock and proximity probes. Methods Cell Biol 75:787–797 Larsson C, Koch J, Nygren A, Janssen G, Raap AK, Landegren U, Nilsson M (2004) In situ genotyping individual DNA molecules by target-primed rolling-circle amplification of padlock probes. Nat Methods 1:227–232 Lizardi PM, Huang X, Zhu Z, Bray-Ward P, Thomas DC, Ward DC (1998) Mutation detection and single-molecule counting using isothermal rolling-circle amplification. Nat Genet 19:225–232 Nilsson M, Baner J, Mendel-Hartvig M, Dahl F, Antson DO, Gullberg M, Landegren U (2002) Making ends meet in genetic analysis using padlock probes. Hum Mutat 19:410–415 Nilsson M, Barbany G, Antson DO, Gertow K, Landegren U (2000) Enhanced detection and distinction of RNA by enzymatic probe ligation. Nat Biotechnol 18:791–793 Nilsson M, Malmgren H, Samiotaki M, Kwiatkowski M, Chowdhary BP, Landegren U (1994) Padlock probes: Circularizing oligonucleotides for localized DNA detection. Science 265:2085–2088 Wienholds E, Kloosterman WP, Miska E, Alvarez-Saavedra E, Berezikov E, de Bruijn E, Horvitz HR, Kauppinen S, Plasterk RH (2005) MicroRNA expression in zebrafish embryonic development. Science 309:310–311

Chapter 17

Invader Assay

Abstract The invader miRNA assay was developed by Allawi et al. in Third Wave Technologies, Inc., Madison, Wisconsin, USA (RNA 10:1153–1161, 2004; Methods Mol Biol 488:279–318, 2008), based on a similar method – the invader mRNA assay – established earlier by the same group for detecting mRNAs (Nat Biotechnol 19:673–676, 2001; Expert Rev Mol Diagn 2:487–496, 2002). The invader miRNA assay has the ability to detect and quantitate as few as 20,000 molecules of an individual miRNA. It distinguishes between miRNAs and their precursors, as well as between closely related miRNA isotypes. This assay has been used in the analysis of several miRNAs, using as little as 50–100 ng of total cellular RNA or as few as 1,000 lysed cells. Its specificity allowed for discrimination between miRNAs differing by a single nucleotide, and between precursor and mature miRNAs. The invader miRNA assay, which can be performed in unfractionated detergent lysates, uses fluorescence detection in microtiter plates and requires only 2–3 h incubation time, allowing for parallel analysis of multiple samples in highthroughput screening analyses (de Arruda et al. 2002).

17.1

Introduction

The invader miRNA detection technology can be used to directly detect specific RNA molecules in preparations of pure total cellular RNA (1–100 ng) or in crude cell lysate (103–104 cells) samples using an isothermal signal amplification process with a fluorescence resonance energy transfer (FRET)-based fluorescence read-out. Roughly, the invader miRNA assay includes two consecutive reactions and a final fluorescence measurement. (1) Primary reaction – capturing the target miRNA: Annealing of the invasive and miRNA probe oligonucleotides to an miRNA target forms an overlap-flap structure that is a substrate for the structure-specific 50 nuclease, Cleavase. The noncomplementary 50 flap of the miRNA probe oligonucleotide is the site of cleavage by Cleavase, which releases the 50 flap. The invasive and miRNA probe oligonucleotides are so designed as to include stretches forming Z. Wang and B. Yang, MicroRNA Expression Detection Methods, DOI 10.1007/978-3-642-04928-6_17, # Springer-Verlag Berlin Heidelberg 2010

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a dumbbell-like structure or the stem–loop hairpin regions for better capturing the short size of an miRNA target. (2) Secondary reaction – generating quantifiable signals: A secondary overlap-flap structure is formed by hybridizing both the 50 flap that had been released in the primary reaction and a FRET oligonucleotide to a secondary reaction template (SRT). The FRET oligonucleotide is labeled with a fluorophore and a quencher so cleavage between them generates a fluorescence signal. An arrestor oligonucleotide complementary to the miRNA probe is added to the secondary reaction, to sequester the uncleaved probes to stop their binding to the SRT. (3) Detect the fluorescence signal using a fluorescence plate reader at 485/ 20 nm excitation and 530/25 nm emission filters (Fig. 17.2).

17.2

Protocol

17.2.1 Materials 1. 96-well microplates (MJ Research, Inc.) 2. Chill-out 14 liquid wax (MJ Research, Inc.)

17.2.2 Instruments 1. Thermal cycler 2. CytoFluor 4000 fluorescence plate reader (Applied Biosystems).

17.2.3 Reagents 1. 2. 3. 4. 5. 6. 7. 8. 9. 10.

Lysis buffer: 20 mM Tris–HCl (pH8.5), 0.5% NP-40, and 20 mg/mL tRNA Trizol reagent (Invitrogen) PBS 10 mM MOPS (pH7.5) 100 mM KCl Invader miRNA assay generic reagent kit (#91–287; Third Wave Technologies, Inc.) Yeast tRNA (Sigma) FAM detection oligonucleotide (#91–288; Third Wave Technologies, Inc.) Invader GAPDH mRNA and U6 RNA kits (#94–002; Third Wave Technologies. Inc.) Redmond Red dye (Epoch Biosciences)

17.2 Protocol

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17.2.4 Procedures The protocols described in this section are essentially the same as reported in the study by Allawi et al. (2004) (see Figs. 17.1 and 17.2).

miR-1 5’-uggaauguaaagaaguauggag-3’ 5’-GGCAGCUUUUGCUGCCCTCCATACTTCT-3’ Invader Oligonucleotide

miRNA Capturing

’ -5 AA GC 1 A G R mi be GCG o C Pr CCA Invader Oligo UCCGAGCACCTTACATTTCTTCATACCTCCCGUCGU 5’-UGGAAUGUAAAGUGCGC-3’ UU UU 5’ Flap Cleaving Arrestor Oligo UGGCUCGuggaauguaaagaaguauggagGGCAGCU miR-1 Cleavase Uncleaved miR-1 Probe 5’-AACGAGGCGCACTTTACATTCCACGAGCCUUUUGGCUCG-3’ 5’ Flap Cleaving

5’-AACGAGGCGCACTTTACATTCCACGAGCCUUUUGGCUCG-3’ miR-1 Probe

3’-CGCGUGAAAUGUAAGGU-5’ Arrestor Oligo

Cleavage site UCCGAGCACCTTACATTTCTTCATACCTCCCGUCGU UU UU UGGCUCGuggaauguaaagaaguauggagGGCAGCU miR-1

5’-AACGAGGCGCACT-3’ Released 5’ Flap 5’ Flap Cleaving

Fa

m

Cleavage site -A 5’-AACGAGGCGCACCACQTGCTTCGTGG-3’

5’-CCAGGCAGCAAGTGGTGCGCCTCGTTT-3’ Secondary Reaction Template Fam-AACQTGCTTCGTGG-3’ FRET Oligo

3’-TTGCTCCGCGTGGTGAACGACGGACC-5’ Secondary Reaction Template Fam-A Cleaving

Fam-A (Unquenched dye)

Detect Fluorescence Signal 485/20 nm excitation

530/25 nm emission

Fig. 17.1 Schematic depiction of the Invader miRNA assay, using miR-1 as an example. The Invader miRNA assay includes two consecutive reactions and a final fluorescence measurement. (1) Primary reaction – capturing the target miRNA (including steps & ): Annealing of the invasive and miRNA probe oligonucleotides to a miRNA target forms an overlap-flap structure that is a substrate for the structure-specific 50 nuclease, Cleavase. The noncomplementary 50 flap of the miRNA probe oligonucleotide is the site of cleavage by Cleavase, which releases the 50 flap. The invasive and miRNA probe oligonucleotides are so designed as to include stretches forming a dumbbell-like structure or the stem-loop hairpin regions for better capturing the short size of a miRNA target. (2) Secondary reaction – generating quantifiable signals (including steps to ): A secondary overlap-flap structure is formed by hybridizing both the 50 flap that had been released in the primary reaction and a FRET oligonucleotide to a secondary reaction template (SRT). The FRET oligonucleotide is labeled with a fluorophore and a quencher so that cleavage between them generates a fluorescence signal. An arrestor oligonucleotide complementary to the miRNA probe is added to the secondary reaction, to sequester the uncleaved probes to stop their binding to the SRT. (3) Finally (including step ), detect the fluorescence signal using a fluorescence plate reader at 485/20 nm excitation and 530/25 nm emission filters. Modified from Allawi et al. (2004)

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17 Invader Assay Select miRNAs of your interests

Design miRNA capture probe; Design invader oligo

Isolate RNA from tissues or cells

Hybridization of the oligos with miRNA to caputre target miRNA

Design: Arrestor oligonucleotides

Arrest reaction of uncleaved miRNA probe

5' Flap cleavage on miRNA probe to release 5' Flap

Design: Secondary reaction template; FRET oligonucleotides

Hybridization of FRET oligo & 5' FLAP with secondary reaction template

Fam A cleaving to generate unquenched dye

Detect the fluorescence signal using a fluorescence plate reader

Fig. 17.2 Flowchart of the Invader miRNA detection technology for miRNA expression detection, according to Allawi et al. (2004)

17.2.4.1

Design of miRNA Probe and Invader Oligonucleotide

1. Obtain sequences for the miRNAs of interest from the miRNA registry (microrna.sanger.ac.uk). 2. The miRNA probe and invasive oligonucleotides required for the primary reaction of the invader miRNA assay should be designed to base pair to equal halves of the miRNA of interest (i.e., for a 22-mer miRNA, both the probe and invasive oligonucleotides should form 11 bp duplexes with the miRNA). According to Allawi et al. (2004), for miRNAs with an odd number of nucleotides, the

17.2 Protocol

3. 4.

5.

6.

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miRNA region complementary to the probe should be 1 nt longer than the region complementary to the invasive oligonucleotide. An exception to this rule was applied to miR-135, a 24-nt AU-rich (71% AU) miRNA, a 14-nt probe region was found to be optimal. The invasive oligonucleotide contains a 30 terminal overlapping nucleotide that is not complementary to the miRNA sequence. The miRNA probe should have a 50 flap sequence 50 -AACGAGGCGCAC-30 that, when cleaved from the probe in the primary reaction, is to be used in the secondary reaction (Fig. 17.1). When the 50 flap sequence would be partially complementary to the miRNA sequence, thereby extending the length of the probe–miRNA heteroduplex, an alternative 50 flap sequence (50 -CCGTCGCTG CGT) should be used. For example, miR-1 has the sequence 50 -uggaauguaaagaaguauggag-30 , its probe sequence is 50 -AACGAGGCGCACTTTACATTCC ACGAGCCUUUUGGCUCG-30 , and the invader oligonucleotide is GGCAGCUUUUGCUGCCCTCCATACTTCC. The miRNA probe and invasive oligonucleotides should be modified to include 20 -O-methylated stem-loops at their 50 and 30 ends to stabilize hybrids containing the target miRNA stacked between the two short base-pair duplexes (Fig. 17.1). According to Allawi et al. (2004), the presence of an extra, overlapping nucleotide at each end of the RNA heteroduplex reduced the maximum signal by about 50%. Similarly, a single nucleotide gap at one or both ends of the miRNA reduced the optimal temperature by several degrees and also reduced the signal. The presence or absence of a 50 phosphate on the miRNA was without effect on the performance of the assay. Thus, base stacking between the ends of the miRNA and the invasive and probe oligonucleotides appears to be important, but not essential for detection. Synthesize the antisense oligos using the service provided by IDT (Integrated DNA Technologies, Coralville, IA, USA).

17.2.4.2

Design of Secondary Reaction template, FRET Oligonucleotide and Arrestor Oligonucleotide

The following oligodeoxynucleotides sequences required for the Invader assay are from the study reported by Allawi et al. (2004). 1. SRT Fluorescein (FAM)- specific secondary reaction template 1: FAM-specific secondary reaction template 2: Redmond Red dye (RED-specific secondary reaction template 1:

50 -CCAGGCAGCAAGTGGTGCGCC TCGTTT-30 50 -CCAGGAAGCAAGTGACGCAGCG ACGGT-30 50 -CGCAGTGAGAATGAGGTGATC TCGGCGGT-30

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2. FRET oligonucleotides FRET oligonucleotide is labeled with a fluorophore (F) and a quencher (Q), which form a FRET pair that is separated upon cleavage. Cleavage between them generates a fluorescence signal. Fluorescein (FAM) FRET probe: Redmond Red dye (RED) FRET probe:

Fam-CAC-Q-TGCTTCGTGG Red-CTC-Q-TTTCTCAGTGCG

3. Arrestor oligonucleotides Arrestor oligonucleotides are designed to be complementary to their miRNA probe sequences to sequester the uncleaved probes so that they cannot bind to the SRT. For example, the arrestor oligonucleotide for miR-1 is 50 -UGGAAUGUAAAGUGCGC-30 .

17.2.4.3

RNA preparation

Follow the procedures described in Chap. 2 for Northern Blotting analysis.

17.2.4.4

Invader miRNA Assay

1. Invader reactions are performed in triplicate in 96-well microplates using an invader miRNA assay generic reagent kit. To determine optimal temperature of the primary reaction, perform invader reactions using 250-aM synthetic miRNA in a heated-lid gradient thermal cycler over a temperature range of 40–60 C for 30 min, in 10-mL volumes containing 10 pM of each of the miRNA probes and invasive oligonucleotides. To determine the miRNA level in a sample, use 5 mL aliquots of cell lysates or 50–100 ng of total RNA (in 5 mL) in the 10mL primary reaction. 2. For no-target controls, use 10 ng/mL yeast tRNA to substitute for samples or synthetic miRNA. To prevent evaporation, add 10 mL of clear chill-out 14 liquid wax to each reaction. 3. Upon the completion of the primary reaction, add 5 mL of a secondary reaction mixture including a FAM detection oligonucleotide and 40 pM arrestor oligonucleotide to the primary reaction. 4. Perform the secondary reaction for 15 min at 60 C. 5. Detect the fluorescence signal using a CytoFluor 4000 fluorescence plate reader using 485/20 nm excitation and 530/25 nm emission filters for the FAM dye. 6. Subtract the no-target signal from the miRNA sample signal to determine the net fluorescence signal. The optimal reaction temperature is defined as the temperature at which the highest net signal can be observed. 7. Use 5 mL samples of synthetic miRNA at known concentrations in the range 5 fM–5 pM (25 zM–25 aM) in the invader assay to obtain a dose response curve for miRNA quantitation in total RNA or cell lysate samples.

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8. For biplex invader miRNA assays, invader GAPDH mRNA and U6 RNA kits can be used, and their redmond red dye signals can be detected using 560/20 nm excitation and 620/40 nm emission filters.

17.3

Application and Limitation

The invader miRNA assay offers a bunch of advantages for miRNA exprersison detection. (1) It is quantitative, specific, and sensitive able to detect 1–10 RNA molecules per cell, to discriminate between miRNAs that differ by a single base, distinguishes between miRNAs and their precursors, and to precisely measure 1.2fold changes in RNA expression. (2) It is simple, rapid (requiring only 2–3 h incubation), does not use radioactivity, and is readily performed directly in cell lysates. Further, an isothermal format and the ability to detect two different miRNA molecules with a biplex format make the invader assay suitable for high-throughput screening applications. (3) Two different miRNAs can be assayed simultaneously by performing the invader assay in a biplex format, in which the probe oligonucleotide for each miRNA has a unique 50 flap that can be detected in the secondary reaction (Eis et al. 2001; Lund et al. 2003; Wagner et al. 2003). (4) Costs are reduced by purifying probe oligonucleotides in-house by gel electrophoresis, and by the use of a standard secondary reaction so that the same FRET pair oligonucleotide is used in all reactions. This assay has been successfully used in the analysis of several miRNAs (let-7a, let-7c, miR-1, miR-15, miR-16, miR-125b, and miR-135) using as little as 50– 100 ng of total cellular RNA or as few as 1,000 lysed cells (Allawi et al. 2004; Eis and Garcia-Blanco 2008).

References Allawi HT, Dahlberg JE, Olson S, Lund E, Olson M, Ma WP, Takova T, Neri BP, Lyamichev VI (2004) Quantitation of microRNAs using a modified Invader assay. RNA 10:1153–1161 de Arruda M, Lyamichev VI, Eis PS, Iszczyszyn W, Kwiatkowski RW, Law SM, Olson MC, Rasmussen EB (2002) Invader technology for DNA and RNA analysis: Principles and applications. Expert Rev Mol Diagn 2:487–496 Eis PS, Garcia-Blanco MA (2008) Quantification of microRNAs, splicing isoforms, and homologous mRNAs with the invader assay. Methods Mol Biol 488:279–318 Eis PS, Olson MC, Takova T, Curtis ML, Olson SM, Vener TI, Ip HS, Vedvik KL, Bartholomay CT, Allawi HT, Ma WP, Hall JG, Morin MD, Rushmore TH, Lyamichev VI, Kwiatkowski RW (2001) An invasive cleavage assay for direct quantitation of specific RNAs. Nat Biotechnol 19:673–676 Lund E, Guttinger S, Calado A, Dahlberg JE, Kutay U (2003) Nuclear export of microRNA precursors. Science 303:95–98 Wagner EJ, Curtis ML, Robson ND, Baraniak AP, Eis PS, Garcia-Blanco MA (2003) Quantification of alternatively spliced FGFR2 RNAs using the RNA invasive cleavage assay. RNA 9:1552–1561

Chapter 18

Single Molecule Method

Abstract The single molecule technique for miRNA quantitation is a method utilized to distinguish among different molecules in solution based on their unique spectral properties with a microfluidic, multicolor laser system capable of counting individual molecules as they flow at a high velocity through the system. This assay is homogeneous in nature requiring no target capture, enrichment, or clean-up steps and does not require any reverse transcription or amplification. The technique was developed by Neely et al. in US Genomics (Woburn, Massachusetts, USA) (Nat Methods 3:41–46, 2006). The single molecule method has demonstrated several important advantages for miRNA detection and quantification: fairly sensitive with a low limit of around 250 fM; highly specific with the ability to distinguish single nucleotide differences; simple and yields reproducible data in less than 2 h; and medium throughput with 96-well and 384-well plate compatible enabling for quantifying expression of hundreds of miRNAs per day.

18.1

Introduction

Several advances in single molecule detection (SMD), laser-induced fluorescence (LIF), and fluorescence correlation spectroscopy (FCS) have all provided highly sensitive approaches to study individual macromolecules under physiological conditions (Chan et al. 2004; Chirico et al. 2001; Castro and Williamson 1997; Korn et al. 2003; Hesse et al. 2002; Goodwin et al. 1996; Yanagida et al. 2000;and Schwille et al. 1997). These single fluorophore, single molecule techniques have been routinely employed to quantitatively measure properties of molecules in dynamic systems, such as protein folding, DNA transcription, DNA-binding proteins and molecules in flowing fluid systems (Schwille et al. 1997; Haab and Mathies 1995; Anazawa et al. 2002; Bennink et al. 1999; and Yamasaki et al. 1999). These methods distinguish among different molecules in the solution based on their unique spectral properties (Schwille et al. 1997; Soper et al. 1992).

Z. Wang and B. Yang, MicroRNA Expression Detection Methods, DOI 10.1007/978-3-642-04928-6_18, # Springer-Verlag Berlin Heidelberg 2010

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Typically, individual molecules are tagged with a single fluorophore and analyzed using a confocal microscope with laser illumination and photonburst detection to enhance signal-to-noise (Chirico et al. 2001; Haab and Mathies 1995; and Schwille and Kettling 2001). In dual-color FCS experiments, individual molecules diffuse into an interrogation volume and a time-dependent cross-correlation provides single molecule analysis with sensitivities of 10–100 nM (Schwille et al. 1997). More recently, two-color coincidence fluorescence detection was found capable of ultra-sensitive quantitation (50 fM) when measuring individual synthetic 40 bp DNA molecules in a nonflowing system (Li et al. 2003). On the basis of the above studies, Neely et al. (2006, 2008) developed a single molecule method for detecting and quantifying miRNAs with great sensitivity and specificity. To achieve this aim, they built a microfluidic, multicolor laser system capable of counting individual molecules as they flow at a high velocity through the system. In addition, they developed a rapid solution-based hybridization assay with femtomolar sensitivity for quantitation of miRNAs using fluorescently labeled locked nucleic acid (LNA) DNA chimeric probes. This assay is homogeneous in nature requiring no target capture, enrichment, or clean-up steps and does not require any reverse transcription or amplification. The inclusion of LNA bases within the probe increases the thermal stability of the LNA/RNA hybrids enabling hybridization at higher temperatures (55 C), thereby minimizing nonspecific hybridization. The use of these short probes also maximizes specificity as a single base mismatch can have a profound effect on the stability of the duplex (Christensen et al. 2001). Following hybridization, the temperature is decreased to 40 C and synthetic DNA oligonucleotides end-labeled with quencher molecules are hybridized to the remaining non-hybridized LNA probes. This quenching reaction reduces the background fluorescence thereby increasing the sensitivity of the assay. Quenched reactions are then diluted two- to ten-fold (depending upon LNA probe concentration) and subjected to single molecule interrogation. A dual-color coincident event detection strategy was employed to detect and quantify molecules of interest. Target molecules bearing two fluorescent probes, one labeled with Oyster 556 (green) and the other labeled with Oyster 656 (red), pass through the laser excitation/detection volumes (Red1, Green, and Red2) and emit photons.

18.2

Protocol

18.2.1 Materials 1. 2. 3. 4. 5.

Capillary 20% native PAGE gel (Invitrogen, Carlsbad, CA) Whatman blotting paper (Florham Park, NJ) Microseal lids (MJ Research, Waltham, MA) The ImageQuant program

18.2 Protocol

259

18.2.2 Instruments 1. 2. 3. 4.

TrilogyTM 2020 confocal laser-induced fluorescence detector (US Genomics) Cary Varian UV/vis spectrophotometer (Palo Alto, CA) or equivalents Thermocycler (MJ Research) or equivalents ImageQuant program (GE Healthcare, Piscataway, NJ)

18.2.3 Reagents 1. RNAse Inhibitor (US Biologicals, Swampscott, MA) 2. Fluorescently modified chimeric DNA/LNA probes (Proligo, Boulder, CO or Exiqon, Vedbaek, Denmark) 3. DNA quencher probes containing A-quenchers (Exiqon) 4. Synthetic miRNA templates (Integrated DNA Technologies, Coralville, IA) 5. T4 PNK 6. Human universal RNA (Ambion, Austin, TX) 7. Conditioning solution (US Genomics (USG),Woburn, MA) 8. Running buffer (USG)

18.2.4 Procedures The protocols described in this section are essentially the same as reported in the study by Neely et al. (2006).

18.2.4.1

Preparation of Single Molecule Detection Platform

In the study reported by Neely et al. (2006), a TrilogyTM 2020 confocal laserinduced fluorescence detector was used to perform the single molecule counting experiments. The instrument enables three-color fluorescent detection in a microfluidic flow stream as previously described (Chan et al. 2004), with engineering modifications to automate sample handling and delivery. Neely et al. modified the experimental platform to include an automated 96-well plate compatible sample delivery system. The sample is pulled by vacuum through a 15 cm long customfused silica microfluidic capillary, which they sheath at one end with a needle. The needle is required to pierce the heat seals covering the 96-well plates. The automated delivery system allows the sample injection and capillary wash routine to be pre-programed. To prepare the SMD platform, perform the following: 1. Flush the capillary with conditioning solution for 10 min and then with 1 running buffer for 20 min.

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2. Following this wash step, the samples are queued up for interrogation. Collect data for 36 s for all samples. 3. Upon completion of each sample draw, the tube is automatically flushed for 10 s with 1 running buffer. 4. The instrument contains a rack that can hold eight 200-mL tubes. Load three tubes containing 100 mL of conditioning solution into this rack and rinse the capillary for 10 s with conditioning buffer followed by 10 s of running buffer every third sample. 18.2.4.2

Dual-color Coincident Event Counting

1. To detect and quantify molecules of interest, use a dual-color coincident event detection strategy: target molecules bearing two fluorescent probes, one labeled with Oyster 556 (green) and the other labeled with Oyster 656 (red), passing through the laser excitation/detection volumes (Red1, Green, and Red2) and emit photons. 2. Record the photon bursts emanating from the laser interrogation volume in 1 ms time intervals. 3. Use cross-correlation between the two red channels to measure the flow velocity of the fluorescently labeled molecules using a method previously described by others (Brinkmeier et al. 1999). Focus the 633 nm laser 2 mm upstream of the 532 nm laser. Count the number of time periods during which the signal from each of two fluorescence channels exceeds some fixed value for that channel. The thresholds are used to eliminate the background signal from ambient fluorescence. 4. Coincidence in two (or more) colors is used to mitigate the effect of free probes and the residual background fluorescence. Set a detection threshold, representing the minimum level of photon burst counted as signal from a fluorescently labeled probe, for each color using the average signal intensity plus five times the standard deviation of the signal observed from mock hybridization reactions containing one microgram of total RNA diluted two- to ten-fold in 1 running buffer. 5. Estimate the number of random coincidences based on the raw data. Subtract this estimate from the raw coincidence count to give an estimate of the number of coincidences caused by dual-tagged molecules, and by inference, of the concentration of the analyte. A detailed description of this correction method is presented in D’Antoni et al. (2006). 18.2.4.3

Synthetic Oligonucleotide RNAs, DNAs, and LNAs

1. Design chimeric DNA/LNA probes complementary to publicly available miRNA target sequences using publicly available LNA design tools at www.exiqon.com. 2. Iteratively adjust the number and placement of the locked nucleotides to achieve LNA probes with similar melting temperatures, minimal LNA homodimer and heterodimer formation, and minimal hybridization to background RNAs.

18.2 Protocol

261 Select miRNAs of your interests

Synthesize fluorescently labeled DNA/LNA miRNA capture probe

Isolate RNA from tissues or cells

Hybridization of miRNA and LNA probe at 55oC

Synthesize DNA quencher probes containing A-quenchers

Synthesize miRNA template

Hybridization of DNA quencher probes and non-hybridized LNA probes at 40oC in the presence of miRNA template

Single molecule interrogation: A microfluidic, multi-color laser system capable of counting individual molecules

Fig. 18.1 Flowchart of the single molecule method for miRNA expression detection, according to Neely et al. (2006)

3. Synthesize and purify chimeric DNA/LNA probes with fluorescent modifications, DNA quencher probes containing A-quenchers, and miRNA templates, using the services provided by companies like IDT1 (Integrated DNA Technologies, Coralville, IA, USA). 4. Determine the concentrations and stoichiometry of dye-label to oligo by measuring the oligonucleotide’s absorbance from 650 nm to 230 nm on a Cary Varian UV/vis spectrophotometer. Dye extinction coefficients and correction factors will be provided by Molecular Probes (Eugene, OR) or Denovo (Munster, Germany). 5. Determine the quencher probe labeling stoichiometry by reverse phase high performance liquid chromatography (Fig. 18.1).

18.2.4.4

Validating Hybridization of miRNA Probe Pairs

To independently verify that LNA probes hybridize efficiently to their miRNA targets and to determine the minimum concentration of probe required to drive the

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18 Single Molecule Method

hybridization reactions to completion, conduct an electrophoretic mobility shift assay using 1 pM of a radiolabeled synthetic miRNA target to various concentrations of LNA probes (100 pM to 5 nM). 1. Radiolabel the miRNA targets with 32P using T4 PNK as previously described (Sambrook et al. 1989). 2. Incubate hybridization reactions in 1 USG hybridization buffer (USG) in a thermocycler at 80 C for 5 min followed by a 1 h incubation at 55 C. 3. Add quencher probes to determine if their addition decreased the amount of hybridized product by competing with the miRNA target for binding of the LNA probes. 4. Cool the reactions to 4 C and electrophorese at 100 V through a 20% native PAGE gel for 3 h. 5. Vacuum-dry the gels on Whatman blotting paper and expose to a phosphor screen. The percent hybridized miRNA target is quantified using the ImageQuant program.

18.2.4.5

Absolute miRNA Quantitation

To quantify miRNA gene expression, construct standard curves containing a range of synthetic miRNA concentrations spiked into a complex RNA background. In the study reported by Neely et al. (2006), the authors prepared two to three independent standard curves consisting of 0.25, 0.5, 1, 2, 5, 10, 25, and 50 pM of synthetic miRNA spiked into either human tissue total RNA depleted for small molecular weight RNAs, E. coli total RNA, or human universal RNA. They suggest that when screening for expression in a few tissues, it is best to match the background RNA to the tissue being studied, and recommend including 6–20 no-target controls (probes and RNA background) for each miRNA. 1. Tissue total RNA hybridization reactions consist of 50–100 ng of tissue total RNA, 200–500 pM LNA probes, and 0.5 ml of RNAse Inhibitor in 1 USG hybridization buffer. 2. Seal the 96-well plates with microseal lids and incubate at 80 C for 5 min followed by incubation at 55 C for 1 h. 3. The plate is chilled and DNA quencher probes (2 with regard to LNA probe concentration) are added. The quenching reactions are incubated at 40 C for 30 min. 4. Dilute the samples to a final LNA probe concentration of 100 pM prior to single molecule interrogation. 5. Calculate 95% confidence intervals based on a one-tailed student’s t-test assuming unequal variances for the negative controls and for each data point in the calibration curve. The lower limit of quantitation is defined as the first data point on the calibration curve that lies above the upper confidence limit of the 95% confidence interval for the no-target controls and has a cv (standard deviation/ mean of n 3) of 20% or less. Fit the data with standard linear regression analyses using ordinary least squares estimations.

18.3 Application and Limitation

263

6. If the coincident event numbers counted in the tissue total RNA from three independent experiments have a CV less than 20% and a lower confidence limit of the 95% confidence interval above the upper limit for the 95% confidence interval of the zero target control, the miRNA (or a closely related family member) is considered expressed. Then calculate the concentration of the miRNA within that tissue.

18.3

Application and Limitation

1. The single molecule method has demonstrated several important advantages for miRNA detection. 2. The assay is highly sensitive, being able to detect as little as 100 fM (~1.4 fg or 0.2 amol of miRNA). The lower limit of quantification for the assay is around 250 fM run concentration (corresponding to 500 fM in the hybridization reaction). The assay has also been tested to quantify miRNA expression in 50 and 100 ng of tissue total RNA (Neely et al. 2006). mir-16, mir-9, mir-22, mir-126, mir-143, mir-145, mir-191, and mir-205 were selected for this test to represent a range of expression at low, moderate, and high levels. The authors observed a good agreement between the amount of each target miRNA measured with 50, 100, and 500 ng of total RNA with CVs less than 20% for 12 out of 14 experiments and less than 22% for all experiments. This data suggest that for most miRNAs 50 ng of tissue total RNA is sufficient for accurate and precise quantitation. 3. The assay has a good specificity. These probes are capable of discrimination amongst the four let-7 family members (let-7a, let-7b, let-7c and let-7d). 4. The assay is quantitative. The quantitative nature of the data is unique yielding an accurate and precise amount of miRNA present in tissue total RNA. This amount is highly reproducible for a given miRNA in a particular tissue as we are able precisely and reproducibly to quantify miRNA gene expression over multiple experiments spanning several months. The accuracy of the method is evidenced by a strong correlation between measured miRNA levels and Northern blot data (Neely et al. 2006). The ability to measure the precise amount of a particular miRNA opens up the exciting possibility of detecting subtle changes in miRNA expression levels that may not be observable by other techniques which measure bulk signal, such as Northern blots, microarrays, and RT-PCR. Neely et al. (2008) quantified the expression of miRNAs in cancer tissues. The expression of 11 human miRNAs was quantified in breast, ovary, and prostate normal tissues and adenocarcinomas, cervix normal tissue and squamous cell carcinoma, and skin normal tissue and melanoma. Tissue total RNAs were procured from Ambion Inc. and were isolated from a single tumor with no pooling of normal or tumor samples. All individual standard curves showed a strong correlation between the number of detected coincident events and the miRNA concentration with the coefficients of determination for 10 of the 11

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miRNAs at 0.99 and one at 0.97. The goodness of fit of the linear data was measured not only by calculating the coefficient of determination but also by calculating the confidence intervals for the regression coefficients. Corroborating published results (Michael et al. 2003; Iorio et al. 2005), they observed significant down-regulation (>two-fold) of miR-143 and miR-145 in the cancerous tissues. They also observed a greater than two-fold decrease in miR-126 expression in breast tumor compared to normal breast tissue. miR-16, miR-22, miR-24, miR-100, miR-126, miR-191, and miR-195 were downregulated in cervical tumor versus normal tissue. miR-16, miR-22, miR-24, miR-100, miR126, and miR-195 are down-regulated by greater than three-fold in ovarian tumors versus normal tissue while mir-181a expression remains constant in normal versus tumor tissue. miR-205 expression was decreased from 13 pg/ mg skin total RNA to undetectable amounts in melanoma and miR-181a expression was increased by three-fold in melanoma versus normal skin. 5. The assay is simple and yields reproducible data in less than 2 h. No preparative enrichment, ligation, or target/signal amplification steps are required, and there is no sample clean-up step. This lack of sample manipulation limits the points at which variability could be introduced into the assay thereby hindering the accuracy of quantitation. 6. The assay is 96-well and 384-well plate compatible enabling for quantifying expression of hundreds of miRNAs per day. After all, the technique requires special equipment that is likely unavailable in most of the molecular biology laboratories, which makes the method difficult to be widely applied.

References Anazawa T, Matsunaga H, Young ES (2002) Electrophoretic quantitation of nucleic acids without amplification by single-molecule imaging. Anal Chem 74:5033–5038 Bennink ML, Schaerer OD, Kanaar R, Sakata-Sogawa K (1999) Single-molecule manipulation of double-stranded DNA using optical tweezers: interaction studies of DNA with RecA and YOYO-1. Cytometry 36:200–208 Brinkmeier M, Dorre K, Stephan J, Eigen M (1999) Two beam cross correlation: A method to characterize transport phenomena in micrometer-sized structures. Analytical Chemistry 71:609–616 Castro A, Williamson JGK (1997) Single molecule detection of specific nucleic acid sequences in unamplified genomic DNA. Anal Chem 69:3915–3920 Chan EY, Goncalves NM, Haeusler RA, Hatch AJ, Larson JW, Maletta AM, Yantz GR, Carstea ED, Fuchs M, Wong GG, Gullans SR, Gilmanshin R (2004) DNA mapping using microfluidic stretching and single-molecule detection of fluorescent site-specific tags. Genome Res 6:1137–1146 Chirico G, Cannone F, Beretta S, Baldini G, Diaspro A (2001) Single molecule studies by means of the two-photon fluorescence distribution. Microsc Res Tech 55: 359–364

References

265

Christensen U, Jacobsen N, Rajwanshi VK, Wengel J, Koch T (2001) Stopped-flow kinetics of locked nucleic acid (LNA)-oligonucleotide duplex formation: studies of LNA-DNA and DNA-DNA interactions. Biochem J 354:481–484 D’Antoni CM, Fuchs M, Harris JL, Ko HP, Meyer RE, Nadel ME, Randall JD, Rooke JE, Nalefski EA (2006) Rapid quantitative analysis using a single molecule counting method. Anal Biochem 352:97–109 Goodwin PM, Ambrose WP, Keller RA (1996) Single molecule detection in liquids by laser induced fluorescence. Acc Chem Res 29:603–613 Haab BB, Mathies RA (1995) Single molecule fluorescence burst detection of DNA fragments separated by capillary electrophoresis. Anal Chem 67:3253–3260 Hesse J, Wechselberger C, Sonnleitner M, Schindler H, Schutz GJ (2002) Single molecule reader for proteomics and genomics. J Chromatogr B Analyt Technol Biomed Life Sci 782:127–135 Iorio MV, Ferracin M, Liu CG, Veronese A, Spizzo R, Sabbioni S, Magri E, Pedriali M, Fabbri M, Campiglio M, Me´nard S, Palazzo JP, Rosenberg A, Musiani P, Volinia S, Nenci I, Calin GA, Querzoli P, Negrini M, Croce CM (2005) MicroRNA gene expression deregulation in human breast cancer. Cancer Res 65:7065–7070 Korn K, Gardellin P, Liao B, Amacker M, Bergstro¨m A, Bjo¨rkman H, Camacho A, Do¨rho¨fer S, Do¨rre K, Enstro¨m J, Ericson T, Favez T, Go¨sch M, Honegger A, Jaccoud S, Lapczyna M, Litborn E, Thyberg P, Winter H, Rigler R (2003) Gene expression analysis using single molecule detection. Nucleic Acids Res 31:e89 Li H, Ying L, Green JJ, Balasubramanian S, Klenerman D (2003) Ultrasensitive coincidence fluorescence detection of single DNA molecules. Anal Chem 75:1664–1670 Michael MZ, O’Connor SM, van Holst Pellekaan NG, Young GP, James RJ (2003) Reduced accumulation of specific microRNAs in colorectal neoplasia. Mol Cancer Res 12:882–891 Neely LA, Patel S, Garver J, Gallo M, Hackett M, McLaughlin S, Nadel M, Harris J, Gullans S, Rooke J (2006) A single-molecule method for the quantitation of microRNA gene expression. Nat Methods 3:41–46 Neely LA, Rieger-Christ KM, Neto BS, Eroshkin A, Garver J, Patel S, Phung NA, McLaughlin S, Libertino JA, Whitney D, Summerhayes IC (2008) A microRNA expression ratio defining the invasive phenotype in bladder tumors. Urol Oncol 2008 Sep 15 [Epub ahead of print] Sambrook J, Fritsch E, Maniatis T (1989) Molecular Cloning: A Laboratory Manual. Cold Spring Harbor, NY: Cold Spring Harbor Laboratory Press Schwille P, Bieschke J, Oehlenschlager F (1997) Kinetic investigations by fluorescence correlation spectroscopy: the analytical and diagnostic potential of diffusion studies. Biophys Chem 66:211–228 Schwille P, Kettling U (2001) Analyzing single protein molecules using optical methods. Curr Opin Biotechnol 12:382–386 Soper SA, Davis LM, Shera EB (1992) Similtaneous detection of two colors by two collinear laser beams at different wavelengths. J Opt Soc Am B 9:1761–1769 Yamasaki R, Hoshino M, Wazawa T, Ishii Y, Yanagida T, Kawata Y, Higurashi T, Sakai K, Nagai J, Goto Y (1999) Single molecular observation of the interaction of GroEL with substrate proteins. J Mol Biol 292:965–972 Yanagida T, Kitamura K, Tanaka H, Hikikoshi Iwane A, Esaki S (2000) Single molecule analysis of the actomyosin motor. Curr Opin Cell Biol 12:20–25

Chapter 19

Enzymatic Method

Abstract The enzymatic miRNA detection method is an enzyme-linked immunosorbent assay (ELISA) procedure specifically modified for the detection and quantification of miRNAs. The technology was developed by Mora and Getts [(Biotechniques 41:420–424, 2006); Genisphere, Inc., Hatfield, PA, USA] based on a previous study designed for miRNA profiling with microarray technology reported by Goff et al. (RNA Biol 2:e9–e16, 2005). The technology consists of three parts: miRNA probes, labeled miRNAs, and horseradish peroxidase (HRP)conjugated DNA dendrimers as detection molecules for signal amplification. The method offers several advantages for miRNA analysis over microarrays: (1) it takes shorter experimental time, (2) it is less expensive, and (3) it has high sample throughput for studying the expression of the same miRNA in many different samples. The disadvantages of the method are: (1) a given experiment can only investigate as many sequences as the number of wells analyzed and (2) in most cases, each well requires approximately 3 ng labeled LMW RNA to generate signal/ noise ratio > 3. Moreover, the study of some miRNAs, such as let-7f might require more than 30 ng/well. The technique has been validated with tissue-specific expression of miR-1 in heart tissue, miR-122 in liver, and miR-124a in brain (Biotechniques 41:420–424, 2006).

19.1

Introduction

The enzymatic miRNA detection technique, belonging to the enzyme-linked immunosorbent assay (ELISA), offers a rapid and quantitative method for the detection of miRNAs. It consists of three parts: miRNA probes, labeled miRNAs, and horseradish peroxidase (HRP)-conjugated DNA dendrimers as detection molecules for signal amplification. miRNA probes, the antisense oligodeoxynucleotides to the miRNAs, are spotted onto microtiter plates. The probes will recognize the labeled miRNAs by full complementarity during the hybridization step. Z. Wang and B. Yang, MicroRNA Expression Detection Methods, DOI 10.1007/978-3-642-04928-6_19, # Springer-Verlag Berlin Heidelberg 2010

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Low molecular weight RNA sample and control oligonucleotides are poly(A) tailed and labeled at the 30 -end with a 31-base capturing oligonucleotide complementary to the outer arms of the dendrimer in the presence of bridging oligonucleotide (Fig. 19.1). Labeled miRNAs will be base-pairing with the corresponding miRNA probes in miRNA hybridization buffer to be detected and quantified. DNA dendrimers are three-dimensional (3-D)-branched structures of DNA (Nilsen et al. 1997) with multiple arms (single-stranded DNA) that can be functionalized or labeled with a diverse range of molecules, such as fluorophores, biotins, antibodies, and enzymes. DNA dendrimers are ideally suited for signal amplification and have been commercially adapted for the amplification of fluorescent signal of microarrays (Stears et al. 2000). Two types of dendrimers can be evaluated as detection molecules: (1) biotinylated dendrimers for detection with streptavidin horseradish peroxidase (SA-HRP) and (2) HRP dendrimer conjugates requiring no additional steps except washing before detection.

miR-1 5’-UGGAAUGUAAAGAAGUAUGUAU-3’ 5'-ACCTTACATTTCTTCATACATTGCGAATACT-3’ Cy3 or Cy 5 Capturing Oligonucleotide 3'-TTTTTTTTTTACCTTACAT-5’ Bridging Oligonucleotide

AAAAAAAAAAOH

PO4

Poly(A) Polymerase

Poly(A) Tailing ATP

5’-UGGAAUGUAAAGAAGUAUGUAUAAAAAAAAAA-3’ Bridging miRNA & Capturing Oligo

Ligation Mix

5'-ACCTTACATTTCTTCATACATTGCGAATACT-3’ ||||||||| 3'-TTTTTTTTTTACCTTACAT-5’

T4 DNA Ligase

5’-UGGAAUGUAAAGAAGUAUGUAUAAAAAAAAAAACCTTACATTTCTTCATACATTGCGAATACT-3’ ||||||||||||||||||| Labeled miR-1 3'-TTTTTTTTTTACCTTACAT-5’ 3'-ACCTTACATTTCTTCATACATA-5’ miR-1 Probe Oligonucleotide

3’-XXXXXXXXXXXXXXXXXXXXXX-----DNA Dentrimer Hybridization

5’-UGGAAUGUAAAGAAGUAUGUAUAAAAAAAAAAACCTTACATTTCTTCATACATTGCGAATACT-3’ ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||| 3'-ACCTTACATTTCTTCATACATATTTTTTTTTTACCTTACATXXXXXXXXXXXXXXXXXXXXXX---------5’ ELISA in a Microplate Reader miRNA Detection by OD Value at 450 nm

Fig. 19.1 Schematic diagram illustrating the principle and procedures of the Enzymetic miRNA Detection Technology. The method consists of three main parts: miRNA probes, labeled miRNAs, and horseradish peroxidase (HRP)-conjugated DNA dendrimers as detection molecules for signal amplification. Modified from Mora and Getts (2006)

19.2 Protocol

19.2

269

Protocol

19.2.1 Materials 1. Microtiter plates (CoStar1 EIA plates; Corning Life Sciences, Acton, MA, USA) 2. Microcon1 YM-100 column (Millipore, Billerica, MA, USA) 3. Microcon1 YM-100 microconcentrator-enriched RNA/well (Millipore, Billerica, MA, USA)

19.2.2 Instruments 1. Microplate reader (with a filter covering 450 nm)

19.2.3 Reagents 1. ELISA wash buffer I (WBI; 50 mM Tris–HCl, pH7.4, 0.2% Tween1 20) 2. Blocking buffer [4% bovine serum albumin (BSA) and 10 mM PBS] 3. miRNA hybridization buffer A: [60.0% formamide, 2.0% dextran sulfate, and12.5% sodium dodecyl sulfate (SDS)-based buffer (Genisphere)] 4. miRNA hybridization buffer B: [40% formamide, 0.04% N-lauroyl-sarcosine (Sigma-Aldrich, St. Louis, MO, USA), 1.125 M tetramethylammonium chloride (TMAC; Sigma-Aldrich), 1.5 mM EDTA, pH8.0, and 18.75 mM Tris– HCl, pH8.0] 5. Washing buffer I: [2 SSC/0.2% SDS, 2 SSC, and 0.2 SSC] 6. Washing buffer II: [50 mM Tris–HCl; pH7.4, 0.2% Tween 20, and 50 mM NaCl 7. 50% SDS-based buffer: [2 Denhardt’s, 1 mM EDTA, 0.5% SDS, 0.25 M Na3PO4, 1 SSC] 8. 12.5% SDS-based buffer (Genisphere) 9. Binding buffer I: [BBI; 48% SA-HRP buffer, 48% WBI, 50 mM Tris, and 0.2% Tween 20; pH7.9–8.1] 10. 2% dextran sulfate (Eppendorf, Hamburg, Germany) 11. Streptavidin horseradish peroxidise (SA-HRP; Pierce, Rockford, IL, USA) 12. Tetramethylbenzidine (TMB) solution (Pierce) 13. H2SO4 14. RiboGreen1 RNA Quantitation kit (Molecular Probes, Eugene, OR, USA) 15. 3DNA Array900 miRNA direct kit (Genisphere, Hatfield, PA, USA) 16. MinElute1 PCR purification kit (Qiagen, Valencia, CA, USA)

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19.2.4 Procedures The protocols described in this section are essentially the same as reported in the study by Mora and Getts (2006) (See Figs. 19.1 and 19.2).

19.2.4.1

Design of Various Oligonucleotides

miRNA Probe Oligonucleotide The initial miRNA probe design incorporated several concepts (Goff et al. 2005), including: (1) trimming of miRNA sequences to adjust for an inherently wide variance in melting temperatures, (2) constructing reverse-complement probes to allow direct hybridization to labelled miRNAs, and (3) comparing monomer, dimer, and trimer probe sequences to maximize sensitivity.

Isolate RNA from tissues or cells

Poly(A) tailing

Synthesize Bridging probe with T30

Synthesize Cy3 capture probe

Hybridization of Capture probe and Bridging probe

Bridge adenylated miRNAs with Capture prove Select miRNAs of your interests

Ligation using T4 ligase

Synthesize miRNA-specific probe

Synthesize miRNA-specific probe

Hybridization of the three construct

miRNA detection using a microplate reader

Fig. 19.2 Flowchart of the single molecule method for miRNA expression detection, according to Mora and Getts (2006)

19.2 Protocol

271

1. Obtain sequences for the miRNAs of interest from the miRNA Registry (microrna.sanger.ac.uk). 2. Design the oligodeoxynucleotides exactly antisense to the selected miRNAs for study. Connect two identical antisense units into a single oligomer for each miRNA. Synthesize the antisense oligos using the service provided by IDT1 (Integrated DNA Technologies, Coralville, IA, USA). 3. Synthesize artificial miRNAs (IDT, Inc., Coralville, IA), as positive controls for each of the miRNAs to be detected. 4. Synthesize negative control probes for each miRNA, with C to A or G to C mutations introduced to create mismatches. Generate a 1-nt mismatch, a 2-nt mismatch, a random sequence, a shuffled sequence, and a monomer probe for each selected control spot to serve as control. Shuffled sequences are randomized using the same base composition and need to be tested for a lack of matches in GenBank by BLAST. Capture Oligonucleotide The capture sequence tag is a 31-base oligonucleotide complementary to an oligonucleotide attached to a 3DNA dendrimer labeled with either Cy3 or Cy5. Bridging Oligonucleotide The bridging oligonucleotide (19 nt) consists of 9 nt that are complementary to the capture sequence tag and 10 nt complementary to the added poly A tail (dT10). 19.2.4.2

Spotting/Coating Oligonucleotides

1. Reconstitute the miRNA probe oligonucleotides in water to a final concentration of 1 mg/mL, and then dilute to 1 mg/mL in 1 phosphate-buffered saline (PBS) for plate coating. 2. Incubate each well with 100 mL diluted probe overnight at room temperature (20–25 C). 3. The next day, remove the solution, wash twice with ELISA wash buffer I, and dry on paper towels. 4. Block unbound sites in the wells with 20–30 mL/well blocking buffer for 1–2 h at room temperature. Wash the wells three times with WBI and blot dry on paper towels.

19.2.4.3

Labeling of miRNAs

1. Extract total RNA and enrich low molecular weight RNA. Purify the sample with a Microcon1 YM-100 column prewashed with 50 mL 10 mM Tris buffer

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(pH8.0). Determine the concentration of enriched RNA with RiboGreen1 RNA Quantitation kit according to manufacturer’s instructions. 2. Label the enriched RNA at the 30 -end with an oligonucleotide complementary to some of the outer arms (capture sequences) of the dendrimers to be used for detection, using a 3DNA Array900 miRNA direct kit. Briefly, poly(A) tail enriched miRNAs at the 30 -end. The poly(A) tail facilitates ligation of the capture sequence, by use of a ligation mixture that contains a dT bridging oligonucleotides (Fig. 18.1). 3. Stop all enzymatic reactions with 0.5 M EDTA and purify the tagged miRNAs using MinElute1 PCR purification kit.

19.2.4.4

Detection of miRNAs

Biotinylated Dendrimer and Streptavidin HRP 1. Dilute control oligonucleotides and labelled miRNA samples in miRNA hybridization buffers A and B. 2. Incubate 10 ng or more labeled sample per well at room temperature for 3 h, followed by two consecutive washes with washing buffers I and II at 65 C. 3. Obtain different size DNA dendrimers from Genisphere. For the evaluation of biotinylated conjugates, use any one of the following: (1) two-layer dendrimer with 45 biotins attached to the outer core,or (2) four-layer with 350 biotins, or (3) hyper-four-layer with 960 biotins. 4. Dilute the biotinylated dendrimer conjugate to 1.2 ng/mL in 50% SDS-based buffer and water. 5. Prewarm the 3DNA:labelled miRNA mixture for 10 min at 80 C prior to incubation (30–50 mL/well) at 65 C for 1 h. 6. Remove unbound dendrimer conjugates by SSC washes as described above. 7. Incubate wells with 100 mL/well SA-HRP diluted to 1:500 in SA-HRP dilution buffer for 20 min at 25 C. 8. Wash 4–5 times with wash buffer II. 9. Dry wells on paper towels.

HRP-Labeled 3DNA Hybridization 1. The DNA core of the dendrimer conjugated with 30 HRP molecules (two-layer) has the same size as the dendrimer core of the conjugate with 45 biotins. 2. Dilute the dendrimer conjugate to 0.6 ng/mL in binding buffer I and 2% dextran sulfate. 3. Incubate the wells with 50 mL hybridization mixture (3DNA:labelled miRNA) at room temperature for 1 h.

19.2 Protocol

273

4. Remove unbound 3DNA conjugates by washing 4–5 times with wash buffer II as described above. 5. Blot dry the wells on paper towels. 6. Add 100 mL HRP substrate/well [TMB solution] and let the reaction proceed for 10 min at room temperature in the dark. Color will be developed by this step (Connors et al 1997; Wood et al 1985). 7. Stop the reaction with 100 mL 0.18 M H2SO4, and read the absorbance at 450 nm. 8. Conduct negative control experiments to determine the background contribution of all processing steps and components: no spotted oligonucleotide and no input miRNA sample. The background values ranged from 0.041 to 0.061 and 0.045 to 0.066, respectively, depending on the assay stringency, and are typical for this type of enzymatic assay.

19.2.4.5

Discrimination of Single-Nucleotide Mutations

1. Spot microtiter wells with a miRNA probes designed for the wild-type sequence and with mutations at several bases. 2. Dilute equal amounts of labeled miRNA control oligonucleotide in a hybridization buffer formulated with four different formamide percentages – 40, 50, 60%, and 70% – all at 2% dextran sulfate and 12.5% SDS-based buffer. This formamide titration allows discrimination of single-nucleotide mutations at 60% formamide. At this percentage, the signal from the single-nucleotide mutant should be only 10% the signal from the wild-type sequence, and the signal from the double mutant should be lower than 1%. 3. Use of dextran sulfate as a volume excluder in the miRNA hybridization buffer at a concentration of 2–3% may increase the output signal, presumably by favoring the hybridization of sample to target probe. According to Mora and Getts (2006), the methods could be optimized with three options. (1) The optimal mass of dendrimer/well is 60 ng/well. HRP-labeled dendrimers (30 HRPs/dendrimer) give a signal over noise ratio (S/N) about fivefold greater than biotinylated dendrimers (960 biotins/dendrimer) chased with SAHRP. (2) Optimization of sample hybridization buffer (concentration of formamide, TMAC, dextran sulfate, and surfactants) allows better discrimination of single-base mutations among miRNAs. Salt concentration, dextran sulfate percentage, and overall conjugate concentration in the dendrimer hybridization buffer may all be important parameters in the optimization of UltraAmp reagent (Genisphere, Hatfield, PA, USA) hybridizing to target. The UltraAmp Biotin (960) gives a greater signal amplification than UltraAmp Biotin reagents. And (3) having the HRP directly attached to minimize extra steps and to outperform any biotinylated dendrimer detected with SA-HRP.

274

19.3

19 Enzymatic Method

Application and Limitation

The enzymatic miRNA detection method has several advantages for miRNA analysis over microarrays: (1) it takes shorter experimental time, (2) it is less expensive, and (3) it has high sample throughput for studying the expression of the same miRNA in many different samples. The disadvantages of the method are: (1) a given experiment can only investigate as many sequences as the number of wells analyzed and (2) in most cases, each well requires approximately 3 ng labeled LMW RNA to generate signal/noise ratio > 3. Moreover, the study of some miRNAs, such as let-7f might require more than 30 ng/well. The technique has been validated with tissue-specific expression of miR-1 in heart tissue, miR-122 in liver, and miR-124a in brain [Mora & Getts 2006]. The smallest discernible analytical signal was determined according to the International Union of Pure and Applied Chemistry’s definition (Long and Winefordner 1983) to be 0.048 absorbance units, which corresponds to 4.9 pg oligonucleotides/ well, for the control oligonucleotides under more stringent conditions. The same determination was done using LMW total RNA from rat liver and brain, and the smallest discernible analytical signal corresponds to 2.150 and 0.909 ng, respectively, using the Microcon1 YM-100 microconcentrator-enriched RNA/well (Mora and Getts 2006).

References Connors TD, Burn TC, VanRaay, Germino GG, Klinger KW, Laudes GM (1997) Evaluation of DNA sequencing ambiguities using tetramethylammonium chloride hybridization conditions. BioTechniques 22:1088–1090 Goff LA, Yang M, Bowers J, Getts RC, Padgett RW, Hart RP (2005) Rational probe optimization and enhanced detection strategy for microRNAs using microarrays. RNA Biol 2:e9–e16 Long GL, Winefordner JD (1983) Limit of detection a closer look at the IUPAC definition. Anal Chem 55:712A–724A Mora JR, Getts RC (2006) Enzymatic microRNA detection in microtiter plates with DNA dendrimers. Biotechniques 41:420–424 Nilsen TW, Grayzel J, Prensky W (1997) Dendritic nucleic acid structures. J Theor Biol 187: 273–284 Stears RL, Getts RC, Gullans SR (2000) A novel, sensitive detection system for high-density microarray using dendrimer technology. Physiol Genomics 3:93–99 Wood WI, Gitschier J, Lasky LA, Lawn RM (1985) Base composition-independent hybridization in tetramethylammonium chloride: a method for oligonucleotide screening of highly complex gene libraries. Proc Natl Acad Sci USA 82:1585–1588

Chapter 20

Surface-Enhanced Raman Spectroscopy Method

Abstract Surface-enhanced Raman scattering (SERS) in combination with advanced multivariate methods for pattern recognition has been used for rapid, sensitive, and accurate identification of miRNAs. The strength of the SERS-based sensor is its sensitivity to detect extremely low levels of analyte and specificity to provide the molecular fingerprint of the analyte. The SERS spectra of related and unrelated miRNAs can be detected in near-real time; that detection is sequence dependent, and that SERS spectra can be used to classify miRNA patterns with high accuracy. This technique requires oblique-angle deposition (OAD) fabrication of nanostructured SERS substrate, spotting of RNA or miRNA onto the SERS substrate, and measurement of SERS spectra, followed by classification of the SERS spectra using partial least squares discriminate analysis (PLS–DA). The first application of this technique to miRNA profiling was made recently by Driskell et al. from the Department of Infectious Diseases, Center for Disease Intervention, University of Georgia (AHRC, USA) (Biosens Bioelectron 24:923–928, 2008). This technology is well suited for routine miRNA expression profiling in clinic laboratory. However, the SERS technique requires sophisticated read-out system, complicated analysis skill, and convoluted data interpretation and verification, which requires highly trained professionals to handle.

20.1

Introduction

Raman Spectroscopy is a powerful technique used to investigate the chemical states of the bonds in carbon materials. Surface-enhanced Raman spectroscopy (SERS), which enables the collection of Raman signals from a single molecule, makes the technique more useful for analysis of molecules deposited onto metal surfaces. SERS has become a powerful technique for analyzing biological samples as it can rapidly and nondestructively provide chemical and, in some cases, structural information about molecules in aqueous environments. In the Raman scattering process, both visible and near-infrared (NIR) wavelengths of light can be used to Z. Wang and B. Yang, MicroRNA Expression Detection Methods, DOI 10.1007/978-3-642-04928-6_20, # Springer-Verlag Berlin Heidelberg 2010

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induce polarization of Raman-active molecules, leading to inelastic light scattering that yields specific molecular vibrational information. SERS is a surface-sensitive technique that results in the enhancement of Raman scattering by molecules adsorbed on rough metal surfaces. The enhancement factor can be as much as 1014–1015, which allows the technique to be sensitive enough to detect single molecules. Surface-enhanced Raman scattering allows for the detection of molecules attached to the surface of a single metallic nanoparticle, typically a gold or silver nanoparticle. Existing SERS nanoparticles, also referred to as nanotags, generally include the metallic nanoparticle having a reporter molecule in close ˚ ), which produces a strong Raman signal proximity thereto (typically less than 50 A because of a surface-enhanced effect. Bringing reporter molecules in close proximity to the metal surfaces is typically achieved by adsorption of the Raman-active molecule onto suitably roughened metal nanoparticles, e.g., gold, silver, copper, or other free electron metals. The characteristic signal of the reporter molecule is used to determine the presence and amount of the SERS nanoparticles. Consequently, SERS nanoparticles have utility as spectroscopic and optical tags and are often used in assays. The development of surface enhancement has enabled Raman scattering to be an effective tool for qualitative as well as quantitative measurements with high sensitivity and specificity. Recent advances have led to many novel applications of SERS for biological analyses, resulting in new insights for biochemistry and molecular biology, the detection of biological warfare agents, and medical diagnostics for cancer, diabetes, and other diseases. This chapter highlights many of these recent investigations and provides a brief outlook in order to assess possible future directions of SERS as a bioanalytical tool. SERS in combination with advanced multivariate methods for pattern recognition has been used for rapid, sensitive, and accurate identification of miRNAs (Driskell et al. 2008). This technique requires oblique-angle deposition (OAD) fabrication of nanostructured SERS substrate, spotting of RNA or miRNA onto the SERS substrate, and measurement of SERS spectra, followed by classification of the SERS spectra using partial least squares discriminate analysis (PLS–DA). The applicability of this technique to miRNA expression detection is based upon the following results from the study conducted by Driskell et al. (2008). (1) A miRNA sample can be reproducibly spotted onto a nanostructured SERS substrate and the miRNA binds to the surface in the same orientation or distribution of orientations as to yield reproducible vibrational spectra; (2) OAD is a reliable means of producing SERS substrates. The reproducibility of this fabrication technique suggests that these substrates can be used to confidently identify spectral differences among different miRNAs resulting from sequence-dependent structural differences; (3) The SERS spectra are directly related to the miRNA sequence; SERS detection and differentiation is sensitive to differences in miRNA composition, and this is the basis for miRNA classification using the SERS spectra and PLS–DA analysis. In previous studies, a large number of bands (>20) are typically reported for each base between 200 and 1700/cm, with many of the bands overlapping making interpretation difficult. Nonetheless, two bands have been identified with minimal overlap for interpretation of the miRNA spectra reported here. A ring-breathing

20.2 Protocol

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mode for adenine (A) produces an isolated band at 731 cm 1, and ring-breathing modes of both cytosine (C) and uracil (U) provide a band at 793 cm 1 (Shanmukh et al. 2006). It is worth noting that these two bands are not solely responsible for the PLS–DA classification of the miRNA sequences. The spectra clearly display multiple bands of significant miRNA-dependent variance, and this type of bivariate spectral interpretation is insufficient for fully identifying the sequence. The challenge is that the bands are convoluted contributions from the various bases. Thus, methods of multivariate analysis, such as PLS–DA, are required to fully identify the miRNA sequences.

20.2

Protocol

20.2.1 Materials 1. Glass microscope slides 2. 20 nm film of titanium (Ti)

20.2.2 Instruments 1. 2. 3. 4. 5. 6. 7. 8.

Computer-controlled power supply Electron-beam/sputtering evaporation system Renishaw inVia Raman microscope system Concave rubber band algorithm (OPUS, Bruker Optics, Inc., Billerica, MA) PLS Toolbox v4.0 (Eigen Vector Research Inc., Wenatchee, WA) MATLAB environment (v7.2, The Mathworks Inc., Natick, MA) GRAMS A/I spectral software package (Galactic Industries, Nashua, NH) Nine-point, 2nd-order polynomial Savitzky–Golay algorithm

20.2.3 Reagents 1. 2. 3. 4.

Piranha solution (80% sulfuric acid, 20% hydrogen peroxide) RNase-free Milli-Q water Reagents for total RNA isolation Piranha solution: a 3:1 mixture of concentrated sulfuric acid (H2SO4) with hydrogen peroxide (H2O2)

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20 Surface‐Enhanced Raman Spectroscopy Method

20.2.4 Procedures The protocols described in this section are essentially the same as reported in the study by Driskell et al. (2008) (see Fig. 20.1).

20.2.4.1

Substrate Preparation

OAD fabrication of aligned silver nanorod arrays is used as SERS substrates the SERS method. 1. Prepare colloidal silver by the aqueous reduction of silver nitrate (10–3 M, 200 mL) with trisodium citrate (1%, 4 mL). 2. Cut glass microscope slides into 1 1 cm portions, clean with hot piranha solution to remove organic residues from substrates (the handling of Piranha solutions requires special protection equipment including: a full face shield, heavy duty rubber gloves (regular Nitrile gloves will not provide sufficient protection), as well as an acid apron to wear on top of the lab coat.) (Seu et al. 2007). 3. Rinse the slides with distilled water. 4. Dry the substrates with N2(g) and load into a custom-designed electron-beam/ sputtering evaporation system (Chaney et al. 2005). 5. Deposit a 20-nm film of Ti as an adhesion layer. 6. Evaporate a film of Ag (500 nm) onto the substrate at an angle normal to the ˚ /s. surface at a rate of 3.0 A 7. Rotate the substrates to 86 C with respect to the surface normal. ˚ /s for 100 min. 8. Allow Ag nanorods to grow at an oblique angle at a rate of 3.0 A Each deposition step is automated using a feedback loop integrated QCM to Prepare colloidal silver by aqueous reduction of silver nitrate

Isolate RNA from tissues or cells

Prepare siver nanorod as SERS substrate

Spot RNA sample onto the substrates

Collect and analyze SERS spectra

Fig. 20.1 Flowchart of the surface-enhanced Raman scattering (SERS) procedures for miRNA expression detection. According to Driskell et al. (2008)

20.2 Protocol

279

record the deposition rate and thickness, and a computer-controlled power supply to adjust the e-beam current. As reported elsewhere (Shanmukh et al. 2006), these deposition conditions result in optimal SERS substrates with overall nanorod lengths of ~900 nm, diameters of ~100 nm, and densities of ~13 nanorods/mm2 (Shanmukh et al. 2006).

20.2.4.2

Total RNA Extraction

See Section II for detailed protocols.

20.2.4.3

SERS Measurements

1. Acquire SERS spectra using a Renishaw inVia Raman microscope system. Use a 785-nm near-IR diode laser as the excitation source. Focus the laser into ~115 11 mm spot using a 5 objective. Set the laser power to 10% (the power at the sample surface is ~15 mW). Extend the scan spectra with a spectral range of 400–1,800/cm using a 10-s exposure. 2. Spot RNA sample onto the substrates, 1 mL/miRNA and allow to dry. Each RNA sample should be spotted in duplicate on a single substrate with 3 spectra recorded for each spot for a total of 6 spectra per miRNA/substrate. To ensure substrate-to-substrate reproducibility, each miRNA should be applied to three different substrates. 3. Use DEPC-treated water as a control.

20.2.4.4

Data Analysis

1. In order to evaluate the reproducibility of the method, SERS spectra should be collected from different locations on a single substrate, from different substrates, and for different miRNA sequences. 2. The spectra need to be baseline corrected using a concave rubber band algorithm with 10 iterations and 64 points, and then vector normalized. These steps allow for comparison of Raman band locations and relative peak intensities in the figures. 3. Perform classification and identification of different miRNAs using PLS Toolbox v4.0, operating in the MATLAB environment. Process SERS spectra for statistical analysis by taking the first derivative of each spectrum using a ninepoint, 2nd-order polynomial Savitzky–Golay algorithm followed by normalization to unit vector length. Then mean-center the normalized first derivative spectra and analyze with partial least squares discriminate analysis (PLS–DA). PLS–DA is an established and statistically robust method for objective and blind data classification. Training spectra of known origin should be used to build a

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20 Surface‐Enhanced Raman Spectroscopy Method

PLS–DA model with a class reserved for each miRNA sequence. The spectra also need to be analyzed manually to strengthen the PLS–DA interpretation of the SERS results, and to independently validate the PLS–DA methodology. 4. Individually analyze raw SERS spectra using the GRAMS A/I spectral software package by measuring the peak heights for the bands located at 731 and 793 cm 1 and comparing the relative intensities for each sample.

20.3

Application and Limitation

The work presented by Driskell et al. demonstrates the utility of SERS for sensitive and rapid (10 s) detection of miRNA members and family members. The SERS platform based on OAD-fabricated silver nanorod arrays can be used for the detection and classification of miRNAs. The SERS method uses limited volumes of the specimen for miRNA analysis without RNA labeling and/or amplification steps, which are dependent on the intrinsic stability and specificity of the reagents. The SERS spectra of related and unrelated miRNAs can be detected in near-real time, that detection is sequence dependent, and that SERS spectra can be used to classify miRNA patterns with high accuracy. This technology is well suited for routine miRNA expression profiling in clinic laboratory. The SERS technique requires sophisticated read-out system, complicated analysis skill, and convoluted data interpretation and verification, rendering it less practical in most of molecular biology laboratories.

References Chaney SB, Shanmukh S, Zhao Y-P, Dluhy RA (2005) Aligned silver nanorod arrays produce high sensitivity surface-enhanced Raman spectroscopy substrates. Appl Phys Lett 87:31908–31910 Driskell JD, Seto AG, Jones LP, Jokela S, Dluhy RA, Zhao YP, Tripp RA (2008) Rapid microRNA (miRNA) detection and classification via surface-enhanced Raman spectroscopy (SERS). Biosens Bioelectron 24:923–928 Seu KJ, Pandey AP, Haque F, Proctor EA, Ribbe AE, Hovis JS (2007) Effect of surface treatment on diffusion and domain formation in supported lipid bilayers. Biophys J 92:2445–2450 Shanmukh S, Jones L, Driskell J, Zhao Y, Dluhy R, Tripp RA (2006) Rapid and sensitive detection of respiratory virus molecular signatures using a silver nanorod array SERS substrate. Nano Lett 6:2630–2636

Chapter 21

RAKE Assay

Abstract The RNA-primed, array-based Klenow enzyme (RAKE) assay is a new method for high-throughput miRNA detection. It involves the on-slide application of the Klenow fragment of DNA polymerase I to extend unmodified miRNAs hybridized to immobilized DNA probes. RAKE offers unique advantages for specificity over Northern blots or other microarray-based expression profiling platforms. An oligo with a 5’ spacer is covalently linked onto a glass platform. The spacer sequence is followed by a miRNA antisense capture probe with three thymidine residues in between. RNA samples are hybridized to this array. miRNAs in the sample would bind to their specific probe and form a double stranded structure. The addition of exonuclease I will only degrade unbound single stranded oligos. The miRNA that have latched onto its probe will act as a primer. Subsequent PCR will result in the addition of biotin-conjugated dATPs onto the spacer template, which emits an augmented signal without PCR amplification of the original RNA sample. The technique was invented by Mourelatos and colleagues from the Department of Pathology and Laboratory Medicine, School of Medicine, University of Pennsylvania (Philadelphia, Pennsylvania, USA) (Nelson et al. 2006). The RAKE technique was initially applied to studying human cell lines and brain tumors, proving that RAKE assay is a sensitive and specific method for miRNA detection and an ideal approach for rapid expression profiling of all known miRNAs. Later, the same group used RAKE to profile miRNAs from normal human adult and fetal brains and from reactive astrocytosis and oligodendroglial tumors (RNA 12:187–191, 2006).

21.1

Introduction

The RNA-primed, array-based Klenow enzyme (RAKE) assay is a new method for high-throughput miRNA detection. It involves on-slide application of the Klenow fragment of DNA polymerase I to extend unmodified miRNAs hybridized to immobilized DNA probes. RAKE offers unique advantages for specificity over Z. Wang and B. Yang, MicroRNA Expression Detection Methods, DOI 10.1007/978-3-642-04928-6_21, # Springer-Verlag Berlin Heidelberg 2010

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Northern blots or other microarray-based expression profiling platforms. An oligo with a 5’ spacer is covalently linked onto a glass platform. The spacer sequence is followed by a miRNA antisense capture probe with three thymidine residues in between. RNA samples are hybridized to this array. miRNAs in the sample would bind to their specific probe and form a double stranded structure. The addition of exonuclease I will only degrade unbound single stranded oligos. The miRNA that have latched onto its probe will act as a primer. Subsequent PCR will result in the addition of biotin-conjugated dATPs onto the spacer template, which emits an augmented signal without PCR amplification of the original RNA sample (Nelson et al. 2004). RAKE is a new tool with high sensitivity and specificity for miRNA profiling. A major advantage of RAKE is that there is no sample RNA manipulation. Certain possible biases that may be introduced during enzymatic labeling, or during cDNA generation or amplification of the sample RNA before hybridization to the glass microarray, are thus avoided. RAKE allows for rapid and simultaneous detection of all known miRNAs from the same sample. Another advantage of RAKE is the ability to completely automate all steps from sample hybridization to detection. This is achieved by using existing technologies and equipment used for traditional mRNA microarrays, and allows for highly consistent performance. Northern blotting is considered the standard method for miRNA validation and quantification (Ambros et al. 2003); it offers both “quantitative” and “qualitative” information and, unlike a microarray experiment, confirms the length of the hybridized transcripts. In contrast to RAKE, however, Northern blotting is laborious and hence less well suited for high-throughput expression profiling. The RAKE assay also gives unique qualitative data, because the 3’ end of the miRNA ‘primer’ should hybridize specifically to the oligonucleotide ‘template’. For this reason, RAKE can discriminate the exact 3’ end of miRNAs. This is significant because of the many mature miRNAs for which paralogs differ at the 3’ end. These miRNAs, derived from different genes, would be predicted to cross-react adversely in northern blots (and in standard microarray methods using labeled target pools) but not usually on RAKE assay. According to Nelson et al. (2004), the RAKE assay is devised to exploit the known ability of the Klenow enzyme fragment to act as a DNA polymerase using an RNA primer on a DNA oligonucleotide template (Huang and Szostak 1996; Huang and Alsaidi 2003). Earlier studies have demonstrated on-slide enzymatic reactions and primer extension (Nikiforov et al. 1994; Head et al. 1997). However, direct detection of RNA hybridization (using RNA-primed DNA polymerase) has not been reported on a microarray, nor has the special properties of the Klenow enzyme been used in microarray studies. It is also necessary to use exonuclease I, a 3’-5’, single-stranded DNA–specific exonuclease that is highly processive (Brody et al. 1986). It is important to note that the activities of both Klenow enzyme and exonuclease I are independent of the sequence of their substrates (Brody et al. 1986). Systematic bias is therefore not introduced. RNA ligases, in contrast, are prone to bias because the enzyme kinetics change with substrate sequence (Ohtsuka et al. 1997; Romaniuk et al. 1982), producing an inaccurate representation of the miRNAs present in a target pool labeled by RNA ligase methods. The results

21.2 Protocol

283

from the study by Nelson et al. (2004) demonstrate sensitivity to the level of 10 pg of target miRNA, which is comparable to that of the Northern blots (Lim et al. 2003).

21.2

Protocol

21.2.1 Materials 1. 2. 3. 4. 5.

Cell culture materials RNA isolation materials 20% urea-PAGE gel 384-well plates (Qiagen) CodeLink slides (Amersham)

21.2.2 Instruments 1. 2. 3. 4. 5.

Storm 860 Phosphorimager (Molecular Dynamics) or an equivalent GeneMachines OmniGrid 100 robot or an equivalent Genepix 4000B laser scanner (Axon) or an equivalent Genepix Pro5.0 software package (Axon) or an equivalent Excel and Genespring 6.2 or equivalents

21.2.3

Reagents

1. MirVana kit designed to isolate low molecular weight RNA (Ambion) 2. Trizol LS reagent (Ambion) 3. 150 mM sodium phosphate buffer (pH8.5; 200 U/ml print buffer) with 0.0005% Sarkosyl 4. 2 and 5 SSC 5. 10% formamide 6. Hybridization buffer 7. Exonuclease I (NEB; fresh buffer at pH7.5; 4 U/mL) 8. 0.05% SDS 9. Exo(–) Klenow (Promega; 0.15 U/ml) 10. DNA polymerase buffer (Promega) 11. biotin-7-dATP (Invitrogen; 4 mM) 12. Streptavidin-conjugated Alexa-fluor-547 (Molecular probes; 15 ng/ml)

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21.2.4 Procedures The procedures described herein are based on the study reported by Nelson et al. (2004) (See Figs. 21.1 and 21.2).

21.2.4.1

RNA Isolation and Northern Blots

1. Isolate total RNA using MirVana kit designed to isolate low molecular weight RNA. 2. If Northern blot analysis needs to be performed to verify the RAKE data, then the total RNA can be isolated from the cells or tissues using the Trizol LS Triple Thymidines Spacer

Anti-miR-1

5’-GTCGTGACTGGGAATAGCCTGTTTATACATACTTCTTTACATTCCA-3’

3’

3’ TTT TTT

5’

3’

3’

TTT

TTT

TTT Hybridization

TTT

TTT

Glass microarray slide Immobilization

TTT

+

TTT

miR-1

TTT

TTT TTT

Oligo probes

3’

3’

3’

3’

miRNA Capture Oligodeoxynucleotide Probe

EndoI

Endonuclease I

Laser scanning Fluorophore-conjugated streptavidin

Klenow dsDNA/RNA Klenow

XXX

Biotin Conjugation

XXX

PolyI PolyI

TTT

TTT

B

Fluorophore Conjugation

AAA

B

TTT

FS

AAA

B

XXX

Biotin-dATP

FS

AAA

Image Analysis

ssDNA Degradation

Fig. 21.1 Schematic diagram of RNA-primed, array-based Klenow enzyme (RAKE) assay. The sample probe at the top of the figure illustrates the generic structure of the DNA oligonucleotides used on the microarray. The nucleotides at the 5’ half comprise a spacer, which is constant for all the probes, followed by three thymidine nucleotides. The variable portion of each probe is at the 3’ end, which is the antisense sequence of various miRNAs. An RNA sample, containing miRNAs, is hybridized to a glass microarray on which all the DNA probes have been spotted. Next, the slide is washed and exonuclease I is applied to degrade unhybridized (single-stranded) DNA probes. The slide is washed again and the Klenow fragment of DNA polymerase I is applied to catalyze the addition of biotin-conjugated dATPs using the miRNA as a primer and the spotted probe as template. Finally, the biotins are labeled with streptavidin-conjugated fluorophore to specifically highlight spots (here, spot ‘X’) where miRNA hybridization occurred. Modified from Nelson et al. (2004)

21.2 Protocol

285 Select miRNAs of your interests

Synthesize miRNA capture probes

Isolate RNA from tissues or cells

Immobilization of probes onto a glasss microarray slide

Hybridization of miRNAs and probes to form miRNA/DNA duplex on the microarray slide

Digest non-hybridized single-stranded probes

Klenow-meidated biotin conjugation

Fluorophore conjugation

Laser scanning

Fig. 21.2 Flowchart of the RAKE assay for miRNA expression detection. According to Nelson et al. (2004)

reagent. Run RNA on 20% urea-PAGE gels, blot, and probe using 5’-end radiolabeled probes against the target miRNAs, as described in detail in Sect. 3. Expose blots on phosphorimager screens overnight and scan signals and quantitate bands using a Storm 860 Phosphorimager.

21.2.4.2

Design of Oligos for Microarray

1. An oligo contains three parts: a 5’ spacer GTCGTGACTGGGAATAGCCTG, three thymidine residues, and a miRNA antisense capture probe. Taking miR-1 as an example, the full oligo sequence then should be 5’-GTCGTGACTGGGAATAGCCTG-TTT-ATACATACTTCTTTACATTCCA-3’ (Fig. 24.1).

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2. Synthesize the oligo in the form of DNA, at 600 pmol on 384-well plates, each containing a 5’-C6-amino modified linker, using the services provided by qualified companies.

21.2.4.3

Microarray Platform

1. Suspend the probes at 40 mM in 150 mM sodium phosphate buffer (pH8.5; 200 U/ml print buffer) with 0.0005% Sarkosyl. 2. Use a GeneMachines OmniGrid 100 robot to print the probes onto CodeLink slides at 30–35% humidity at 24–27 C, so that the oligo is covalently linked onto a glass platform. 3. Each spot element is B120 mm in diameter and the center-to-center spacing is 400 mm. Each glass slide contain six spots (three spatially separated pairs) corresponding to each probe, for a total 1,422 spots including controls.

21.2.4.4 1. 2. 3. 4.

5. 6. 7. 8. 9. 10. 11. 12. 13.

RAKE Protocol

1 min wash in 2 SSC at 25 C 5 min rinse in 5 SSC with 10% formamide at 25 C 3 30 s rinse in 2 SSC at 25 C 18 h target/probe hybridization (35 mL concentrated hybridization buffer, 65 mL small RNA preparation containing 4 mg low molecular weight RNA, and 10 mL plant DNA spike-in solution, which are together heated to 75 C and allowed to cool at room temperature prior to hybridization) at 25 C 3 1 min rinse in 2 SSC at 37 C 3 h incubation with Exonuclease I (4 U/mL) at 27 C 3 1 min rinse in 2 SSC at 27 C 10 min rinse in 2 SSC with 0.05% SDS at 27 C 4 1 min rinse in 2 SSC at 37 C 60 min incubation with Exo(–) Klenow (0.15 U/ml) in 1 DNA polymerase buffer (Promega) with biotin-7-dATP (4 mM) at 27 C 2 1 min rinse in 2 SSC at 25 C 30 min incubation with streptavidin-conjugated Alexa-fluor-547 (15 ng/ml) at 25 C 3 1 min rinse in 2 SSC at 25 C

21.2.4.5

Validation Steps

1. Generate a concentration curve using a synthetic target RNA oligonucleotide (e.g., miR-1) in the background of a complex RNA mixture (e.g., low-molecularweight RNA isolated from HeLa cells, a cell line that does not contain miR-1).

References

21.2.4.6

287

Image Analysis and Data Processing

1. Scan slides using a Genepix 4000B laser scanner (Axon) at a constant power level and sensitivity (550 PMT) using a single color channel (532-nm wavelength). 2. Eliminate nonhybridizing and artifact-associated spots by both visual- and software-guided flags. 3. Measure image intensities as a function of the median of foreground minus background. Normalize negative values to zero. 4. Analyze images using the Genepix Pro5.0 software package (Axon). Use Excel and Genespring 6.2 for further data analysis.

21.3

Application and Limitation

The RAKE assay is a sensitive, specific technique for assessing DNA and RNA targets, offering unique advantages for specificity over Northern blots or other microarray-based expression profiling platforms, and may have broader applications than for miRNA detection (Nelson et al. 2004). The RAKE assay allows for sensitive, specific and high-throughput miRNA expression profiling. The sensitivity of RAKE is very similar to the sensitivity of the technique reported by Miska et al. (2004). However, in contrast to other microarray techniques, RAKE does not involve the generation of a cDNA library or amplification of the RNA sample and avoids sample RNA manipulation prior to hybridization. RAKE should be superior at discriminating paralogous miRNAs that differ at their 3’ ends, because these other techniques rely solely on hybridization to detect and discriminate between miRNA paralogs (Miska et al. 2004; Liu et al. 2004).

References Ambros V, Bartel B, Bartel DP, Burge CB, Carrington JC, Chen X, Dreyfuss G, Eddy SR, Griffiths-Jones S, Marshall M, Matzke M, Ruvkun G, Tuschl T (2003) A uniform system for microRNA annotation. RNA 9:277–279 Brody RS, Doherty KG, Zimmerman PD (1986) Processivity and kinetics of the reaction of exonuclease I from E. coli with polydeoxyribonucleotides. J Biol Chem 261:7136–7143 Head SR, Rogers YH, Parikh K, Lan G, Anderson S, Goelet P, Boyce-Jacino MT (1997) Nested genetic bit analysis (N-GBA) formutation detection in the p53 tumor suppressor gene. Nucleic Acids Res 25:5065–5071 Huang Z, Alsaidi M (2003) Selective labeling and detection of specific mRNA in a total-RNA sample. Anal Biochem 322:269–274 Huang Z, Szostak JW (1996) A simple method for 3’-labeling of RNA. Nucleic Acids Res 24:4360–4361 Lim LP, Lau NC, Weinstein EG, Abdelhakim A, Yekta S, Rhoades MW, Burge CB, Bartel DP (2003) The microRNAs of Caenorhabditis elegans. Genes Dev 17:991–1008

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Liu CG, Calin GA, Meloon B, Gamliel N, Sevignani C, Ferracin M, Dumitru CD, Shimizu M, Zupo S, Dono M, Alder H, Bullrich F, Negrini M, Croce CM (2004) An oligonucleotide microchip for genome-wide microRNA profiling in human and mouse tissues. Proc Natl Acad Sci USA 101:9740–9744 Miska EA, Alvarez-Saavedra E, Townsend M, Yoshii A, Sestan N, Rakic P, Constantine-Paton M, Horvitz HR (2004) Microarray analysis of microRNA expression in the developing mammalian brain. Genome Biol 5:R68 Nelson PT, Baldwin DA, Kloosterman WP, Kauppinen S, Plasterk RH, Mourelatos Z (2006) RAKE and LNA-ISH reveal microRNA expression and localization in archival human brain. RNA 12:187–191 Nelson PT, Baldwin DA, Scearce LM, Oberholtzer JC, Tobias JW, Mourelatos Z (2004) Microarray-based, high-throughput gene expression profiling of microRNAs. Nat Methods 1:106–107 Nikiforov TT, Rendle RB, Goelet P, Rogers YH, Kotewicz ML, Anderson S, Trainor GL, Knapp MR (1994) Genetic Bit Analysis: a solid phase method for typing single nucleotide polymorphisms. Nucleic Acids Res 22:4167–4175 Ohtsuka E, Nishikawa S, Fukumoto R, Tanaka S, Markham AF (1997) Joining of synthetic ribotrinucleotides with defined sequences catalyzed by T4 RNA ligase. Eur J Biochem 81:285–291 Romaniuk E, McLaughlin LW, Neilson T, Romaniuk PJ (1982) The effect of acceptor oligoribonucleotide sequence on the T4 RNA ligase reaction. Eur J Biochem 125:639–643

Chapter 22

Bead-Based Flow Cytometric miRNA Expression Profiling

Abstract The bead-based flow cytometric miRNA expression profiling method involves coupling of miRNA capture probes to carboxylated 5-micron polystyrene beads, RT-PCR amplification of miRNAs, hybridization of miRNAs to the capture beads, staining with streptavidin-phycoerythrin, and and final read out using a flow cytometer to measure bead color (denoting miRNA identity) and phycoerythrin intensity (denoting miRNA abundance). The technique was invented by Lu et al in Broad Institute of MIT and Harvard (Cambridge, Massachusetts, USA) (Lu et al. 2005). The bead-based method has been applied to conduct a systematic expression analysis of 217 mammalian miRNAs from 334 samples, including multiple human cancers. The data proved it is able to distinguish tumors of different developmental origin, to distinguish tumors from normal tissues, and to distinguish tumors of different stages. The technique is feasible, and has the attractive properties of improved accuracy, high speed and low cost. Moreover, it is an efficient detection platform for high throughput miRNA profiling in a quantitative manner. The beadbased miRNA detection method is also easy to implement in a routine clinical setting.

22.1

Introduction

The bead-based flow cytometric miRNA expression profiling method takes the advantages of bead-anchored hybridization. The procedures involve several straightforward steps: (1) Coupling of oligonucleotide-capture probes complementary to miRNAs of interest to carboxylated 5-micron polystyrene beads impregnated with variable mixtures of two fluorescent dyes (that can yield up to 100 colors), each representing a single miRNA; (2) Adaptor ligations using both the 5’-phosphate and the 3’-hydroxyl groups of miRNAs, (3) RT-PCR amplification of miRNAs using a common biotinylated primer; (4) Hybridization to the capture beads; and (5) Staining with streptavidin-phycoerythrin; and (6) Analyze the beads

Z. Wang and B. Yang, MicroRNA Expression Detection Methods, DOI 10.1007/978-3-642-04928-6_22, # Springer-Verlag Berlin Heidelberg 2010

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22 Bead-Based Flow Cytometric miRNA Expression Profiling

using a flow cytometer capable of measuring bead color (denoting miRNA identity) and phycoerythrin intensity (denoting miRNA abundance). Bead-based hybridization is superior to glass microarray hybridization in which it is more closely approximate hybridization in solution. In addition, the bead method has a linear relationship for detection over a hundred-fold range of expression. The method meets the requirements of both high throughput and quantitativeness.

22.2

Protocol

22.2.1 Materials 1. 2. 3. 4. 5.

MAP beads (Luminex Corporation) 96-well plates All materials for cell culture All materials for RNA exaction All materials for RT-PCR

22.2.2 Instruments 1. Thermocycler 2. Flow cytometer, Luminex 100IS machine

22.2.3 Reagents 1. TE buffer (10 mM Tris–HCl pH8.0, 1 mM EDTA) 2. T4 RNA ligase (Amersham Biosciences) 3. 1.5 TMAC (4.5 M tetramethylammonium chloride, 0.15% sarkosyl, 75 mM Tris–HCl pH 8.0, 6 mM EDTA) 4. Streptavidin-phycoerythrin (Molecular Probes) 5. All reagents for cell culture 6. All reagents for RNA exaction 7. All reagents for RT-PCR

22.2.4 Procedures The procedures described herein are based on the study reported by Lu et al. (2007) (See Fig. 22.1).

22.2 Protocol

291 Isolate RNA from tissues or cells

Synthesize Adaptor -specific primer

Synthesize Adaptor probe

Ligation of adaptor probe to 3' and 5' ends of RNAs using T4 ligase

Revser transcription

Select miRNAs of your interests

PCR

Synthesize miRNA capture probes

Precipitate PCR products

Conjugate the capturee probes to carboxylated xMAP beads in 96-well plates

Hybridization

Measure median fluorescence intensity

Computational analysis

Fig. 22.1 Flowchart of the bead-based flow cytometric miRNA expression profiling method for miRNA expression detection. According to Lu et al. (2007)

22.2.4.1

miRNA Labeling

1. Prepare total RNA samples from tissues or cells from control and diseased subjects. 2. Use two synthetic pre-labeling-control RNA oligonucleotides (5’-pCAGUCAGUCAGUCAGUCAGUCAG-3’, and 5’-pGACCUCCAUGUAAACGUACAA-3’) to control for target preparation efficiency. 3. Spike them each at 3 fmoles per mg total RNA. Small RNAs (18–26 nucleotides) are recovered from 1–10 mg total RNA through denaturing polyacrylamide gel purification. 4. Adaptor-ligate small RNAs sequentially on the 3’-end and 5’-end using T4 RNA ligase. 5. After reverse-transcription using adaptor-specific primers, PCR-amplify the products (95 C for 40 s; 50 C for 30 s; 72 C for 30 s; 18 cycles for 10 mg starting

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22 Bead-Based Flow Cytometric miRNA Expression Profiling

total RNA) using a 3’-primer 5’-TACTGGAATTCGCGGTTA-3’ and 5’ primer 5’-biotin-CAACGGAATTCCTCACTAAA-3’ (IDT). 6. Precipitate the PCR products and dissolve in 66 mL TE buffer containing two biotinylated post-labeling-control oligonucleotides (100 fmoles of FVR506, 25 fmoles PTG20210).

22.2.4.2

Design of miRNA Capture Probes

1. Obtain sequences for the miRNAs of interest from the miRNA Registry (microrna. sanger.ac.uk). 2. Design the miRNA capture oligonucleotides probes exactly antisense to the selected miRNAs for study with a 6-carbon linker at 5’end. Synthesize the antisense oligos using the service provided by IDT (Integrated DNA Technologies, Coralville, IA, USA).

22.2.4.3

Bead-Based Detection

1. Conjugate the capture probes to carboxylated xMAP beads in 96-well plates, following the manufacturer’s protocol. For each probe set, mix 3 mL of every probe–bead conjugate into 1 mL of 1.5 TMAC. 2. Hybridize the samples in a 96-well plate, with two mock PCR samples (using water as template) in each plate as a background control. Hybridization should be carried out overnight at 50 C with 33 mL of the bead mixture and 15 mL of labeled material. 3. Spin down beads, resuspend in 1 TMAC containing 10 mg/mL streptavidinphycoerythrin and incubate at 50 C for 10 min before data acquisition. 4. Use Luminex 100IS machine to measure median fluorescence intensity values.

22.2.4.4

Computational Analyses

1. To eliminate bead-specific background, subtract the average readings of that particular bead in the two-embedded mock-PCR samples in each plate. Scale profiling data according to the post-labeling-controls and then the pre-labelingcontrols, in order to normalize readings from different probe/bead sets for the same sample, and to normalize for the labeling efficiency, respectively. Scaling should be done in two steps: (1) well-to-well scaling – scale the reading from each well such that the total of the two post-labeling controls, in that well, become the median value based on a pilot study and (2) sample scaling – scale the normalized readings such that the total of the six pre-labeling controls in each sample reach the median value based on a pilot study. 2. Set a threshold of data at 32 and log2-transformed.

22.3 Application and Limitation

293

3. Before clustering, filter data to eliminate genes with expression lower than 7.25 (on a log2 scale) in all samples. A miRNA is regarded as “not expressed” or “not detectible”, if in none of the samples, that particular miRNA has an expression value above a minimal cutoff. This cutoff value is determined based on noise analyses of target preparation and bead detection. Any feature that is not expressed under this criterion will be filtered out before clustering. 4. Perform hierarchical clustering with average linkage and Pearson correlation. 5. Next, center all features and normalize to a mean of 0 and a standard deviation of 1. Perform k-nearest-neighbor classification of normal versus disease samples with k ¼ 3 in the selected feature space using euclidean distance measure. 6. Quality control should be performed as part of the preprocessing by requiring that the reading from each control probe exceeds some minimal probe-specific threshold. These thresholds are determined by identifying a natural lower cutoff, i.e., a dip, in the distribution of each control probe. The cutoff values should be chosen based on a set of samples in a pilot study. The lower post-control should be greater than 500 and the higher post-control must exceed 2,450. The lower and higher pre-controls should exceed 1,400 and 2,000 respectively (after wellto-well scaling).

22.3

Application and Limitation

Utilizing the bead-based flow cytometric miRNA expression profiling method, Lu et al. (2007) carried out a systematic expression analysis of 217 mammalian miRNAs from 334 samples, including multiple human cancers. They observed a general down regulation of miRNAs in tumors compared with normal tissues and were able to successfully classify poorly differentiated tumors using miRNA expression profiles. In particular, the data from the bead-based miRNA profiling is able to distinguish tumors of different developmental origin, to distinguish tumors from normal tissues, and to distinguish tumors of different stages. By comparison, messenger RNA (mRNA) profiles were found highly inaccurate when applied to the same samples. These findings highlight the potential of miRNA profiling as a better diagnostic tool for cancer. Their observation that miRNA expression seems globally higher in normal tissues compared with tumors led us to the hypothesis that global miRNA expression reflects the state of cellular differentiation. These experiments support the hypothesis that global changes in miRNA expression are associated with differentiation, the abrogation of which is a hallmark of all human cancers. These findings are also consistent with the recent observation that mouse embryonic stem cells lacking Dicer, an enzyme required for miRNA maturation, fail to differentiate normally. miRNAs can function to prevent cell division and drive terminal differentiation. An implication of this hypothesis is that down regulation of some miRNAs might play a causal role in the generation or maintenance of tumors.

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22 Bead-Based Flow Cytometric miRNA Expression Profiling

Their results demonstrate that the bead-based miRNA detection is feasible, and has the attractive properties of improved accuracy, high speed and low cost. Moreover, it is an efficient detection platform for high throughput miRNA profiling in a quantitative manner. The bead-based miRNA detection method is also easy to implement in a routine clinical setting.

References Lu J, Getz G, Miska EA, Alvarez-Saavedra E, Lamb J, Peck D, Sweet-Cordero A, Ebert BL, Mak RH, Ferrando AA, Downing JR, Jacks T, Horvitz HR, Golub TR (2005) MicroRNA expression profiles classify human cancers. Nature 435:745–746

Chapter 23

Bioluminescence miRNA Detection Method

Abstract The Bioluminescence miRNA detection technology is a competitive solid-phase hybridization-based method using the bioluminescent protein Renilla luciferase (Rluc) as a label for detection and quantification of miRNAs. The method is an alternative ELISA for the detection of target miRNAs using the free synthetic miRNAs and Rluc-labeled miRNAs that compete to bind to an immobilized antimiRNA probe. The technology was developed by Cissell et al. (Anal Chem 80:2319–2325, 2008) (Department of Chemistry and Chemical Biology, Indiana University Purdue University Indianapolis; Anal Chem 80:2319–2325, 2008). It is rapid, requiring a microplate format, a total assay time of 1.5 h without the need for sample PCR amplification, and it is a highly sensitive method for miRNA detection with a detection limit of 1 fmol. The assay offers the advantage of parallel analysis in a 96-well microtiter plate and makes it suitable for application in clinical diagnostics and drug discovery. Moreover, the high signal-to-noise ratio afforded by bioluminescence and minimal sample preparation of the technology, enhances the suitability of this method for adapting to miniaturized analytical platforms providing for high sample throughput. This technology has been applied for the determination of miR-21 in both human breast adenocarcinoma MCF-7 and nontumorigenic epithelial MCF-10A cellular extracts (Anal Chem 80:2319–2325, 2008).

23.1

Introduction

The Bioluminescence miRNA detection technology is a competitive solid-phase hybridization-based method using the bioluminescent protein Renilla luciferase (Rluc) as a label for detection and quantification of miRNAs. The method is an alternative ELISA for the detection of target miRNAs using the free synthetic miRNAs and Rluc-labeled miRNAs that compete to bind to an immobilized antimiRNA probe (Fig. 23.1). Rluc, a small 38-kDa protein which is usually employed as a reporter for gene expression analysis, can catalyze the oxidative decarboxylation of coelenterazine in Z. Wang and B. Yang, MicroRNA Expression Detection Methods, DOI 10.1007/978-3-642-04928-6_23, # Springer-Verlag Berlin Heidelberg 2010

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296

23 Bioluminescence miRNA Detection Method

Anti-miR-1

Biotin

5’-ATACATACTTCTTTACATTCCA-Biotin

NH2

miR-1

5’-UGGAAUGUAAAGAAGUAUGUAU-NH2

miR-1

5’-UGGAAUGUAAAGAAGUAUGUAU-3’

Sulfo-SMCC

Rluc

5’-UGGAAUGUAAAGAAGUAUGUAU-NH2-Rluc

Compete for binding 5’-UGGAAUGUAAAGAAGUAUGUAU-3’ 5’-ATACATACTTCTTTACATTCCA-Biotin

Neutravidin-coated microtiter plate

miR-1

5’-UGGAAUGUAAAGAAGUAUGUAU-NH2-Rluc

5’-ATACATACTTCTTTACATTCCA-Biotin

5’-ATACATACTTCTTTACATTCCA-Biotin

Immobilize onto plate

Reduced Rluc

Coelenterazine

Measure luminescence

Fig. 23.1 Schematic representation of the Bioluminescence-based miRNA detection technology using miR-1 as an example. Modified from Cissell et al. (2008)

the presence of molecular oxygen to coelenteramide (Hart et al. 1979; Matthews et al. 1977a, b; Srikantha et al. 1996). This process leads to emission of light that follows glow-type kinetics and has an emission wavelength a maximum of 485 nm (Matthews et al. 1977a). Several coelenterazine analogs have been synthesized with different properties in terms of emission wavelength, quantum efficiency, and cell permeability, thus enhancing the applications of Rluc (Matthews et al. 1977b). Since light generation occurs due to a chemical reaction, there is no requirement for external excitation light providing for the detection of Rluc activity with a high signal-to-noise ratio yielding a detection limit in the attomolezeptomole range. Additionally, due to availability of the Rluc gene for cloning, the protein can be reproducibly produced in unlimited amounts and fused to any desired molecule.

23.2

Protocol

23.2.1 Materials 1. 96-well neutravidin-coated white microtiter plates 2. High-performance nickel-bound Sepharose column 3. Plasmid phRl-CMV (Promega, WI)

23.2 Protocol

4. 5. 6. 7.

297

Plasmid pRSetB (Invitrogen) Escherichia coli cells strain 2566 (NEB Biolabs) All materials for cell culture All materials for RNA exaction

23.2.2 Instruments 1. 2. 3. 4. 5.

Fisher Scientific orbital shaker Beckman J2-MI centrifuge (Palo Alto, CA) Varian Cary Eclipse fluorescence spectrophotometer (Palo Alto, CA) Genios workstation luminometer (Tecan, NC) Perkin-Elmer UV/vis/NIR Lambda spectrophotometer

23.2.3 Reagents 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14.

15. 16. 17. 18.

Luria Bertani (LB) broth Agar Ampicillin Bovine serum albumin (BSA) Coomasie Brilliant Blue R250 stain High performance Ni+2-Sepharose beads Coelenterazine Chemically synthesized oligonucleotide probes Sulfosuccinimidyl 4-(N-maleimidomethyl)cyclohexane-1-carboxylate (sulfoSMCC) Trizol reagent Trypsin-EDTA DEPC Buffer A: [10 mM HEPES, pH7.9 containing 1.5 mM MgCl2, 10 mM KCl, and 0.5 mM DTT] Buffer B: [20 mM HEPES, pH7.9 containing 25% (v/v) glycerol, 0.42 M NaCl, 1.5 mM MgCl2, 0.2 mM EDTA, 0.5 mM phenylmethylsulfonyl fluoride (PMSF), 0.5 mM DTT, and 0.1% Nonidet P-40] Buffer C: [20 mM sodium phosphate, pH7.5 containing 20% (v/v) glycerol, 0.1 M KCl, 0.2 mM EDTA, 0.5 mM PMSF, and 0.5 mM DTT] Binding buffer: [100 mM potassium phosphate buffer containing 250 mM NaCl, 0.6 mM NaN3, and 20 mM imidazole, pH7.6] Wash buffer: [100 mM phosphate buffer containing 250 mM NaCl, 1 mM EDTA, and 0.5% BSA, pH7.4] Elution buffer: [100 mM potassium phosphate buffer containing 250 mM NaCl, 0.6 mM NaN3, and 100 mM imidazole, pH7.6]

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19. Rluc buffer: [100 mM phosphate buffer containing 250 mM NaCl, 1 mMEDTA, and 0.1% BSA, pH7.4] 20. 100 mM potassium phosphate buffer containing 100 mM NaCl, pH7.4 21. Binding buffer: [100 mM potassium phosphate buffer containing 250 mM NaCl, 0.6 mM NaN3, and 20 mM imidazole, pH7.6] 22. Wash buffer I: [100 mM potassium phosphate buffer containing 250 mM NaCl, 0.6 mM NaN3, and 50 mM imidazole, pH7.6] 23. Dialysis buffer: [100 mM potassium phosphate containing 250 mM NaCl, 0.1% BSA, 0.6 mM NaN3 pH7.4] 24. Wash buffer II: [100 mM phosphate buffer containing 250 mM NaCl, 1 mM EDTA, and 0.5% BSA, pH7.4] 25. All reagents for cell culture 26. All reagents for RNA exaction

23.2.4 Procedures The protocols described in this section are essentially the same as reported in the study by Cissell et al. (2008) (See Figs. 23.1 and 23.2). Construct Rluc (Renilla luciferase ) plasmid

Amplify and purify the Rluc plasmid

Isolate RNA from tissues or cells

Select miRNAs of your interests

Synthesize the selected miRNAs with an amino modification at its 3' end

Synthesize biotinylated anti-miRNA probes

Reduce the Rluc plasmid using TCEP

miRNA-Rluc conjugate

Hybridization (miRNAs in the RNA sample compete with the anti-miRNA probes for binding miRNA-Rluc conjugate)

Measure luminescence with a microplate reader

Fig. 23.2 Flowchart of the Bioluminescence method for miRNA expression detection. According to Cissell et al. (2008)

23.2 Protocol

23.2.4.1

299

Rluc Plasmid Construction

1. Isolate the Rluc gene from the plasmid phRl-CMV using polymerase chain reaction (PCR) employing primers (Rlucfor-50 GGTGGTGGATCCGATGGC TTCCAAGGTGTACGACCCCGAG30 , RlucRev-50 GGTGGTGAATTCTTAC TGCTCGTTCTTCAGCACGCGCTCC30 ). 2. Digest the Rluc gene and the plasmid pRsetB with restriction enzymes BamHI and EcoRI and subcloned into pRSetB to construct the plasmid pSKD2. 3. Transform the ligated plasmid into Escherichia coli cells strain 2566. 4. Perform Gene sequencing and restriction analysis to confirm the presence of the gene for Rluc.

23.2.4.2

Rluc Expression

1. Grow the cells containing the plasmid pSKD2 in LB broth containing ampicillin (100 mg/ml) at 37 C to an optical density of 0.5 measured at 600 nm. 2. Induce protein expression using IPTG (0.5 mM final concentration), and grow the cells for an additional 3 h at 37 C. 3. Harvest the cells by centrifugation at 16 C, 8,600 g, for 15 min and sonicate for 5 min with a 20 s on and 20 s off cycle. 4. Obtain the crude protein by centrifugation at 16 C, 8600 g, for 15 min.

23.2.4.3

Rluc Purification

1. Prepare high-performance nickel-bound Sepharose column by washing with distilled water followed by equilibration with the Binding buffer. 2. Load the crude Rluc protein in the Binding buffer onto the nickel-bound Sepharose column and incubate overnight at 4 C. 3. Wash the column with 20 mL of the Binding buffer, followed by 20 mL of Wash buffer. 4. Elute the purified Rluc with 5 mL of the Elution buffer. Collect the eluted fractions that show luminescent activity, and determine the purity by SDSPAGE using Coomassie staining solution. 5. Determine protein concentration by measuring absorbance at 280 nm employing an extinction coefficient of 62 520/M/cm.

23.2.4.4

Labeling of Oligonucleotide Probes

1. Obtain the sequence of a miRNA of interest from the miRNA Registry (microrna.sanger.ac.uk).

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23 Bioluminescence miRNA Detection Method

2. Design a “miRNA” oligonucleotide probe that has the same sequence as the selected miRNA but contains an amino modification at its 30 end (50 -miRNANH2-30 ). 3. Incubate this “miRNA” oligonucleotide probe with equimoles of sulfo-SMCC (1:1) at room temperature for 30 min. 4. Add TCEP to the purified Rluc descibed in 23.2.4.3 to reduce the Rluc. 5. Then add the reduced Rluc to the probe mixture at a ratio of Rluc to probe 1:5. 6. Incubate for 30 min at room temperature, and add BSA to the reaction mix to achieve a final amount of 0.1% BSA. 7. Remove the unconjugated probe and sulfo-SMCC using the Dialysis buffer to eliminate interference from unlabeled oligonucleotide. Removal of unconjugated probe is also necessary for the determination of concentration of miRNA-Rluc concentration. 8. Determine the concentration of miRNA-Rluc conjugate spectrophotometrically at 260 nm [48]. 9. Place a volume of 200 mL of miRNA-Rluc probe into a microtiter plate, and obtain the emission scan of the protein after the addition of 0.5 mL of coelenterazine (1 mg/mL).

23.2.4.5

Hybridization Study

1. Design an antisense oligonucleotide probe complementary to the selected miRNA containing a biotin moiety at the 30 end (50 -anti-miRNA-biotin-30 ). 2. Prepare biotinylated anti-miRNA probe a 20 pM/mL concentration of in Rluc buffer. 3. Make a serial dilution of the miRNA-Rluc probe using the Rluc buffer. 4. Mix 50 mL anti-miRNA-biotin probe and 50 mL of varying concentrations of miRNA-Rluc probe and incubate at 37 C for 30 min. 5. At the end of 30 min, place the probe mixture in a neutravidin-coated microtiter plate prewashed three times with the Wash buffer II and incubate with shaking at room temperature for 1 h, followed by three washes using the Wash buffer II. 6. Next, add 200 mL Rluc buffer and 0.5 mL coelenterazine to the plate, and measure luminescence with a 100-ms integration time.

23.2.4.6

Standard Curve of miRNA

1. Incubate a mixture of anti-miRNA-biotin probe (50 mL, 20 pM), miRNA-Rluc probe (25 mL, 20 pM), and 25 mL of varying concentrations of unlabeled miRNA DNA probe at 37 C for 30 min. 2. Next, add the mixture to the neutravidin-coated plate and perform the immobilization step at room temperature for 1 h shaking it well. 3. After 3-times of the wash step, add 200 mL Rluc buffer and 0.5 mL coelenterazine to the wells and measure luminescence.

23.3 Application and Limitation

301

4. Generate a dose-response curve for the miRNA in the form of DNA by plotting luminescence intensity against the concentration of that miRNA. 5. Generate a dose-response curve using miRNA in the form of RNA.

23.2.4.7

Quantification of miRNA

1. Detach adherent cells by trypsin treatment and wash with autoclaved and DEPCtreated 100 mM potassium phosphate buffer containing 100 mM NaCl, pH7.4. 2. Prepare the cellular extract using previously published protocol. Wash the cell pellet with the Buffer A, resuspend the cells in the Buffer B, and incubate on ice for 15 min. 3. Vortex the cell suspension and centrifuge for 10 min at 4 C. 4. Dilute the supernatant with the Buffer C and extract the total RNA using Trizol reagent, followed by chloroform/2-propanol precipitation. 5. Use a total of 10 mg of isolated RNA in the assay to determine the amount of the miRNA of interest by generating a dose-response curve using standard amounts of that synthetic miRNA.

23.3

Application and Limitation

The Bioluminescence miRNA detection technology employs Rluc as a direct label for hybridization assays; the bioluminescence emission, small size and requirement for the addition of only coelenterazine make Rluc an efficient label for hybridization assays. It is rapid, requiring a microplate format a total assay time of 1.5 h without the need for sample PCR amplification, and it is a highly sensitive method for miRNA detection with a detection limit of 1 fM. The assay offers the advantage of parallel analysis in a 96-well microtiter plate and makes it suitable for application in clinical diagnostics and drug discovery. Moreover, the high signal-to-noise ratio afforded by bioluminescence and minimal sample preparation of the technology enhances the suitability of this method for adapting to miniaturized analytical platforms providing for high sample throughput. Furthermore, recent findings in miRNAs research area indicate that miRNAs can be isolated from biofluids such as serum, plasma, saliva, and urine (Tricoli and Jacobson 2007). The assay developed in this work can be easily applied to the determination of miRNAs in these sample matrixes if desired since bioluminescent proteins have been employed previously in saliva and blood analysis of physiological molecules (Desai et al. 2002; Mirasoli et al. 2002). This technology has been applied for the determination of miR-21 in both human breast adenocarcinoma MCF-7 and nontumorigenic epithelial MCF-10A cellular extracts (Cissell et al. 2008). The method allows sensitive, accurate, and precise measurement of miR-21 in vitro as well as in cells and is able to discriminate levels

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23 Bioluminescence miRNA Detection Method

of miR-21 in cancerous versus noncancerous cells, which can be a significant advantage in early cancer diagnosis.

References Cissell KA, Rahimi Y, Shrestha S, Hunt EA, Deo SK (2008) Bioluminescence-based detection of microRNA, miR21 in breast cancer cells. Anal Chem 80:2319–2325 Desai UA, Deo SK, Hyland KV, Poon M, Daunert S (2002) Determination of prostacyclin in plasma through a bioluminescent immunoassay for 6-keto-prostaglandin F1alpha: implication of dosage in patients with primary pulmonary hypertension. Anal Chem 74:3892–3898 Hart RC, Matthews JC, Hori K, Cormier MJ (1979) Renilla reniformis bioluminescence: luciferase-catalyzed production of nonradiating excited states from luciferin analogues and elucidation of the excited state species involved in energy transfer to Renilla green fluorescent protein. Biochemistry 18:2204–2210 Matthews JC, Hori K, Cormier MJ (1977a) Purification and properties of Renilla reniformis luciferase. Biochemistry 16:85–91 Matthews JC, Hori K, Cormier MJ (1977b) Substrate and substrate analogue binding properties of Renilla luciferase. Biochemistry 16:5217–5220 Mirasoli M, Deo SK, Lewis JC, Roda A, Daunert S (2002) Bioluminescence immunoassay for cortisol using recombinant aequorin as a label. Anal Biochem 306:204–211 Srikantha T, Klapach A, Lorenz WW, Tsai LK, Laughlin LA, Gorman JA, Soll DR (1996) The sea pansy Renilla reniformis luciferase serves as a sensitive bioluminescent reporter for differential gene expression in Candida albicans. J Bacteriol 178:121–129 Tricoli JV, Jacobson JW (2007) MicroRNA: potential for cancer detection, diagnosis, and prognosis. Cancer Res 67:4553–4555

Chapter 24

Molecular Beacon Method

Abstract Molecular beacons are single-stranded oligonucleotide hybridization probes with a stem-and-loop structure that recognize and report the presence of specific nucleic acids in homogeneous solutions (Tyagi et al. 1996). The loop contains a probe sequence that is complementary to a target sequence, and the stem is formed by the annealing of complementary arm sequences that are located on either side of the probe sequence. Paiboonskuwong and Kato (Nucleic Acids Symp Ser (Oxf) 50:327–328, 2006) from the Research Institute for Cell Engineering, National Institute of Advanced Industrial Science and Technology (AIST; Tsukuba Science City, Japan) modified the molecular beacon approach with an aim to detect miRNAs in the mature form. The probe is modified so that when hybridizing with either pre-miRNA or pri-miRNA, its fluorescence is quenched by the guanine in the sequences of pre-miRNA or pri-miRNA complementarily to the probe. Hybridization of the probe with a mature miRNA which has no complementary guanine can result in fluorescent emission. In this way, the modified molecular beacon method can distinguish the mature from the precursor miRNAs.

24.1

Introduction

Molecular beacons are single-stranded oligonucleotide hybridization probes with a stem-and-loop structure that recognize and report the presence of specific nucleic acids in homogeneous solutions (Tyagi and Kramer 1996; Marras 2006; Marras et al. 2006; Bratu 2006; Broude 2002). The loop contains a probe sequence that is complementary to a target sequence, and the stem is formed by the annealing of complementary arm sequences that are located on either side of the probe sequence. A fluorophore is covalently linked to the end of one arm and a quencher is covalently linked to the end of the other arm. Molecular beacons do not fluoresce when they are free in solution or in the absence of targets, because the stem places

Z. Wang and B. Yang, MicroRNA Expression Detection Methods, DOI 10.1007/978-3-642-04928-6_24, # Springer-Verlag Berlin Heidelberg 2010

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24 Molecular Beacon Method

the fluorophore so close to the nonfluorescent quencher that they transiently share electrons, eliminating the ability of the fluorophore to fluoresce. However, when they hybridize to a nucleic acid strand containing a target sequence, they form a probe-target hybrid that is longer and more stable than the stem hybrid. Consequently, the molecular beacons undergo a spontaneous conformational reorganization that forces the stem hybrid to dissociate and the fluorophore and the quencher to move away from each other, restoring fluorescence (Figs. 24.1 and 24.2). The rigidity and length of the probe-target hybrid precludes the simultaneous existence of the stem hybrid. Only perfectly complementary targets elicit this response, as hybridization does not occur when the target contains a mismatched nucleotide or a deletion (Kramer http://www.molecular-beacons.org/Introduction.html). Molecular beacons can be synthesized that possess differently colored fluorophores, enabling assays to be carried out that simultaneously detect different targets

Probe

a

miRNA miRNA

+

Probe

Q

F

Q F Quenched

F G

Pre-miRNA

b

Probe

Pre-miRNA

Probe

F G Probe

G F Quenched

Mature miRNA

Mature miRNA

G

Fig. 24.1 Schematic depiction of molecular beacon probe and the mechanism for miRNA detection. The 3’ terminal cytosine of of a molecular beacon probe is covalently linked to BODIPY1 FL (F), which is quenched by the adjacent guinine base (G) through base-pairing. In this inactivated state, molecular beacons do not fluoresce. (a) When they hybridize to a nucleic acid strand containing a target sequence, they form a probe-target hybrid that is longer and more stable than the stem hybrid. Consequently, the molecular beacons undergo a spontaneous conformational reorganization that forces the stem hybrid to dissociate and the fluorophore and the quencher to move away from each other, restoring fluorescence. (b) When the probe hybridizdes with the precursor miRNA (pre-miRNA), it stays inactivated due to the quench by the guanine base in the pre-miRNA. However, when a molecular beacon probe is hybridized with a mature miRNA that does not have guanine bases at the position to basepair with the cytosine in the probe, then it fluoresces. Modified from Paiboonskuwong and Kato (2006)

24.1 Introduction

305 Select miRNAs of your interests

Design and synthesize stem-loop molecular beacon probes

Isolate RNA from tissues or cells

Label the probes with fluorophore BODIPY® FL at 3'end

Hybridization of miRNAs and probes

Detect fluorescence and analyze data

Fig. 24.2 Flowchart of the Molecular Beacon method for miRNA expression detection. According to Paiboonskuwong and Kato (2006)

in the same reaction (Vet and Marras 2004; Kramer http://www.molecular-beacons. org/Introduction.html). For example, multiplex assays can contain a number of different primer sets, each set enabling the amplification of a unique gene sequence from a different pathogenic agent, and a corresponding number of molecular beacons can be present, each containing a probe sequence specific for one of the amplicons, and each labeled with a fluorophore of a different color. The color of the resulting fluorescence, if any, identifies the pathogenic agent in the sample, and the number of amplification cycles required to generate detectable fluorescence provides a quantitative measure of the number of target organisms present. Molecular beacons are extraordinarily specific. They easily discriminate target sequences that differ from one another by a single nucleotide substitution (Tyagi et al. 1998; Marras et al. 1999; Kramer http://www.molecular-beacons.org/ Introduction.html). The reason that molecular beacons are so “finicky” is that they can exist in two different stable physical states. In one state, the molecular beacons are hybridized to their targets, and energy is stored in the probe-target helix. In the second state, the molecular beacons are free in solution, and energy is stored in their stem helix. Molecular beacons are designed so that their probe sequence is just long enough for a perfectly complementary probe-target hybrid to be more stable than the stem hybrid. Consequently, the molecular beacons spontaneously form fluorescent probe-target hybrids. However, if as little as a single nucleotide in the target is not complementary to the probe sequence of the molecular beacon, the probetarget helix would be less stable. In this situation, the stem helix of the molecular beacon is more stable than the mismatched probe-target helix, and the molecular

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24 Molecular Beacon Method

beacons remain unhybridized. Thus, molecular beacons can be thought of as “molecular switches” that are on their targets and brightly fluorescent when the targets are perfectly complementary to the probe, but remain off the targets and dark if the targets contain a mutation (Kramer http://www.molecular-beacons.org/ Introduction.html). The probes are particularly suited for monitoring the synthesis of specific nucleic acids in real/actual time. When used in nucleic acid amplification assays, gene detection is homogeneous and sensitive, and can be carried out in a sealed tube. When introduced into living cells, these probes should enable the origin, movement, and fate of specific mRNAs to be traced. This provides a novel nonradioactive method for detecting specific sequences of nucleic acids. They are useful in situations where it is either not possible or desirable to isolate the probe-target hybrids from an excess of the hybridization probes (Kramer http://www.molecularbeacons.org/Introduction.html). Molecular beacons can be used as amplicon detector probes in diagnostic assays. As nonhybridized molecular beacons are dark, it is not necessary to isolate the probe-target hybrids to determine the number of amplicons synthesized during an assay. Molecular beacons are added to the assay mixture before carrying out gene amplification and fluorescence is measured in real/actual time. The assay tube remains sealed. Consequently, the amplicons cannot escape to contaminate untested samples. Furthermore, the use of molecular beacons provides an additional level of specificity. Since it is very unlikely that false amplicons or primer-dimers possess target sequences for the molecular beacons, the generation of fluorescence is exclusively due to the synthesis of the intended amplicons (Kramer http://www. molecular-beacons.org/Introduction.html). Paiboonskuwong and Kato used the molecular beacon approach with a small, elegant modification to detect miRNAs in the mature form (Paiboonskuwong and Kato 2006). The probe is modified so that when hybridizing with either pre-miRNA or pri-miRNA, its fluorescence is quenched by the guanine in the sequences of premiRNA or pri-miRNA complementarily to the probe. Hybridization of the probe with a mature miRNA which has no complementary guanine can result in fluorescent emission. In this way, the modified molecular beacon method can distinguish the mature from the precursor miRNAs (Fig. 24.1).

24.2

Protocol

24.2.1 Materials 1. Zuker DNA folding program (available on the internet at http://frontend.bioinfo. rpi.edu/applications/mfold/cgi-bin/dna-form1.cgi) 2. Beacon Designer software (available from Premier Biosoft International at www.premierbiosoft.com)

24.2 Protocol

307

24.2.2 Instruments 1. Real-time PCR system 750 (Applied Biosystems) or an equivalent 2. Any instrument able to detect fluorescence

24.2.3 Reagents 1. Reverse transcription reaction kit 2. PCR reaction kit 3. RNA isolation kit and reagents

24.2.4 Procedures The protocols described in this section are essentially the same as reported in the study by Kramer posted in the website http://www.molecular-beacons.org/PA_ design.html (see Figs. 24.1 and 24.2). 24.2.4.1

Design of Molecular Beacon Probes

According to Tyagi (Tyagi and Kramer 1996; Marras et al. 2003), in order to design molecular beacons that function optimally under a given set of assay conditions, it is important to understand how their fluorescence changes with temperature in the presence and in the absence of their targets. As shown by the green fluorescence versus temperature trace below, at lower temperatures molecular beacons exist in a closed state, the fluorophore and the quencher are held in close proximity to each other by the hairpin stem, and there is no fluorescence. However, at high temperatures the helical order of the stem gives way to a random-coil configuration, separating the fluorophore from the quencher and restoring fluorescence. The temperature at which the stem melts depends upon the GC content and the length of the stem sequence. If a target is added to a solution containing a molecular beacon at temperatures below the melting temperature of its stem, the molecular beacon spontaneously binds to its target, dissociating the stem, and turning on its fluorescence. How the fluorescence of the probe-target hybrid varies with the temperature is indicated by the red fluorescence versus temperature trace. At low temperatures, the probe-target hybrid remains brightly fluorescent, but as the temperature is raised the probe dissociates from the target and tends to return to its hairpin state, diminishing the fluorescence significantly. The temperature, at which the probe-target hybrid melts apart, depends upon the GC content and the length of the probe sequence. The longer the probe and the higher its GC content, the higher the melting temperature of the probe-target hybrid. It is important to note that the probe-target hybrid melting temperature can be adjusted independently

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24 Molecular Beacon Method

from the melting temperature of the stem by selecting a target region of appropriate length. The fluorescence versus temperature profiles of the molecular beacon that were used in this example indicate that the molecular beacon is suitable for assays that are performed below 55˚C, because below 55˚C the free molecular beacons remain dark, yet the probe-target hybrids form spontaneously and are stable. Molecular beacons can also be designed with the help of a dedicated software package called “Beacon Designer”, which is available from Premier Biosoft International (www.premierbiosoft.com). In addition, companies, such as NYtor (www. nytor.nl), are specialized in designing real-time PCR assays that utilize molecular beacons for the detection of single nucleotide polymorphisms and high-throughput multiplex diagnostic assays. 1. The process of molecular beacon design begins with the selection of the probe sequence or selection of target miRNAs of interest. If you are designing molecular beacons to detect the synthesis of products during polymerase chain reactions, you can select any region within the amplicon that is outside the primer binding sites. The probe sequence of the molecular beacon should be so long that at the annealing temperature of the PCR it is able to bind to its target. In order to discriminate between amplicons that differ from one another by as little as a single nucleotide substitution, the length of the probe sequence should be such that it dissociates from its target at temperatures 7–10 C higher than the annealing temperature of the PCR. If, on the other hand, single-nucleotide allele discrimination is not desired, longer and more stable probes can be chosen. The melting temperature of the probe-target hybrid can be predicted using the “percent-GC” rule or “nearest neighbor” rules (available in most probe or primer design software packages). The prediction should be made for the probe sequence alone before choosing the stem sequences. In practice, the length of the probe sequence usually falls in the range between 15 and 30 nucleotides [Kramer http://www.molecular-beacons.org/PA_design.html]. 2. A molecular beacon probe is a stem-loop structure. The loop portion is a probe sequence that is complementary to the predetermined sequence in the target miRNA. For example, miR-1 has the sequence of 5’-UGGAAUGUAAAGAAGUAUGUAU-3’. Accordingly, the loop portion of a molecular beacon probe should be: 5’-AUACAUACUUCUUUACAUUCCA-3’. 3. After selecting the probe sequence, two complementary arm sequences are added on either side of the probe sequence. The sequence of 3’-stem should be designed to be complementary to the sequences of pri-miRNA or pre-miRNA but not that of mature miRNA (Fig. 24.1). We therefore have for miR-1: 5’gcacgAUACAUACUUCUUUACAUUCCAcgugc-3’. 4. The cytosine base at one end of the probe is covalently linked with BODIPY1 FL (Molecular Probes). The fluorescent emission from the probe modified with BODIPY1 FL should be diminished after hybridization with the adjacent guanine base. The length and the GC content of the stem sequence is designed in such a way that at the annealing temperature of the PCR, and in the absence of the target,

24.2 Protocol

309

the molecular beacons remain closed and non-fluorescent. This is ensured by choosing a stem that melts 7–10 C higher than the annealing temperature of the PCR. Usually the stems are 5–7 basepairs long and have a very high GC content (75–100%). The melting temperature of the stem cannot be predicted by the percent-GC rule, since the stem is created by intramolecular hybridization. In general, 5 basepair-long GC-rich stems will melt between 55 and 60 C, 6 basepair-long GC-rich stems will melt between 60 and 65 C, and 7-basepair long GC-rich stems will melt between 65 and 70 C. Although any arbitrary sequence can be used in designing the stems, don’t use guanosine residues near the end to which the fluorophore is attached (instead, use them at the end where the quencher is attached), as guanosine residues tend to quench the fluorophore. Longer stems can be used to enhance the specificity of the molecular beacons (Kramer http://www.molecular-beacons.org/PA_design.html). 5. It is important that the conformation assumed by the free molecular beacons be the intended hairpin structure, rather than other structures that either do not place the fluorophore in the immediate vicinity of the quencher, or that form longer stems than intended. The former will cause high background signals, and the latter will make the molecular beacons sluggish in binding to the targets. A folding of the selected sequence by the Zuker DNA folding program will reveal such problems. If unexpected secondary structures result from the choice of the stem sequence, a different stem sequence can be chosen. If, on the other hand, unexpected secondary structures arise from the identity of the probe sequence, the frame of the probe can be moved along the target sequence to obtain a probe sequence that is not self-complementary. Small stems within the probe’s hairpin loop that are 2- to 3-nucleotides long do not adversely affect the performance of molecular beacons. 6. As with PCR primers, the sequence of the molecular beacon should be compared with the sequences of the primers, using a primer design software program to make sure that there are no regions of substantial complementarities that may cause the molecular beacon to bind to one of the primers, causing primer extension. Also, the primers that are used should be designed to produce a relatively short amplicon. In general, the amplicons should be less than 150basepairs long. Molecular beacons are internal probes that must compete with the other strands of the amplicon for binding to the strand that contains their target sequence. Having a shorter amplicon allows the molecular beacons to compete more efficiently, and therefore produces stronger fluorescence signals during real-time PCR. In addition, smaller amplicons result in more efficient amplification. And finally, the magnitude of the molecular beacon signal can be increased by performing asymmetric PCR, in which the primer that makes the strand that is complementary to the molecular beacon is present at a slightly higher concentration than the other primer. 7. A 3’ terminus of the probe is labeled with 4,4-defluoro-5,7-dimethyl-4-bora3a,4a-diaza-s-indacene-3-propionic acid (BODIPY1 FL), whose fluorescence is quenched by the adjacent guanine base(s).

310

24.2.4.2

24 Molecular Beacon Method

Total RNA Extraction

See Section II for detailed protocols.

24.2.4.3

Detect Fluorescence

See Fig. 24.2

24.3

Application and Limitation

Molecular beacons have three key properties that enable the design of new and powerful diagnostic assays: (1) they only fluoresce when bound to their targets, (2) they can be labeled with a fluorophore of any desired color, and (3) they are so specific that they easily discriminate single-nucleotide differences in miRNAs. Now that a number of new and versatile spectrofluorometric thermal cyclers are available to clinical diagnostic and research laboratories, assays that simultaneously utilize as many as seven differently colored molecular beacons can be designed. This enables cost-efficient multiplex assays to be developed for miRNA expression profiling. Molecular beacons are also ideal probes for use in diagnostic assays designed for genetic screening and SNP detection for miRNAs. Molecular beacon probes in conjunction with real-time RT-PCR provide quantitative analysis of miRNA expression in a cell. The method is straightforward and simple to perform.

References Bratu DP (2006) Molecular beacons: fluorescent probes for detection of endogenous mRNAs in living cells. Methods Mol Biol 319:1–14 Broude NE (2002) Stem-loop oligonucleotides: a robust tool for molecular biology and biotechnology. Trends Biotechnol 20:249–256 Marras SAE (2006) Selection of fluorophore and quencher pairs for fluorescent nucleic acid hybridization probes. Methods Mol Biol 335:3–16 Marras SAE, Kramer FR, and Tyagi S (1999) Multiplex detection of single-nucleotide variations using molecular beacons. Genet Anal 14:151–156 Marras SAE, Kramer FR, and Tyagi S (2003) Genotyping single nucleotide polymorphisms with molecular beacons. In Kwok, P. Y. (ed.), Single nucleotide polymorphisms: methods and protocols. The Humana Press Inc., Totowa, NJ, Vol. 212, pp. 111–128 Marras SAE, Tyagi S, and Kramer FR (2006) Real-time assays with molecular beacons and other fluorescent nucleic acid hybridization probes. Clin Chim Acta 363:48–60 Paiboonskuwong K, Kato Y (2006) Detection of the mature, but not precursor, RNA using a fluorescent DNA probe. Nucleic Acids Symp Ser (Oxf) 50:327–328

References

311

Tyagi S, Kramer FR (1996) Molecular beacons: probes that fluoresce upon hybridization. Nat Biotechnol 14:303–308 Tyagi S, Bratu DP, and Kramer FR (1998) Multicolor molecular beacons for allele discrimination. Nat Biotechnol 16:49–53 Vet JAM, Marras SAE (2004) Design and optimization of molecular beacon real-time polymerase chain reaction assays. In Herdewijn, P. (ed.), Oligonucleotide synthesis: Methods and Applications. Humana Press, Totowa, NJ, Vol. 288, pp. 273–290

Chapter 25

Ribozyme Method

Abstract The hairpin ribozyme belongs to a family of small catalytic RNAs that cleave the RNA substrates in a reversible reaction, generating 20,30-cyclic phosphate and 50-hydroxyl termini. The ribozyme method takes advantage of their ability to perform RNA-cleavage reactions under multiple turnovers and their potential to be regulated by external oligonucleotides. The rational and straightforward design of the hairpin ribozymes can be sequence-specifically induced by external oligonucleotides and cleave a short RNA substrate labeled with a fluorophor at the 30 -end and a quencher at the 50 -end, as a function of the presence or absence of a miRNA effector. This design enables real-time monitoring of ribozyme activity via FRET read-out. This technique was first introduced to detect miRNAs by Hartig et al., Kekule´ Institut fu¨r Organische Chemie und Biochemie, Universita¨t Bonn, Bonn, Germany (J Am Chem Soc 126:722–723, 2004). The ribozyme method has been tested with nine examples of ribozymes, which are activated by nine different miRNAs from Drosophila. Due to intrinsic signal amplification, the sensitivity of ribozyme method for the detection of miRNAs is increased at least an order of magnitude compared to that of standard molecular beacons. These probes may be useful in applications that require direct detection of nucleic acids within their natural environment.

25.1

Introduction

Ribozymes are small and versatile catalytic RNA molecules that cleave RNAs at specific sites in a Watson–Crick-based specific manner. The rapidly developing field of RNA catalysts is of current interest not only because of their intrinsic catalytic properties but also because of their potential utility as therapeutic agents and specific regulators of gene expression (Rossi and Sarver 1990; Sarver et al. 1990; Erickson and Izant 1992; Rossi 1995; Eckstein and Lilley 1996; Turner 1997; Krupp and Gaur 2000; Sioud 2004).

Z. Wang and B. Yang, MicroRNA Expression Detection Methods, DOI 10.1007/978-3-642-04928-6_25, # Springer-Verlag Berlin Heidelberg 2010

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25 Ribozyme Method

Two types of ribozymes, hammerhead and hairpin ribozymes according to their tertiary structure, have been extensively investigated and proven to be effective in repressing the expression of various genes (Kawasaki et al. 2004; Akashi et al. 2005; Doherty and Doudna 2001; Lilley 2005). The hammerhead ribozyme consists of two binding regions (stems I and III), which are complementary to the target mRNA sequences flanking the cleave site, and a catalytic core region that includes a mutually complementary stem II (Doherty and Doudna 2001). The target mRNA, on the other hand, requires NUX triplet (N: any base; X: any base except for G) to be present as a substrate of the ribozyme (Shimayama et al. 1995). The hairpin ribozyme belongs to the family of small catalytic RNAs that cleave RNA substrates in a reversible reaction that generates 20,30-cyclic phosphate and 50-hydroxyl termini. The ribozyme method takes advantage of their ability to perform RNA-cleavage reactions under multiple turnover and their potential to be regulated by external oligonucleotides. The rational and straightforward design of hairpin ribozymes can be sequence-specifically induced by external

c g ug c UU C CA U UC GU U A AG C C A U A A UUU UGU Active A UC AA U G CA AC AA Q F C C AU G 3’-AUC CGUGC AC UAU U 5’-UAG A gcacg A UG A A UA AG

Inactive

C

CA Q F C C 3’-AUC CGUGC-5’ 5’-UAGAGAAgcacg |||||||||||| Anti-miRNA AUCUCUUcgugc DA-HR

miRNA

B

Pseudo half-knot formation

B

Fig. 25.1 Schematic illustration of a hairpin ribozyme responsive to a target miRNA. A hairpin ribozyme consists of three domains: domains A, B, and C. Domain C contains a region complementary to the target miRNA (anti-miRNA region in red) and a region that can hybridize to domain A (DA-HR in purple). Normally, domain C is inserted between domains A and B, preventing A and B from docking and rendering the ribozyme inactive (left). When the target miRNA (red) binds to its complementary region in domain C, the catalytic activity of ribozyme is activated to cleave the fluorophor (F)- and quencher (Q)-labeled substrate (Modified from Hartig et al. 2004)

25.2 Protocol

315

oligonucleotides and cleave a short RNA substrate labeled with a fluorophor at the 30 -end and a quencher at the 50 -end, as a function of the presence or absence of a miRNA effector (Fig. 25.1). This design enables real-time monitoring of ribozyme activity via FRET read-out (Jenne et al. 1999, 2001; Hartig et al. 2002, 2004; Vitiello et al. 2000). Catalysis of RNA cleavage by hairpin ribozyme depends on its conformational flexibility during the docking of two helical domains, A and B (Fedor 2000; Rupert and Ferre´-D’Amare´ 2001). On the basis of this mechanism, variants of the hairpin ribozyme can be induced by external effector oligonucleotides (e.g., miRNAs) that interfere with the docking process. This is done by incorporating domain C, which is complementary to the target miRNA, and also contains a short sequence that can partially pair with domain A, thus rendering the ribozyme inactive. When the complementary target sequence is added, it hybridizes with domain C and forms a pseudo-half-knot structure (Ecker et al. 1992). Domains A and B can dock again, resulting in the cleavage of the substrate and the generation of a fluorescence signal (Komatsu et al. 2002; Vaish et al. 2003). Due to intrinsic signal amplification, their sensitivity is increased at least an order of magnitude compared to standard molecular beacons. Properly designed ribozymes exhibit very low cleavage activity in the absence of the corresponding miRNA and are activated when it is added. Significant increase in fluorescence should be well detectable at miRNA concentrations as low as 5 nM, corresponding to a detection limit of 50 fmol miRNA in the reaction mixture (Hartig et al. 2004).

25.2

Protocol

25.2.1 Materials 1. 384-well-plates (Corning)

25.2.2 Instruments 1. Fluorscan (Ascent FL) or an equivalent 2. NanoDrop spectrophotometer

25.2.3 Reagents 1. MgCl2 2. MAXIscript1 T7 Kit (Ambion) 3. Trizol Reagent (Invitrogen, Carlsbad, CA)

316

4. 5. 6. 7. 8.

25 Ribozyme Method

mirVana miRNA Isolation Kit (Ambion) Phenol/chloroform DNase I (Invitrogen) Absolute ethanol DMSO (Invitrogen)

25.2.4 Procedures The protocols described in this section are essentially the same as reported in the study by Hartig et al. in 2004 (see Figs. 25.1 and 25.2). A hairpin ribozyme contains two domains (A and B), with each domain containing an unpaired loop and two basepaired helices (H1-H2 or H3-H4). Thus, a hairpin ribozyme has two loops (loops A and B) and four stems (H1–H4), with the reactive

Domain C 5’-UAGAGAAgcacgAUACAUACUUCUUUACAUUCCAcgugcUUCUCUA-3’ Anti-miR-1 (or other miRNAs) DA-HR

5’-UAGAGAAgcacg |||||||||||| Anti-miR-1 3’-AUCUCUUcgugc DA-HR

+

Ribozyme substrate 5’-(Q)-CGUGCCCACCUA-(F)-3’

miR-1 5’-UGGAAUGUAAAGAAGUAUGUAU-3’

AC C C (F)-AUC CGUGC-(Q) 5’-UAGAGAAgcacgAUACAUACUUCUUUACAUUCCAcgugcUUCUCUA-3’ 3’-UAUGUAUGAAGAAAUGUAAGGU-5’

Fig. 25.2 Schematic illustration of the design of domain C and ribozyme substrate using miR-1 as a target example. The sequences of domain C and ribozyme are shown. Domain C contains a region complementary to miR-1 (anti-miR-1 region in red), a region that can hybridize to domain A (DA-HR in purple), and a linker sequence that is partially complementary to the ribozyme substrate. The ribozyme substrate contains from 50 - to 30 -end a region partially complementary to the linker sequence in domain C, an uncomplimentary region (CCAC), and a region complementary to the domain A-binding site in domain C

25.2 Protocol

317

phosphodiester located within loop A (Feldstein et al. 1989; Hampel and Tritz 1989; Haseloff and Gerlach 1989). Assembly of the functional structure imposes no obvious constraints on the maximum length of H1 or H4. And H2 and H4 have strictly four and five basepairs, respectively. Domain C contains a region complementary to the target miRNA, and a region that can hybridize to domain A of the ribozyme, preventing A and B from docking and rendering the ribozyme inactive. The ribozyme cleaves the fluorophor (F)- and quencher (Q)-labeled substrate when the miRNA for detection binds to its complementary region in domain C (Fig. 25.1).

25.2.4.1

Design of Ribozyme Domain B and Domain C

1. Obtain sequences for the miRNAs of interest from the miRNA Registry (microrna.sanger.ac.uk). 2. Design a hairpin ribozyme domain B for the selected miRNA using the computer software pcFOLD or other compatible softwares. 3. Domain C, a stem-loop structure, should contain a miRNA-hybridizing region, domain A-hybridizing region, and 30 - and 50 -flanking regions. Design an oligonucleotide fragment antisense to the selected miRNA for the detection of a miRNA-hybridizing region. 4. Add linker sequences “gcacg” to the 50 -end and “cgugc” to the 30 -end of the miRNA-hybridizing region, respectively. 5. Add a sequence “UAGAGAA” upstream (to the 50 -end) and a sequence UUCUCUA (the domain A-hybridizing region) downstream (to the 30 -end), the above fragment. Considering domain C for miR-1 as an example, we have: 50 -UAGAGAAgcacgAUACAUACUUCUUUACAUUCCAcgugcUUCUCUA-30 . Note that the region UAGAGAAgcacg is complementary to cgugcUUCUCUA, which forms a stem leaving miR-1 sequence in a loop structure (Fig. 25.2). 6. Connect domains C and B, with domain C upstream of domain B.

25.2.4.2

Synthesis of Ribozyme

1. Attach the sequence of T7 promoter to the 50 -end of the above-designed ribozyme to form a template for in vitro transcription. Synthesize the T7-ribozyme fragment in the form of DNA using the service provided by IDT1 (Integrated DNA Technologies, Coralville, IA, USA) or PCR amplify the fragment. 2. Perform in vitro transcription using T7-ribozyme fragment as a template and MAXIscript1 T7 Kit (Ambion) according to the manufacture’s instruction. 3. Digest the template with RNAse A free DNase I and purify the ribozyme. 4. Dissolve the ribozyme in DEPC H2O and measure the concentration using a NanoDrop spectrophotometer.

318

25.2.4.3

25 Ribozyme Method

Designing of Ribozyme Domain A Substrate

1. A ribozyme substrate should contain in its 50 -end a region partially complementary to the 50 -end linker sequence (gcacg) in domain C and in its 30 -end region partially complementary to the most 50 -end 3 nts in domain C. The middle region should contain 4 nts that are not complementary to the domain A-hybridizing region in domain C. Considering domain C for miR-1 as an example, we have: 50 -AUGUAUCCACCUA-30 , in which the bold letters represent the regions complementary to the linker and the domain A-hybridizing region, respectively (Fig. 25.2). 2. Label the substrate sequence with a quencher at the 50 -end and a fluorophor at the 30 -end using the service provided by Dharmacon (Lafayette, CO).

25.2.4.4

Ribozyme Reaction

1. Mix ribozyme and miRNA in 50 mM Tris/HCl (pH 7.5), 30 mM MgCl2 and 200 nM (20-fold excess over ribozyme) of labeled ribozyme substrate in 384-well-plates in a volume of 50 mL. Incubate for 10 min at 37 C. 2. Start the cleavage reaction by adding MgCl2. 3. Monitor the cleavage reaction by measuring the fluorescence of the FAMgroup, with excitation at 485 nm and emission at 518 nm using Fluorscan (Ascent FL).

25.3

Application and Limitation

The catalytic activity increases in a concentration-dependent manner. The ribozyme method has been tested with nine examples of ribozymes, activated by nine different miRNAs from Drosophila: iHP-miR-1, iHP-miR-2, iHP-miR-4, iHPmiR-5, iHP-mi-R7, iHP-miR-10, iHP-miR-34, iHP-miR-79, and iHP-let-7 (Hartig et al. 2004). Their results demonstrated that the hairpin ribozymes can be designed in a rational and straightforward manner that can be induced by external oligonucleotides sequence-specifically. This design enables real-time monitoring of ribozyme activity via FRET read-out. Due to intrinsic signal amplification, their sensitivity is increased at least an order of magnitude compared to that of standard molecular beacons. These probes may be useful in applications that require direct detection of nucleic acids within their natural environment. When ribozymes work inside the cells, they must be internalized into individual cells and access the target mRNA. However, cellular uptake of ribozymes and other naked nucleic acids is usually inefficient, due to their charged composition and large molecular size. To overcome this problem, liposomes and charged lipids are commonly used as delivery systems for ribozymes. Complexes of nucleic acids

References

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with cationic lipids are usually internalized into the cells by endocytosis. For application of ribozymes in vivo, chemically synthesized ribozymes can be directly administered, or a plasmid vector encoding ribozyme genes can be introduced into the cells, where ribozymes can be transcribed by transcriptional factors in the host. Since naked nucleic acids are rapidly degraded by nucleases in cells, especially in the gastrointestinal tract and blood, ribozymes synthesized in vitro should be protected by chemical modification, such as thio-modification or alkylation at the 20 position of the ribose ring.

References Akashi H, Matsumoto S, Taira K (2005) Gene discovery by ribozyme and siRNA libraries. Nat Rev Mol Cell Biol 6:413–422 Doherty EA, Doudna JA (2001) Ribozyme structures and mechanisms. Annu Rev Biophys Biomol Struct 30:457–475 Ecker DJ, Vickers TA, Bruice TW, Freier SM, Jenison RD, Manoharan M, Zounes M (1992) Pseudo–half-knot formation with RNA. Science 257:958–961 Erickson RP, Izant J, eds (1992) Gene Regulation: Biology of Antisense RNA and DNA. RavenPress, New York Eckstein F, Lilley DMJ, eds (1996) Nucleic Acids and Moleculad Biology: Catalytic RNA Vol. 10, Spling-Verlag, Berlin Fedor MJ (2000) Structure and function of the hairpin ribozyme. J Mol Biol 297:269–291 Feldstein PA, Buzayan JM, Bruening G (1989) Two sequences participating in the autolytic processing of satellite tobacco ringspot virus complementary RNA. Gene 82:53–61 Hampel A, Tritz R (1989) RNA catalytic properties of the minimum (-)sTRSV sequence. Biochemistry 28:4929–4933 Hartig JS, Gru¨ne I, Najafi-Shoushtari SH, Famulok M (2004) Sequence-specific detection of MicroRNAs by signal-amplifying ribozymes. J Am Chem Soc 126:722–723 Hartig JS, Najafi-Shoushtari SH, Gru¨ne I, Yan A, Ellington AD, Famulok M (2002) Proteindependent ribozymes report molecular interactions in real time. Nat Biotechnol 20:717–722 Haseloff J, Gerlach WL (1989) Sequences required for self-catalysed cleavage of the satellite RNA of tobacco ringspot virus. Gene 82:43–52 Jenne A, Gmelin W, Raffler N, Famulok M (1999) Real-time characterization of ribozymes by fluorescence resonance energy transfer (FRET). Angew Chem Int Ed 38:1300–1303 Jenne A, Hartig JS, Piganeau N, Tauer A, Samarsky DA, Green MR, Davies J, Famulok M (2001) Rapid identification and characterization of hammerhead-ribozyme inhibitors using fluorescence-based technology. Nat Biotechnol 19:56–61 Kawasaki H, Wadhwa R, Taira K. (2004) World of small RNAs: from ribozymes to siRNA and miRNA. Differentiation 72:58–64 Komatsu Y, Nobuoka K, Karino-Abe N, Matsuda A, Ohtsuka E (2002) In vitro selection of hairpin ribozymes activated with short oligonucleotides. Biochemistry 41:9090–9098 Krupp G, Gaur RK, eds (2000) Ribozyme, Biochemistry and Biotechnology. Lilley DM (2005) Structure, folding and mechanisms of ribozymes. Curr Opin Struct Biol 15:313–323 Rossi JJ (1995) Controlled, targeted, intracellular expression of ribozymes: progress and problems. Trends Biotechnol 13:301–306 Rossi JJ, Sarver N (1990) RNA enzymes (ribozymes) as antiviral therapeutic agents. Trends Biotechnol 8:179–183

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Rupert PB, Ferre´-D’Amare´ AR (2001) Crystal structure of a hairpin ribozyme-inhibitor complex with implications for catalysis. Nature 410:780–786 Sarver N, Cantin EM, Chang PS, Zaida JA, Ladne PA, Stephenes DA, Rossi JJ (1990) Ribozymes as potential anti-HIV-1 therapeutic agents. Science 247:1222–1225 Shimayama T, Nishikawa S, Taira K (1995) Generality of the NUX rule: kinetic analysis of the results of systematic mutations in the trinucleotide at the cleavage site of hammerhead ribozymes. Biochemistry 34:3649–3654 Sioud M, ed (2004) Methods in Molecular Biology: Ribozymesand siRNA Protocols. Vol. 252, Humana Press, Totowa Turner PC, ed (1997) Methods in Molecular Biology: Ribozyme Protocols. Vol. 74, Humana Press, Totowa Vaish NK, Jadhav VR, Kossen K, Pasko C, Andrews LE, McSwiggen JA, Polisky B, Seiwert SD (2003) Zeptomole detection of a viral nucleic acid using a target-activated ribozyme. RNA 9:1058–1072 Vitiello D, Pecchia DB, Burke JM. Intracellular ribozyme-catalyzed trans-cleavage of RNA monitored by fluorescence resonance energy transfer. RNA. 2000;6:628–637

Chapter 26

Electrocatalytic Moiety Labeling Technique for High-Sensitivity miRNA Expression Analysis

Abstract Chemical labeling of miRNAs for electrochemical assay has become a useful approach for miRNA detection. It is believed that due to the extremely small size of miRNAs, direct labeling on miRNAs may be more advantageous. Recently, Gao and Yu from the Institute of Microelectronics (Singapore) presented a novel labeling procedure that utilizes a chemical ligation to directly label miRNA with a redox active and catalytic moiety (Biosens Bioelectron 22:933–940, 2007). The miRNA is labeled in the total RNA mixture in a one-step non-enzymatic reaction under very mild conditions with a redox active and electrocatalytic moiety, Ru (PD)2Cl2 (PD ¼ 1,10-phenanthroline-5,6-dione), through coordinative bonds with purine bases in the miRNA molecule. The excellent electrocatalytic activity of the Ru(PD)2Cl2 towards the oxidation of hydrazine makes it possible to conduct ultrasensitive miRNA detection. Under optimized experimental conditions, the assay allows the detection of miRNAs in the range of 0.50–400 pM with a detection limit of 0.20 pM in 2.5 microl (0.50 amole). miRNA quantitation is, therefore, performed in as little as 10 ng of total RNA, providing a handy platform for miRNA expression analysis. The amplification from the electrocatalytic oxidation of hydrazine greatly enhances the detectability of the assay, thereby lowering the detection limit to 0.2 pM. The electrochemical miRNA assay described here is rapid, ultrasensitive, non-radioactive, and is able to directly detect miRNA without involving biological ligation. The method allows us to identify miRNAs with less than twofold difference in expression levels under two conditions.

26.1

Introduction

Chemical labeling of miRNAs for electrochemical assay has become a useful approach for miRNA detection. It is believed that due to the extremely small size of miRNAs, direct labeling on miRNAs may be more advantageous. Recently, Babak and co-workers proposed a cisplatin-based chemical labeling procedure for miRNAs (Babak et al. 2004). One binding site of cisplatin is covalently bound to a Z. Wang and B. Yang, MicroRNA Expression Detection Methods, DOI 10.1007/978-3-642-04928-6_26, # Springer-Verlag Berlin Heidelberg 2010

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26 Electrocatalytic Moiety Labeling Technique

fluorophore and the other site is a labile nitrate ligand. Incubation in an aqueous solution with miRNAs at elevated temperatures results in a ligand exchange between the labile nitrate of the cisplatin complex and the more strongly coordinating purine moiety, forming a new complex between cisplatin and the N7 position of G base. The miRNA was, therefore, directly labeled with the cisplatin–fluorophore conjugate through a coordinative bond with G base in miRNA. Another direct labeling procedure at the 3’ end was recently developed by Liang et al. in which miRNAs were first tagged with biotin. After the introduction of quantum dots to the hybridized miRNAs through a reaction with quantum dots–avidin conjugates, the miRNAs were detected fluorescently with a dynamic range from 156 pM to 20 nM (Liang et al. 2005). Thomson’s group used T4 RNA ligase to couple the 3’ end of miRNA to a fluorophore-tagged nucleotide (Thomson et al. 2004). Overall, these direct ligation procedures do not offer the expected sensitivity for miRNA expression analysis. To further enhance the sensitivity and lower the detection limit, Gao and Yu proposed that a chemical or biological amplification scheme must be employed in the direct ligation procedures. It has been demonstrated that the sensitivity of the amplified electrochemical detection of nucleic acids is comparable to that of PCR-based fluorescent assays (Zhang et al. 2003; Xie et al. 2004a, b; Piunno and Krull 2005). Recently, Gao and Yu presented a novel labeling procedure that utilizes a chemical ligation to directly label miRNA with a redox active and catalytic moiety (Gao and Yu 2007). The miRNA is labeled in total RNA mixture in a one-step non-enzymatic reaction under very mild conditions with a redox active and electrocatalytic moiety, Ru(PD)2Cl2 (PD ¼ 1,10-phenanthroline-5,6-dione), through coordinative bonds with purine bases in the miRNA molecule. The excellent electrocatalytic activity of the Ru(PD)2Cl2 towards the oxidation of hydrazine makes it possible to conduct ultrasensitive miRNA detection. Under optimized experimental conditions, the assay allows the detection of miRNAs in the range of 0.50–400 pM, with a detection limit of 0.20 pM in 2.5 microl (0.50 amole). miRNA quantitation is, therefore, performed in as little as 10 ng of total RNA, providing a much-needed platform for miRNA expression analysis. The amplification from the electrocatalytic oxidation of hydrazine greatly enhances the detectability of the assay, thereby lowering the detection limit to 0.2 pM (Fig. 26.1).

26.2

Protocol

26.2.1 Materials 1. Indium tin oxide (ITO)-coated glass slides (Delta Technologies Limited, Stillwater, MN) 2. Montage spin column YM-50 column (Millipore Corporation)

26.2 Protocol

323

miRNA miRNA capture probes

+

ITO-coated glass slide

Immobilization

Hybridization

hydrazine

N2 Electrochemical Measurement to Detect miRNAs

Electrocatalysis

Fig. 26.1 Diagram illustrating the principle of electrocatalytic moiety labeling technique for highsensitivity miRNA expression analysis. Modified from Gao and Yu (2007)

3. Non-leak Ag/AgCl (3.0 M NaCl) reference electrode (Cypress Systems, Lawrence, KS) 4. Platinum wire counter electrode

26.2.2 Instruments 1. CH Instruments model 660A electrochemical workstation (CH Instruments, Austin, TX) 2. Conventional three-electrode system, consisting of an ITO working electrode, was used in all electrochemical measurements 3. Finnigan/MAT LCQ Mass Spectrometer (ThermoFinnigan, San Jose, CA) 4. Elan DRC II ICP-MS spectrometer (PerkinElmer, Wellesley, MA) 5. V-570 UV/VIS/NIR spectrophotometer (JASCO Corp., Japan) 6. UV–vis spectrophotometry 7. An environmental chamber

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26 Electrocatalytic Moiety Labeling Technique

26.2.3 Reagents 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14.

RuCl3 A phosphate buffered-saline (PBS, pH8.0): [0.15 M NaCl, 20 mM phosphate] Diethyl pyrocarbonate RNaseZap (Ambion, TX) 0.1 M pH6.0 acetate buffer 3 M KCl Trizol Reagent (Invitrogen, Carlsbad, CA) mirVana miRNA Isolation Kit (Ambion) Phenol/chloroform DNase I (Invitrogen) Absolute ethanol 0.1% SDS Sodium borohydride solution. 11-Aminoundecanoic acid (AUA, > 99%)

26.2.4 Procedures The protocols described in this section are essentially the same as reported in the study by Gao and Yu in 2007.

26.2.4.1

Synthesis of Ru(PD)2Cl2

Synthesize Ru(PD)2Cl2 (PD ¼ 1,10-phenanthroline-5,6-dione) according to the procedures described in literature (Goss and Abrun˜a 1985; Rivera et al 1994).

26.2.4.2

Total RNA Extraction and Labeling

1. Extract total RNA from cultured cells or tissues using TRIzol reagent according to the manufacturer’s recommended protocol 2. Enrich miRNAs in the total RNA using a Montage spin column YM-50 column 3. Determine the RNA concentration by UV–vis spectrophotometry. Typically, 1.0 mg of total RNA is used in each of the labeling reactions 4. Add 20 mL of 0.25 mM Ru(PD)2Cl2 in 0.1 M pH 6.0 acetate buffer to 5 mL of total RNA solution 5. Incubate the mixture for 30–40 min at 80 C and cool on ice 6. Add 5 mL of 3 M KCl into the labeled RNA sample and store at 20 C Since the labeling process is only effective to G and A bases, the label/base ratio is normally in the range of 1/3 to 1/4, depending on the sequence of

26.2 Protocol

325

individual miRNA molecule. Theoretically, if this ratio remains unchanged for all miRNAs, the same current sensitivity per base should be obtained for all miRNAs. At the same molar concentration, the sensitivity should be roughly proportional to the number of base in the miRNA, but this trend was not observed in our experiments. It is noteworthy that the sensitivity per base is dependent on miRNA sequence and (G þ A) content. However, no straightforward relation between (G þ A) content and current sensitivity was observed. This is probably due to the fact that G and A are not evenly distributed. Owing to the steric hindrance and three-dimensional packing of the miRNA molecules on the electrode surface, it would be impossible to label some of the G and A bases when they are in a cluster; hence, a low labeling efficiency is expected. For example, the (G þ A) content (78%) in miR-320 is more than doubled as compared to that of miR-92, but the sensitivity for miR-320 was merely 35% higher than that of miRr-92 (Gao and Yu 2007).

26.2.4.3

Design of miRNA Capture Oligonucleotide Probes

1. Obtain sequences for miRNA detection from the miRNA Registry (microrna. sanger.ac.uk) 2. Design miRNA capture oligonucleotide probes exactly antisense to the selected miRNAs 3. Synthesize and aldehyde-modify the probes by Invitrogen (Carlsbad, CA)

26.2.4.4

Electrode Preparation and Capture Probes Immobilization

1. Silanize the ITO electrodes with the bifunctional reagent 1,12-dodecanedicarboxylic acid (DDCA) to form a carboxylic acid-terminated monolayer (Hedges et al. 2004; Gao and Tansil 2005). 2. Denature aldehyde-modified capture probes for 10 min at 90 C and dilute to a concentration of 0.5 mM in 0.1 M acetate buffer (pH6.0). 3. Dispense a 25 mL aliquot of the capture probes solution onto the silanized electrode and incubate for 2–3 h at 20 C in an environmental chamber. 4. After incubation, rinse the electrode successively with 0.1% SDS and water. 5. Conduct reduction of the imines by a 5 min incubation of the ITO electrode in a 2.5 mg/ml sodium borohydride solution made of PBS/ethanol (3/1). 6. Soak the electrode in vigorously stirred hot water (9095 C) for 2 min, copiously rinse with water and blow dry with a stream of nitrogen. 7. Immerse the capture probe-coated electrode in an ethanolic solution of 2.0 mg/ ml AUA for 3–5 h. 8. Rinse off unreacted AUA molecules and wash the electrode by immersion in stirred ethanol for 10 min, followed by a thorough rinsing with ethanol and water. The surface density of the immobilized capture probes should be around (6.0–8.5) 1012 mol/cm2.

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26 Electrocatalytic Moiety Labeling Technique

The amount of capture probes immobilized on the electrode surface and hybridization efficiency determine the amount of target miRNA bound to the surface and thereby the amount of labels. However, in the model proposed by Gao and Yu (2007), multiple Ru(PD)2Clþ labels on a single miRNA strand greatly increases the label loading, proportionally increasing the response from electrocatalytic oxidation, and hence the sensitivity and detection limit of the miRNA assay is substantially improved.

26.2.4.5

miRNA Assay

1. Place the treated electrode in a moisture saturated environmental chamber maintained at 30 C 2. Uniformly spread a 2.5 mL aliquot of hybridization solution, containing the desired amount of labeled miRNA, onto the electrode 3. Rinse the electrode thoroughly with a blank hybridization solution at 30 C after a 60-min hybridization period 4. Measure the hydrazine electrooxidation current amperometrically at 0.1 V in vigorously stirred PBS containing 5 mM hydrazine 5. At low miRNA concentrations, smoothing need to be applied after each amperometric measurement to remove random noise and electromagnetic interference 6. Electrochemical experiments should be carried out using a CH Instruments model 660A electrochemical workstation 7. Utilize a conventional three-electrode system (consisting of an ITO working electrode, a non-leak Ag/AgCl (3 M NaCl) reference electrode, and a platinum wire counter electrode) in all electrochemical measurements. All potentials should be referred to the Ag/AgCl electrode 8. Perform electrospray ionization mass spectrometric (ESI-MS) experiments with a Finnigan/MAT LCQ Mass Spectrometer 9. Conduct inductively coupled plasma mass spectrometry (ICP–MS) with an Elan DRC II ICP–MS spectrometer 10. Record UV–vis spectra on a V-570 UV/VIS/NIR spectrophotometer 11. Carry out all experiments at room temperature, unless otherwise stated

26.2.4.6

Calibration Curves for miRNAs

1. For quantitative detection of miRNAs, it is necessary to establish a standard curve for each of the miRNAs in question. Synthesize miRNAs using the services provided by companies. 2. Solutions of different concentrations of labeled miRNAs, ranging from 0.10 to 1,000 pM, should be tested, following the procedures described above. 3. For the control experiments, non-complementary capture probes were used in the electrode preparation (Fig. 26.2).

26.3 Application and Limitation

327

Silanize the ITO electrodes Select a miRNA of interest Preparation of tissue or cells total RNA samples

Design and synthesize RNA labelling with Ru(PD)2Cl2 of miRNA capture probes

Coat electrodes with miRNA capture probes

Immobilize electrodes on indium tin oxide (ITO) coated glass slides

Hybridization of labelled miRNAs with miRNA capture probes

Electrooxidation with hydrazine

Electrocatalysis

Electrochemical measurements

Fig. 26.2 Flowchart of the electrocatalytic moiety labeling technique for miRNA expression detection. According to Gao and Yu (2007)

26.3

Application and Limitation

The Electrocatalytic Moiety Labeling technique for miRNA detection possesses several advantages. 1. The electrochemical miRNA assay described here is rapid, ultrasensitive, nonradioactive and is able to directly detect miRNA without involving biological ligation. 2. By employing Ru(PD)2Cl2, miRNA is directly labeled with redox and electrocatalytic moieties under very mild conditions. 3. Specific miRNAs are detected amperometrically at sub-picomolar levels with high specificity. This electrochemical miRNA assay is easily extendable to a

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26 Electrocatalytic Moiety Labeling Technique

low-density array of 50–100 electrodes. The relatively limited number of miRNA offers excellent opportunity for low-density electrochemical arrays in miRNA assays. The advantages of low-density electrochemical biosensor arrays are: (a) more cost-effective than optical biosensor arrays; (b) ultrasensitive when coupled with catalysis; (c) rapid, direct, turbid, and light absorbing-tolerant detection; and (d) portable, robust, low cost, and easy-to-handle electrical components suitable for field tests and homecare use. Such a tool would be of great scientific value and may open the door to routine miRNA expression profiling and molecular diagnostics. 4. The method allows us to identify miRNAs with less than two-fold difference in expression levels under two conditions. The major drawbacks of this technique are the requirement of many instruments and chemical synthesis of reagents, which are not frequently seen in a regular molecular biology laboratory. And since the labeling process is only effective to G and A bases, the labeling intensity of a miRNA will depend on the GA content and the clustering of the GAs owing to the steric hindrance and three-dimensional packing of the miRNA molecules on the electrode surface. This property makes quantitative analysis of miRNAs difficult.

References Babak T, Zhang W, Morris Q, Blencowe BJ, Hughes TR (2004) Probing microRNAs with microarrays: tissue specificity and functional inference. RNA 10:1813–1819 Gao Z, Tansil NC (2005) An ultrasensitive photoelectrochemical nucleic acid biosensor. Nucleic Acids Res 33:e123 Gao Z, Yu YH (2007) Direct labeling microRNA with an electrocatalytic moiety and its application in ultrasensitive microRNA assays. Biosens Bioelectron 22:933–940 Goss CA, Abrun˜a HD (1985) Inorg Chem 24:4263 Hedges DH, Richardson DJ, Russell DA (2004) Electrochemical control of protein monolayers at indium tin oxide surfaces for the reagentless optical biosensing of nitric oxide. Langmuir 20:1901–1908 Liang RQ, Li W, Li Y, Tan CY, Li JX, Jin YX, Ruan KC (2005) An oligonucleotide microarray for microRNA expression analysis based on labeling RNA with quantum dot and nanogold probe. Nucleic Acids Res 33:e17 Piunno PA, Krull UJ (2005) Trends in the development of nucleic acid biosensors for medical diagnostics. Anal Bioanal Chem 381:1004–1011 Rivera N, Colo´n Y, Guadalupe AR (1994) Bioelectrochem Bioenerg 34:169 Thomson JM, Parker J, Perou CM, Hammond SM (2004) A custom microarray platform for analysis of microRNA gene expression. Nat Methods 1:47–53 Xie H, Yu YH, Xie F, Lao YZ, Gao Z (2004a) A nucleic acid biosensor for gene expression analysis in nanograms of mRNA. Anal Chem 76:4023–4029 Xie H, Zhang C, Gao Z (2004b) Amperometric detection of nucleic acid at femtomolar levels with a nucleic acid/electrochemical activator bilayer on gold electrode. Anal Chem 76:1611–1617 Zhang Y, Kim HH, Heller A (2003) Enzyme-amplified amperometric detection of 3000 copies of DNA in a 10-microL droplet at 0.5 fM concentration. Anal Chem 75:3267–3269

Part IX

Circulating miRNA Detection Methods

Chapter 27

Serum and Plasma miRNA Detection

Abstract Blood-based miRNA detection is not a method but an approach. This approach relies on the facts that (1) miRNAs are present in the serum and plasma of humans and animals such as mice, rats, bovine fetuses, calves, and horses; (2) miRNAs are present in human plasma in a remarkably stable form that is protected from endogenous RNase activity; and (3) the levels of miRNAs in serum are reproducible and consistent among individuals of the same species. The bloodbased miRNA detection enables the development of miRNAs as novel biomarkers for diagnosis and prognosis of human diseases and of noninvasive diagnostic and prognostic approaches as practical tools in clinical use. In theory, most of the miRNA detection methods described above can be used to detect serum and plasma miRNAs. The feasibility of blood-based miRNA detection has recently been verified by several groups with the studies documented by Chen et al. (Cell Res 18:997–1006, 2008) and Mitchell et al. (Proc Natl Acad Sci USA 105:10513– 10518, 2008) representing the first successful examples of such efforts. These studies clearly indicate the value of blood-based miRNA detection as an invaluable approach for identifying aberrantly expressed miRNAs under specific pathological conditions and for realizing these miRNAs as novel biomarkers for diagnosis and prognosis of human diseases.

27.1

Introduction

Besides being recognized as key molecules in intracellular regulatory networks for gene expression, the spectra and levels of some miRNAs are emerging as biomarkers for various pathological conditions (Waldman and Terzic 2008; Lee and Dutta 2008; Dillhoff et al. 2008). Recent findings suggest that circulating miRNAs may be plasma biomarkers for the diagnosis of the lung (Chen et al. 2008), colorectal (Chen et al. 2008), and prostate cancers (Mitchell et al. 2008). Indeed, miRNAs are present in the serum and plasma of humans and animals such as mice, rats, bovine fetuses, calves, and horses (Ai et al. 2009; Chen et al. 2008; Hunter et al. 2008; Mitchell et al. 2008; Chim et al. 2008; Gilad et al. 2008; Lawrie et al. 2008; Z. Wang and B. Yang, MicroRNA Expression Detection Methods, DOI 10.1007/978-3-642-04928-6_27, # Springer-Verlag Berlin Heidelberg 2010

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Resnick et al. 2009; Taylor and Gercel-Taylor 2008; Wang et al. 2009). miRNAs are present in human plasma in a remarkably stable form that is protected from endogenous RNase activity. The levels of miRNAs in serum are reproducible and consistent among individuals of the same species. Strikingly, Gilad et al. (2008) confirmed that miRNAs are also detectable in other body fluids, such as urine, saliva, amniotic fluid, and pleural fluid. Of note, serum and urine display different miRNA abundance profiles as might be expected for two dissimilar biological fluids, further supporting the hypothesis that bodily fluid microRNA profiles reflect physiology. Challenges for developing protein-based biomarkers from body fluids, such as plasma, serum, and urine, include the complexity of protein composition, the assorted posttranslational modifications, the low abundance of proteins of interest, the difficulty of developing suitable high-affinity capture agents, the complexities of proteolysis and protein denaturation, and potentially complex assay methods (Cowan and Vera 2008; Ebert et al. 2006). All of these make the discovery and development of protein-based biomarkers with proper specificity, sensitivity, and predictive value an expensive, time consuming, and difficult task. The biological function of circulating miRNA is largely unknown; however, unlike proteins, there are far fewer known miRNA species, so obtaining a complete profile is relatively easy. Currently, there are 866 known miRNAs for human and 627 for mouse (based on the latest miRBase release 12.0 http://microrna.sanger.ac.uk/sequences/index. shtml) compared with perhaps a million or more serum proteins, including various processing variants and posttranslationally modified proteins. mRNA must be translated into protein to have a biological effect whereas miRNAs are themselves the active moiety, often influencing the expression of multiple other genes, and thus likely reflect altered physiology more directly. In addition, miRNAs do not have known post-processing modifications, and with their size, their chemical composition is much less complex than most other biological molecules. Detecting specific miRNA species, although somewhat challenging, is inherently a much easier task than detecting proteins. A synthetic complementary oligonucleotide should deliver sufficient specificity in most cases, and a standard PCR assay can be used to increase the detection sensitivity. It has also been demonstrated that the circulating miRNAs are stable and can be reliably extracted and assayed in either serum or plasma (Mitchell et al. 2008). Both real-time RT-PCR and microarray methods have been applied to detect blood miRNAs. Theoretically speaking, most of the methodologies described in earlier chapters can be applied for miRNA expression profiling and detection.

27.2

Protocol

27.2.1 Materials 1. 18 gage needle 2. 2 mL syringe

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3. 2 mL microfuge tubes 4. TaqMan Array Human MicroRNA Panel (v.1, Applied Biosystems, Foster City, CA)

27.2.2 Instruments 1. Surgical blade 2. Centrifuge 3. Nanodrop instrument (ND-1000 Spectrophotometer, NanoDrop Technologies) or an equivalent 4. Agilent1 2100 bioanalyzer (Agilent Technologies) or an equivalent 5. A Real-Time PCR System 6. ABI Prism 7900HT Sequence detection system (Applied Biosystems) 7. ThermoScientific NanoDrop1000 (Thermo Fisher Scientific, Inc., Waltham, MA) or an equivalent

27.2.3 Reagents 1. RNAlater1 Tissue Collection:RNA Stabilization Solution (Ambion, Austin TX) 2. Trizol Reagent (Invitrogen, Carlsbad, CA) 3. mirVana miRNA Isolation Kit (Ambion) 4. Ambion Mouse RiboPureTM -Blood RNA Isolation kit (AB cat #AM1951) 5. Phenol/chloroform 6. DNase I (Invitrogen) 7. Absolute ethanol 8. TaqMan1 MicroRNA Assays (Applied Biosystems) 9. AB TaqMan1 microRNA Reverse Transcription Kit (AB cat #4366597) 10. DMSO (Invitrogen) 11. TaqMan Universal PCR Master Mix (Applied Biosystems) 12. Tri-Reagent BD (Molecular Research Center, Inc., Cincinnati, OH)

27.2.4 Procedures The protocols described in this section are essentially the same as reported in the studies by Chen et al. (2008) and by Mitchell et al. (2008).

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27.2.4.1

27 Serum and Plasma miRNA Detection

Blood Collection

Mouse Blood 1. To perform cardiac puncture, euthanize mice using carbon dioxide. Then access the ventricle with an 18 gage needle and aspirate 400–500 mL blood into a 2 mL syringe. Discharge the blood immediately into a 2 mL microfuge tube preloaded with 1.3 mL RNAlater1 Tissue Collection:RNA Stabilization Solution, mix by inversion, and store at 20 C. 2. To perform blood collection by tail vein, place mice under a heat lamp for 5 min, then make a small peripheral tail incision. Collect 2–10 drops of blood directly into a 2 mL microfuge tube preloaded with 1.3 ml RNALater1 Solution, mix by inversion, and store at 20 C. An average drop of blood approximates 24 mL (Fan et al. 2008).

Human Blood 1. Obtain written consent and ethics permission from blood donors 2. Separate whole blood into serum and cellular fractions within 2 h after blood has been derived, by clotting blood sample 3. Freeze cellular fractions immediately in liquid nitrogen and store sera at 80 C (Chen et al. 2008)

27.2.4.2

Separation of Plasma and Blood Cells

1. To harvest cell-free plasma, centrifuge blood samples twice at 4 C: first at 1,600 g for 10 min and collect the supernatant, followed by a second centrifuge of the supernatant at 16,000 g for 10 min to remove blood cells. 2. To harvest blood cells (including leukocytes and erythrocytes), centrifuge the blood cells obtained in the first centrifugation at 2,300 g for 5 min to remove residual plasma (Chim et al. 2008).

27.2.4.3

RNA Extraction

Whole Blood RNA 1. The Ambion Mouse RiboPureTM -Blood RNA Isolation kit can be used for extraction of RNA. Centrifuge samples and remove the RNAlater1 Solution prior to disruption of the blood pellet in a guanidinium-based lysis solution, followed by organic extraction and purification of the total RNA fraction (including small RNA) by solid phase extraction onto a silica matrix.

27.2 Protocol

335

2. Use the Alternative Protocol described in the kit instruction manual for samples less than 250 ml from the mouse tail vein. 3. Determine RNA yields by UV absorbance using a Nanodrop instrument and examine the intactness on an Agilent1 2100 bioanalyzer. 4. Dilute the cardiac puncture samples 1:10 before running (Fan et al. 2008).

Serum RNA 1. Extract RNA from 250 mL of serum using the Tri-Reagent BD as described by the manufacturer 2. Or use Trizol Reagent and add three steps of phenol/chloroform purification since serum is full of proteins. In general, the yield is expected to be around 5–10 mg RNA/50 mL serum (Chen et al. 2008) 3. Or use the mirVana miRNA Isolation Kit (Ambion) 4. To minimize DNA contamination, treat the eluted RNA preparation with DNase I 5. Assess RNA quality with the ThermoScientific NanoDrop1000 (Resnick et al. 2009)

Blood Cell RNA 1. Isolate blood cell total RNA using Trizol, according to the manufacturer’s instructions.

27.2.4.4

miRNA Profiling and Quantification

1. Reverse transcription reactions are carried out for 65 min using the AB TaqMan1 microRNA Reverse Transcription Kit which includes M-MLV reverse transcriptase. 2. Conduct miRNA analysis using the mirVana qRT-PCR primer sets or the TaqMan1 MicroRNA Assays. The mirVana qRT-PCR assays use 10 ng of input total RNA that is analyzed using target-specific primers for reverse transcription with M-MLV reverse transcriptase, followed by PCR amplification with a pair of miRNA target-specific primers and detection with SYBR1 Green I nucleic acid gel stain 10,000 concentrate in DMSO. Melting curve analysis is carried out for each target to assess amplification specificity; for some targets, nonspecific amplification is observed in the no-template negative controls, which cannot be discriminated by melt-curve analysis. 3. The TaqMan MicroRNA Assays use 10 ng of input total RNA with miRNAtarget-specific reverse transcription primers and target-specific internal hybridization probes (“TaqMan probes”), and are run in 96-well or 384-well formats. qRT-PCR assays of similar design need to be carried out for constitutively

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expressed small RNAs of similar size to miRNAs (e.g., snoRNAs) and used for normalization of input RNA amount (analogous to use of constitutive mRNAs such as GAPDH for normalization of protein-coding genes). 4. Amplification reactions consist of a hold of 10 min at 95 C and 40 cycles (15 s/ 95 C, 60 s/60 C) on an Applied Biosystems 7900HT Real-Time PCR System and require about 1.5 h to complete. The assays should be carried out in duplicate or triplicate and the geometric average Ct value is used to calculate relative expression for each data point. Within each experiment, the endogenous control that has the highest Ct should be set as the baseline, and the Ct between the baseline and the Ct of the small RNA control in each sample should be used as a normalization factor that is added to the raw Ct for each sample. Normalized Ct values larger than 35 were reported as 35 (Fan et al. 2008) (Fig. 27.1).

Collect blood sample from mice by cardiac puncture or via tail vein

Collect blood sample from human donors

Two centrifuges to separate plasma and blood cells in the presence of anti-coagulant

Collect supernatant as plasma

Let blood sample coagulate without anti-coagulant

Centrifuge again

Remove supernatant to collect blood cells

Collect fluid as serum

Isolate total RNA from various blood components using Tri-Reagent BD

Real-time RT-PCR to detect circulating miRNAs

Fig. 27.1 Flowchart of the use of blood samples for miRNA expression detection. According to Chen et al. (2008), Mitchell et al. (2008)

References

27.3

337

Application and Limitation

The diagnostic and prognostic utility of circulating RNAs in both benign and malignant conditions has recently been revealed. Placental-associated circulating miRNAs correlate with pregnancy progression (Chim et al. 2008). In malignant states, circulating mRNAs in renal cell carcinoma patients (Feng et al. 2008) as well as miRNAs from the serum of patients with diffuse large B cell lymphoma (Lawrie et al. 2008) have been shown to be stable and highly predictive of malignancy as well as survival. miRNAs originating from human prostate cancer xenografts enter the circulation, are readily measured in plasma, and can robustly distinguish xenografted mice from controls. This concept extends to cancer in humans, where serum levels of miR-141 (a miRNA expressed in prostate cancer) can distinguish patients with prostate cancer from healthy controls (Mitchell et al. 2008). It has been demonstrated that the miRNA signature of circulating tumor exosomes of ovarian cancer patients demonstrates high correlation with miRNA expression of the primary tumor (Taylor and Gercel-Taylor 2008). Resnick et al. (2009) described miRNA extraction from the serum of ovarian cancer patients, the differential expression of a number of these miRNAs between patients and healthy controls as well as a novel real-time PCR microarray detection method (Resnick et al. 2009). Wang et al. (2009) recently established specific circulating microRNAs as sensitive and informative biomarkers for drug-induced liver injury. More recently, Ai et al. (2009) described a study designed to establish circulating miR-1 as a novel biomarker for acute myocardial infarction. miR-1 level was found significantly higher in both whole blood and plasma from patients with acute myocardial infarction relative to healthy subjects and the level was dropped to normal on discharge following medication. Increased circulating miR-1 was not associated with age, gender, blood pressure as well as diabetes mellitus, or established biomarkers. However, miR-1 level was well correlated with the width of QRS complex in ECG, consistent with the ability of miR-1 to slow cardiac conduction and promotes arrhythmogenesis (Yang et al. 2007, 2008; Wang et al. 2008). Use of blood samples (serum, plasma, or blood cells) for miRNA profiling and detection holds a promising future for using circulating miRNAs as biomarkers for diagnosis and prognaosis of human diseases. With the recent rapid development in this field, the age of miRNA-based biomarkers is soon to come.

References Ai J, Zhang R, Pu J, Li Y, Lu Y, Jiao J, Li K, Yu B, Li Z, Wang R, Wang L, Li Q, Wang N, Shan H, Yang B (2009) Circulating microRNA-1 as a novel biomarker for acute myocardial infarction. Cardiovasc Res (in revision) Chen X, Ba Y, Ma L, Cai X, Yin Y, Wang K, Guo J, Zhang Y, Chen J, Guo X, Li Q, Li X, Wang W, Zhang Y, Wang J, Jiang X, Xiang Y, Xu C, Zheng P, Zhang J, Li R, Zhang H, Shang X, Gong T, Ning G, Wang J, Zen K, Zhang J, Zhang CY (2008) Characterization of

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microRNAs in serum: a novel class of biomarkers for diagnosis of cancer and other diseases. Cell Res 18:997–1006 Chim SS, Shing TK, Hung EC, Leung TY, Lau TK, Chiu RW, Lo YM (2008) Detection and characterization of placental microRNAs in maternal plasma. Clin Chem 54:482–490 Cowan ML, Vera J (2008) Proteomics: Advances in biomarker discovery. Exp Rev Proteomics 5:21–23 Dillhoff M, Wojcik SE, Bloomston M (2008) MicroRNAs in solid tumors. J Surg Res, in press Ebert MP, Korc M, Malfertheiner P, Rocken C (2006) Advances, challenges, and limitations in serum-proteome-based cancer diagnosis. J Proteome Res 5:19–25 Fan AC, Goldrick MM, Ho J, Liang Y, Bachireddy P, Felsher DW (2008) A quantitative PCR method to detect blood microRNAs associated with tumorigenesis in transgenic mice. Mol Cancer 7:74 Feng G, Li G, Gentil-Perret A, Tostain J, Genin C (2008) Elevated serumcirculating RNA in patients with conventional renal cell cancer. Anticancer Res 28:321–326 Gilad S, Meiri E, Yogev Y, Benjamin S, Lebanony D, Yerushalmi N, Benjamin H, Kushnir M, Cholakh H, Melamed N, Bentwich Z, Hod M, Goren Y, Chajut A (2008) Serum microRNAs are promising novel biomarkers. PLoS ONE 3:e3148 Hunter MP, Ismail N, Zhang X, Aguda BD, Lee EJ, Yu L, Xiao T, Schafer J, Lee ML, Schmittgen TD, Nana-Sinkam SP, Jarjoura D, Marsh CB (2008) Detection of microRNA expression in human peripheral blood microvesicles. PLoS ONE 3:e3694 Lawrie CH, Gal S, Dunlop HM, Pushkaran B, Liggins AP, Pulford K, Banham AH, Pezzella F, Boultwood J, Wainscoat JS, Hatton CS, Harris AL (2008) Detection of elevated levels of tumourassociated microRNAs in serum of patients with diffuse large B-cell lymphoma. Br J Haematol 141:672–675 Lee YS, Dutta A (2008) MicroRNAs in cancer. Annu Rev Pathol 4:199–227 Mitchell PS, Parkin RK, Kroh EM, Fritz BR, Wyman SK, Pogosova-Agadjanyan EL, Peterson A, Noteboom J, O’Briant KC, Allen A, Lin DW, Urban N, Drescher CW, Knudsen BS, Stirewalt DL, Gentleman R, Vessella RL, Nelson PS, Martin DB, Tewari M (2008) Circulating microRNAs as stable blood-based markers for cancer detection. Proc Natl Acad Sci USA 105:10513–10518 Resnick KE, Alder H, Hagan JP, Richardson DL, Croce CM, Cohn DE (2009) The detection of differentially expressed microRNAs from the serum of ovarian cancer patients using a novel real-time PCR platform. Gynecol Oncol 112:55–59 Taylor DD, Gercel-Taylor C (2008) MicroRNA signatures of tumor-derived exosomes as diagnostic biomarkers of ovarian cancer. Gynecol Oncol 110:13–21 Waldman SA, Terzic A (2008) MicroRNA signatures as diagnostic and therapeutic targets. Clin Chem 54:943–944 Wang K, Zhang S, Marzolf B, Troisch P, Brightman A, Hu Z, Hood LE, Galas DJ (2009) Circulating microRNAs, potential biomarkers for drug-induced liver injury. Proc Natl Acad Sci USA 106:4402–4407 Wang Z, Luo X, Lu Y, Yang B (2008) miRNAs at the heart of the matter. J Mol Med 86:771–783 Yang B, Lin H, Xiao J, Lu Y, Luo X, Li B, Zhang Y, Xu C, Bai Y, Wang H, Chen G, Wang Z (2007) The muscle-specific microRNA miR-1 causes cardiac arrhythmias by targeting GJA1 and KCNJ2 genes. Nat Med 13:486–491 Yang B, Lu Y, Wang Z (2008) Control of cardiac excitability by microRNAs. Cardiovasc Res 79:571–580

Chapter 28

miRNA Detection from Peripheral Blood Microvesicles

Abstract Microvesicles are small exosomes/vesicles of endocytic origin released by normal healthy or damaged cell types or activated platelets. These cell particles are enriched in bioactive molecules and contain nucleic acid and/or protein, playing a role in growth, differentiation, cancer progression, and cell signal transduction. Changes of microvesicles concentration have been noticed under several pathological conditions. The miRNAs expressed in the microvesicles from the blood may play a role in homeostasis, and the miRNA expression profiles may also reflect certain pathophysiological conditions. The first efforts to measure miRNA expression in microvesicles were made by Valadi and colleagues (Nat Cell Biol 9:654–659, 2007) from the Department of Internal Medicine and Department of Respiratory Medicine and Allergology, the Sahlgrenska Academy, Go¨teborg University (Sweden) and by Hunter et al. (PLoS ONE 3:e3694, 2008) from the Division of Pulmonary, Allergy, Critical Care, Sleep Medicine, College of Medicine, The Ohio State University (Columbus, OH, USA).

28.1

Introduction

It has recently been recognized that microvesicles play an important role in genetic exchange of mRNA and miRNA between cells (Valadi et al. 2007). Microvesicles are small exosomes/vesicles (from 50 nm to 1 mm) of endocytic origin released by normal healthy or damaged cell types or activated platelets (Wieckowski and Whiteside 2006; Ratajczak et al. 2006b; Valenti et al. 2007). They can be considered as shed from the plasma membrane into the extracellular environment to facilitate communication between cells. These cell particles also play a role in growth, differentiation, and cancer progression (Ratajczak et al. 2006a). Microvesicles are enriched in bioactive molecules and contain nucleic acid and/or protein. Concentration of microvesicle in the peripheral blood of healthy individuals is estimated to be in the range of 5–50 mg/mL. In the peripheral blood, two-thirds of microvesicles are derived from platelets; a small percentage of microvesicles are derived Z. Wang and B. Yang, MicroRNA Expression Detection Methods, DOI 10.1007/978-3-642-04928-6_28, # Springer-Verlag Berlin Heidelberg 2010

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from endothelial cells (Hunter et al. 2008). Elevation of endothelial-derived microvesicles has been reported in patients with pulmonary arterial hypertension (Bakouboula et al. 2008). Platelet-derived microvesicles play a role in angiogenesis and the metastatic spread of cancers, such as lung cancer (Janowska-Wieczorek et al. 2005), and in immune response upon the regulation of gene expression in hematopoietic, endothelial, and monocytic cells (Setzer et al. 2006; Majka et al. 2007). Notably, platelet-derived microvesicle subpopulations are increased in patients with sepsis (Janiszewski et al. 2004; Meziani et al. 2008), whereas patients with pulmonary arterial hypertension have increased endothelial-derived microvesicles (Bakouboula et al. 2008). Intriguingly, Valadi and colleagues reported that vesicles released from human and murine mast cell lines contain over 1,200 mRNA and ~121 miRNA molecules (Valadi et al. 2007). Hunter et al. (2008) defined miRNA expression profile in circulating plasma microvesicles of normal human subjects, providing a basis for future studies to determine the predictive role of peripheral blood miRNA signatures in human diseases. Hierarchical clustering of the data sets indicated significant differences in miRNA expression between peripheral blood mononuclear cells (PBMC) and plasma microvesicles, with 33 and 4 miRNAs significantly differentially expressed in the plasma microvesicles and mononuclear cells, respectively. The majority of the miRNAs expressed in the microvesicles from the blood were predicted to regulate cellular differentiation of blood cells and metabolic pathways and a few miRNAs were predicted to be important modulators of immune function (Hunter et al. 2008). The detection of tissue specific miRNAs and microvesicles in the peripheral blood may become a frequent event upon tissue damage.

28.2

Protocol

28.2.1 Materials 1. 2. 3. 4. 5.

19-gage needle Syringe (50 cc) EDTA tubes Ficoll-hypaque (d = 1.077) (Mediatech, Inc.) 2 micron bead standards (BD Biosciences)

28.2.2 Instruments 1. Agilent 2100 Bioanalyzer (Agilent Technologies, Inc, Santa Clara, CA) or an equivalent 2. Applied Biosystems 7900HT real-time PCR instrument or an equivalent

28.2 Protocol

341

3. Liquid-handling robots 4. Zymak Twister robot 5. BD Aria flow cytometer (BD Biosciences)

28.2.3 Reagents 1. 2. 3. 4. 5.

Sterile low endotoxin PBS (Mediatech, Inc. Manassas, VA) PBS 3.8% sodium citrate tubes 1 mM PGE1 (Sigma-Aldrich, St. Louis, MO) Tyrodes buffer: [138 mM NaCl, 2.9 mM KCl, 12 mM NaHCO3, 0.36 mM NaHPO4, 5.5 mM glucose, 1.8 mM CaCl2, and 0.49 mM MgCl2, pH 6.5 or pH7.4] 6. Trizol (Invitrogen, Carlsbad, CA) 7. Stem-looped primers (Mega Plex kit, Applied Biosystems, Foster City, CA)

28.2.4 Procedures The protocols described in this section are essentially the same as reported in the study by Hunter et al. (2008).

28.2.4.1

Blood Collection

1. Collect peripheral blood: discard the first 2 cc, then draw 40 cc of blood through a 19-gage needle in EDTA tubes from diseased and healthy subjects following informed consent, between morning and early afternoon. 2. Dilute the peripheral blood 1:1 with sterile low endotoxin PBS, layer over ficollhypaque (d = 1.077), and centrifuge at 1,000 g. 3. Wash the mononuclear cell fraction once in PBS.

28.2.4.2

Microvesicle Isolation

1. To purify the microvesicles from the plasma, concentrate the vesicles by centrifugation at 160,000 g for 1 hr at 4 C (Nieuwland et al. 2000). 2. To isolate platelets, collect blood from donors in 3.8% sodium citrate tubes in 1:9 volume ratio. 3. Following centrifugation of the blood at 1000 g for 15 min at room temperature, incubate the platelet rich plasma with 1 mM PGE1.

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4. Centrifuge again and wash the platelets twice by resuspending the platelet pellet in Tyrodes buffer (pH6.5) and centrifuged again. 5. Then wash the platelets one additional time in Tyrodes buffer (pH7.4) (Hunter et al. 2008; Walkowiak et al. 2000).

28.2.4.3

RNA Extraction

1. Isolate total RNA by Trizol extraction method 2. To increase the yield of small RNAs, precipitate the RNA overnight 3. Determine RNA concentration and RNA integrity by capillary electrophoresis on an Agilent 2100 Bioanalyzer or a Nanodrop 4. For RNA isolated from mononuclear cells, only a RNA integrity number (RIN) 9 should be used along with its matched plasma sample for profiling

28.2.4.4

miRNA Profiling by Quantive PCR

1. Perform real-time PCR using an Applied Biosystems 7900HT real-time PCR instrument (or an equivalent) equipped with a 384 well reaction plate. 2. Convert RNA (500 ng) to cDNA by priming with a mixture of looped primers using previously published reverse transcription conditions as described in Section V. 3. Primers to the internal controls, small nucleolar (sno)RNA U38B, snoRNA U43, small nuclear (sn)RNA U6 as well as 18 S and 5 S rRNA should be included in the mix of primers. 4. Use Liquid-handling robots and the Zymak Twister robot to increase throughput and reduce error.

28.2.4.5

Flow Cytometry

1. Directly immunostain peripheral blood microvesicles from plasma without concentration by centrifugation 2. To confirm that the microvesicles were the correct size, set flow cytometry gates using 2 micron bead standards 3. Analyze the samples on BD Aria flow cytometer or an equivalent. Data can be expressed as percent of gated events

28.2.4.6

Other Analyses

1. Since Ct scores of real-time RT-PCR >35 are considered non-specific (Schmittgen et al. 2008), miRNAs in which 80% of individual observations have a raw Ct

References

343

score >35 should not be included in the final data analysis. However, based on our experience, Ct value >30 can be considered as null expression. 2. To reduce the bias caused by the use of an arbitrary miRNA as a normalization correction factor, the miRNAs should be compared between plasma microvesicles and PBMC based on their relative expression to the overall miRNA expression on each array using median normalization analysis (Wang et al. 2007). 3. To test the difference of specific miRNA between plasma microvesicles and PBMC, linear mixed models should be used and p-values should be generated from the model based on the estimated difference and sample variation. miRNAs should be subjected to hierarchical clustering using Euclidean distance based on their relative mean expression. miRNAs should also be ranked based on their raw Ct score for each plasma microvesicles and PBMC (Hunter et al. 2008).

28.3

Application and Limitation

In the study reported by Hunter et al. (2008), the miRNAs circulating in plasma microvesicles, platelets, and PBMC of normal human volunteers in the peripheral blood were detected and analyzed. They characterized peripheral blood miRNA patterns in healthy humans, and found significant differences in miRNA expression between plasma microvesicles, platelets, and PBMC. The data indicate that the miRNAs are also contained in plasma microvesicles, and these microvesiclecontaining miRNAs on target cell mRNA expression in their target cells influence homeostasis. The detection of tumors exosomes (microvesicles) in the peripheral blood has been found to contain miRNAs (Taylor and Gercel-Taylor 2008). It has been found that the internal controls (18 S, 5 S, snoRNA U38B, snoRNA U43, and snRNA U6) were highly variable in the plasma microvesicles and were significantly different in plasma microvesicles versus PBMC (Hunter et al. 2008). Careful selection of reasonable controls therefore becomes a significant factor influencing the quantification of microvesicle-containing miRNAs.

References Bakouboula B, Morel O, Faure A, Zobairi F, Jesel L, Trinh A, Zupan M, Canuet M, Grunebaum L, Brunette A, Desprez D, Chabot F, Weitzenblum E, Freyssinet JM, Chaouat A, Toti F (2008) Procoagulant membrane microparticles correlate with the severity of pulmonary arterial hypertension. Am J Respir Crit Care Med 177: 536–543 Hunter MP, Ismail N, Zhang X, Aguda BD, Lee EJ, Yu L, Xiao T, Schafer J, Lee ML, Schmittgen TD, Nana-Sinkam SP, Jarjoura D, Marsh CB (2008) Detection of microRNA expression in human peripheral blood microvesicles. PLoS ONE 3:e3694

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Janiszewski M, Do Carmo AO, Pedro MA, Silva E, Knobel E, Laurindo FR (2004) Plateletderived exosomes of septic individuals possess proapoptotic NAD(P)H oxidase activity: A novel vascular redox pathway. Crit Care Med 32:818–825 Janowska-Wieczorek A, Wysoczynski M, Kijowski J, Marquez-Curtis L, Machalinski B, Ratajczak J, Ratajczak MZ (2005) Microvesicles derived from activated platelets induce metastasis and angiogenesis in lung cancer. Int J Cancer 113:752–760 Majka M, Kijowski J, Lesko E, Goz´dizk J, Zupanska B, Ratajczak MZ (2007) Evidence that platelet-derived microvesicles may transfer platelet-specific immunoreactive antigens to the surface of endothelial cells and CD34+ hematopoietic stem/progenitor cells–implication for the pathogenesis of immune thrombocytopenias. Folia Histochem Cytobiol 45:27–32 Meziani F, Tesse A, Andriantsitohaina R (2008) Microparticles are vectors of paradoxical information in vascular cells including the endothelium: role in health and diseases. Pharmacol Rep 60:75–84 Nieuwland R, Berckmans RJ, McGregor S, Boing AN, Romijn FP, Westendorp RG, Hack CE, Sturk A (2000) Cellular origin and procoagulant properties of microparticles in meningococcal sepsis. Blood 95:930–935 Ratajczak J, Miekus K, Kucia M, Zhang J, Reca R, Dvorak P, Ratajczak MZ (2006a) Embryonic stem cell-derived microvesicles reprogram hematopoietic progenitors: evidence for horizontal transfer of mRNA and protein delivery. Leukemia 20:847–856 Ratajczak J, Wysoczynski M, Hayek F, Janowska-Wieczorek A, Ratajczak MZ (2006b) Membrane-derived microvesicles: important and underappreciated mediators of cell-to-cell communication. Leukemia 20: 1487–1495 Schmittgen TD, Lee EJ, Jiang J, Sarkar A, Yang L, Elton TS, Chen C (2008) Real-time PCR quantification of precursor and mature microRNA. Methods 44:31–38 Setzer F, Oberle V, Bla¨ss M, Mo¨ller E, Russwurm S, Deigner HP, Claus RA, Bauer M, Reinhart K, Lo¨sche W (2006) Platelet-derived microvesicles induce differential gene expression in monocytic cells: a DNA microarray study. Platelets 17:571–576 Taylor DD, Gercel-Taylor C (2008) MicroRNA signatures of tumor-derived exosomes as diagnostic biomarkers of ovarian cancer. Gynecol Oncol 110:13–21 Valadi H, Ekstro¨m K, Bossios A, Sjo¨strand M, Lee JJ, Lo¨tvall JO (2007) Exosomemediated transfer of mRNAs and microRNAs is a novel mechanism of genetic exchange between cells. Nat Cell Biol 9:654–659 Valenti R, Huber V, Iero M, Filipazzi P, Parmiani G, Rivoltini L (2007) Tumorreleased microvesicles as vehicles of immunosuppression. Cancer Res 67:2912–2915 Walkowiak B, Kralisz U, Michalec L, Majewska E, Koziolkiewicz W, Ligocka A, Cierniewski CS (2000) Comparison of platelet aggregability and P-selectin surface expression on platelets isolated by different methods. Thromb Res 99:495–502 Wang Y, Zeigler MM, Lam GK, Hunter MG, Eubank TD, Khramtsov VV, Tridandapani S, Sen CK, Marsh CB (2007) The role of the NADPH oxidase complex, p38 MAPK, and Akt in regulating human monocyte/macrophage survival. Am J Respir Cell Mol Biol 36: 68–77 Wieckowski E, Whiteside TL (2006) Human tumor-derived vs dendritic cellderived exosomes have distinct biologic roles and molecular profiles. Immunol Res 36:247–254

Chapter 29

Detection of Placental miRNAs in Maternal Plasma

Abstract The discovery of fetal nucleic acids in the plasma of pregnant women (Lancet 350:485–487, 1997, Nat Rev Genet 8:71–77, 2007; Proc Natl Acad Sci USA 100:4748 –4753, 2003) has led to the development of a number of noninvasive prenatal diagnostic tests. Nucleic acids of placental origin were previously shown to be released into maternal plasma (Proc Natl Acad Sci USA 100:4748– 4753, 2003; Proc Natl Acad Sci USA 102:14753–14758, 2005). The recent work documented by Lo’s group from the Centre for Research into Circulating Fetal Nucleic Acids, Li Ka Shing Institute of Health Sciences, The Chinese University of Hong Kong (Hong Kong SAR, China) (Clin Chem 54:482–490, 2008) clearly demonstrated that placental miRNAs exist in maternal plasma in readily detectable quantities. The successful application of the approach indicates that placental miRNAs represent a novel class of fetal nucleic acid markers in maternal plasma. Because miRNAs are exceptionally stable in plasma, they hold promise as markers in the clinical setting. The measurement of miRNAs in maternal plasma may become a useful and practical strategy for prenatal monitoring and diagnosis.

29.1

Introduction

The discovery of fetal nucleic acids in the plasma of pregnant women (Lo et al. 1997; Lo and Chiu 2007; Ng et al. 2003) has led to the development of a number of noninvasive prenatal diagnostic tests. Nucleic acids of placental origin were previously shown to be released into maternal plasma (Ng et al. 2003; Chim et al. 2005). The recent work documented by Lo and colleagues (Chim et al. 2008) clearly demonstrated that placental miRNAs exist in maternal plasma in readily detectable quantities. By systematically searching a panel of 157 miRNA assays, these authors have identified 17 placental miRNAs as candidate biomarkers for monitoring pregnancy in maternal plasma. Furthermore, they were able to identify four placental miRNAs (miR-141, miR-149, miR-299-5p, and miR-135b) at higher rates in the maternal plasma before delivery than after. Hence, the authors concluded that these Z. Wang and B. Yang, MicroRNA Expression Detection Methods, DOI 10.1007/978-3-642-04928-6_29, # Springer-Verlag Berlin Heidelberg 2010

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miRNA species are associated with pregnancy. Their data lhave also shown that the plasma concentration of a placental miRNA, miR-141, increases as the pregnancy progresses into the third trimester. This increase in miR-141 in maternal plasma may reflect an increase in the size of the placenta or an increased concentration of miR-141 in the third-trimester placenta. The quantification of placental miRNAs in maternal plasma may offer a noninvasive means for monitoring gene regulation in the placenta. Further, this study also revealed that miRNAs themselves are intrinsically more stable in plasma than mRNAs (Chim et al. 2008). The biological significance of placental miRNAs in maternal plasma requires further elucidation. An intriguing possibility is that these small molecules are taken up by cells exposed to the maternal circulation and may modulate gene expression of the maternal compartment. These findings suggest that the detection of circulating fetal miRNAs holds much promise for noninvasive prenatal diagnosis.

29.2

Protocol

29.2.1 Materials 1. Filter with a pore size of 5 mm, 0.45 mm, or 0.22 mm (Millex-GV; Millipore)

29.2.2 Instruments 1. ABI Prism 7300 or 7900 Sequence Detector (Applied Biosystems)

29.2.3 Reagents 1. 2. 3. 4.

EDTA Trizol LS reagent (Invitrogen) mirVana miRNA Isolation Kit (Ambion) TaqMan Array Human MicroRNA Panel v1.0 (Early Access) (Applied Biosystems), which contains 157 TaqMan MicroRNA Assays, including the respective reverse-transcription primers, PCR primers, and TaqMan probe 5. TaqMan MicroRNA Reverse Transcription Kit (Applied Biosystems) 6. TaqMan Universal PCR Master Mix (Applied Biosystems) 7. DNase I (Invitrogen)

29.2 Protocol

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29.2.4 Procedures The protocols described in this section are essentially the same as reported in the study by Lo’s laboratory (Chim et al. 2008).

29.2.4.1

Maternal Peripheral Blood Sample

1. Collect samples of maternal peripheral blood (12 mL) into tubes containing EDTA. 2. To harvest cell-free plasma, centrifuge the maternal blood samples twice at 4 C. After the first centrifugation at 1600 g for 10 min, centrifuge again the supernatant at 16,000 g for 10 min to remove blood cells (Chiu et al. 2001). 3. To harvest maternal blood cells (including leukocytes and erythrocytes), centrifuge the blood cells obtained in the first centrifugation at 2,300 g for 5 min to remove residual plasma. 4. Add Trizol LS reagent in volumetric ratios of 1:0.8 and 3:1 to the harvested maternal plasma and maternal blood cells, respectively. Specifically, add 0.4 mL of chloroform to 1.6 mL of plasma preserved in 2 mL of Trizol LS reagent. Add 0.24 mL of chloroform to 0.3 mL of processed blood cells preserved in 0.9 mL Trizol LS reagent.

29.2.4.2

RNA Extraction

1. Extract total RNA containing small RNA molecules with the Trizol LS or Trizol reagent and the mirVana miRNA Isolation Kit, according to manufacturer’s protocols. 2. After the chloroform-addition steps and phase separation, mix the aqueous layer with 1.25 volumes of absolute ethanol, load the solution onto the cartridge provided in the mirVana miRNA Isolation Kit, and process the sample. 3. To minimize DNA contamination, treat the eluted RNA preparation with DNase I. For miRNA profiling, further dilute RNA preparations obtained from samples of placentas, maternal blood cells, or postdelivery maternal plasma to 1 mg/L, according to absorbance readings at 260 nm.

29.2.4.3

Real-Time Quantitative RT–PCR Analysis

See Section V for detail.

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29.2.4.4

29 Detection of Placental miRNAs in Maternal Plasma

miRNA Profiling

1. Use the TaqMan Array Human MicroRNA Panel v1.0 for miRNA profiling 2. Use the TaqMan MicroRNA Reverse Transcription Kit for reverse transcription in a 25 mL of total reaction volume with 2.5 mL (2.5 ng) of the total RNA sample 3. Use the TaqMan Universal PCR Master Mix for the PCR

29.2.4.5

Filtration Studies of Placental miRNA and mRNA in Maternal Plasma

1. To ensure that the pregnancy-specific miRNA molecules in maternal plasma are not associated with subcellular particles (Ng et al. 2003), filter samples of maternal plasma through a filter with a pore size of 5 mm, 0.45 mm, or 0.22 mm 2. Extract RNA from the plasma samples with 1 mL of Trizol LS 3. Perform qPCR as for the earlier steps

29.3

Application and Limitation

Chim et al systematically searched for placental miRNAs in maternal plasma to identify miRNAs that are at high concentrations in placentas compared with maternal blood cells and then investigated the stability and filterability of this novel class of pregnancy-associated markers in maternal plasma (Chim et al. 2008). In a panel of TaqMan MicroRNA Assays available for 157 well-established miRNAs, 17 occurred at concentrations >10-fold higher in the placentas than in maternal blood cells and were undetectable in postdelivery maternal plasma. The four most abundant of these placental miRNAs (miR-141, miR-149, miR-299-5p, and miR-135b) were detectable in maternal plasma during pregnancy and showed reduced detection rates in postdelivery plasma. The plasma concentration of miR-141 increased as pregnancy progressed into the third trimester. The successful application of the approach indicates that placental miRNAs represent a novel class of fetal nucleic acid markers in maternal plasma. Because miRNAs are exceptionally stable in plasma, they hold promise as markers in the clinical setting. The measurement of miRNAs in maternal plasma may become a useful and practical strategy for prenatal monitoring and diagnosis. The biological significance of placental miRNAs in maternal plasma requires further elucidation, but an intriguing possibility is that these small molecules are taken up by cells exposed to the maternal circulation and may modulate gene expression of the maternal compartment.

References

349

References Chim SS, Shing TK, Hung EC, Leung TY, Lau TK, Chiu RW, Lo YM (2008) Detection and characterization of placental microRNAs in maternal plasma. Clin Chem 54:482–490 Chim SS, Tong YK, Chiu RW, Lau TK, Leung TN, Chan LY, Oudejans CB, Ding C, Lo YM (2005) Detection of the placental epigenetic signature of the maspin gene in maternal plasma. Proc Natl Acad Sci USA 102:14753–14758 Chiu RWK, Poon LLM, Lau TK, Leung TN, Wong EM, Lo YMD (2001) Effects of bloodprocessing protocols on fetal and total DNA quantification in maternal plasma. Clin Chem 47:1607–1613 Lo YMD, Chiu RWK (2007) Prenatal diagnosis: progress through plasma nucleic acids. Nat Rev Genet 8:71–77 Lo YM, Corbetta N, Chamberlain PF, Rai V, Sargent IL, Redman CW, Wainscoat JS (1997) Presence of fetal DNA in maternal plasma and serum. Lancet 350:485–487 Ng EK, Tsui NB, Lau TK, Leung TN, Chiu RW, Panesar NS, Lit LC, Chan KW, Lo YM (2003) mRNA of placental origin is readily detectable in maternal plasma. Proc Natl Acad Sci USA 100:4748–4753

Part X

Single-cell miRNA Detection Methods

Chapter 30

Quantitative LNA-ELF-FISH Method for miRNA Detection in Single Mammalian Cell

Abstract A method for single-cell miRNA detection has recently been reported, combining the unique recognition properties of locked nucleic acid (LNA) probes with enzyme-labeled fluorescence (ELF) signal amplification. Using this approach, individual miRNAs are identified as bright, photostable fluorescent spots. This technique was developed by Lu and Tsourkas (Nucleic Acids Res doi:10.1093/ nar/gkp482, 2009) from the Department of Bioengineering, University of Pennsylvania School of Engineering and Applied Sciences (Philadelphia, PA, USA) to tackle the problem of cell-to-cell fluctuations in gene expression on phenotypic diversity encountered in the analyses of miRNA expression of cell populations. The LNA-ELF-FISH (fluorescence in situ hybridization) approach has been used to quantify miR-15a in MDA-MB-231 and HeLa cells and miR-155 in MCF-7 cells (Nucleic Acids Res doi:10.1093/nar/gkp482, 2009). The results verified that LNAELF-FISH is a highly sensitive and specific method for miRNA detection at the single molecule level in individual cells. With this technique, single miRNAs could be visualized and counted to yield quantitative information on miRNA expression. LNA-ELF-FISH is extremely simple and yields reproducible data.

30.1

Introduction

Considering recent reports on the complex stochastic nature of gene expression in mammalian cells and the impact of these fluctuations on phenotypic diversity, it is likely that looking at the average miRNA expression of cell populations could result in the loss of important information connecting miRNA expression and cell function. Therefore, it is predicted that insight into the physiologic function of miRNA will require miRNA abundance to be quantified at the single-cell level. Profiling of miRNAs in individual cells is a prerequisite in some instances where there is inherent variability among the cells or because only a few such cells are available for analysis such as the case of early developing embryos. However, because of the

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technical difficulties, the methods suitable for single-cell analysis of miRNA expression have been sparse thus far. With the recent introduction of locked nucleic acid (LNA) oligonucleotides as hybridization probes, miRNA-FISH has become a powerful technique for imaging the spatial localization of miRNA at the tissue, cellular, and even subcellular level (Kloosterman et al. 2006; Nelson et al. 2005; Politz et al. 2006; Silahtaroglu et al. 2007; Wienholds et al. 2005). LNA probes exhibit a remarkable affinity for and specificity against RNA targets, allowing for the discrimination of even single-base mismatches (Chou et al. 2005; Johnson et al. 2004; Valoczi et al. 2004; You et al. 2006). Unfortunately, miRNA-FISH generally cannot be used to provide accurate quantitative measures of miRNA expression, but rather is typically limited to providing a qualitative assessment of miRNA localization patterns and tissue distribution. One of the methods for single-cell miRNA detection has recently been reported by Lu and Tsourkas (2009), combining the unique recognition properties of LNA probes with enzyme-labeled fluorescence (ELF) signal amplification (Paragas et al. 1997). ELF is a process whereby cleavage of a pro-luminescent substrate by phosphatase yields a brilliant, yellow-green fluorescent product at the site of enzymatic activity. The ELF precipitate is not only photostable compared to commonly used fluorophores, but also results in labeling that is up to 40 times brighter than signals achieved with probes directly labeled with fluorophores (Paragas et al. 1997). Using this approach, individual miRNAs are identified as bright, photostable fluorescent spots.

30.2

Protocol

30.2.1 Materials 1. Cell culture materials 2. Multi-chambered coverglass slides (Lab-Tek, Nalge Nunc, Rochester, New York, United States)

30.2.2 Instruments 1. Olympus IX81 motorized inverted fluorescence microscope equipped with a back-illuminated EMCCD camera (Andor) 2. X-cite 120 excitation source (EXFO) 3. Sutter excitation and filter wheels 4. IPLab acquisition software 5. AutoQuant plug-in software 6. Particle analysis counter program

30.2 Protocol

355

30.2.3 Reagents 1. Cy3 dyes (Amersham Biosciences) 2. 4% formaldehyde 3. Pre-hybridization buffer: [25% formamide, 0.05 M EDTA, 4 SSC, 10% dextran sulfate, 1 Denhardt’s solution, 0.5 mg/mL Escherichia coli tRNA, and 0.5 mg/ml RVC) 4. 4 SSC, 2 SSC and 1 SSC 5. PBS 6. ELF 97 mRNA In Situ Hybridization Kit (Molecular Probes, Inc, Eugene, OR, USA) 7. Anti-DIG antibody (Jackson Immuno Research) 8. Post-fixation solution: [2% formaldehyde, 20 mg/mL BSA in 1 PBS] 9. 0.16 M l-ethyl-3-(3-dimethylaminopropyl) carbodiimide (EDC) (Sigma) 10. 0.2% (w/v) glycine/TBS 11. Wash solution: [0.13 M 1-methylimidazole, 300 mM NaCl, pH8.0 adjusted with HCl] 12. 70% ethanol

30.2.4 Procedures The protocols described in this section are essentially the same as reported in the study by Lu and Tsourkas (2009).

30.2.4.1

Design of LNA-FISH Probes

1. Obtain sequences for the miRNAs of interest from the miRNA Registry (microrna.sanger.ac.uk) 2. Design oligonucleotide probes exactly antisense to the selected target miRNAs. Synthesize the probes from Exiqon with LNA modification and a digoxigenin at the 30 -end

30.2.4.2

Enzyme-Labeled Fluorescence (ELF) Treatment

1. Seed cells onto multi-chambered coverglass slides and incubate under normal growth conditions overnight, reaching 50–70% confluency. 2. Fix the cells with 4% formaldehyde for 30 min at room temperature, wash three times with 1 PBS, and permeabilize at 4 C in 70% ethanol overnight. 3. Perform hybridization with the LNA probe (10 nM) at 20–22 C below the melting temperature of the LNA-FISH probe for 1 h after incubation in the

356

4. 5.

6.

7.

30 Quantitative LNA-ELF-FISH Method for miRNA Detection

pre-hybridization buffer for 2 h at 60 C. The optimal level of formamide to be used during hybridization and washing for maximal signal-to-background is empirically determined to be 25%. Perform three stringent washes in 4 SSC, 2 SSC, and 1 SSC. After three stringent washes, as described above, treat the cell samples using the ELF 97 mRNA In Situ Hybridization Kit, according to the manufacturer’s instructions. Briefly, (1) incubate the cells in blocking buffer from the ELF 97 mRNA In Situ Hybridization Kit for 1 h at room temperature. Then, (2) add 2 mg/mL anti-DIG antibody in blocking buffer to the cells and incubate at room temperature for 1 h. (3) After three washes in 1 wash buffer, amplify signals in ELF 97 phosphatase substrate working solution for 10–15 min. For signal preservation, wash the cell samples with 1 wash buffer and postfix the cells by incubating the slides in the post-fixation solution for 30 min at room temperature. Counterstain the slides in 1 mg/ mL Hoechst 33342 and mount in mounting solution. Conduct control experiments using the identical procedure as described above, except for a single hybridization step, were performed with the LNA probes (i.e., LNA-ELF-FISH). ELF should be also performed only as deemed necessary.

30.2.4.3

LNA-ELF-FISH with EDC Treatment

1. After having fixed the cells in 4% paraformaldehyde for 30 min at room temperature and permeabilized in 70% ethanol at 4 C overnight, rehydrate the cells and wash three times with 1 PBS 2. To remove residual phosphate from the PBS washes, incubate slides twice for 10 min in a freshly prepared wash solution 3. Add l-ethyl-3-(3-dimethylaminopropyl) carbodiimide (EDC) to the cells and incubate for 1 h at 25 C 4. Wash the slides in 0.2% (w/v) glycine/TBS and then wash twice in PBS for prehybridization 5. Carry out subsequent LNA hybridization and ELF signal amplifications steps as described above

30.2.4.4

Image Acquisition and Analysis

1. Following in situ hybridization, image cells using an Olympus IX81 motorized inverted fluorescence microscope equipped with a back-illuminated EMCCD camera (Andor), an X-cite 120 excitation source (EXFO), and Sutter excitation and filter wheels. Use a UPLN 60 oilimmersion objective, N.A. 0.9, or an equivalent, for all imaging experiments. Use IPLab acquisition software to acquire the 2D and 3D images.

30.2 Protocol

357

2. After randomly selecting cells in a field, take a 3D stack viewed image with 0.3 mm increments in the z-direction and a total of 35 sections. 3. After 3D deconvolution of the images in IPLab using AutoQuant plug-in software, a 2D image was constructed in IPLab using a maximum intensity merged image. 4. Open images in ImageJ and process using the following commands: (1) Process ->Sharpen, (2) Image ->type ->8-bit, and (3) Process ->binary ->make binary. 5. Count the total number of isolated signals in ImageJ using the particle analysis counter program (Analyze ->analyze particles) (Fig. 30.1).

Select a miRNA of interest

Synthesize LNA-FISH miRNA capture probes with LNA modification and a 3'digoxigenin

Fix cells with 4% formaldehyde

Hybridization of cells with LNA-FISH probe

Treatment with enzyme-labeled fluorescence (ELF) 97 mRNA In Situ Hybridization Kit

Treatment with anti-DIG antibody

Counterstain the slides in Hoechst 33342 and mount in mounting solution.

Fluorescence microscope for image acquisition and analysis

Fig. 30.1 Flowchart of the LNA-ELF-FISH approach for miRNA expression detection. According to Lu and Tsourkas (2009)

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30 Quantitative LNA-ELF-FISH Method for miRNA Detection

Application and Limitation

The LNA-ELF-FISH approach has been used to quantify miR-15a in MDA-MB231 and HeLa cells and miR-155 in MCF-7 cells (Lu and Tsourkas 2009). The results verified that LNA-ELF-FISH is a highly sensitive and specific method for miRNA detection at the single molecule level in individual cells; by combining LNA hybridization probes with ELF signal amplification, single miRNAs could be visualized and counted to yield quantitative information on miRNA expression. The dynamic range of this approach spans more than three orders of magnitude (i.e., 1 to ~1,000 miRNAs per cell) directly and through the construction of standardization curves could also yield quantitative measurements on cells with higher miRNA copy numbers (Lu and Tsourkas 2009). LNA-ELF-FISH is extremely simple and yields reproducible data. Further, in contrast to RT-PCR, no cell lysis, miRNA purification, or sample enrichment steps are required and spatial information is retained. One important advantage is the need for only a single hybridization probe, a large savings in cost and elimination of the need to identify a large number of probes with similar melting temperatures (Lu and Tsourkas 2009). In addition, LNA-ELF-FISH includes the long stokes shift and high photostability of the fluorescent precipitate, conferring low autofluorescence; the high photostability allows for repeated imaging. Moreover, the fluorescent precipitate is extremely bright and thus only short exposure times are needed (i.e., ~10 ms). However, the current availability of only a single ELF substrate limits this approach to imaging only a single RNA per cell sample.

References Chou LS, Meadows C, Wittwer CT, Lyon E (2005) Unlabeled oligonucleotide probes modified with locked nucleic acids for improved mismatch discrimination in genotyping by melting analysis. Biotechniques 39:644, 646, 648 passim Johnson MP, Haupt LM, Griffiths LR (2004) Locked nucleic acid (LNA) single nucleotide polymorphism (SNP) genotype analysis and validation using real-time PCR. Nucleic Acids Res 32:e55 Kloosterman WP, Wienholds E, de Bruijn E, Kauppinen S, Plasterk RH (2006) In situ detection of miRNAs in animal embryos using LNA-modified oligonucleotide probes. Nat Methods 3:27–29 Lu J, Tsourkas A (2009) Imaging individual microRNAs in single mammalian cells in situ. Nucleic Acids Res doi:10.1093/nar/gkp482 Nelson PT, Baldwin DA, Kloosterman WP, Kauppinen S, Plasterk RH, Mourelatos Z (2005) RAKE and LNA-ISH reveal microRNA expression and localization in archival human brain. RNA 12:187–191 Paragas VB, ZhangYZ, Haugland RP, Singer VL (1997) The ELF-97 alkaline phosphatase substrate provides a bright, photostable, fluorescent signal amplification method for FISH. J Histochem Cytochem 45:345–357 Politz JC, Zhang F, Pederson T (2006) MicroRNA-206 colocalizes with ribosome-rich regions in both the nucleolus and cytoplasm of rat myogenic cells. Proc Natl Acad Sci USA 103: 18957–18962

References

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Silahtaroglu AN, Nolting D, Dyrskjot L, Berezikov E, Moller M, Tommerup N, Kauppinen S (2007) Detection of microRNAs in frozen tissue sections by fluorescence in situ hybridization using locked nucleic acid probes and tyramide signal amplification. Nat Protoc 2:2520–2528 Valoczi A, Hornyik C, Varga N, Burgyan J, Kauppinen S, Havelda Z (2004) Sensitive and specific detection of microRNAs by northern blot analysis using LNA-modified oligonucleotide probes. Nucleic Acids Res 32:e175 Wienholds E, Kloosterman WP, Miska E, Alvarez-Saavedra E, Berezikov E, de Bruijn E, Horvitz HR, Kauppinen S, Plasterk RH (2005) MicroRNA expression in zebrafish embryonic development. Science 309:310–311 You Y, Moreira BG, Behlke MA, Owczarzy R (2006) Design of LNA probes that improve mismatch discrimination. Nucleic Acids Res 34:e60

Chapter 31

Single Cell Stem-Looped Real-Time PCR

Abstract Profiling of miRNAs in individual cells is prerequisite in some instances where there is inherent variability among the cells, or because only a few such cells are available for analysis, as in the case of early developing embryos. It is also the case that seemingly uniform cells, such as stem cells, may indeed differ from each other, which can only be judged by analysis of single cells. Surani and colleagues from Wellcome Trust/Cancer Research UK Gurdon Institute of Cancer and Developmental Biology, University of Cambridge (Cambridge, UK) (Nucleic Acids Res 34:e9 2006a, Nat Protoc 1:1154–1159, 2006b) developed a Single Cell StemLooped Real-Time PCR (SC-SL-RT-PCR) protocol for the detection of the miRNA expression profile in a single cell by stem-looped real-time PCR. A single cell is first lysed by heat treatment without further purification. Then, 220 known miRNAs are reverse transcribed into corresponding cDNAs by stem-looped primers. This is followed by an initial PCR step to amplify the cDNAs and generate enough material to permit separate multiplex detection. The diluted initial PCR product is used as a template to check individual miRNA expression by real-time PCR. This sensitive technique permits miRNA expression profiling from a single cell, and allows analysis of a few cells from early embryos as well as individual cells (such as stem cells). The method is extremely sensitive and can be used routinely for the analysis of single cells presumably with 0.015 ng total RNA, which is thousands of times more sensitive than other commonly used miRNA profiling methods; thus, it can be used when only nanogram amounts of rare samples are available. And this method covers a linear dynamic range of six logs.

31.1

Introduction

In many previous studies on miRNA expression detection, total RNA extracted from a large number of possibly heterogeneous cells has been used for analysis. Even cultured cells can show inherent variations. The approaches used previously have hitherto been necessary since nearly all known miRNA profiling methods need Z. Wang and B. Yang, MicroRNA Expression Detection Methods, DOI 10.1007/978-3-642-04928-6_31, # Springer-Verlag Berlin Heidelberg 2010

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31 Single Cell Stem‐Looped Real‐Time PCR

microgram amounts of total RNA (Nelson et al. 2004; Babak et al. 2004; Barad et al. 2004; Krichevsky et al. 2003; Liang et al. 2005; Miska et al. 2004; Sun et al. 2004; Liu et al. 2004), which is required for the fractionation of small RNAs before subsequent analysis (Nelson et al. 2004; Babak et al. 2004; Barad et al. 2004; Krichevsky et al. 2003; Liang et al. 2005; Miska et al. 2004). As already addressed in the previous chapter, profiling of miRNAs in individual cells is prerequisite in some instances where there is inherent variability among the cells, or because only a few such cells are available for analysis, as in the case of early developing embryos. It is also the case that seemingly uniform cells, such as stem cells, may indeed differ from each other, which can only be judged by analysis of single cells. It has been indeed realized that because of the complex stochastic nature of gene expression in mammalian cells and the impact of these fluctuations on phenotypic diversity, looking at the average miRNA expression of cell populations could result in the loss of important information connecting miRNA expression and cell function. For this reason, it is highly desirable to develop efficient methods that provide unambiguous miRNA profile of individual specific cell types. In this regard, the recently developed method by Chen et al. (2005) using a looped real-time PCR-based technique to detect expression of miRNAs is potentially helpful. With this approach they can cover at least seven log of expression range that is accurate and specific for mature miRNA, which can clearly discriminate between mature miRNA and corresponding primary miRNA and precursor miRNA. On top of this approach, Surani and colleagues developed a Single Cell StemLooped Real-Time PCR (SC-SL-RT-PCR) protocol for the detection of the miRNA expression profile in a single cell by stem-looped real-time PCR (Tang et al. 2006a, b). A single cell is first lysed by heat treatment without further purification. Then, 220 known miRNAs are reverse transcribed into corresponding cDNAs by stem-looped primers. This is followed by an initial PCR step to amplify the cDNAs and generate enough material to permit separate multiplex detection. The diluted initial PCR product is used as a template to check individual miRNA expression by real-time PCR (Fig. 31.1). In this study the authors validated the technique with the following experiments. (1) They tested the sensitivity of the looped real-time PCRbased miRNA expression profiling method in multiplex format, and found that this approach works accurately for 8 log range of expression for miR-16, which is effective with 1 mg–0.01 pg of total RNA. As the usual amount of total RNA in a cell is around 15 pg, this strongly suggests that the method is sensitive enough for single cell miRNA expression profiling. (2) They confirmed that the method is sensitive and accurate for the analysis of total RNA from single cells. (3) They then verified that the method can work on the whole cell lysate rather than on purified total RNA. (4) They further demonstrated that the approach can indeed work directly on individual handpicked cells. To determine the extent to which individual cells may differ from each other, they picked 15 single embryonic cells (ES) that were analyzed separately. We observed ~2-fold difference in miR-16 among these 15 single ES cell samples. And (5) finally, the workers examine the ability of the technique for use to obtain a comprehensive miRNA expression profile of single cells. In principle, their single cell miRNA profiling data correlate well with the

31.2 Protocol

363

Lysis Reverse Transcription A single cell

Cell lysate containing miRNAs 1st Strand cDNAs of miRNAs

Real-time Quantification

Pre-PCR miRNA-specific forward primer

Universal reverse primer

PCR products PCR Amplification 40 cycles

Fig. 31.1 Schematic representation of real-time PCR-based multiplex miRNA expression profiling method

cloning and Northern blot data although the published cloning frequencies are very low. This further proved that the Single-cell stem-loop expression profiling method works reliably for single whole ES cells (Tang et al. 2006a, b).

31.2

Protocol

31.2.1 Materials 1. 2. 3. 4. 5.

Thin-walled PCR Eppendorf tube Micropipette Mouth tubes Heat-polished Pasteur pipette 96-well plate

31.2.2 Instruments 1. Prism 7000 SDS (ABI) or any other real-time PCR instruments compatible with TaqMan probe-directed real-time PCR assays 2. Dissection microscope

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31 Single Cell Stem‐Looped Real‐Time PCR

31.2.3 Reagents 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15. 16.

BSA (Sigma) PBS (Gibco; pH 7.2) Molecular biology grade H2O (Eppendorf) BSA–PBS solution: [0.1% (wt/vol) or 1 mg/mL BSA in PBS] EDTA–PBS solution: [0.038% (wt/vol) or 0.38 mg/mL EDTA in PBS] 1 trypsin–EDTA solution: [Gibco; 0.25% (wt/vol) or 2.5 mg/mL trypsin, 0.038% (wt/vol) or 0.38 mg/mL EDTA] 220-plex forward and reverse primers (synthesized by Integrated DNA Technologies Inc) Universal RP (URP; 100 mM; the sequence is: 50 -CTCAAGTGTCGTGGAGTCGGCAA-30 ; Integrated DNA Technologies Inc) 100 mM dNTP (ABI) 100 mM MgCl2 (ABI) 2 TaqMans Universal PCR Master Mix (2 UMM) without AmpErases UNG (ABI) RNase inhibitor (20 U/mL; ABI) Moloney murine leukemia virus (MMLV) reverse transcriptase (50 U/mL ABI: high capacity cDNA achieve kit) AmpliTaqGold DNA polymerase (5 U/ml; ABI) 1-plex FP (5 mM) plus TaqMan probe (1 mM) mix (synthesized by Integrated DNA Technologies Inc)

31.2.4 Procedures The protocols described in this section are essentially the same as reported in the studies by Surani and colleagues (Tang et al. 2006a, b).

31.2.4.1

Preparation of Single Cells

1. In order to pick up and transfer individual cells, ideally use a micropipette attached to, and controlled by, a mouth tube, which is commonly used for manipulating early mouse embryos under a dissection microscope. To commence, pick up an ES cell colony with a micropipette. 2. Transfer it to a drop of EDTA–PBS and incubate at room temperature (20–30 C) for 10 min. 3. Transfer the ES cell cluster to a drop of trypsin–EDTA solution for 10 min at 37 C. 4. Transfer the cell cluster to a drop of BSA–PBS, and then to another drop of BSA–PBS. 5. Gently pipette 20–30 times with a heat-polished Pasteur pipette or microcapillary until the cell cluster dissociates into single cells. Transfer a proportion

31.2 Protocol

365

(i.e., numbers that are sufficient for later analysis) of the single cells to another drop of BSA–PBS. 31.2.4.2

Lysis of Single Cells and Reverse Transcription

1. Prepare an RT master mix containing the following and mix gently but thoroughly:

H2O 10 cDNA archiving kit buffer 220-plex RPs (200 nM) RNase inhibitor (20 U/mL) Total volume

X1 3.61 mL 0.5 mL 0.125 mL 0.065 mL 4.3 mL

X20X1.1 79.42 mL 11 mL 2.75 mL 1.43 mL 94.6 mL

2. Prepare an RT reaction medium by adding 4.3 mL RT master mix to each thinwalled PCR Eppendorf tube on ice 3. Pick and transfer a single cell into each tube. It is critical to ensure that the amount of BSA–PBS carry over with each cell is minimal 4. Centrifuge at 9,000g for 10 s, followed by treatment of the samples at 95 C for 5 min. Place on ice 5. Prepare an enzyme mix containing the following: RNase inhibitor (20 U/mL) MMLV RT (50 U/mL) dNTP (100 mM; with dTTP) Total volume

X1 0.065 mL 0.335 mL 0.25 mL 0.65 mL

X20X1.1 1.43 mL 7.37 mL 5.5 mL 14.3 mL

6. Add 0.65 mL enzyme mix into each tube, mix evenly by vortexing the samples briefly, and then spin down the samples by centrifuging at 9,000g for 10 s 7. Perform the RT reaction by performing the following cycles: first, 16 C for 30 min; then, 20 C for 30 s, 42 C for 30 s, and 50 C for 1 s for 60 cycles; then, 85 C for 5 min (to inactivate RT); and, finally, 4 C prior to the next step 31.2.4.3

Pre-PCR

1. Prepare a pre-PCR master mix by combining the following: 2 UMM 220-plex FPs (450 nM) H2O MgCl2 (100 mM) dNTP (100 mM) URP (100 mM) AmpliTaqGold polymerase (5 U/mL) Total volume

X1 12.5 mL 2.78 mL 0.72 mL 0.5 mL 1 mL 1.25 mL 1.25 mL 20 mL

X20X1.1 275 mL 61.16 mL 15.84 mL 11 mL 22 mL 27.5 mL 27.5 mL 440 mL

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31 Single Cell Stem‐Looped Real‐Time PCR

2. Add 20 mL pre-PCR master mix into each RT reaction tube. Mix evenly by vortexing and then spin down the samples by centrifuging at 9,000g for 10 s 3. Perform the pre-PCR reaction by the following cycles: first, 95 C for 10 min (to activate the AmpliTaqGold polymerase); then, 55 C for 2 min; then, 95 C for 1 s and 65 C for 1 min for 18 cycles; and, finally, save at 4 C 4. Dilute the pre-PCR product to give a 1:4 dilution (i.e., 25 mL pre-PCR product plus 75 mL H2O)

31.2.4.4

Real-Time PCR for Quantification

1. Prepare a TaqMan master mix containing the following: 2 UMM (no UNG) URP (100 mM) 1:4 diluted pre-PCR product H2O Total volume

X1 5 mL 0.1 mL 0.1 mL 2.8 mL 8 mL

X220X1.1X1.1 1,331 mL 26.62 mL 26.62 mL 745.36 mL 2,129.6 mL

2. For the PCR reaction, combine 17.6 mL TaqMan master mix and 4.4 mL 1-plex FP (5 mM) plus TaqMan probe (1 mM), to give a total volume of 22 mL. 3. For the standard, prepare the following PCR master mix: 2 UMM (no UNG) H2O URP (100 mM) 1-plex FP (5 mM)/TaqMan probe (1 mM) Total volume

X1 5 mL 1.9 mL 0.1 mL 2 mL 9 mL

X40X1.1 220 mL 83.6 mL 4.4 mL 88 mL 396 mL

4. For the PCR reaction of standards, combine 19.8 mL PCR master mix and 2.2 mL standard cDNA sample to give a total volume of 22 mL. 5. Mix thoroughly and add 10 mL to each well of a 96-well plate. In the plate, samples S1–S4 represent the standard samples with serial tenfold dilution. NTC represents the no template control reaction. Samples A1–A43 represent the samples for assaying the miRNA expression. Each assay is duplicated. Ideally, assays for the same miRNA in different samples should be run on the same plate to reduce potential variations between different plates. 6. Load the plate into the ABI Prism 7000 SDS, or any other real-time PCR instruments compatible with the TaqMan probe-directed real-time PCR assay. 7. Run the following real-time PCR program: 95 C for 10 min; 95 C for 15 s and 60 C for 1 min for 40 cycles.

References

31.3

367

Application and Limitation

Due to their extremely small size, most current miRNA profiling methods are not highly sensitive, and they usually require microgram quantities of total RNA for analysis, which correspond to hundreds of thousands of cells. For a typical hybridization-based miRNA profiling method, it is usually necessary to have 5–20 mg total RNA (Babak et al. 2004; Barad et al. 2004; Krichevsky et al. 2003; Liang et al. 2005; Miska et al. 2004; Sun et al. 2004; Liu et al. 2004). Moreover, most methods also have a relatively narrow linear dynamic range, usually < 3 logs. By comparison, the SC-SL-RT-PCR is sufficiently sensitive to generate a miRNA expression profile of single cells. In contrast, our method is extremely sensitive and can be used routinely for the analysis of single cells (0.015 ng total RNA), which is thousands of times more sensitive than other commonly used miRNA profiling methods (Chen et al. 2005; Lao et al. 2006; Tang et al. 2006a). And it is easy to cover a linear dynamic range of six logs by this method. The SC-SL-RT-PCR method should prove useful in many cases. For example, specification of cells, such as primordial germ cells in very early embryos, occurs in just 40 cells (Saitou et al. 2002). To understand the role that miRNA may play in this process requires analysis at the single cell level. Other relatively rare cells such as some stem cells in adults also require analysis of single cells. Tumors also often consist of a heterogeneous group of cells, so it is desirable to analyze them individually to determine what role miRNA plays in cancers (Wang and Dick 2005). We propose that the method we have described here will help to advance an understanding of the functions of miRNAs generally.

References Babak T, Zhang W, Morris Q, Blencowe BJ, Hughes TR (2004) Probing microRNAs with microarrays: tissue specificity and functional inference. RNA 10:1813–1819 Barad O, Meiri E, Avniel A, Aharonov R, Barzilai A, Bentwich I, Einav U, Gilad S, Hurban P, Karov Y, Lobenhofer EK, Sharon E, Shiboleth YM, Shtutman M, Bentwich Z, Einat P (2004) MicroRNA expression detected by oligonucleotide microarrays: system establishment and expression profiling in human tissues. Genome Res 14:2486–2494 Chen C, Ridzon DA, Broomer AJ, Zhou Z, Lee DH, Nguyen JT, Barbisin M, Xu NL, Mahuvakar VR, Andersen MR, Lao KQ, Livak KJ, Guegler KJ (2005) Real-time quantification of microRNAs by stem-loop RT-PCR. Nucleic Acids Res 33:e179 Krichevsky AM, King KS, Donahue CP, Khrapko K, Kosik KS (2003) A microRNA array reveals extensive regulation of microRNAs during brain development. RNA 9:1274–1281 Lao K, Xu NL, Yeung V, Chen C, Livak KJ, Straus NA (2006) Multiplexing RT-PCR for the detection of multiple miRNA species in small samples. Biochem Biophys Res Commun 343:85–89 Liang RQ, Li W, Li Y, Tan CY, Li JX, Jin YX, Ruan KC (2005) An oligonucleotide microarray for microRNA expression analysis based on labeling RNA with quantum dot and nanogold probe. Nucleic Acids Res 33:e17

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Liu C-G, Calin GA, Meloon B, Gamliel N, Sevignani C, Ferracin M, Dumitru CD, Shimizu M, Zupo S, Dono M, Alder H, Bullrich F, Negrini M, Croce CM (2004) An oligonucleotide microchip for genome-wide microRNA profiling in human and mouse tissues. Proc Natl Acad Sci USA 101:9740–9744 Miska EA, Alvarez-Saavedra E, Townsend M, Yoshii A, Sestan N, Rakic P, Constantine-Paton M, Horvitz HR (2004) Microarray analysis of microRNA expression in the developing mammalian brain. Genome Biol 5:R68 Nelson PT, Baldwin DA, Scearce LM, Oberholtzer JC, Tobias JW, Mourelatos Z (2004) Microarray-based, high-throughput gene expression profiling of microRNAs. Nat Methods 1:155–161 Saitou M, Barton SC, Surani MA (2002) A molecular programme for the specification of germ cell fate in mice. Nature 418:293–300 Sun Y, Koo S, White N, Peralta E, Esau C, Dean NM, Perera RJ (2004) Development of a microarray to detect human and mouse microRNAs and characterization of expression in human organs. Nucleic Acids Res 32:e188 Tang F, Hajkova P, Barton SC, Lao K, Surani MA (2006a) MicroRNA expression profiling of single whole embryonic stem cells. Nucleic Acids Res 34:e9 Tang F, Hajkova P, Barton SC, O’Carroll D, Lee C, Lao K, Surani MA (2006b) 220-plex microRNA expression profile of a single cell. Nat Protoc 1:1154–1159 Wang JC, Dick JE (2005) Cancer stem cells: lessons from leukemia. Trends Cell Biol 15:494–501

Chapter 32

miRNA Function-Reporter Expression Assay

Abstract The detection of microRNAs (miRNAs) at single-cell resolution is crucial for acquiring knowledge about the role of these post-transcriptional regulators. Huttner and colleagues from the Max Planck Institute of Molecular Cell Biology and Genetics (Dresden, Germany) developed a miRNA Function-Reporter Expression assay for this purpose. It is a relatively simple and reliable system that allows the detection of miRNAs with cellular resolution in vivo without the need to generate transgenic animals (Biotechniques 41:727–732, 2006). The system is based on the acute administration of a dual-fluorescence GFP-reporter/mRFPsensor (DFRS) plasmid for a specific miRNA into the organism of interest. In their DFRS plasmids, both GFP and mRFP are under the control of identical constitutive promoters. The GFP-reporter is used to identify the cells actually expressing the plasmid, given that the sensor-based strategy relies on the silencing of a transcript. The mRFP-sensor contained a 30 untranslated region (30 UTR) with a tandem cassette complementary to the miRNA of interest. To establish a system allowing the monitoring of miRNA dynamics in defined cell lineages during mammalian embryonic development, the group explored the use of DFRS plasmids in conjunction with a combination of methods previously used to achieve acute expression of transgenes and RNA interference in developing mouse embryos. This combination consists of the topical release of nucleic acids in the proximity of a specific tissue of a mouse embryo developing either in culture or in utero and their delivery into this tissue by directed electroporation. This strategy provides a simple approach to study a specific miRNA in the tissue and cell lineage of interest.

32.1

Introduction

The detection of microRNAs (miRNAs) at single-cell resolution is crucial for acquiring knowledge about the role of these post-transcriptional regulators. To detect miRNAs in tissues microscopically, two strategies have been used: in situ hybridization using locked nucleic acid (LNA)-modified DNA oligonucleotide probes, which detect the presence of miRNAs irrespective of their potential activity Z. Wang and B. Yang, MicroRNA Expression Detection Methods, DOI 10.1007/978-3-642-04928-6_32, # Springer-Verlag Berlin Heidelberg 2010

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32 miRNA Function-Reporter Expression Assay

(Wienholds et al. 2005), and the expression or administration of target mRNAs (sensors), which detect miRNAs via their degradation-triggering activity toward the sensor (Giraldez et al. 2005; Brennecke et al. 2003; Mansfield et al. 2004). Although both approaches are powerful, certain limitations remain. In situ hybridization using LNA probes requires tissue fixation, which prevents the monitoring of miRNA appearance/disappearance in a given cell lineage during cell fate change. While this limitation could potentially be overcome by in vivo expression of a sensor mRNA encoding a fluorescent protein, the latter approach has typically involved the generation of transgenic animals (Brennecke et al. 2003; Mansfield et al. 2004). Moreover, in the sensor approach, a lack of signal is interpreted as being indicative of the presence of a miRNA, which calls for some means of verification that the sensor mRNA is actually being transcribed in the cell lacking sensor protein. To overcome these limitations, Huttner and colleagues (De Pietri Tonelli et al. 2006) developed a miRNA Function-Reporter Expression assay, which is a relatively simple and reliable system that allows the detection of miRNAs with cellular resolution in vivo without the need to generate transgenic animals. The system is based on the acute administration of a dual-fluorescence GFP-reporter/mRFPsensor (DFRS) plasmid for a specific miRNA into the organism of interest. In their DFRS plasmids, both GFP and mRFP are under the control of identical constitutive promoters. The GFP-reporter is used to identify the cells actually expressing the plasmid, given that the sensor-based strategy relies on the silencing of a transcript. The mRFP-sensor contained a 30 untranslated region (30 UTR) with a tandem cassette (Brennecke et al. 2003; Mansfield et al. 2004) complementary to the miRNA of interest. To establish a system allowing the monitoring of miRNA dynamics in defined cell lineages during mammalian embryonic development, De Pietri Tonelli et al explored the use of DFRS plasmids in conjunction with a combination of methods previously used to achieve acute expression of transgenes and RNA interference in developing mouse embryos (De Pietri Tonelli et al. 2006). This combination consists of the topical release of nucleic acids in the proximity of a specific tissue of a mouse embryo developing either in culture or in utero and their delivery into this tissue by directed electroporation. This combination of methods has been successfully applied, in particular, to the developing mouse brain (Calegari et al. 2002; Takahashi et al. 2002). This strategy provides a simple approach to study a specific miRNA in the tissue and cell lineage of interest.

32.2

Protocol

32.2.1 Materials 1. Primers (F-mRFP-Nhe and R-mRFP-Eco, Sigma-Genosys Ltd., Pampisford, Cambrigdeshire, UK) 2. pCMS-EGFP vector (Clontech Laboratories, Mountain View, CA, USA)

32.2 Protocol

3. 4. 5. 6.

371

pGEMT (Promega GmbH, Mannheim, Germany) Platinum electrodes (2 mm diameter) Tissue-Tek1 LNA-modified DNA oligonucleotides (Exiqon A/S, Vedbaek, Denmark)

32.2.2 Instruments 1. BTX1-ECM1830 electroporator (Harvard Apparatus, Holliston, MA, USA) 2. Standard upright microscope (Olympus1 Optical, Europe GmbH, Hamburg, Germany) 3. Model SZX12 dissecting microscope equipped with epifluorescence (Olympus, Hamburg, Germany) 4. IPlab software version 3.5.1 (Scanalytics, Rockville, MD, USA) or the Zeiss LSM Image Examiner software version 3.2.0.70 (Carl Zeiss GmbH)

32.2.3 Reagents 1. 4% paraformaldehyde 2. 120 mM phosphate buffer (pH7.4) 3. DIG oligonucleotide 30 end labeling kit (Roche Diagnostic GmbH, Mannheim, Germany) 4. Alkaline phosphatase-conjugated anti-DIG antibody 5. 5-bromo-4-chloro-3-indoxylphosphate/nitro blue tetrazolium (BCIP/NBT; Sigma-Aldrich Chemie GmbH, Taufkirchen, Germany)

32.2.4 Procedures The procedures described here primarily follow the study reported by De Pietri Tonelli et al. (2006).

32.2.4.1

Construction of Dual-Fluorescence GFP-reporter/mRFP-Sensor (DFRS) Plasmids

1. Use a plasmid containing the cDNA for monomeric red fluorescent protein (mRFP) (HHMI-UCSD La Jolla, CA, USA) (Campbell et al. 2002) as a template to obtain an mRFP cDNA by PCR, using the primers (F-mRFP-Nhe and R-mRFP-Eco). 2. Digest the PCR fragment with NheI and NotI, and ligate into the NheI and NotI sites of pCMS-EGFP vector to yield pCMS-EGFP-mRFP. 3. Amplify the simian virus 40 (SV40) promoter by PCR from pCMS-EGFP, using the oligonucleotides Bgl-SV40-F1 and Nhe-SV40-R1.

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4. Ligate the PCR products into BglII-NheI-digested pCMS-EGFP-mRFP to obtain the plasmid pDSV-GmR. 5. After removal of the polylinker 50 to the mRFP coding sequence by NheI-SacII digestion of pDSV-G-mR, fill in the overhangs by Klenow DNA polymerase-I, and relegate the plasmid to yield the plasmid pDSV2-G-mR. 6. Obtain two fragments of the Unc54 30 untranslated region (UTR) by reverse transcription PCR (RT-PCR) of Caenorhabditis elegans total RNA (mixed stage of development), using the sense/antisense oligonucleotides Eco-UncF/R-Unc-PacXho and F-Unc-XhoFse/R-Unc-Not for the 50 and the 30 fragment, respectively. 7. Ligate the 50 fragment of the unc54 30 UTR into pGEMT plasmid, yielding pGEMT-Unc-50 F. 8. Digest the 30 fragment of the Unc54 30 UTR with XhoI and EagI and ligate into XhoI-EagI-digested pGEMT-Unc-50 F to obtain pGEMTUnc-30 UTR. 9. Prepare the tandem cassettes complementary to the target miRNAs for detection, and the tandem cassettes containing the control sequence or the mutated miRNA complementary sequences by annealing synthetic oligonucleotides containing an overhang to generate a PacI-restricted 50 end, followed by two sequences complementary to the miRNAs of interest (separated by an AscI restriction site) and an overhang to generate a FseI-restricted 30 end. 10. Ligate the annealed product into PacI-FseI-digested pGEMT-Unc-30 UTR, yielding pGEMT-UncSh-SmiR-X (X represents the target miRNA or control or mutated sequence). 11. Digest the latter plasmids with EcoRI and NotI, and ligate the tandem cassette, containing modified Unc54 30 UTR into EcoRI-NotI-digested pDSV2-G-mR, yielding the dual-fluorescent green fluorescent protein (GFP)-reporter/mRFPsensor (DFRS) plasmids for the target miRNAs, the DFRS control plasmid, and the mutated DFRS plasmids (De Pietri Tonelli et al. 2006).

32.2.4.2

Mouse Embryo Electroporation

1. Determine the topology of the embryos using illumination and a dissecting microscope rather than ultrasound microscopy. Then perform in utero electroporation of mouse embryos (Takahashi et al. 2002), as below. 2. Anesthetize pregnant mice 13 days postcoitum with isofluorane vapor and expose their uteri. 3. Using a glass capillary, inject 1–3 mL PBS containing 3–5 mg/mL DFRS plasmid through the uterine wall into the lumen of the telencephalic vesicles or release in proximity of the ectoderm of the embryo. 4. Immediately after injection, deliver 6 square electrical pulses of 30 V, 50 ms each at 1-s intervals through platinum electrodes (2 mm diameter) using an electroporator. 5. Use the orientation of the electric field to direct the uptake of the plasmid to specific regions of the developing brain or ectoderm.

32.3 Application and Limitation

373

6. After electroporation, relocate the uterus into the peritoneal cavity, and suture the abdomen. 7. Kill the mice either 24 or 72 h after in utero electroporation, and collect the embryos for further analyses. 8. Perform ex utero electroporation of DFRS plasmids into telencephalic vesicles of E10 mouse embryos followed by 24 h of whole-embryo culture as described previously (Calegari et al. 2002).

32.2.4.3

In situ Hybridization on Cryosections Using LNA-Modified Oligonucleotide Probes

1. Fix whole-mount E11 and E14 mouse embryos overnight at 4 C in 4% paraformaldehyde in 120 mM phosphate buffer equilibrated in 30% sucrose in PBS 2. Embed in Tissue-Tek1 3. Prepare 10-mm cryosections 4. Perform in situ hybridization according to standard protocols with the following modifications. (1) Label LNA-modified DNA oligonucleotides with digoxygenin (DIG)-ddUTP using the DIG oligonucleotide 30 end labeling kit according to manufacturer’s instructions. (2) Prehybridize the cryosections by incubating overnight at 50–60 C in hybridization buffer containing 200 pmol/mL of DIG-labeled LNA-oligonucleotide. (3) Following incubation with alkaline phosphatase-conjugated anti-DIG antibody at 4 C overnight, stain with 5-bromo-4-chloro-3-indoxylphosphate/nitro blue tetrazolium (BCIP/NBT) at 37 C for 2 h and then either at 4 C for 1–2 days or at room temperature for 6–12 h 5. Acquire images with a standard upright microscope

32.2.4.4

Fluorescence Microscopy

1. Image whole-mount mouse embryos using a Model SZX12 dissecting microscope equipped with epifluorescence 2. Process images using the IPlab software version 3.5.1 or the Zeiss LSM Image Examiner software version 3.2.0.70

32.3

Application and Limitation

The a miRNA Function-Reporter Expression assay through acutely administering a DFRS plasmid for a specific miRNA offers a convenient method to detect these important post-transcriptional regulators with single-cell resolution and to monitor their dynamics in vivo. The approach, which presumably is applicable to a wide variety of species, circumvents the need to generate transgenic organisms, which is

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much more labor intensive (Mansfield et al. 2004; Brennecke et al. 2003). Moreover, the topical administration of DFRS plasmids, followed by their directed electroporation (Osumi and Inoue, 2001; Calegari et al. 2002), provides a simple approach to study a specific miRNA in the tissue and cell lineage of interest.

References Brennecke J, Hipfner DR, Stark A, Russell RB, Cohen SM (2003) bantam encodes a developmentally regulated microRNA that controls cell proliferation and regulates the proapoptotic gene hid in Drosophila. Cell 113:25–36 Calegari F, Haubensak W, Yang D, Huttner WB, Buchholz F (2002) Tissue-specific RNA interference in postimplantation mouse embryos with endoribonuclease-prepared short interfering RNA. Proc Natl Acad Sci USA 99:14236–14240 Campbell RE, Tour O, Palmer AE, Steinbach PA, Baird GS, Zacharias DA, Tsien RY (2002) A monomeric red fluorescent protein. Proc Natl Acad Sci USA 99:7877–7882 De Pietri Tonelli P, Calegari F, Fei JF, Nomura T, Osumi N, Heisenberg CP, Huttner WB (2006) Single-cell detection of microRNAs in developing vertebrate embryos after acute administration of a dual-fluorescence reporter/sensor plasmid. Biotechniques 41:727–732 Giraldez AJ, Cinalli RM, Glasner ME, Enright AJ, Thomson JM, Baskerville S, Hammond SM, Bartel DP, Schier AF (2005) MicroRNAs regulate brain morphogenesis in zebrafish. Science 308:833–838 Mansfield JH, Harfe BD, Nissen R, Obenauer J, Srineel J, Chaudhuri A, Farzan-Kashani R, Zuker M, Pasquinelli AE, Ruvkun G, Sharp PA, Tabin CJ, McManus MT (2004) MicroRNAresponsive ‘sensor’ transgenes uncover Hox-like and other developmentally regulated patterns of vertebrate microRNA expression. Nat Genet 36:1079–1083 Osumi N, Inoue T (2001)Gene transfer into cultured mammalian embryos by electroporation. Methods 24:35–42 Takahashi M, Sato K, Nomura T, Osumi N (2002) Manipulating gene expressions by electroporation in the developing brain of mammalian embryos. Differentiation 70:155–162 Wienholds E, Kloosterman WP, Miska E, Alvarez-Saavedra E, Berezikov E, de Bruijn E, Horvitz HR, Kauppinen S, Plasterk RH (2005) MicroRNA expression in zebrafish embryonic development. Science 309:310–311

Part XI

Whole Mount In Situ Analysis

Chapter 33

Whole Mount In Situ Hybridization (WM-ISH) for miRNA Expression Profiling During Vertebrate Development

Abstract Knowledge of tissue-specific and cell-specific expression patterns of miRNAs can directly inform functional studies. However, detailed analysis of spatial patterns of miRNA expression has been technically challenging. While the regular ISH technique has been extensively used for characterizing cellular localization and tissue distribution of miRNAs in tissue and cell preparations, it cannot be efficiently applied to monitor miRNA expression in whole animals. Utilization of locked nucleic acids (LNAs) in ISH, however, has allowed for the whole mount in situ hybridization techniques (WM-ISH) to monitor miRNA expression in whole animals or animal embryos. The WM-ISH techniques have been tested in Drosophila, zebrafish, chicken, and mouse embryos by several laboratories (Proc Natl Acad Sci USA 102:18017–18022, 2005; Science 309:310–311, 2005; Nat Methods 3:27– 29, 2006; Dev Dyn 235:3156–3165, 2006). These studies led us to tremendous insight into the spatial and temporal patterns of miRNA expression and control of embryonic development by miRNAs. The major features and advantages of the WM-ISH techniques are whole mount analysis and high-throughput profiling of miRNAs. The data from WM-ISH highlight cell-type, organ or structure-specific expression, localization within germ layers and their derivatives, and expression in multiple cell and tissue types and within subregions of structures and tissues, the information which is otherwise inaccessible with other miRNA expression detection methods. This chapter mainly introduces the methods provided in the study by Darnell et al. (Dev Dyn 235:3156–3165, 2006) from the Department of Cell Biology and Anatomy, University of Arizona (Tucson, Arizona, USA).

33.1

Introduction

Knowledge of tissue-specific and cell-specific expression patterns of miRNAs can directly inform functional studies (Aboobaker et al. 2005). For example, murine miR-181 was isolated on the basis of its predominant expression in the thymus and Z. Wang and B. Yang, MicroRNA Expression Detection Methods, DOI 10.1007/978-3-642-04928-6_33, # Springer-Verlag Berlin Heidelberg 2010

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proved to regulate cell fate choice in the hematopoietic lineage (Chen et al. 2004). miR-375 is specifically expressed in pancreatic islet cells, where it regulates genes involved in insulin secretion (Poy et al. 2004). miR-1 is found exclusively in muscles, where it regulates cardiomyocyte proliferation in vertebrates (Zhao et al. 2005) and muscle physiology in flies (Sokol and Ambros 2005) of ASE neurons, and they control the identity of these two neurons by inhibiting different transcription factors that regulate ASE left/right cell fate (Chang et al. 2004; Johnston and Hobert 2003). Although temporal expression of miRNAs can be assessed by Northern analysis, methods to analyze their spatial expression have been limited. The strategy most widely used has been Northern analysis using RNA from dissected vertebrate organs (Lagos-Quintana et al. 2002). However, this process provides coarse spatial and cell type resolution as most organs have many cell types, which may or may not all express a given miRNA. Detailed analysis of spatial patterns of miRNA expression has been technically challenging (Aboobaker et al. 2005). For these reasons, methods to directly visualize miRNAs in situ are desirable. The application of the in situ hybridization (ISH) technique to miRNA detection has been described in detail in Section IV. While the regular ISH technique has been extensively used for characterizing cellular localization and tissue distribution of miRNAs in tissue and cell preparations, it cannot be efficiently applied to monitor miRNA expression in whole animals. With the development of locked nucleic acids (LNAs), however, it is now possible to design RNA probes that can hybridize to target RNA sequences with extremely high specificity and stability (Wahlestedt et al. 2000; Elmen et al. 2005). And utilization of LNAs in ISH directly the whole mount in situ hybridization techniques (WM-ISH) has been successfully employed to monitor miRNA expression in whole animals or animal embryos (Aboobaker et al. 2005; Wienholds et al. 2005; Kloosterman et al. 2006). WM-ISH has proven effective for WM-ISH detection of miRNA expression (Wienholds et al. 2005; Kloosterman et al. 2006). Aboobaker et al reported the spatial patterns of miRNA transcription during Drosophila embryonic development using WM-ISH methods (Aboobaker et al. 2005). A WM-ISH overview of miRNA expression in zebrafish detected the majority of miRNAs and demonstrated dynamic spatial and temporal expression patterns (Wienholds et al. 2005). By contrast, a WM-ISH screen in mouse embryo established the first comprehensive set of miRNA expression patterns in animal development (Kloosterman et al. 2006). The study found these patterns to be remarkably specific and diverse, which suggests highly specific and diverse roles for miRNAs. Most miRNAs are expressed in a tissue-specific manner during segmentation and later stages but are not present during early development. Using their improved WM-ISH techniques, Darnell et al. (2006) presented a comprehensive view of miRNA expression during embryogenesis in chicken to map expression of 135 miRNA genes including five miRNAs that had not been previously reported in chicken. They detected 84 miRNAs before day 5 of embryogenesis, of which 75 miRNAs showed differential expression. Whereas few miRNAs were expressed during formation of the primary germ layers, the number of miRNAs detected increased rapidly during organogenesis.

33.2 Protocol

379

The chicken embryo provides an advantageous alternative to mouse for miRNA expression analyses. As an amniote, developmental processes in chicken closely mimic those in mammalian species, including mouse and humans. Whole mount in situ hybridization protocols have also been optimized for chick to give outstanding sensitivity and low background (Nieto et al. 1996; Bell et al. 2004), and embryos are easily and inexpensively obtained.

33.2

Protocol

33.2.1 Materials 1. 2. 3. 4.

Fertile chicken eggs (HyLine, Iowa; not a commercially available source) 6- or 12-well plates 24-well plates 15- or 24-mm Netwell Inserts with 74-mM polyester mesh bottoms (Corning, Inc) 5. Paraplast (Kendall)

33.2.2 Instruments 1. 2. 3. 4. 5.

Nutator Leica PlanApo stereomicroscope Digital acquisition system Cytoseal XYL (Richard-Allan Scientific) Leica DMRXE microscope

33.2.3 Reagents 1. 2. 3. 4.

5. 6. 7. 8.

Chick saline (123 mM NaCl in nanopure water) PBS DIG Oligonucleotide 3’ end labeling kit (Roche) Prehybridization solution (50% formamide, 5x SSC, 2% blocking powder, 0.1% Tween-20, 0.1% CHAPS, 50 mg/mL yeast RNA, 5 mM EDTA, 50 mg/ mL heparin, DEPC water) 2 SSC 0.1% Chaps 20% sheep serum KTBT (50 mM Tris, pH7.5, 150 mM NaCl, 10 mM KCl, 1% Tween-20)

33 Whole Mount In Situ Hybridization

380

9. NTMT (two solutions changes 10 min; 100 mM NaCl, 100 mM Tris of pH9.5, 50 mM MgCl2, 0.1% Tween-20) 10. 0.1% sodium azide 11. Xylene

33.2.4 Procedures The procedures described herein are based on the studies reported by Kloosterman et al. (2006) and by Darnell et al. (2006).

33.2.4.1

Embryo Collection and Preparation

1. Incubate fertile chicken eggs in a forced-draft, humidified incubator at 37.5 C for 0.5–5 days, depending on the stages desired. 2. Collect embryos into chilled chick saline, remove from the vitelline membrane, and clean of yolk. 3. Open extraembryonic membranes and large body cavities (brain vesicles, atria, allantois, eye) to minimize trapping of the in situ reagents. 4. Fix embryos in fresh, cold 4% paraformaldehyde in PBS overnight at 4 C. Maintain them at 4 C during collection, because significant loss of labeling is correlated with increased time at room temperature during collection. 5. Rinse the embryos in PBS, then in PBS plus 1% Tween-20 (PBT). 6. Dehydrate the embryos by steps (25, 50, 75, 100, 100%) into methanol before being cooled to 20 C overnight (or up to 10 days). 7. Rehydrate to reverse this series. 8. Rinse the embryos twice in PBS and treat older embryos with proteinase K: stages 8–13 and 14–18 at 10 mg/mL of proteinase K for 10 and 20 min, respectively; stages 19 and older at 20 mg/mL of proteinase K for 20 min. 9. Rinse the embryos repeatedly in PBT to stop the digestion. 10. Place them into prehybridization. 11. Store the embryos until use either at the methanol step or in prehybridization at 20 C for fewer than 10 days. (Embryos stored for more than 10 days will have a considerable decrease in hybridization signal and increase in background.)

33.2.4.2

Probe Preparation

1. Obtain sequences for the miRNAs of interest from the miRNA Registry (microrna.sanger.ac.uk). 2. Design Locked Nucleic Acid modified DNA oligonucleotides (LNAs) capture probes complementary to the mature miRNAs and synthesize by Exiqon A/S.

33.2 Protocol

381

Add digoxigenin-labeled UTP to the 3’ end of the LNAs using a DIG Oligonucleotide 3’ end labeling kit.

33.2.4.3

In Situ Hybridization

1. Transfer the prepared embryos into a standard prehybridization solution and incubate for 2 h in 24-well plates (1 ml/well) in a shaking hybridization oven at a temperature between 21 and 23 C below the reported melting temperature of the LNAs. 2. Add probe to 1 ml fresh prehyb buffer and perform hybridization overnight at the prehybridization temperature. 3. Transfer embryos after hybridization to 6- or 12-well plates containing 15- or 24-mm Netwell Inserts, respectively, with attached 74-mM polyester mesh bottoms in 2 SSC, 0.1% Chaps prewarmed to the hybridization temperature. Prewarming the wash solutions to the hybridization temperature before washing is crucial for maximum signal-to-background ratio and is not available in some robots. Embryos in the Netwell inserts could be moved quickly into plates filled with prewarmed wash buffer, minimizing cooling for high throughput processing. 4. Wash embryos 3 20 min in the high salt wash, then 3 20 min in 0.2 SSC, 0.1% Chaps. 5. Rinse the embryos twice in KTBT and transfer back into clean 24-well plates to minimize volume for the antibody step. 6. Pretreat the embryos in 20% sheep serum in KTBT at 4 C for 2–3 h or longer. 7. Perform anti-DIG antibody binding (1:2,000–1:4,000) in 24-well plates at 4 C on a nutator. Final washes were in KTBT in large Netwell inserts at room temperature for a minimum of 5 changes over 5 h, but often including overnight at 4 C. 8. Transfer the embryos back to 24-well plates into fresh NTMT. 9. Color reactions (NBT/BCIP in NTMT) for 1–6 h at room temperature on a nutator until signal or background becomes visible, followed by overnight washing in KTBT. 10. Perform a second or third round of color reaction until each probe yields a strong signal, or until the negative control begin to show background label. 11. Stop reactions with KTBT and wash the embryos in PBS. 12. Dehydrate by a methanol series described earlier to remove background and enhance signal, then rehydrate and store in PBS plus 0.1% sodium azide.

33.2.4.4

Imaging and Histology

1. Photograph the embryos on a Leica PlanApo stereomicroscope using a digital acquisition system. 2. Transmit lateral and/or direct illumination.

33 Whole Mount In Situ Hybridization

382

Select a miRNA of interest

Collect embryos from mouse or chicken

Synthesize LNA-FISH miRNA capture probes with LNA modification and a 3'digoxigenin

Fix the embryos with 4% formaldehyde

Dehydrate the embryos with methanol

Rehydrate the embryos

Hybridization of cells with LNA-FISH probe

Treatment with anti-DIG antibody

Perform 3 runs of color reactions (NBT/BCIP in NTMT)

Dehydrate the preparation with methanol

Photograph the embryos

Fig. 33.1 Flowchart of the LNA-ELF-FISH approach for miRNA expression detection. According to Darnell et al. (2006)

3. Dehydrate some embryos into methanol (25, 50, 75, 100, 100% in PBS), and transfer to xylene (2 changes for 10 min), and embed them in paraplast. 4. Cut sections at 14 mm, mount the sections using Cytoseal XYL. 5. Photograph the sections using DIC optics on a Leica DMRXE microscope.

33.3 Application and Limitation

383

Modify the images using Adobe Photoshop only to correct brightness, contrast, color balance, and to remove particulates (Fig. 33.1).

33.2.4.5

Northern Blot Analysis

1. Heart, limb, head, and trunk tissues were dissected from stage-24 embryos, suspended in Trizol (Invitrogen Corp), and total RNA was isolated by following the manufacturer’s protocol. Forty micrograms of total RNA and 5.8 pmol of 22mer DNA control oligos were fractionated by 15% PAGE and transferred to Nytran N_membrane. Nucleic acids were UV crosslinked to membranes and baked at 80 for 30 min. Blots were pehybridized in 5_SSC, 20 mM Na2HPO4, pH 7.2, 7% SDS, 40 _g/ml yeast tRNA, and 2_Denhardt’s solution at 50 C for 2 h, followed by hybridization overnight in 25 ml of fresh hybridization solution containing 25 pmol of DIG-labeled LNA probe at 50 C. Blots were rinsed 2_in 45 ml of buffer (3_SSC, 25 nM NaH2PO4, pH 7.5, 5% SDS, and 10_Denhardt’s solution) at 25 C, followed by 30 min at 50 C, 30 min at 50 C in 1_SSC and 1% SDS, and 2_for 30 min in 0.1_SSC and 0.1% SDS at 65 C. To visualize bound probe, blots were washed briefly with 150 ml of KTBT at room temperature, then in 200 ml blocking solution (5% Skim Milk Powder, 20% sheep serum in KTBT) overnight at 4 C, then incubated in blocking solution containing a 1:2,500 dilution of anti-Digoxigenin-AP Fab Fragment (Roche) for 2 h at room temperature. Blots were washed 3_at room temperature on a nutator in KTBT, followed by two washes in NTMT. Blots were stained with BOLD APB Chemiluminescent substrate (Molecular Probes) according to the manufacturer’s protocol. Chemiluminescent signal was detected by exposing on BioMax Light X-ray film 15 min to 3 h.

33.3

Application and Limitation

The WM-ISH techniques have been tested in Drosophila, zebrafish, chicken, and mouse embryos by several laboratories (Aboobaker et al. 2005; Wienholds et al. 2005; Kloosterman et al. 2006; Darnell et al. 2006). These studies led us to tremendous insight into the spatial and temporal patterns of miRNA expression and control of embryonic development by miRNAs. The major features and advantages of the WM-ISH techniques are whole mount analysis and high-throughput profiling of miRNAs. The data from WM-ISH highlight cell-type, organ or structure-specific expression, localization within germ layers and their derivatives, and expression in multiple cell and tissue types and within subregions of structures and tissues, the information which is otherwise inaccessible with other miRNA expression detection methods.

384

33 Whole Mount In Situ Hybridization

References Aboobaker AA, Tomancak P, Patel N, Rubin GM, Lai EC (2005) Drosophila microRNAs exhibit diverse spatial expression patterns during embryonic development. Proc Natl Acad Sci USA 102:18017–18022. Bell GW, Yatskievych TA, Antin PB (2004) GEISHA, A whole-mount in situ hybridization gene expression screen in chicken embryos. Dev Dyn 229:677–687. Chang S, Johnston RJ Jr, Frøkjaer-Jensen C, Lockery S, Hobert O (2004) MicroRNAs act sequentially and asymmetrically to control chemosensory laterality in the nematode. Nature 430:785–789. Chen C-Z, Li L, Lodish HF, Bartel DP (2004) MicroRNAs modulate hematopoietic lineage differentiation. Science 303:83–86. Darnell DK, Kaur S, Stanislaw S, Konieczka JH, Yatskievych TA, Antin PB (2006) MicroRNA expression during chick embryo development. Dev Dyn 235:3156–3165. Elmen J, Thonberg H, Ljungberg K, Frieden M, Westergaard M, Xu YH, Wahren B, Liang ZC, Urum H, Koch T, Wahlestedt C (2005) Locked nucleic acid (LNA) mediated improvements in siRNA stability and functionality. Nucleic Acids Res 33:439–447. Johnston RJ, Hobert O (2003) A microRNA controlling left/right neuronal asymmetry in Caenorhabditis elegans. Nature 426:845–849. Kloosterman WP, Wienholds E, de Bruijn E, Kauppinen S, Plasterk RHA (2006) In situ detection of miRNAs in animal embryos using LNA-modified oligonucleotide probes. Nat Methods 3:27–29. Lagos-Quintana M, Rauhut R, Yalcin A, Meyer J, Lendeckel W, Tuschl T (2002) Identification of tissue-specific microRNAs from mouse. Curr Biol 12:735–739. Nieto MA, Patel K, Wilkinson DG (1996) In situ hybridization analysis of chick embryos in whole mount and tissue sections. In: Methods in cell biology. New York: Academic Press, Inc. Poy MN, Eliasson L, Krutzfeldt J, Kuwajima S, Ma XS, MacDonald PE, Pfeffer B, Tuschl T, Rajewsky N, Rorsman P, Stoffel M (2004) A pancreatic islet-specific microRNA regulates insulin secretion. Nature 432:226–230. Sokol NS, Ambros V (2005) Mesodermally expressed Drosophila microRNA-1 is regulated by Twist and is required in muscles during larval growth. Genes Dev 19:2343–2354. Wahlestedt C, Salmi P, Good L, Kela J, Johnsson T, Hokfelt T, Broberger C, Porreca F, Lai J, Ren KK, Ossipov M, Koshkin A, Jakobsen N, Skouv J, Oerum H, Jacobsen MH, Wengel J (2000) Potent and nontoxic antisense oligonucleotides containing locked nucleic acids. Proc Natl Acad Sci USA 97:5633–5638. Wienholds E, Kloosterman WP, Miska E, Alvarez-Saavedra E, Berezikov E, de Bruijn E, Horvitz HR, Kauppinen S, Plasterk RHA (2005) MicroRNA expression in zebrafish embryonic development. Science 309:310–311. Zhao Y, Samal E, Srivastava D (2005) Serum response factor regulates a muscle-specific microRNA that targets Hand2 during cardiogenesis. Nature 436:214–220.

Index

A Array fabrication, 74, 212

B Bead, 11, 40, 174, 176, 177, 180, 183, 185, 289–294, 297, 340, 342 Bioluminescence, 40, 295–302 Biomarker, 21, 28, 37–38, 131, 134, 139, 331, 332, 337, 345 Biosensor, 52, 192, 208–210, 212–214, 328

F FISH. See Fluorescence in situ hybridization (FISH) Fluorescence correlation spectroscopy (FCS), 118, 257, 258 Fluorescence in situ hybridization (FISH), 70, 104, 110, 353–358, 382 FRET oligonucleotide, 250, 251, 253–254

G Gold, 39, 40, 52, 53, 85, 99, 132, 156, 163, 164, 167, 175, 178, 192, 199–201, 203–205, 212, 214, 217–224, 276, 364

C Cancer, 20–31, 37, 38, 126, 154, 214, 263, 264, 276, 293, 302, 331, 337, 339, 340, 367 Cardiac, 11, 16, 31–34, 334, 335, 337 Cardiovascular, 31–35, 38 Cell lysis, 222–223, 245, 358 Circulating miRNAs, 38, 331, 332, 337 Conducting polymer nanowires, 207–215

E Electrocatalytic, 40, 52, 191–197, 321–328 Electrocatalytic nanoparticle tags, 191–197 End-Point Stem-Loop Real-Time RT-PCR, 131–139 ENT enzyme-labeled fluorescence (ELF), 335–336 Enzymatic miRNA detection, 267, 274

H High-throughput, 27, 39, 51, 139, 153–156, 160, 169, 200, 205, 239, 255, 281, 282, 287, 290, 294, 308 Hybridization, 27, 39, 51, 52, 68–71, 73, 76–78, 84, 85, 88–89, 91, 92, 95–99, 133, 135, 137, 191–194, 196–200, 204, 205, 208–210, 213, 217, 218, 223, 224, 230, 238, 245, 247, 258, 260–263, 267–270, 272–273, 282–284, 286, 287, 289, 290, 292, 295, 300, 301, 303, 304, 306, 308, 309, 326, 354–356, 358, 367, 369, 373

I IDT. See Integrated DNA Technologies (IDT)

385

386

In situ hybridization (ISH), 9, 10, 18, 39, 40, 97, 103–127, 247, 355, 356, 373, 377–383 Integrated DNA Technologies (IDT), 73, 124, 137, 155, 195, 202, 203, 212, 253, 259, 261, 271, 292, 317, 364 Invader miRNA, 249–252, 254–255 In vitro transcription, 97, 99, 244, 317

L Laser induced fluorescence (LIF), 257, 259 Ligation, 35, 40, 52, 160, 163–165, 167, 168, 177, 178, 184, 186, 192, 197, 214, 229–239, 241, 242, 244, 245, 264, 272, 289, 322, 327 LNA. See Locked nucleic acid (LNA) Locked nucleic acid (LNA), 10, 51, 67, 69, 70, 73, 77, 91–96, 105, 106, 116–123, 126, 133, 192, 199, 200, 202–204, 258–262, 353–358, 369–371, 373, 378, 380–383 LongSage, 176, 178, 180, 183, 184, 187

M Microarray, 9–11, 15, 17, 19, 20, 22, 24, 27–30, 33, 35–37, 39, 40, 51, 52, 67–78, 132, 180, 192, 199, 200, 203–206, 214, 239, 263, 268, 274, 282, 284–287, 290, 332, 337 miRAGE, 39, 40, 162, 173–188 miRNA amplification profiling (mRAP), 159–169, 249 miRNA serial analysis of gene expression (miRAGE/SAGE), 173–188 miR-Q RT-PCR, 141–146 Molecular beacons, 303–310, 315, 318 mRAP. See miRNA amplification profiling (mRAP) Multiplexing RT-PCR, 153–156

N Nanoparticle, 39, 40, 52, 53, 191–197, 199–206, 213, 214, 217–224, 276 Nanoparticle-amplified surface plasmon resonance imaging (Nanoparticleamplified SPRI), 192, 199–206

Index

Neuronal, 35–36 Northern blot, 10, 25, 30, 39, 40, 51, 52, 83–99, 116, 132, 148, 151, 197, 238, 239, 243–246, 254, 263, 282–285, 287, 363, 383

O Oligonucleotide capture probe (OCP), 191, 195, 196, 289 Oligonucleotides, 15, 32, 34, 51–53, 68–70, 73, 85, 91–99, 105–109, 111–112, 114, 122, 124, 126, 141–144, 155, 166, 167, 176, 191, 194, 195, 202–204, 208, 209, 219–222, 230, 232, 235–238, 243, 247, 249–255, 258, 260–261, 268, 270–274, 282, 284, 286, 289, 291, 292, 297, 299–300, 303, 314, 315, 325, 332, 354, 355, 369, 371–373, 379, 380

P Padlock-probes and rolling-circle amplification, 241–247 PCR. See Polymerase chain reaction (PCR) Peptide nucleic acid (PNA), 52, 67, 69, 208–210, 212, 213 Plasma, 18, 24, 38, 301, 326, 331–337, 339–343, 345–348 Polymerase chain reaction (PCR), 10, 12, 17, 19, 21–25, 28–30, 36–40, 51–53, 70, 97, 99, 123, 125, 126, 131–139, 141–151, 153–156, 159, 161, 164–169, 173, 174, 176–178, 180, 182–184, 187, 214, 224, 230, 237, 239, 263, 269, 272, 282, 289–292, 299, 301, 307–310, 317, 322, 332, 333, 335–337, 340, 342, 346–348, 358, 361–367, 371, 372 Poly(A)-Tailed Universal Reverse Transcription, 147–151 Probe, 19, 68, 84, 104, 134, 154, 160, 175, 191, 199, 208, 217, 238, 241–247, 249, 258, 267, 281, 289, 295, 303, 318, 325, 335, 346, 354, 363, 369, 378

Q qRT-PCR, 19, 22, 25, 28, 29, 37, 51, 52, 133, 141–146, 180, 239, 335

Index

R Real-time RT-PCR, 10, 12, 17, 23, 40, 131–139, 310, 332, 342 Ribozyme, 40, 313–319 Rluc plasmid, 299 RNA-primed, array-based Klenow enzyme (RAKE), 40, 52, 123, 281–287 RT-PCR, 25, 29, 30, 36, 38–40, 51, 52, 147, 148, 151, 153–156, 224, 263, 289, 290, 310, 332, 342, 358, 362, 367, 372

S SAGE. See Serial analysis of gene expression (SAGE) Serial analysis of gene expression (SAGE), 40, 173–188 Serum, 34, 38, 39, 107–109, 115, 164, 167, 269, 297, 301, 331–337, 379, 381, 383

387

Silver, 39, 192, 214, 217, 218, 223, 224, 276, 278, 280 Single-cell, 51, 126, 131, 133, 139, 154, 353, 354, 361–367, 369, 373 Single molecule, 40, 257–264, 270, 275, 353, 358 Single molecule detection (SMD), 40, 257, 259–260 Splinted-ligation, 229–239 Stem-loop Real-time RT-PCR, 131–139 Surface-enhanced Raman scattering (SERS), 275, 276, 278–280

W Whole mount in situ hybridization (WM-ISH), 377–383