RNAi: Design and Application [1 ed.] 9781588298744, 1588298744

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RNAi

M E T H O D S

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B I O L O G YTM

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

RNAi Design and Application

Edited by

Sailen Barik Department of Biochemistry and Molecular Biology, University of South Alabama, College of Medicine, Mobile, Alabama

Editor Sailen Barik Department of Biochemistry and Molecular Biology College of Medicine University of South Alabama Mobile, Alabama

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

ISBN: 978-1-58829-874-4 ISSN:1064-3745

e-ISBN: 978-1-59745-191-8 e-ISSN: 1940-6029

Library of Congress Control Number: 2007940759 © 2008 Humana Press, a part of Springer Science+Business Media, LLC All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Humana Press, 999 Riverview Drive, Suite 208, Totowa, NJ 07512 USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. While the advice and information in this book are believed to be true and accurate at the date of going to press, neither the authors nor the editors nor the publishers can accept any legal responsibility for any errors or omissions that may be made. The publisher makes no warranty, express or implied, with respect to the material contained herein. Cover illustration: Figure 3, Chapter 16, “Temporal Control of Gene Silencing by in ovo Electroporation” by Thomas Baeriswyl, Olivier Mauti, and Esther T. Stoeckli. Figure 3, Chapter 17, “Altering Flower Color in Transgenic Plants by RNAi-Mediated Engineering of Flavonoid Biosynthetic Pathway,” by Yoshikazu Tanaka, Noriko Nakamura, and Junichi Togami. Printed on acid-free paper 987654321 springer.com

In memory of my Mother, Promilla Barik (1932–2006)

Preface

RNA interference (RNAi), in which RNA silences RNA, is the most recent discovery to revolutionize biology and to be recognized by a Nobel Prize (in 2006, to Andrew Fire and Craig Mello). It is a story that began with historic observations in plants and fungi and eventually worked its way up to humans. If one were to describe the major steps of RNAi very briefly, it would read as follows: RNAi is triggered by double-stranded RNA (dsRNA), produced endogenously or introduced by scientists –> Long dsRNA is trimmed into short interfering RNA or microRNA (siRNA or miRNA) by Dicer –> The individual strands of the si/miRNA then guide the assembly of a multiprotein complex, known as RISC, the key constituent of which is Argonaute –> Depending on the extent of homology of the guide RNA to the target, RISC either destroys the target RNA or suppresses its translation, leading to gene silencing. The chapters in RNAi: Design and Application, contributed by leaders in the field, sum up the state-of-the-art methods on practical, everyday use of RNAi in biological research. Although multiple books and monographs have been published on RNAi, there is a noticeable dearth of bench protocols that can be used quickly and easily by beginners aspiring to enter this new field. This volume aims to fill that void. RNAi: Design and Application is divided into two parts. The first and smaller part (chapters 1–4) covers the fundamentals including designs of RNAi, biochemical assay protocols for the major components of RNAi, and study of potential off-target effects. The larger second part (chapters 5–18) covers various applications of RNAi in diverse model organisms and systems, from antiviral and anticancer applications to altering flower color in plants. Armed with this volume, a researcher with standard molecular biological training should be able to perform today’s major RNAi-related experiments and carry out gene knock-down analyses in virtually any cell line or species of interest. In the established tradition of the Methods in Molecular BiologyTM series, each chapter contains step-by-step protocols, extra notes, and problem-solving tips, which are usually not found in original research papers. As the horizon of RNAi application is rapidly broadening, we have strived to offer the most recent protocols in each area so that they remain useful for years to come.

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My sincere thanks go to all the authors and the Humana staff for bringing it all together, and to Professor John M. Walker for his guidance. I remain indebted to my wife, Kumkum, and my children, Titus and Tiasha, for their immeasurable support and encouragement. Sailen Barik

Contents

Preface . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Contributors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Part I: Designing Optimal RNAi Tools 1.

Principles of Dicer Substrate (D-siRNA) Design and Function Mohammed Amarzguioui and John J. Rossi . . . . . . . . . . . . . . . . . . . . . . . .

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2.

Expression, Purification, and Analysis of Recombinant Drosophila Dicer-1 and Dicer-2 Enzymes Xuecheng Ye and Qinghua Liu . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11

3.

In vitro RNA Cleavage Assay for Argonaute-Family Proteins Keita Miyoshi, Hiroshi Uejima, Tomoko Nagami-Okada, Haruhiko Siomi, and Mikiko C. Siomi . . . . . . . . . . . . . . . . . . . . . . . . . . . 29

4.

Identifying siRNA-Induced Off-Targets by Microarray Analysis Emily Anderson, Queta Boese, Anastasia Khvorova, and Jon Karpilow . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 45

Part II:

Application of RNAi in Diverse Organisms

5.

Hydrodynamic Delivery of siRNA in a Mouse Model of Sepsis Doreen E. Wesche-Soldato, Joanne Lomas-Neira, Mario Perl, Chun-Shiang Chung, and Alfred Ayala . . . . . . . . . . . . . . . . . . . . . . . . . . 67

6.

Nasal Delivery of siRNA Vira Bitko and Sailen Barik . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 75

7.

RNA Interference as a Genetic Tool in Trypanosomes Vivian Bellofatto and Jennifer B. Palenchar . . . . . . . . . . . . . . . . . . . . . . . . 83

8.

Lentivirus-Mediated RNA Interference in Mammalian Neurons Scott Q. Harper and Pedro Gonzalez-Alegre . . . . . . . . . . . . . . . . . . . . . . 95

9.

Silencing Genes by RNA Interference in the Protozoan Parasite Entamoeba histolytica Carlos F. Solis and Nancy Guillén . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 113

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Contents 10. Use of RNAi in C. elegans Tsuyoshi Ohkumo, Chikahide Masutani, Toshihiko Eki, and Fumio Hanaoka . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 129 11. Application of siRNA Against SARS in the Rhesus Macaque Model Qingquan Tang, Baojian Li, Martin Woodle, and Patrick Y. Lu . . . . . 139 12. siRNA and shRNA as Anticancer Agents in a Cervical Cancer Model Wenyi Gu, Lisa Putral, and Nigel McMillan . . . . . . . . . . . . . . . . . . . . . . . . 159 13. Identification and Expression Analysis of Small RNAs During Development Toshiaki Watanabe, Hiroshi Imai, and Naojiro Minami. . . . . . . . . . . . . 173 14. Screening and Identification of Virus-Encoded RNA Silencing Suppressors Sumona Karjee, Mohammad Nurul Islam, and Sunil K. Mukherjee . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 187 15. Application of RNA Interference in Functional Genomics Studies of a Social Insect Michael E. Scharf, Xuguo Zhou, and Margaret A. Schwinghammer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 205 16. Temporal Control of Gene Silencing by in ovo Electroporation Thomas Baeriswyl, Olivier Mauti, and Esther T. Stoeckli . . . . . . . . . . . 231 17. Altering Flower Color in Transgenic Plants by RNAi-Mediated Engineering of Flavonoid Biosynthetic Pathway Yoshikazu Tanaka, Noriko Nakamura, and Junichi Togami . . . . . . . . . 245 18. Transgenic RNA Interference in Mice Pumin Zhang . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 259

Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 267

Contributors

Mohammed Amarzguioui • The Biotechnology Centre of Oslo, Oslo, Norway Emily Anderson • Dharmacon, ThermoFisher Scientific, Lafayette, CO Alfred Ayala • Division of Surgical Research, Department of Surgery, Rhode Island Hospital/Brown University School of Medicine, Providence, RI Thomas Baeriswyl • Institute of Zoology, University of Zürich, Zürich, Switzerland Sailen Barik • Department of Biochemistry and Molecular Biology, University of South Alabama, College of Medicine, Mobile, AL Vivian Bellofatto • Department of Microbiology and Molecular Genetics, UMDNJ-NJ Medical School, International Center for Public Health, Newark, NJ Vira Bitko • Department of Biochemistry and Molecular Biology, University of South Alabama, College of Medicine, Mobile, AL Queta Boese • Dharmacon, ThermoFisher Scientific, Lafayette, CO Chun-Shiang Chung • Division of Surgical Research, Department of Surgery, Rhode Island Hospital/Brown University School of Medicine, Providence, RI Toshihiko Eki • Department of Ecological Engineering, Toyohashi University of Technology, Toyohashi, Japan Pedro Gonzalez-Alegre • Department of Neurology, Carver College of Medicine at The University of Iowa, Iowa City, IA Wenyi Gu • Cancer Biology Program, Centre for Immunology and Cancer Research, Princess Alexandra Hospital, University of Queensland, Brisbane, Australia Nancy Guillén • Unité de Biologie Cellulaire du Parasitisme, Institut Pasteur, Paris, France Fumio Hanaoka • Graduate School of Frontier Biosciences, Osaka University, and SORST, Japan Science and Technology Agency, Osaka, Japan

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Contributors

Scott Q. Harper • Center for Gene Therapy, Department of Pediatrics, The Ohio State University, Columbus, OH Hiroshi Imai • Laboratory of Reproductive Biology, Department of Agriculture, Kyoto University, Kyoto, Japan Mohammad Nurul Islam • International Center for Genetic Engineering and Biotechnology, PMB Lab, New Delhi, India Sumona Karjee • International Center for Genetic Engineering and Biotechnology, PMB Lab, New Delhi, India Jon Karpilow • Dharmacon, ThermoFisher Scientific, Lafayette, CO Anastasia Khvorova • Dharmacon, ThermoFisher Scientific, Lafayette, CO Baojian Li • Top Genomics, Ltd., and College of Life Sciences, Sun Yat-sen University, Guangzhou, China Qinghua Liu • Department of Biochemistry, University of Texas Southwestern Medical Center, Dallas, TX Joanne Lomas-Neira • Division of Surgical Research, Department of Surgery, Rhode Island Hospital/Brown University School of Medicine, Providence, RI Patrick Y. Lu • Sirnaomics, Inc., Rockville, MD Chikahide Masutani • Graduate School of Frontier Biosciences, Osaka University; and SORST, Japan Science and Technology Agency, Osaka, Japan Olivier Mauti • Institute of Zoology, University of Zürich, Zürich, Switzerland Nigel McMillan • Cancer Biology Program, Centre for Immunology and Cancer Research, Princess Alexandra Hospital, University of Queensland, Brisbane, Australia Naojiro Minami • Laboratory of Reproductive Biology, Department of Agriculture, Kyoto University, Kyoto, Japan Keita Miyoshi • Institute for Genome Research, University of Tokushima, Tokushima, Japan Sunil K. Mukherjee • International Center for Genetic Engineering and Biotechnology, PMB Lab, New Delhi, India Tomoko Nagami-Okada • Institute for Genome Research, University of Tokushima, Tokushima, Japan Noriko Nakamura • Institute for Advanced Core Technology, Suntory Ltd., Osaka, Japan Tsuyoshi Ohkumo • Graduate School of Frontier Biosciences, Osaka University; and SORST, Japan Science and Technology Agency, Osaka, Japan

Contributors

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Jennifer B. Palenchar • Department of Chemistry, Villanova University, Villanova, PA Mario Perl • Universitätsklinikum Ulm, Zentrum für Chirurgie, Klinik für Unfallchirurgie, Hand-, Plastische- und Wiederherstellungschirurgie, Ulm, Germany Lisa Putral • Cancer Biology Program, Centre for Immunology and Cancer Research, Princess Alexandra Hospital, University of Queensland, Brisbane, Australia John J. Rossi • Division of Molecular Biology, Beckman Research Institute of the City of Hope, Duarte, CA Michael E. Scharf • Molecular and Applied Insect Toxicology, Entomology and Nematology Department, University of Florida, Gainesville, FL Margaret A. Schwinghammer • Department of Entomology, Purdue University, West Lafayette, IN Haruhiko Siomi • Keio University School of Medicine, Tokyo, Japan Mikiko C. Siomi • Institute for Genome Research, University of Tokushima, JST, CREST, Tokushima, Japan Carlos F. Solis • Unité de Biologie Cellulaire du Parasitisme, Institut Pasteur, Paris, France Esther T. Stoeckli • Institute of Zoology, University of Zürich, Zürich, Switzerland Yoshikazu Tanaka • Institute for Advanced Core Technology, Suntory Ltd., Osaka, Japan Qingquan Tang • OriGene Technologies, Inc., Rockville, MD Junichi Togami • Institute for Advanced Core Technology, Suntory Ltd., Osaka, Japan Hiroshi Uejima • Institute for Genome Research, University of Tokushima, Tokushima, Japan Toshiaki Watanabe • Division of Human Genetics, Department of Integrated Genetics, National Institute of Genetics, Research Organization of Information and Systems; and Department of Genetics, School of Life Science, The Graduate University for Advanced Studies (SOKENDAI), Mishima, Japan Doreen E. Wesche-Soldato • Division of Surgical Research, Department of Surgery, Rhode Island Hospital/Brown University School of Medicine, Providence, RI Martin Woodle • Nanotides Pharmaceuticals, Inc., Rockville, MD Xuecheng Ye • Department of Biochemistry, University of Texas Southwestern Medical Center, Dallas, TX

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Contributors

Pumin Zhang • Department of Molecular Physiology and Biophysics, Baylor College of Medicine, Houston, TX Xuguo Zhou • Molecular and Applied Insect Toxicology, Entomology and Nematology Department, University of Florida, Gainesville, FL

I Designing Optimal RNAi Tools

1 Principles of Dicer Substrate (D-siRNA) Design and Function Mohammed Amarzguioui and John J. Rossi

Summary An efficient RNAi largely depends on optimal design of the siRNA. In recent studies, Dicer substrates were found to be more potent than classical synthetic 21-mer siRNAs, suggesting a coupling of the Dicer-mediated processing step to the efficient assembly of the silencing complex, RISC. We describe the fundamental principles and experimental results leading to optimal Dicer substrates.

Key Words: siRNA; shRNA; RNAi; Dicer.

1. Introduction The formal description of RNAi as a biological response to double-stranded RNA resulted from a desire to understand a number of intriguing observations arising from the use of antisense RNAs in C. elegans (1). This ultimately led to the discovery that worms could be programmed to silence genes by exposing animals to homologous dsRNAs (termed triggers) (1). It is now clear that an RNAi pathway is present in many if not most eukaryotes (2). A biochemical understanding of the RNAi pathway was crucial to realizing that dsRNAs shorter than 30 bp could be used to specifically trigger an RNAi response in mammals. Tuschl and colleagues demonstrated that transfection of mammalian cells with small interfering RNAs (siRNAs) could specifically induce RNAi and thus cracked the barrier to the use of RNAi as a genetic tool in mammals (3). It took a remarkably short period of time for siRNAs to be adopted as a standard component of the molecular biology toolkit. From: Methods in Molecular Biology, vol. 442: RNAi: Design and Application Edited by: S. Barik © Humana Press, Totowa, NJ

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The introduction of siRNAs into mammalian cells can be achieved through standard transfection. The strength and duration of the silencing response are determined by several factors. On a population basis, the overall efficiency of transfection is a major determinant, which must be addressed by optimizing conditions. In each individual cell, silencing depends upon a combination of the amount of siRNA that is delivered and upon the potential of that individual siRNA to suppress its target (the potency). Even a relatively poor siRNA can silence its target provided that sufficient quantities are delivered. However, essentially “forcing” the system with such reagents is likely to lead to numerous undesired effects (4). Most investigators utilize siRNAs that consist of 19-base duplexes and 2-base 3′ -overhangs on each strand. Our laboratory and that of Greg Hannon have both reported improved efficacy of longer-than-standard RNAi effectors. During investigation of cellular interferon induction caused by in vitro transcribed siRNAs, it was observed that at limiting concentrations some siRNAs of length 25–27 appeared to have greater potency than 21-mer siRNAs that could potentially be generated from the larger duplex (5). Hannon and colleagues reported a similar phenomenon for shRNA (6). They found synthetic shRNAs with 29base-pair stems and 2-nucleotide 3′ overhangs to be more potent inducers of RNAi than shorter hairpins (6). Their studies further demonstrated that in vitro processing by Dicer was directional, starting predominantly from the open end of the stem and generating a mixture of 21- and 22-mer cleavage products. In both above cases, the increased potency can confidently be attributed to Dicer processing, which is thought to promote more efficient incorporation into RISC through the physical association of Dicer with the effectors of RNAi, the Argonaute proteins. This interpretation is supported by biochemical evidence in the fruit fly indicating a role for Dicer in the initial stages of RISC assembly (7) and by recent reports that Dicer-mediated processing of miRNA precursors in human cells is functionally coupled to miRISC assembly and improves subsequent silencing (8,9).

2. Materials 1. An algorithm for the prediction of relevant parameters of siRNA such as secondary structure, thermodynamic property, etc. (See the algorithm provided in Section 3). 2. In-house RNA synthetic facility or a commercial source (such as Dharmacon, Ambion, IDT, Sigma-Proligo, etc.) for obtaining siRNA of various lengths and compositions. 3. An assay for siRNA function, primarily a cultured cell line with a target gene. 4. If stable knockdown is desired, express the Dicer substrate from recombinant lentiviral or other suitable vectors (see Chapter 8, for example).

Dicer Substrate Design

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3. Methods 3.1. Design of D-siRNAs While Dicer processing is generally beneficial, the composition and potency of the processing products are also of importance for the overall efficacy. Dicer processing of unmodified 27-mer duplexes is largely unpredictable, sometimes resulting in the generation of siRNAs of poor activity, thereby reducing the activity of the 27-mer to below that of an optimal 21-mer within its target sequence. Consequently, there is no guarantee that a randomly designed 27mer will be more efficacious than the best of the potential 21-mers within its target sequence. The problem of making the processing of Dicer substrate siRNA (D-siRNA) predictable, thereby enabling rational design on the basis of published design algorithms, now appears to have been solved. The new optimal design introduces directionality and uniqueness of processing into the Dicer cleavage step by mimicking the relevant structural features of naturally occurring Dicer substrates, pre-miRNAs. These can be summarized as follows. 1. To select the D-siRNA sequence, identify potentially good siRNA/target combinations using a computer-based algorithm. We have one that analyzes all the possible 21-mer targets in a given mRNA for secondary structure, GC content, and thermodynamic end properties of the 21-mer siRNAs that would be used to target the sequence. The target/siRNAs are ranked top to bottom by a numerical value (13). Access to this algorithm is free using the following Web address: http:// www.cityofhope.org/researchers/RossiJohn/RossiJohnResearch.htm. Provide an E-mail address to which the output, which can be accessed by Microsoft Excel, will be returned. 2. Have bulged stem-loop structures with 2-nt 3′ -overhangs. Recent reports (6,10) suggest that the overhangs in the open end of the stem in such structures are bound by Dicer and determine the direction of processing as well as preferential strand selection. 3. When synthesizing a linear dsRNA substrate, make one end of the duplex bluntended to mimic a natural Dicer substrate in which the corresponding end of the duplex is closed by a loop, precluding binding of Dicer to that end. In addition, this feature introduces two DNA nucleotides in the sense strand in the blunt end of the duplex (Fig. 1) (11). The incorporation of a 3′ -overhang in one end introduces a preference for processing to start from that end, while DNA nucleotides in the opposite blunt end enforce this asymmetry while blocking processing events involving the terminal two phosphodiester linkages. This results in the predictable production of a single or major 21-mer product of processing starting from the overhang terminus, sometimes accompanied by a minor 22-mer product resulting from processing from the same end. This mixture is similar to that previously reported to result from Dicer cleavage (6,12), and this flexibility in Dicer processing may reflect some level of sequence preference near the putative cleavage site.

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Fig. 1. D-siRNA conformations. Dicer entry is from the 2-base 3′ -overhang. N = ribonucleic acid; d = deoxyribonucleic acid. Arrows point to Dicer cleavage sites, and italicized bases are the 21-mer siRNAs produced. 4. The same 21-mer can be generated from Dicer substrate siRNAs of slightly different sequence and opposite “polarity”: one in which the passenger strand carries the 3′ overhang and processing proceeds from right to left (L-form), and the other in which the overhang is on the guide strand, and processing proceeds from left to right (R-form) (Fig. 1). Interestingly, although processing of the two forms of D-siRNA produces the same 21-mer siRNA species (confirmed by mass spectrometry data; see Fig. 2), when considering that the strand with the 2-base 3′ overhang is antisense to the target mRNA, the “R”-versions are consistently more efficacious than the “L”-forms. This comparison was performed for nine different pairs of D-siRNA, targeting four different genes. In seven out of nine cases, the R-form was superior to the L-form (11). We hypothesize that preferential binding to the 3′ -overhang by Dicer during processing favors incorporation of the strand bearing the overhang. Thus, sense target silencing is more efficient with the R-form because that’s the configuration in which the guide strand bears the overhang. This effect can be demonstrated experimentally by cotransfection experiments with reporters in which the two forms of D-siRNA (R and L) are cotransfected with a target gene (Fig. 2). Silencing of the reporter is markedly better when the R-form is used even though the Mass Spec-determined siRNAs are virtually the same. Thus, the polarity of the Dicer entry is an important determinant for the selection of the guide strand, which is generally the strand with the 2-base 3′ -overhang. This polarity also has the advantage of lower off-target possibilities by the passenger strand incorporation into RISC. 5. Due to variable efficacy of even rationally designed siRNAs, it is advisable to design multiple D-siRNAs targeting different sites and to titer their concentration to determine the optimal sequences and concentration for adequate silencing. Irrelevant control siRNA(s) should be included at all concentrations tested. 6. Avoid nonspecific and off-target effects (see Notes 1 and 2): Published data suggest that although near-complete inactivation of siRNA by a single mutation is possible, multiple mutations are generally necessary to ensure that the siRNA will be inactive (14–18). When targeting the 3′ UTR of a transcript, it is also important to consider the possibility that mismatches with the target may abrogate cleavage but can still function in translational repression. Although sequence context, mismatch type, and mismatch position all influence the impact of mutations

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Fig. 2. Polarity of D-siRNA dictates efficacy. An EGFP target was cotransfected into HEK293 cells with either an L-form D-siRNA (#2) or an R-form (#3). The deoxynucleotides (“tt” in #2 and “cg” in #3) are in lowercase. The knockdown efficacy at two different concentrations of D-siRNAs was determined. Note that the R-form is the most potent for this target despite the fact that Dicer produces an identical bottom strand in both cases. The polarity of the Dicer entry is therefore an important determinant for guide strand selection into RISC. (14,15,19), making it difficult to devise clear rules, some general guidelines can be formulated: (a) Ideally, the selected siRNA should have multiple mismatches to all nontarget mRNA sequences. (b) Mismatches located near the cleavage site or within the seed region (positions 2–11 within the putative guide strand and 9–18 within the target) are more disruptive than mismatches within the 5′ end of the target site. Full match within positions 9–18 of the “off-targets” should be avoided even if there are multiple mismatches in the 5′ end. (c) Extensive matches to the self-complement of the target sequence are less critical since they would be targeted by the passenger strand, whose incorporation into RISC is designed to be minimal. For these sequences, the duplex region mismatch sensitivity is reversed (mismatches within positions 2–11 of the query are more critical).

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Amarzguioui and Rossi 7. Delivery of D-siRNA: Formation of lipid-siRNA or D-siRNA complexes should be performed in one batch for all different treatments of the same siRNA, and the complexes diluted appropriately to their final concentration (17). Adherent cells can be transfected either while adherent or in suspension after trypsinmediated detachment (26). The latter procedure is recommended due to its greater flexibility and robustness (confluency of cells is not an issue). For difficultto-transfect adherent cells, the methodology is frequently also associated with improved silencing (26). 8. Summary (see Note 3): Based on the aforesaid, once the optimal 21-mers are chosen using an algorithm, extend the passenger strand to 25 bases and make the last two nucleotides deoxys. Similarly, extend the guide strand to 27 nucleotides with (Fig. 1). Deliver the final D-siRNA using cationic lipids or electroporation and assay for loss of target. Screen various amounts of each D-siRNA for the most efficient knockdown.

4. Notes 1. Off-target effects: siRNA specificity determinations have traditionally been performed using BLAST searches. Recent data, however, have cast serious doubt on the value of BLAST searches for general siRNA specificity determination (19). Experimentally determined siRNA off-target effects were shown to correlate strongly with matches between positions 2–8 within the guide strand (the seed sequence) and sequences in the 3′ UTRs of affected genes (19). These seed matches are too short to be confidently detected by BLAST. These searches are therefore only useful for identifying near-perfect matches. A Web-based search tool is available for identification of all possible seed matches for any given siRNA (http://www.dharmacon.com/seedlocator/default.aspx). Despite the significant correlation of seed matches with off-targets, the predictive value of such a list is at present limited since only a small fraction of seed matches results in actual off-target effects. Thus, while BLAST searches can be used to select away the poorest candidates, and seed match searches can help in deciding among sequences on the basis of the number of potential off-targets, there appears for the foreseeable future no substitute for experimental determination of specificity, preferably by genome-wide gene expression profiling. Thus, for functional genomic studies, verification of phenotypes by the application of a combination of multiple active, target-specific siRNAs as well as inactivated or irrelevant control sequences is of paramount importance. Since D-siRNAs are processed to predictable 21- and 22-mer sequences, the off-target effects are not expected to be substantially different from those resulting from their corresponding 21-mers. 2. Activation of innate immune response: An emerging property of siRNA is their potentially immune-stimulatory effects in vitro and in vivo as a result of engaging members of the Toll-like receptor family following liposome-mediated endosomal trafficking (20–24). No induction of interferon or activation of PKR was observed in HEK293 cells following delivery of 27-mer siRNA (5). It should be pointed

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out that in some cells the D-siRNAs have been shown to activate the antiviral protein RIG1. This activation appears to be eliminated (25) by the design shown in Figs. 1 and 2. 3. The design and application of D-siRNAs do not differ significantly from the standard 21-mer siRNAs. The advantages of the D-siRNA design described above are better selectivity of the guide strand as a consequence of Dicer processing and handoff to RISC and, in many instances, improved potency, which translates into lower effective concentrations. Other potential advantages of D-siRNAs are that they can be physically linked to cell-specific delivery ligands such as aptamers (27) or other nonnucleic acid compounds, and the siRNAs can be processed from the longer molecules via interaction of Dicer with the 2-base 3′ -overhang in the D-siRNA design.

References 1. Fire, A., Xu, S., Montgomery, M. K., Kostas, S. A., Driver, S. E., and Mello, C. C. (1998). Potent and specific genetic interference by double-stranded RNA in Caenorhabditis elegans. Nature 391, 806–811. 2. Hannon, G. J., and Rossi, J. J. (2004). Unlocking the potential of the human genome with RNA interference. Nature 431, 371–378. 3. Elbashir, S. M., Harborth, J., Lendeckel, W., Yalcin, A., Weber, K., and Tuschl, T. (2001). Duplexes of 21-nucleotide RNAs mediate RNA interference in cultured mammalian cells. Nature 411, 494–498. 4. Jackson, A. L., Bartz, S. R., Schelter, J., et al. (2003). Expression profiling reveals off-target gene regulation by RNAi. Nat. Biotechnol. 21, 635–637. 5. Kim, D. H., Behlke, M. A., Rose, S. D., Chang, M. S., Choi, S., and Rossi, J. J. (2005). Synthetic dsRNA Dicer substrates enhance RNAi potency and efficacy. Nat. Biotechnol. 23, 222–226. 6. Siolas, D., Lerner, C., Burchard, J., et al. (2005). Synthetic shRNAs as potent RNAi triggers. Nat. Biotechnol. 23, 227–231. 7. Pham, J. W., Pellino, J. L., Lee, Y. S., Carthew, R. W., and Sontheimer, E. J. (2004). A Dicer-2-dependent 80s complex cleaves targeted mRNAs during RNAi in Drosophila. Cell 117, 83–94. 8. Gregory, R. I., Chendrimada, T. P., Cooch, N., and Shiekhattar, R. (2005). Human RISC couples microRNA biogenesis and posttranscriptional gene silencing. Cell 123, 631–640. 9. Maniataki, E., and Mourelatos, Z. (2005). A human, ATP-independent, RISC assembly machine fueled by pre-miRNA. Genes Dev. 19, 2979–2990. 10. Krol, J., Sobczak, K., Wilczynska, U., et al. (2004). Structural features of microRNA (miRNA) precursors and their relevance to miRNA biogenesis and small interfering RNA/short hairpin RNA design. J. Biol. Chem. 279, 42230–42239. 11. Rose, S. D., Kim, D. H., Amarzguioui, M., et al. (2005). Functional polarity is introduced by Dicer processing of short substrate RNAs. Nucleic Acids Res. 33, 4140–4156.

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12. Elbashir, S. M., Lendeckel, W., and Tuschl, T. (2001). RNA interference is mediated by 21- and 22-nucleotide RNAs. Genes Dev. 15, 188–200. 13. Heale, B. S., Soifer, H. S., Bowers, C., and Rossi, J. J. (2005). siRNA target site secondary structure predictions using local stable substructures. Nucleic Acids Res. 33, e30. 14. Amarzguioui, M., Holen, T., Babaie, E., and Prydz, H. (2003). Tolerance for mutations and chemical modifications in a siRNA. Nucleic Acids Res. 31, 589–595. 15. Du, Q., Thonberg, H., Wang, J., Wahlestedt, C., and Liang, Z. (2005). A systematic analysis of the silencing effects of an active siRNA at all single-nucleotide mismatched target sites. Nucleic Acids Res. 33, 1671–1677. 16. Elbashir, S. M., Martinez, J., Patkaniowska, A., Lendeckel, W., and Tuschl, T. (2001). Functional anatomy of siRNAs for mediating efficient RNAi in Drosophila melanogaster embryo lysate. EMBO J. 20, 6877–6888. 17. Holen, T., Amarzguioui, M., Wiiger, M. T., Babaie, E., and Prydz, H. (2002). Positional effects of short interfering RNAs targeting the human coagulation trigger Tissue Factor. Nucleic Acids Res. 30, 1757–1766. 18. Miller, V. M., Gouvion, C. M., Davidson, B. L., and Paulson, H. L. (2004). Targeting Alzheimer’s disease genes with RNA interference: An efficient strategy for silencing mutant alleles. Nucleic Acids Res. 32, 661–668. 19. Birmingham, A., Anderson, E. M., Reynolds, A., et al. (2006). 3′ UTR seed matches, but not overall identity, are associated with RNAi off-targets. Nat. Meth. 3, 199–204. 20. Hornung, V., Guenthner-Biller, M., Bourquin, C., et al. (2005). Sequence-specific potent induction of IFN- by short interfering RNA in plasmacytoid dendritic cells through TLR7. Nat. Med. 11, 263–270. 21. Judge, A. D., Sood, V., Shaw, J. R., Fang, D., McClintock, K., and MacLachlan, I. (2005). Sequence-dependent stimulation of the mammalian innate immune response by synthetic siRNA. Nat. Biotechnol. 23, 457–562. 22. Kariko, K., Bhuyan, P., Capodici, J., and Weissman, D. (2004). Small interfering RNAsmediate sequence-independent gene suppression and induce immune activation by signaling through Toll-like receptor 3. J. Immunol. 172, 6545–6549. 23. Robbins, M. A., and Rossi, J. J. (2005). Sensing the danger in RNA. Nat. Med. 11, 250–251. 24. Sioud, M. (2005). Induction of inflammatory cytokines and interferon responses by double- stranded and single-stranded siRNAs is sequence-dependent and requires endosomal localization. J. Mol. Biol. 348, 1079–1090. 25. Marques, J. T., Devosse, T., Wang, D., et al. (2006). A structural basis for discriminating between self and nonself double-stranded RNAs in mammalian cells. Nat. Biotechnol. 24, 559–565. 26. Amarzguioui, M. (2004). Improved siRNA-mediated silencing in refractory adherent cell lines by detachment and transfection in suspension. Biotechniques 36, 766–768, 770. 27. Rossi, J. J. (2006). Partnering aptamer and RNAi technologies. Mol. Ther. 14, 461–462.

2 Expression, Purification, and Analysis of Recombinant Drosophila Dicer-1 and Dicer-2 Enzymes Xuecheng Ye and Qinghua Liu

Summary RNA interference (RNAi) is a form of posttranscriptional gene silencing mediated by microRNA (miRNA) and small interfering RNA (siRNA). In Drosophila melanogaster, the RNase III enzymes Dicer-1 and Dicer-2 generate miRNA and siRNA, respectively. We describe the methods for the expression, purification, and analysis of recombinant Dicer-1 and Dicer-2 enzymes. Our studies demonstrate that Dicer-1 and Dicer-2 display different substrate specificities and ATP requirements.

Key Words: RNA interference; Dicer-1; Dicer-2; baculovirus; dsRNA; siRNA; pre-miRNA; miRNA. 1. Introduction RNA interference (RNAi) is a form of posttranscriptional gene silencing mediated by 21- to 25-nucleotide microRNA (miRNA) and small interfering RNA (siRNA). These tiny regulatory RNAs control many important biological processes, such as development, antiviral defense, heterochromatin formation, and maintenance of genomic stability (1–4). It follows that misregulation of miRNAs has been linked to human diseases, such as cancer (5,6). Moreover, siRNAs or miRNAs have been widely used as a powerful gene-silencing tool for functional genomic analyses in multiple model organisms, including humans (1,3). In principle, miRNAs and siRNAs can be viewed as two parallel branches of the RNAi pathway, which consists of initiation and effector steps. In the initiation step, siRNAs are derived from mostly exogenous long dsRNAs, From: Methods in Molecular Biology, vol. 442: RNAi: Design and Application Edited by: S. Barik © Humana Press, Totowa, NJ

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whereas miRNAs originate from endogenous (∼ 60–70 nt) stem-loop precursor miRNAs (pre-miRNAs). Both siRNAs and miRNAs are produced as 21to 25-nt duplexes with 2-nt 3′ -overhangs and 5′ -phosphate and 3′ -hydroxyl termini. While siRNAs are perfectly complementary, miRNAs frequently contain mismatches, bulges, or G:U wobble base pairs. In the effector step, nascent siRNA and miRNA duplexes are assembled into the respective RNAinduced silencing complexes that are simply referred to as siRISC and miRISC. In RISCs, single-stranded siRNA or miRNA functions as the guide RNA to direct sequence-specific cleavage and/or translational repression of complementary mRNA (1,2,7,8). Both siRNAs and miRNAs are produced by Dicers, a family of large (∼ 200 kDa) multidomain RNaseIII enzymes that exist in most eukaryotic organisms (9). A typical Dicer contains a putative RNA helicase domain, a DUF (domain of unknown function) 283 domain, and a PAZ domain at the amino (N)-terminus as well as two RNase III domains and a dsRNA-binding domain at the carboxyl (C)-terminus. The tandem RNase III domains of Dicer form an intramolecular dimer to make a pair of cuts on dsRNA, creating a characteristic 2-nt 3′ -overhang (10,11). In most organisms, such as C. elegans and humans, a single Dicer generates both siRNAs and miRNAs. However, two Dicer enzymes, Dicer-1 and Dicer-2, have been identified in the Drosophila genome (10,12,13). Both genetic and biochemical studies have implicated that Dicer-1 and Dicer-2 are involved in the biogenesis of miRNAs and siRNAs, respectively (14–18). Dicer enzymes do not function alone. We have previously purified the siRNA-generating enzyme from Drosophila S2 cells and found that it consisted of Dicer-2 and R2D2 (15). R2D2 contains two dsRNA-binding domains (R2) and forms a heterodimeric complex with Dicer-2 (D2). Although R2D2 does not regulate siRNA production, R2D2 and Dicer-2 coordinately bind siRNA and facilitate its incorporation into the effector siRISC complex (15,19). An R2D2-like protein, Loquacious (Loqs, also known as R3D1), has recently been identified as a cofactor for Dicer-1 in the miRNA pathway. The loqs gene encodes at least two alternatively spliced proteins, Loqs-L (long) and Loqs-S (short). Both Loqs isoforms contain three putative dsRNA-binding domains. Loqs-L forms a stable complex with Dicer-1 and greatly enhances its miRNAgenerating activity. It remains uncertain if the Dicer-1/Loqs complex facilitates miRNA loading onto the miRISC complex (16–18). Therefore, the Dicer-1/Loqs-L and Dicer-2/R2D2 complexes function in parallel to generate miRNAs and siRNAs in Drosophila cells. The functional differences of the two Dicer complexes can be explained by either that Dicer-1 and Dicer-2 possess distinct biochemical activities or that Loqs-L and R2D2 help define the functional specificities for Dicer-1 and Dicer-2 (16). Here we

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describe the detailed protocols for the expression, purification, and analysis of recombinant Dicer-1 and Dicer-2 enzymes. Our reconstitution studies indicate that Dicer-1 and Dicer-2 are biochemically distinct enzymes with different substrate specificities and ATP requirements. 2. Materials 2.1. For Expression of Dicer 1. The BAC-to-BAC® Baculovirus Expression System (Invitrogen, Carlsbad, CA, Cat# 10359-016), which includes the pFastBacTM 1 vector, E. coli DH10 Bac competent cells, Cellfectin transfection reagent, and an instruction manual. 2. Recombinant Dicer bacmid DNA, prepared from 2 mL of bacteria culture, and resuspended in 40 μL of sterile TE. 3. Complete IPL-41 medium: 1 L of incomplete IPL-41 medium (Invitrogen Cat# 11405-081) supplemented with 100 mL of fetal bovine serum (Gemini BioProducts, West Sacramento, CA, Cat# 100-106, heat-inactivated at 55 ºC for 30 min), 10 mL of Pluronic®-F68 (Invitrogen, Cat# 24040-032), 10 mL of 100X Antibiotic-antimycotic (Invitrogen, Cat# 15240-062), 20 mL of 50X Tryptose Phosphate Broth (130 mg/mL, powder is ordered from Sigma, St. Louis, MO, Cat# T9157), and 20 mL of 50X Yeastolate Ultrafiltrate (Invitrogen, Cat# 18200-048). 4. Buffers and reagents for SDS-Polyacrylamide gel electrophoresis (PAGE): 4X SDS sample buffer (0.25 M Tris-HCl pH 6.8, 8% SDS, 30% glycerol, 0.02% bromophenol blue, and 10% -mercaptoethanol); 10X running buffer (250 mM Tris, 1.92 M glycine, and 1% SDS; do not adjust pH). 5. Buffers and reagents for Western blot: 10X transfer buffer (312.5 mM Tris, 2.4 M glycine); rabbit anti-His antibody (Bethyl, Montgomery, TX, Cat#A190-114A); HRP-conjugated anti-rabbit IgG antibody (Sigma, Cat# A6154); ECL reagents (PerkinElmer, Waltham, MA, Cat# NEL102).

2.2. For Purification of Dicer 1. Preparation of cell lysates: Buffer A [10 mM KOAc, 10 mM HEPES, pH 7.4, 2 mM Mg(OAc)2 , 5 mM -mercaptoethanol]; Buffer B (buffer A plus 1 M NaCl); Protease inhibitors, including Pefabloc SC (Roche, Indianapolis, IN, Cat# 1-585916), Leupeptin (Roche, Cat# 1-034-626), and Pepstatin (Roche, Indianapolis, IN, Cat# 1-359-053). Store 1,000X stocks (1 mg/mL Pefabloc SC, 5 mg/mL Leupeptin, and 0.7 mg/mL Pepstain) at –20 ºC in small aliquots. 2. Nickel purification: Ni-NTA Agarose beads (Qiagen, Valencia, CA, Cat# 30230); 50% slurry in ethanol, store at 4 ºC; Imidazole (Sigma), 2.5 M of stock dissolved in buffer A; Econo-Pac® disposable chromatography column (Bio-Rad, Hercules, CA, Cat# 732-1010). 3. Ion-exchange chromatography: 1 mL of SP-Sepharose column (GE Healthcare, Piscataway, NJ, Cat# 17-1151-01) and 1 mL of Q-Sepharose column (GE Healthcare, Piscataway, NJ, Cat# 17-1153-01).

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4. Recombinant protein stocking: Glycerol (Sigma) and liquid nitrogen. 5. Buffers and reagents for SDS-PAGE and Coomassie blue staining: 4X SDS sample loading buffer and 10X running buffer (see Section 2.1). Coomassie blue staining solution (Bio-Rad, Cat# 161-0436) and destaining solution (10% methanol and 10% acetic acid in water).

2.3. For Assay of Dicer 1. Templates for in vitro transcription of dsRNA: The sense and antisense (250 bp) templates were generated from the firefly luciferase gene by PCR using primers 5′ -GCGTAATACGACTCACTATAGATGCACATATCGAGGTGGA-3′ and 5′ -GAACACCACGGTAGGCTGC-3′ (sense template); 5′ -GATGCACATATCGAGGTGGA-3′ and 5′ -GCTAATACGACTCACTATAGAACACCACGGTAGGCTGC-3′ (antisense template). The two long primers contain T7 polymerase promoter sequence. 2. Synthetic RNA oligonucleotides: 61 nt pre-let7, 5′ -UGAGGUAGUAGGUUGUAUAGUAGUAAUUACACAUCAUACUAUACAAUGUGCUAGCUUUCUU-3′ . 21-nt RNA marker, 5′ -CGUACGCGGAAUACUUCGATT-3′ . Both are customordered from Dharmacon, Chicago, IL. 3. Isotopes: [-32 P] GTP (40 mCi/mL, Cat # 32010XT01) and [-32 P]ATP (10 mCi/mL, Cat # 38101X) from MP Biomedicals, Solon, OH. 4. Reagents for radiolabeling: Riboprobe® System-T7 Kit (Promega, Madison, WI, Cat# P1440) contains T7 polymerase, 5X T7 transcription buffer, 100 mM DTT, 10 mM ATP, UTP, CTP, and GTP, nuclease-free water, RNase inhibitor, and RNase-free DNase I. Polynucleotide kinase (PNK) and 10X PNK buffer (NEB, Ipswich, MA, Cat# M0201S). 5. Buffers for the assays: Buffer A (see Section 2.2), buffer X (500 mM ammonium acetate, 5 mM EDTA, 0.5% SDS); 10X dsRNA processing buffer [500 mM Kacetate, 150 mM HEPES (pH7.4); 18 mM magnesium acetate, 25 mM DTT]; and 10X pre-miRNA processing buffer [1 M K-acetate, 150 mM HEPES (pH 7.4), 100 mM magnesium acetate, and 25 mM DTT]. 6. Other reagents for the assays: 10 mM of ATP (Ambion, Austin, TX, Cat# 8110G), SUPERase•In™ (Ambion, Cat# 2696), and phenol/chloroform (5:1, pH 4.5, Ambion, Cat# 9722), glycogen (5 mg/mL, Ambion), 3 M and 0.3 M sodium acetate (pH5.2), diethyl pyrocarbonate (DEPC)-treated water [add 1 mL DEPC (Simga) in 1 L deionized water, stir overnight at room temperature, then autoclave for 30 min to inactivate DEPC]. 7. Buffers and reagents for denaturing polyacrylamide gel electrophoresis: Formamide load dye (Ambion), 16% denaturing polyacrylamide gel (freshly made with 5 mL 16% Urea gel mix, 40 μL 10% ammonium persulfate, and 4 μL TEMED), and 10X TBE running buffer (Ambion). For 16% urea gel mix, we made a solution containing 7 M Urea, 16% Acrylamide/Bis (from 40% Acrylamide/Bis 19:1 solution, Bio-Rad, Cat# 161-0144), and 1X TBE (from 10X TBE running buffer) and stored at 4 ºC.

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3. Methods 3.1. Expression of Dicer Both recombinant Dicer-1 and Dicer-2 enzymes are expressed in insect cells using the BAC-to-BAC® Baculovirus Expression System (15,16). Thus, for simplicity, we will refer to both Dicer-1 and Dicer-2 as “Dicer” when not specified in the sections below. The cDNAs of Dicer-1 and Dicer-2 are cloned by reverse transcription from total RNA of Drosophila S2 cells using the RLM-RACE kit from Ambion (Cat# 1700). To simplify purification of recombinant Dicers, polyhistidine (His)-tags were added by polymerase chain reaction (PCR) to the N- and C-termini of Dicers. We found that the addition of double His-tags greatly enhanced the expression levels of Dicer viruses. After verified by sequencing, the double His-tagged Dicer cDNAs were subcloned into the pFastBacTM 1 vector. The pFastBac-Dicer plasmids were transformed into E. coli DH10Bac competent cells to generate recombinant bacmids. The recombinant Dicer bacmids were isolated from positive bacteria transformants and verified by PCR that they contained the full-length Dicer cDNA (Fig. 1). The protocols for this section are described in detail in the manufacturer’s instruction manual and will not be repeated here.

Fig. 1. Recombinant Dicer-1 bacmids contain full-length Dicer-1 cDNA. The 5′ and 3 PCR reactions were performed without DNA template (lane 1 and 2) or with 1 μL of recombinant Dicer-1 bacmids prepared from four positive transformants (lanes 3–10). The 5′ PCR reactions use a 5′ bacmid (forward) primer (5′ -GTTTTCCCAGTCACGAC-3′ ) and a 5′ Dicer-1 (reverse) primer (5′ -CCGTCCAGCAATGATCAAAG-3′ ). The 3′ PCR reactions use a 3′ Dicer-1 (forward) primer (5′ -TTCCACAAGTTCTTCCGGCA-3′ ) and a 3′ bacmid (reverse) primer (5′ -CAGGAAACAGCTATGAC-3′ ). The predicted size of 5′ and 3′ PCR products are ∼ 2 kb and 1 kb. ′

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3.1.1. Culture of Sf21 Insect Cell 1. We have had good results with Sf21 cells from Invitrogen (Cat# 10497-013), but the reader is welcome to try insect cells from other sources. Every 3 months, thaw 1 vial of low-passage frozen Sf21 cells in a 25 ºC water bath. Transfer the cells to a 15-mL tube. Add 10 mL (prewarmed to 25 ºC) of complete IPL-41 medium and mix well by pipetting up and down. 2. Spin at 1,000 rpm for 5 min. Remove the supernatant and resuspend the cell pellet with 5 mL of completed IPL-41 medium. Count the cell density with a hemocytometer under a microscope. Transfer the cells into a 250-mL tissue culture flask (Corning, NY). Add complete IPL-41 medium to dilute the cell density to 1 × 106 cells/mL. 3. Incubate the suspension culture in a 27 °C shaker at 125 rpm. The doubling time of Sf21 cells is ∼ 24 h, and the optimal cell density should be kept between 0.5–5 × 106 cells/mL. Healthy Sf21 cells are round in shape and uniform in size. 4. It is best to freeze large numbers of small aliquots of low-passage Sf21 cells at the beginning. Spin 250 mL of mid-log-phase Sf21 cell culture (∼ 4 × 106 cells/mL) in a sterile centrifuge bottle at 1,000 rpm for 5 min. Remove the supernatant, and gently resuspend the cell pellet with 50 mL of complete IPL-41 medium containing 10% DMSO. Aliquot to freezing vials at 2 × 107 cells/mL/vial. 5. Place the vials into a room-temperature Cryo 1 °C freezing container (Nalgene, Rochester, NY, Cat# 5100-0001) and put the container into a –80 °C freezer. The Nalgene container allows the temperature to drop at a rate of 1 ºC per min. After 24 h, thaw one vial for testing and move the remaining frozen vials into a –190 ºC liquid nitrogen freezer for permanent storage (see Notes 4.1).

3.1.2. Generation of Recombinant Dicer Virus (Passage 1 and 2) 1. Seed total 2 × 106 Sf21 cells in 4 mL of complete IPL-41 medium in a T-25 (cm2 )flask (BD Bioscience, San Jose, CA, Cat# 353109). Incubate at 27 °C to allow cells to attach for at least 15 min. We often perform two transfections for each bacmid for backup. 2. Solution A: Dilute 15 μL of bacmid DNA into 200 μL of IPL-41 medium. Solution B: Dilute 12 μL of Cellfectin Reagent (inverting the tube 5 to 10 times before removing) into 200 μL of IPL-41 medium. Add solution B dropwise into solution A, mix gently by inverting the tube 8–10 times, and incubate at room temperature for 30–45 min. 3. Wash the attached Sf21 cells once with 2 mL of incomplete IPL-41 medium (no supplements). Transfer 1.6 mL of incomplete IPL-41 medium and then add 400 μL of lipid-DNA complexes (A and B) dropwise into the T-25 flask. Gently rock the flask several times to mix the content. Incubate at 27 °C for 4–5 h. 4. Remove the transfection mixture, add 5 mL of complete IPL-41 medium, and incubate at 27 ºC for 8–10 days. Check the cells under a microscope for contamination or cell lysis. Since Dicers are large proteins, it takes 8–10 days for more complete cell lysis in order to obtain high-titer viruses.

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5. When 50–60% of the cells are lysed or floating, blow the cells off and transfer to a sterile 15-mL tube. Spin at 1,000 rpm for 5 min and transfer the supernatant into a fresh 15-mL tube. This is called the passage 1 (P1) virus. 6. Transfer the entire P1 virus into a 250-mL flask containing 45 mL of Sf21 cells (2 × 106 cells/mL). Incubate in a 27 ºC shaker (125 rpm) for 5–7 days. Check the cells under a microscope. When more than 90% cells are lysed, harvest the cells by spinning and transfer the supernatant to a 50-mL tube. This is called the passage 2 (P2) virus. 7. Both P1 and P2 viruses can be stored at 4 ºC wrapped with aluminum foil for protection from light. For long-term storage, it is best to store the virus in 7% DMSO at –80 ºC.

3.1.3. Test the Expression Level of Dicer Virus 1. Plate 300 μL (2 × 106 cells/mL) of Sf21 cells in a 24-well dish. Add 3 μL of P2 Dicer virus in each well. Save one well for no virus control. Gently mix by rocking the dish several times. Incubate at 27 ºC for 40–44 h. 2. Remove the medium by aspirating. Place the dish on ice. Add 400 μL of 2% SDS/water to each well. Sit on ice for 1 min. 3. Resuspend the cell lysates and transfer to 1.5-mL microcentrifuge tubes. 4. Vortex vigorously for 30 sec to break the viscous genomic DNA. Transfer 45 μL of each sample to a new microcentrifuge tube containing 15 μL of 4X SDS sample buffer. Boil the samples for 5 min. Spin at 14,000 rpm for 2 min to precipitate insoluble material. Load 20 μL of the top layer on a 6% SDS-PAGE gel and run until the front dye migrates to the bottom of the gel. 5. Transfer the SDS-PAGE gel to a nitrocellulose membrane for 2 h at 400 mA. Perform Western blot using rabbit anti-His (primary, 1:5,000) antibody and HRPconjugated anti-rabbit IgG (secondary, 1:10,000) antibody (Fig. 2).

3.1.4. Amplification of Dicer Virus (P3, P4) and Large-Scale Protein Production 1. Grow 600 mL of Sf21 cells to 2.5 × 106 cells/mL in a 2-L flask. Add 25 mL of Dicer P2 virus and incubate in a 27 ºC shaker (125 rpm) for 5–7 days. 2. Check the cells under a microscope. When more than 90% cells are broken, distribute the culture to multiple 50-mL sterile tubes and spin at 2,000 rpm for 5 min. The supernatant is called the P3 virus, which can be amplified to the P4 virus in the same manner. Store the P3 and P4 viruses at 4 ºC wrapped in aluminum foil. 3. For large-scale protein production, grow 2 × 600 mL of Sf21 cells in 2-L flasks to a cell density of 2.5 × 106 cells/mL. Add 25 mL of the P3 Dicer virus into each of two flasks. Incubate in a 27 ºC shaker (125 rpm) for 42–48 h.

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Fig. 2. Expression test of P2 baculovirus. Western blot was performed with anti-His antibodies using lysates prepared from Sf21 cells that are mock treated (lane 1) or infected with P2 viruses of Dicer-2 (lane 2) or Dicer-1 (lane 3). 4. Spin down the cells in 1-L centrifuge bottles at 2,500 rpm for 20 min at 4 ºC. Remove the supernatant and resuspend each pellet with 20 mL of PBS. Transfer the cell suspension into 50-mL tubes and spin again at 4,000 rpm for 15 min at 4 ºC. Remove the supernatant. The cell pellets can be immediately used to make lysate for purifying recombinant Dicer proteins or can be stored at –80 ºC for future use.

3.2. Purification of Dicer Both Dicer-1 and Dicer-2 recombinant proteins are highly purified by nickel affinity purification followed by SP- and Q-Sepharose chromatography (Fig. 3). Nickel affinity purification is based on the double His-tags of recombinant Dicers, which is the same for Dicer-1 and Dicer-2. However, the column behaviors of Dicer-1 and Dicer-2 are different for the SP- and Q-Sepharose chromatography. On the SP-Sepharose column, Dicer-1 flows through, whereas Dicer-2 binds and is eluted at 180–220 mM of sodium chloride. On the Q-Sepharose column, both Dicer-1 and Dicer-2 bind but are respectively eluted at 350–380 mM and 260–300 mM sodium chloride. All purification steps are performed at 4 ºC on an ACTA FPLC machine purchased from Amersham. After the multistep purification, we can typically get ∼ 0.6–1.0-mg recombinant Dicer proteins with more than 90% purity from 1.2 L of insect cell culture (Fig. 3).

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Fig. 3. Purification of recombinant Dicer-1 and Dicer-2. The His-tagged Dicer-1 and Dicer-2 proteins are both purified by Ni+2 -affinity purification followed by SPSepharose and Q-Sepharose chromatography. (a) Lane 1: 250 mM imidazole elution, also as input of SP column; lane 2: SP flowthrough, also as input of Q column; lanes 3–7: Dicer-1 peak fractions of Q column. (b) Lane 1: 250 mM imidazole elution, also as input of SP column; lane 2: Dicer-2 peak fractions of SP column; also combined as input of Q column; lane 3: Dicer-2 peak fractions of Q column.

3.2.1. Preparation of Cell Lysate and Nickel Purification 1. Quick-thaw the cell pellet by placing the frozen tube in a beaker filled with roomtemperature water. Resuspend the cell pellet thoroughly (from 1.2 L Dicer virusinfected culture, 15–20 mL pellet volume) in three times pellet volume of buffer A freshly supplemented with protease inhibitors and 20 mM of imidazole (see Notes 4.2). Let the cells swell in hypotonic buffer A by staying on ice for 20 min. 2. Pour the cell suspension into a 100-mL glass douncer and make 40 strokes to break the cells completely. Transfer the whole-cell lysates into 50-mL centrifuge tubes and spin at 20,000 x g at 4 ºC for 30 min. The supernatant is called S20. 3. Wash 2-mL nickel beads (4 mL 50% slurry) three times with 10 mL of buffer A containing 20 mM of imidazole in a 15-mL conical tube. Each time, resuspend the beads thoroughly, spin at 1,000 rpm for 3 min, and remove the supernatant.

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4. Transfer S20 of step 2 to a 50-mL conical tube. Use S20 to resuspend the prewashed nickel beads and transfer back to the 50-mL conical tube. Rotate at 4 ºC for 4 h to allow recombinant Dicer proteins to bind with the nickel beads. 5. Pour the lysate/beads mixture into a 25-mL Econo-Pac® disposable chromatography column. Break the bottom plastic cap, let the beads settle down by gravity, and collect the flowthrough. 6. Wash the beads sequentially with 200 mL of buffer B containing 20 mM of imidazole, 50 mL of buffer A containing 20 mM of imidazole, and 10 mL of buffer A containing 50 mM of imidazole. Elute with 10 mL of buffer A (freshly supplemented with protease inhibitors and 250 mM imidazole). Nickel elution can be filtered and used for the next purification steps or stored at −80 ºC with 10% glycerol for future use. 7. Take 15 μL of the elution and mix with 5 μL of the 4X SDS sample buffer. Boil for 5 min and load onto a 6% SDS-PAGE to check the quantity and quality of recombinant Dicer proteins by Coomassie Blue staining.

3.2.2. Ion-Exchange Chromatography for Dicer-1 Purification 1. Filter the Dicer-1 nickel elution with a 0.2-μm syringe filter. Wash the SP- and Q-Sepharose columns and the super-loop of the FPLC system with buffer A. 2. Inject the sample into the super-loop and load it onto the 1-mL SP-Sepharose column. Collect the flowthrough that contains recombinant Dicer-1 proteins. 3. Reinject the SP flowthrough back into the super-loop and load it onto the 1-mL Q-Sepharose column. Perform a 20-mL 0–45% buffer B gradient elution followed by a 5-mL 100% buffer B step wash (fraction volume = 1 mL). The peak of Dicer-1 is at 34–38% buffer B elution. 4. Take 15 μL of SP flowthrough and Q peak elution of Dicer-1, mix with 5 μL of 4X SDS sample loading buffer, boil for 5 min, and load onto a 6% SDSPAGE followed by Coomassie Blue staining. Compare the nickel elution with SP flowthrough and Q peak fractions to see the increasing purity of recombinant Dicer-1. 5. Take 1 μL of Q peak fractions to measure the concentration of purified recombinant Dicer-1 proteins. They typically fall between 0.3 and 0.5 mg/mL. Add 10% glycerol, mix well, and store at −80 ºC in small aliquots.

3.2.3. Ion-Exchange Chromatography for Dicer-2 Purification 1. Filter the Dicer-2 nickel elution with a 0.2-μm syringe filter. Wash the SP- and Q-Sepharose columns and the super-loop with buffer A. 2. Inject the sample into the super-loop and load onto the 1-mL SP-Sepharose column. Perform a 15-mL 0–30% gradient buffer B elution followed by a 5-mL 100% buffer B step wash (fraction volume = 1 mL). The peak of Dicer-2 is at 18–22% buffer B elution.

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3. Combine the SP peak fractions, and dilute with two volumes of buffer A to reduce the salt concentration. Reinject it into the FPLC system and load onto the 1-mL Q-Sepharose column. Perform a 20-mL 0–40% gradient buffer B elution followed by a 5-mL 100% buffer B step wash (fraction volume = 1 mL). The peak of Dicer-2 is at 26–30% buffer B elution. 4. Take 15 μL of the SP and Q peak fractions, mix with 5 μL of 4 x SDS sample loading buffer, boil for 5 min, and load onto a 6% SDS-PAGE followed by Coomassie Blue staining. Compare the nickel elution with SP and Q peak fractions to see the increasing purity of recombinant Dicer-2. 5. Take 1 μL of the Q peak fractions to measure the concentration of purified recombinant Dicer-2. They typically fall between 0.3 and 0.5 mg/mL. Add 10% glycerol, mix well, and store at –80 ºC in small aliquots.

3.3. Assay of Dicer Genetic studies have suggested that Dicer-1 and Dicer-2 are respectively involved in miRNA and siRNA production (14). By biochemical fractionation, it has been shown that the Dicer-1/Loqs-L complex processes pre-miRNA into mature miRNA, whereas the Dicer-2/R2D2 complex cleaves long dsRNA into siRNA (15–18). The functional specificities of the two Dicer complexes can be explained by the fact that Dicer-1 and Dicer-2 possess different biochemical activities or that Loqs-L and R2D2 help define the functional specificities for their respective Dicer partners. Here we compare the abilities of purified recombinant Dicer-1 and Dicer-2 enzymes to process radiolabeled pre-miRNA or long dsRNA in the absence or presence of ATP (see Notes 4.3). Our studies indicate that, despite sharing extensive sequence homology, Drosophila Dicer-1 and Dicer-2 enzymes display different substrate specificities and ATP requirements. While Dicer-1 prefers pre-miRNA as its ideal substrate, Dicer-2 is a much better enzyme for processing dsRNA (Fig. 4). Furthermore, Dicer-1 produces miRNA or siRNA independent of ATP, whereas Dicer-2 absolutely requires ATP hydrolysis for efficient siRNA production (Fig. 5). 3.3.1. Preparation of Radiolabeled dsRNA 1. Set up a 20-μL in vitro transcription reaction using the Riboprobe®-T7 Kit: 4 μL of 5 × T7 transcription buffer, 2 μL of 100 mM DTT, 1 μL of RNase inhibitor, 4 μL of NTP mix (10 μL ATP/UTP/CTP + 2 μL GTP + 8 μL water), 1.5 μL of sense template DNA (250 ng/μL), 1.5 μL of antisense template DNA (250 ng/μL), 5 μL [-32 P] of GTP, and 1 μL of T7 polymerase. Mix well and incubate at 37 ºC for 1.5 h. 2. Add 1 μL of RNase-free DNase I to the reaction tube, mix well, and incubate at 37 ºC for 15 min.

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Fig. 4. Dicer-1 and Dicer-2 display different substrate specificities. (a) The dsRNAprocessing assays were performed with 0.03, 0.1, 0.3, 1 nmol of recombinant Dicer-1 (lanes 3–6) and Dicer-2 (lanes 7–10) enzymes. Dicer-2 is a much better enzyme than Dicer-1 in cleaving dsRNA into siRNA. (b) The pre-miRNA processing assays were performed with 0.03, 0.1, 0.3, 1 nmol of recombinant Dicer-1 (lanes 3–6) and Dicer2 (lanes 7–10) enzymes. While Dicer-1 efficiently processes pre-miRNA into 21 nt mature miRNA (lanes 3–6), Dicer-2 cannot cleave pre-miRNA at these concentrations (lanes 7–10). Lane 1: radiolabeled 21 nt marker, lane 2: buffer control.

3. Stop the reaction with addition of 115 μL of nuclease-free water and 15 μL of 3 M sodium acetate (pH 5.2). Incubate at 90 ºC for 1 min. 4. Add 150 μL of acid phenol/chloroform at room temperature. Close the cap tightly and vortex vigorously for 30 sec. Spin at 13,000 rpm for 5 min at room temperature. Transfer the aqueous phase to a new microcentrifuge tube and repeat this step. The two phenol/chloroform extractions will facilitate annealing of dsRNA substrates. 5. Transfer the aqueous phase to a new microcentrifuge tube. Add 3.5 μL of 5 mg/mL glycogen and 300 μL of –20 ºC 100% ethanol. Invert the tube several times to mix well. Incubate at −20 ºC for 1 h to overnight. 6. Spin at 13,000 rpm for 5 min at 4 ºC. Remove the supernatant and wash the pellet with 0.5 mL of 70% ethanol. Remove all supernatant, air-dry the pellet for 5–10 min, and resuspend in 100 μL of nuclease-free water. 7. Count 1 μL with a scintillation counter and dilute the probe to 1 × 105 cpm/μL. Store the probe in aliquots at –20 ºC, which will be good for 2–3 weeks.

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Fig. 5. Dicer-1 and Dicer-2 have different ATP requirements. (a) Dicer-1 processes pre-miRNA into miRNA in an ATP-independent manner. The pre-miRNA-processing assays were performed with 0.03, 0.1, 0.3, 1 nmol recombinant Dicer-1 in the absence (lanes 1–4) or presence (lanes 5–8) of ATP. (b) Dicer-1 cleaves dsRNA into siRNA in an ATP-independent manner. The dsRNA-processing assays were performed with 0.03, 0.1, 0.3, 1 nmol recombinant Dicer-1 in the absence (lanes 1–4) or presence (lanes 5–8) of ATP. (c) Dicer-2 requires ATP for efficient siRNA production. The dsRNAprocessing assays were performed with 1.25, 2.5, 5, 10 pmol recombinant Dicer-2 in the absence (lanes 1–4) or presence (lanes 5–8) of ATP.

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3.3.2. Preparation of Radiolabeled Pre-miRNA 1. Set up a 20-μL 5′ end-labeling reaction in a microcentrifuge tube with the following: 2 μL of 10X PNK buffer, 8 μL of synthetic pre-let7 RNA (2 μM), 8 μL of [-32 P] ATP, 2 μL of PNK. Mix well and incubate at 37 ºC for 2 h. 2. Add 20 μL of formamide loading dye to stop the reaction. Incubate at 95 ºC for 5 min. Load onto a 5-well 6% denaturing polyacrylamide gel. 3. Run at 300 V for 15 min. Expose to an autoradiography film for 1 min to detect the position of radiolabeled pre-miRNA. Cut the pre-miRNA containing band from the gel and place it into a special microcentrifuge tube. 4. Add 500 μL of buffer X into the tube. Crush the gel slice with an accompanying pestle (20 strokes). Rotate at room temperature for 1 h. 5. Spin the tube at 13,000 rpm for 2 min. Transfer the aqueous phase to a Wizard SV minicolumn (included in Wizard Plus SV Minipreps DNA Purification System, Promega, Cat# A1460), place the minicolumn in a microcentrifuge tube, and spin at 13,000 rpm for 2 min. 6. Transfer the aqueous phase (∼450 μL) to a new microcentrifuge tube. Add 3.5 μL of glycogen and 1 mL of –20 ºC ethanol. Invert the tube several times to mix well. Incubate at –20 ºC for 1 h to overnight. 7. Spin at 13,000 rpm for 5 min at 4 ºC. Remove the supernatant and wash the pellet with 0.5 mL of 70% ethanol. Remove all the supernatant, air-dry the pellet for 5–10 min, and dissolve it in 50 μL of nuclease-free water. 8. Count 1 μL with scintillation counter and dilute the probe to 4 × 104 cpm/μL. Store in aliquots at –20 ºC.

3.3.3. DsRNA-Processing Assays of Dicer-1 and Dicer-2 1. Make 90 μL of the master mix for 10 reactions with the following: 10 μL of 10X dsRNA processing buffer, 10 μL of 10 mM ATP, 2 μL of SUPERase•In™, 10 μL of dsRNA probe (1 × 105 cpm/μL), and 58 μL of nuclease-free water. 2. Distribute 9 μL of the master mix to each microcentrifuge tube, and add 1 μL of buffer A or recombinant Dicer-1 or Dicer-2 proteins of various concentrations. Incubate the reaction tubes at 30 ºC for 30 min. 3. Stop the reactions by the addition of 200 μL of 0.3 M sodium acetate (pH 5.2) and extract with 200 μL of phenol/chloroform. Transfer the aqueous phase to new microcentrifuge tubes containing 3.5 μL of glycogen. Add 0.5 mL of –20 ºC 100% ethanol to each tube. Invert the tube several times. Incubate at –20 ºC for 30 min. 4. Spin at 13,000 rpm for 5 min at 4 ºC. Remove all the supernatant, air-dry the pellet for 5–10 min, and resuspend the pellet in 5 μL of formamide loading dye. Boil for 5 min and load onto a 16% denaturing polyacrylamide gel. Use a radiolabeled synthetic 21-mer RNA oligonucleotide as the size marker. 5. Run at 300 V for 40 min. Expose to an autoradiography film at –80 ºC for 1 to 2 h. Develop the film and compare the dsRNA processing activities of Dicer-1 and Dicer-2.

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3.3.4. Pre-miRNA-Processing Assays of Dicer-1 and Dicer-2 Make 90 μL of the master mixture for 10 reactions with the following: 10 μL of 10X pre-miRNA processing buffer, 10 μL of 10 mM ATP, 5 μL of SUPERase•In™, 10 μL of pre-miRNA probe (4 × 104 cpm/μL), and 55 μL of nuclease-free water. For the rest of the procedure, follow steps 2–5 as above. 4. Notes 4.1. Notes on Expression 1. Frozen stocks of bacmids and viruses should be in small aliquots (each for one use). Repetitive freeze-and-thaw cycles will destroy their structures and activities. 2. Amplification of P2 virus to the P3 and P4 viruses is essential to get large amounts of higher-titer viruses. However, we found that the expression levels of Dicer viruses would decrease with more than four passages. 3. For protein productions, do not let the cells grow more than 48 h after virus infection. Cells will begin to lyse after 48 h, resulting in more degradation products of recombinant Dicer proteins.

4.2. Notes on Purification 4. For nickel affinity purification, beta-mercaptoethanol is used instead of DTT because DTT will destroy the nickel beads. The amount of nickel beads to use is based on the expression levels of Dicer proteins. 5. It is critical to store recombinant Dicer in 10% glycerol at –80 ºC. Freezing without glycerol will kill the enzymes. Fast freezing in liquid nitrogen is recommended.

4.3. Notes on Assay 6. When performing radioactive experiments, always take care to avoid contaminating the environment, and protect yourself and people around you from exposure to the radioactive materials. 7. To minimize RNase contamination, all solutions and buffers used in these experiments should be made with DEPC-treated water. Always wear gloves, and use RNase-free tips and reagents designated to the RNA bench. 8. When preparing radiolabeled dsRNA or pre-miRNA substrates, it is best to verify the quality of the radiolabeled probes by running 1 μL of probe on a denaturing PAGE prior to performing the actual assays. The pre-miRNA substrate needs to be PAGE gel-purified before use. We use a mixture of sense and antisense DNA templates (each contains a single T7 promoter) to generate blunt-ended dsRNA, which has been shown to be the ideal substrate for Dicer enzymes (15,16). 9. For assays of ATP requirements of Dicer-1 and Dicer-2, it is desirable to check if purified Dicer-1 and Dicer-2 proteins have residual ATP contamination. Alternatively, simply perform one set of regular reactions at 4 ºC (ATP won’t be hydrolyzed at this temperature) and compare the activities between 4 ºC and 30 ºC.

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Acknowledgments We thank Dr. Zain Paroo for critical reading of the manuscript and Feng Jiang for technical assistance. Q. L. is a W. A. “Tex” Moncrief Jr. Scholar in Medical Research and a Damon Runyon Scholar supported by the Damon Runyon Cancer Research Foundation (DRS-43). The work is also supported by a Welch grant (I-1608).

References 1. Hannon, G. J. (2002). RNA interference. Nature 418, 244–251. 2. Bartel, D. P. (2004). MicroRNAs: Genomics, biogenesis, mechanism, and function. Cell 116,281–297. 3. Filipowicz, W., Jaskiewicz, L., Kolb, F. A., and Pillai, R. S. (2005). Posttranscriptional gene silencing by siRNAs and miRNAs. Curr. Opin. Struct. Biol. 15, 331–341. 4. Sontheimer, E. J., and Carthew, R. W. (2005). Silence from within: Endogenous siRNAs and miRNAs. Cell 122, 9–12. 5. Esquela-Kerscher, A., and Slack, F. J. (2006). Oncomirs—MicroRNAs with a role in cancer. Nat. Rev. Cancer 6, 259–269. 6. Plasterk, R. H. (2006). MicroRNAs in animal development. Cell 124, 877–881. 7. Tomari, Y., and Zamore, P. D. (2005). Perspective: Machines for RNAi. Genes Dev. 19, 517–529. 8. Tang, G. (2005). siRNA and MiRNA: An insight into RISC. Trends Biochem. Sci. 30, 106–114. 9. Carmell, M. A., and Hannon, G. J. (2004). RNase III enzymes and the initiation of gene silencing. Nat. Struct. Mol. Biol. 11, 214–218. 10. Zhang, H., Kolb, F. A., Jashiewicz, L., Westhof, E., and Filipowicz, W. (2004). Single processing center models for human Dicer and bacterial Rnase III. Cell 118, 57–68. 11. MacRae, I. J., Zhou, K., Li, F., et al. (2006). Structural basis for double-stranded RNA processing by Dicer. Science 311, 195–198. 12. Bernstein, E., Caudy, A. A., Hammond, S. M., and Hannon, G. J. (2001). Role for a bidentate ribonuclease in the initiation step of RNA interference. Nature 409, 363–366. 13. Ketting, R. F., Fischer, S. E., Bernstein, E., Sijen, T., Hannon, G. J., and Plasterk, R. H. (2001). Dicer functions in RNA interference and in synthesis of small RNA involved in developmental timing in C. elegans. Genes Dev. 15, 2654–2659. 14. Lee, Y. S., Nakahara, K., Pham, J. W., et al. (2004). Distinct roles for Drosophila Dicer-1 and Dicer-2 in the RNAi/miRNA silence pathway. Cell 117, 69–81. 15. Liu, Q., Rand, T. A., Kalidas, S., et al. (2003). R2D2, a bridge between the initiation and effector steps of the Drosophila RNAi pathway. Science 301, 1921–1925.

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16. Jiang, F., Ye, X., Liu, X., Fincher, L., McKearin, D., and Liu, Q. (2005). Dicer-1 and R3D1-L catalyze microRNA maturation in Drosophila. Genes Dev. 19, 1674–1679. 17. Saito, K., Ishizuka, A., Siomi, H., and Siomi, M. C. (2005). Processing of premicroRNAs by the Dicer-1-Loquacious complex in Drosophila cells. PLoS Biol. 3, e235. 18. Forstemann, K., Tomari, Y., Du, T., et al. (2005). Normal microRNA maturation and germ-line stem cell maintenance requires Loquacious, a double-stranded RNAbinding domain protein. PLoS Biol. 3, e236. 19. Tomari, Y., Matranga, C., Haley, B., Martinez, N., and Zamore, P. D. (2004). A protein sensor for siRNA asymmetry. Science 306, 1378–1380.

3 In vitro RNA Cleavage Assay for Argonaute-Family Proteins Keita Miyoshi, Hiroshi Uejima, Tomoko Nagami-Okada, Haruhiko Siomi, and Mikiko C. Siomi

Summary Recent studies have revealed that Argonaute proteins are crucial components of the RNA-induced silencing complexes (RISCs) that direct both small interfering RNA (siRNA)- and microRNA (miRNA)-mediated gene silencing. Full complementarity between the small RNA and its target messenger RNA (mRNA) results in RISC-mediated cleavage (“Slicing”) of the target mRNA. A subset of Argonaute proteins directly contributes to the target cleavage (“Slicer”) activity of the RISC. We describe in vitro Slicer assays using endogenous Argonaute protein immunopurified from animal cells and recombinant Argonaute protein produced in and purified from Escherichia coli.

Key Words: RNAi; Argonaute; Slicer; RISC; Drosophila.

1. Introduction Although RNA interference (RNAi) has been enthusiastically viewed as a new therapeutic modality, it is important to keep in mind that RNAi is but one aspect of a larger web of sequence-specific cellular responses to RNA now known collectively as “RNA silencing” (1). In essence, it is a sequence-specific RNA cleavage process triggered by double-stranded (ds) RNA from a variety of sources (see Fig. 1 in Chapter 4) (1,2). One such source is long dsRNA exogenously introduced or endogenously expressed in cells; this is first converted to 21–23-nucleotide (nt) small RNAs termed “small interfering RNAs” (siRNA) by Dicer in the cytoplasm of cells (3–6). Subsequently, the siRNA duplex From: Methods in Molecular Biology, vol. 442: RNAi: Design and Application Edited by: S. Barik © Humana Press, Totowa, NJ

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Fig. 1. Sequences of RNA targets, miR-21 target RNA and luc target RNA, and potential base pairings with miR-21 and luc guide siRNA. Cleavage sites are indicated with lightning bolts. RNA targets that bear complete complementarity to the small RNAs are cleaved at a position across from their middle (i.e., between the 10th and 11th nucleotides of small RNAs).

goes through an unwinding process, and one strand over the other is preferentially loaded onto the RNA-induced silencing complex (siRISC), which in turn endonucleolytically cleaves target RNA at sites completely complementary to the siRNA (7–10). Another source of dsRNA is transcripts of a large family of cellular genes encoding self-complementary RNA molecules that form imperfect hairpins, termed “primary microRNA” (pri-miRNA). Pri-miRNAs are first processed by the Drosha-Pasha/DGCR8 complex in the nucleus to generate pre-miRNA (precursor of miRNA). The pre-miRNA is exported to the cytoplasm by exportin-5 and further processed by Dicer into an approximately 22-nt miRNA duplex; then, one strand over the other is loaded onto RISC (miRISC) as siRNA in RNAi (11). Mature miRNAs residing in the miRISC anneal to target mRNAs at specific sites, and miRISC induce cleavage of the messages or inhibit their translation (12–16). By regulating endogenous gene expression, miRNAs are known to play crucial roles in development (17–21). Chromatographic purification of RISC nuclease or Slicer activity from Drosophila cells revealed several RISC components (2). However, since the discovery of RNAi, the central question of which RISC component actually catalyzes the hydrolytic cleavage reaction has remained unanswered. Argonaute-family proteins, categorized by two characteristic domains, PAZ and PIWI, were the first identified component of RISC. In 2004, JoshuaTor and colleagues solved the crystal structure of the Argonaute protein from Pyrococcus furiosus, which revealed that the PIWI domain shows an RNaseHlike structure and that essential residues for the endonucleolytic activity of

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RNaseH are conserved in the Argonaute-PIWI domain (22). These data strongly indicate that Argonaute (Ago) might be the “Slicer,” the endonucleolytic enzyme residing in RISC. Indeed, Hannon and colleagues showed that epitopetagged human Ago2 (hAgo2) expressed in and purified from 293T cells was able to catalyze target RNA cleavage when the immunopurified hAgo2 was pre-incubated with single-stranded siRNA, but not with single-stranded small DNA (23). Mutagenizing the conserved residues in hAgo2 abolished target RNA cleavage activity (23). Later, Slicer activity was reconstituted with recombinant hAgo2 produced in E. coli and single-stranded siRNA (24). Taking all of the above into consideration, it was concluded that hAgo2 is Slicer in RNAi. In humans, there are four closely related Argonaute-family members, Ago1–4 (hAgo1–4), all of which have been associated with both siRNA and miRNA; however, only hAgo2 mediates RNA cleavage targeted by the small RNAs (23–25). hAgo1, hAgo3, and hAgo4 show high similarities with hAgo2 at peptide sequence levels, and the amino acid residues required for hAgo2 Slicer activity are well conserved among the three other proteins (26): Thus, it is somewhat anomalous that only hAgo2 has Slicer activity. In Drosophila, the Argonaute family consists of five members (AGO1, AGO2, AGO3, Piwi, and Aubergine) (27). Among these, AGO1 and AGO2 are the most closely related to hAgo1–4. Our previous studies have focused on the functional contribution of AGO1 and AGO2 in RNA silencing and revealed that AGO2 is required for the siRNA-directed target RNA cleavage process, while AGO1, the closest relative to AGO2 among the Argonaute members in fly, is dispensable for such reaction (28). Thus, we assessed if AGO2 isolated from Drosophila cells could show target RNA cleavage activity, as hAgo2 did from human cells. Unlike Hannon and colleagues, we decided to utilize endogenous AGO2 protein since we could easily obtain purified AGO2 by immunoprecipitation from Drosophila Schneider 2 (S2) cells using a monoclonal antibody we raised against the N-terminal region of AGO2 (29). We first confirmed by Northern blot analysis that the immunopurified AGO2 from S2 lysate preprogrammed with luc siRNA duplex was indeed associated with luc guide siRNA, as expected. Later, by performing in vitro target RNA cleavage assay, we found that the immunopurified AGO2 associated with luc siRNA was capable of cleaving luc target RNA. These data strongly implicated that in Drosophila, as with hAgo2 in human, AGO2 functions as Slicer. Through the course of our studies, we became aware that miRNA-directed target RNA cleavage occurred even without AGO2. Further investigation revealed that AGO1 is the protein essential for miRNA-directed target RNA cleavage reaction. Analysis of amino acid sequences showed that AGO1 more resembles hAgo2 than it does AGO2. Therefore, we next assessed if AGO1 also functions as Slicer. We prepared AGO1 protein by two independent methods. The first

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was endogenous AGO1 immunopurified from S2 cells using the monoclonal antibody against AGO1. The second was recombinant GST-tagged full-length AGO1 produced in and purified from E. coli. Target RNA used in the assay was an RNA molecule containing a sequence completely matched to miR-bantam. That AGO1 immunopurified from S2 cells was associated with miR-bantam was confirmed in advance by Northern blotting. GST-AGO1 was incubated with miR-bantam prior to the target RNA cleavage assay. Both forms of AGO1 were able to efficiently cleave the target RNA; thus, it was concluded that AGO1 also has Slicer activity. In this chapter, we describe in vitro target RNA cleavage assay using purified Argonaute proteins from both animal cells and E. coli. While this in vitro assay will clearly be useful for elucidating the Slicer function of Argonaute proteins from a variety of organisms, it will also be useful for other applications. Indeed, this assay allowed us to ascertain that AGO2 Slicer function is needed for facilitating siRNA duplex unwinding by cleaving and discarding the passenger strand of siRNA duplex, as if AGO2 cleaves a target RNA annealed to guide siRNA residing in siRISC (29). Finally, we address important issues as well as suggest useful techniques to optimize the assay. 2. Materials 1. RNA extraction reagent (ISOGEN, or ISOGEN-LS, Nippon Gene, Toyama, Japan). 2. Cloning vectors harboring phage promoters: pBluescript SK (Stratagene, La Jolla, CA) or equivalent. 3. MEGAscript kit (Ambion, Austin, TX). 4. Pellet Paint® Co-precipitant (EMD Bioscience, Darmstadt, Germany). 5. 6% acrylamide denaturing gel: 1X Tris-borate-ethylenediaminetetraacetic acid, 6 M of urea, 6% acrylamide. 6. 8% acrylamide denaturing gel: 1X Tris-borate-ethylenediaminetetraacetic acid, 6 M of urea, 8% acrylamide. 7. RNA elution buffer: 0.5 M of ammonium acetate, 1 mM of ethylenediaminetetraacetic acid (EDTA), and 0.2% sodium dodecylsulfate. 8. Guanylyltransferase, 10X Capping reaction buffer, and S-adenosyl methionine (Ambion). 9. [-32 P]GTP (3000 Ci/mmol) (PerkinElmer, Boston, MA). 10. Gel-filtration columns: Micro Bio-Spin Columns P-30 Tris, RNase-Free (BioRad, Hercules, CA). 11. luc siRNA (guide strand: 5′ -UCGAAGUAUUCCGCGUACGUG-3′ , passenger strand: 5′ -CGUACGCGGAAUACUUCGAAA-3′ ). 12. Lithium salt of ATP, 100 mM, pH 7 (Roche, Basel, Switzerland). 13. T4 Polynucleotide Kinase (T4PNK) and 10X PNK reaction buffer (TaKaRa, Shiga, Japan).

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14. HeLa cells. 15. Dulbecco’s modified Eagle’s medium (SIGMA, St. Louis, MO) supplemented with 10% fetal calf serum. 16. Phosphate-buffered saline (PBS). 17. Monoclonal anti-hAgo2 antibody (4G8), against the N-terminus of hAgo2 (120 amino acids). 18. Binding buffer: 30 mM HEPES, pH 7.4, 150 mM KOAc, 2 mM Mg(OAc)2 , 5 mM dithiothreitol (DTT), 0.1% Nonidet P-40, 2 μg/mL pepstatin, 2 μg/mL leupeptin, and 0.5% aprotinin. 19. GammaBind G Sepharose (GE Healthcare Bio-Sciences, Piscataway, NJ). 20. 5X cleavage buffer: 125 mM Hepes-KOH, pH 7.5, 250 mM KOAc, 25 mM Mg(OAc)2 , 25 mM DTT. 21. RNasin® Plus RNase Inhibitor (Promega, Madison, WI). 22. Escherichia coli (E. coli) BL21 (DE3). 23. 1 M isopropyl thiogalactoside (IPTG). 24. Complete, Mini, EDTA-free (Roche). 25. Glutathione SepharoseTM 4B (GE Healthcare Bio-Sciences). 26. GST elution buffer: 10 mM reduced glutathione, 50 mM Tris-HCl, pH 8.0, 10% glycerol, and 5mM Mg(OAc)2 . 27. Protein assay (Bio-Rad). 28. 5 mg/mL yeast RNA (Ambion). 29. Image plates: BAS-MS2040 (Fujifilm, Tokyo, Japan). 30. BAS-2500 imaging system (Fujifilm).

3. Methods 3.1. Preparation of Target RNA Both siRNAs and miRNAs are able to guide mRNA degradation or translational repression depending on the complementarity of the target RNAs to the small RNAs. Namely, even the Slicer function of Argonaute that is naturally associated with miRNAs, for example Drosophila AGO1, can be assessed with this in vitro target RNA cleavage assay when a particular target RNA containing a sequence completely matching the miRNA is prepared. Target RNAs that appear in this chapter are miR-21 target RNA and luc target RNA (Fig. 1). 3.1.1. Preparation of Template DNA of Target RNA To make miR-21 target RNA harboring a sequence fully complementary to miR-21, PCR was performed to amplify a sequence between the T7 and T3 promoter regions of the pBS-miR-21-target plasmid, which was constructed as follows: A KpnI-EcoRI fragment containing a sequence fully complementary to miR-21 was produced by annealing a set of DNA oligos

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(5′ -CTCAACATCAGTCTGATAAGCTAG-3′ and 5′ -AATTCTAGCTTATCAGACTGATGTTGAGGTAC -3′ ), which was then inserted into pBluescript digested with KpnI and EcoRI. To prepare a template DNA to make luc target RNA, PCR was performed to amplify a portion of luciferase cDNA. The forward and reverse primers used for this PCR reaction were forward 5′ -TAATACGACTCACTATAGGGCTATCCTCTAGAGGATGGAAC-3′ , reverse 5′ -AATTAACCCTCACTAAAGGGCATAGCTTCTGCCAACCGAAC-3′ . It should be noted that the forward and reverse primers contain the T7 and T3 promoter sequences, respectively. 3.1.2. Target RNA Transcription Target RNA is transcribed according to the manufacturer’s instructions (MEGAscript, Ambion) with minor modifications. The detailed protocol is as follows: 1. Combine the following transcription reaction mixture in a total volume of 20 μL: a. b. c. d. e.

2 μL each of 75 mM ATP, CTP, GTP, and UTP. 2 μL of 10X reaction buffer. Roughly 0.5 μg of template DNA. 2 μL of enzyme mix. Nuclease-free water.

2. Incubate the transcription reaction solution at 37 ºC overnight. 3. Add 1 μL of DNase I and incubate at 37 ºC for 15 min. 4. Add 15 μL of 3 M NaOAc, 115 μL of nuclease-free water, and 150 μL of phenol/chloroform, and mix thoroughly. Recover the aqueous phase and transfer to a new tube. 5. Precipitate the RNA by adding an equal volume of isopropanol and 1 μL of pellet-paint® co-precipitant and mixing well. 6. Chill the mixture for at least 15 min at –80 ºC. Centrifuge at 4 ºC for 20 min at maximum speed (20,000 x g) to pellet the RNA. 7. Isolate the target RNA by 8% acrylamide denaturing gel electrophoresis as follows: a. b. c. d. e. f. g.

Separate the RNA by 8% acrylamide denaturing gel electrophoresis. Excise the gel piece containing the target RNA from the gel. Crush the gel piece into small pieces with a disposable pipette tip. Add 400 μL (> two gel volumes) of RNA elution buffer. Rotate at 4 ºC overnight to elute the RNA. Remove the gel pieces and collect the supernatant. Purify the target RNA by phenol extraction and isopropanol precipitation in the presence of a precipitation carrier.

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8. Cap labeling of the target RNA (5′ -terminal phosphate of the 5′ -triphosphate target RNA): Combine the following capping reaction mixture in a total volume of 15 μL: a. b. c. d. e.

1.5 μL of 10X Capping reaction buffer. 3 μL of [-32 P]GTP (3000 Ci/mmol). 1 μL of S-adenosyl methionine. 0.5 μg of target RNA. 1 μL of guanylyltransferase enzyme.

9. Incubate the reaction at 37 ºC for 60 min. 10. Gel-filter with a P-30 column, according to the manufacturer’s instructions, to remove any unincorporated radioactive GTP. 11. Isolate the target RNA by 8% acrylamide denaturing gel electrophoresis as below. 12. Dissolve RNA with nuclease-free water to a working concentration of 2,000–5,000 cpm/μL.

3.2. Preparation of Phosphorylated siRNA The processing of dsRNA, pri-miRNA, or pre-miRNA by Dicer or Drosha leaves 5′ -phosphate groups at the end of the resultant small RNAs. This feature has been proposed as an important quality check during RISC assembly (3,4,7, 30), and the function has been attributed to the recognition of those phosphate groups within the RISC loading complex (RLC), in part by the asymmetry determining factor, R2D2 (31). However, in the reconstitution system using recombinant hAgo2, the siRNA 5′ -phosphate was shown to be unnecessary for the Slicer activity, although it was important for the stability and the fidelity of the RISC (24). In this chapter, 5′ -phosphorylated siRNA is used to ensure the cleavage activity for purified Argonaute proteins. When synthetic single-stranded siRNA, purchased commercially, is not ′ 5 -phosphorylated, it can be phosphorylated at the 5′ ends using T4 polynucleotide kinase (PNK) in the presence of ATP: 1. Combine the following reaction mixture in a total volume of 30 μL: a. b. c. d. e.

1 μL of single-stranded siRNA (100 μM). 1 μL of ATP (100 mM). 1 μL of T4PNK. 3 μL of 10X PNK reaction buffer. 24 μL of H2 O.

2. Incubate the reaction at 37 ºC for 60 min. 3. Add 15 μL of 3 M NaOAc, 105 μL of nuclease-free water, and 150 μL of phenol/chloroform, and mix thoroughly. Recover the aqueous phase and transfer to a new tube.

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4. Precipitate the RNA by adding 1 volume of isopropanol and 1 μL of pellet-paint® co-precipitant, mixing well. 5. Chill the mixture for at least 15 min at –80 ºC. Centrifuge at 4 ºC for 20 min at maximum speed (20,000 x g) to pellet the RNA. 6. Wash with 70% ethanol.

3.3. Purification of Argonaute Proteins In this section, we perform in vitro target RNA cleavage assay using two particular types of Argonaute proteins. One is recombinant Drosophila AGO1 produced in E. coli. Although recombinant Argonaute proteins are notorious for their difficulty to produce in E. coli, partially because of their low solubility and high toxicity, we were able to obtain GST-tagged, full-length recombinant Drosophila AGO1. The other is endogenous hAgo2 immunopurified from HeLa cells. The monoclonal antibody against hAgo2 was originally made in our laboratory by immunizing mice with the N-terminal region (about 120 amino acids) of hAgo2 expressed in and purified from E. coli and fusing their lymphocytes with myeloma cells. Details of the expression of GST-AGO1 in E. coli and its purification procedures, and hAgo2 immunoprecipitation from HeLa cells, are indicated below. 3.3.1. Production of Recombinant Argonaute Proteins 1. Inoculate an E. coli BL21(DE3) colony containing the transformed plasmid into 2 mL of LB media with 100 μg/mL of Ampicillin (Amp); grow at 37 ºC overnight. 2. Inoculate 100 mL of LB/Amp in a 500-mL flask. Grow for 6–8 h at 37 ºC. 3. Inoculate 500 mL of LB/Amp in a 2-L flask. Grow for 1–2 h at 37 ºC. Check OD600 . Aim for a final OD600 between 0.8 and 1.0. 4. Add 1 M of IPTG to a 1 mM concentration to induce fusion protein expression, and incubate culture at 16 ºC overnight (see Note 1). 5. Spin down cells for 25 min at 3,500g. 6. Discard media and remove the cells into 50 mL tubes. 7. Spin down cells for 15 min at 4,500g. 8. Discard media strictly. (At this point cells can be stocked at –80 ºC for later use.) 9. Resuspend cells in 25 mL of PBS containing protease inhibitor (Complete, Mini, EDTA-free). Keep cells on ice at all times. 10. Sonicate cells for 15 sec five times using 50% output (Branson Sonifier or equivalent). 11. Add 10% Triton X-100 detergent to 1% final concentration to lysis cells. 12. Place tube in rotary shaker at 4 ºC for 20 min. 13. Spin cells for 30 min at 12,000g. Separate supernatant. 14. Set the column containing Glutathione Sepharose beads and wash with PBS. 15. Load the cell lysate into the column and wash the beads three times with PBS.

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16. Add 250 μL of GST elution buffer to the column, to elute GST proteins, and collect the flowthrough. Repeat this step nine times. 17. Electrophoresis 10 μL of each sample and check for the expression of the fusion protein (Fig. 2a). Assay protein concentration of each fraction using Protein Assay.

Fig. 2. Purification of Argonaute proteins. (a) Purification of bacterially expressed Drosophila AGO1. GST-AGO1 was purified with Glutathione Sepharose beads, resolved on an SDS-PAGE gel, and visualized by Coomassie Brilliant Blue staining. The preparation contained two additional proteins (indicated with * and **) other than GST-AGO1 (indicated by an arrow); identified by mass spectrometry as E. coli proteins, Dnak (*) and GroEL (**) (see Note 1). The left lane represents molecular markers (MW). (b) Purification of hAgo2 from HeLa cell lysates by immunoprecipitation with a specific monoclonal antibody (4G8). Endogenous hAgo2 immunopurified with 4G8 and immunoprecipitates with a control non-immune antibody (n.i.) were resolved on SDS-PAGE and visualized by silver staining. 4G8 itself bound on beads was also visualized (4G8 only). The farthest left lane shows molecular markers (MW). h.c. and l.c. show heavy and light chains, respectively, of the antibodies.

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3.3.2. Immunopurification of hAgo2 from HeLa Cells So far, the expression of the full-length recombinant hAgo2 in E. coli has been unsuccessful in our laboratory. Thus, we instead obtained immunopurified endogenous hAgo2 from HeLa cell lysates as follows and used it in the assay. Culture HeLa cells. Harvest 5–10 × 107 cells and wash twice with PBS. Suspend the cells in 500 μL of Binding buffer. Incubate on ice for 5 min. Sonicate briefly (three times, 5 sec each) using 30% output (Branson Sonifier or equivalent). 5. Centrifuge at maximum speed (20,000g) for 25 min. 6. Incubate supernatant with anti-hAgo2 (4G8) antibody immobilized on GammaBind beads. Add NaCl to the lysates to 1 M just before immunoprecipitation is started (see Note 2). Rock the reaction mixtures at 4 ºC for at least 60 min. 7. Wash the beads four times with binding buffer containing 1 M of NaCl buffer and then twice with 1X cleavage buffer (IP-hAgo2 bound with miRNAs) (Fig. 2b). 1. 2. 3. 4.

3.4. In vitro Target RNA Cleavage (Slicer) Assay Previously, Hannon and colleagues showed that immunopurified siRNAprogrammed hAgo2 has Slicer activity in vitro (23). Several other groups also showed quite similar data from in vitro cleavage assays (25,32). However, these experiments were performed with tagged (mostly FLAG-tagged) proteins overexpressed from transfected DNA constructs. Here, we show for the first time that endogenous hAgo2 exhibits Slicer activity (Fig. 3b). In vitro target RNA cleavage assay was performed according to previously reported methods (23,24,33), but with some modifications. 3.4.1. Slicer Assay Using Recombinant Full-Length Drosophila AGO1 Joshua-Tor and colleagues demonstrated an RISC reconstitution system using recombinant hAgo2 and siRNA (24). In our laboratory, in vitro target RNA cleavage assay with recombinant GST-AGO1 was performed according to their method (24), with some modifications. 1. For a 20-μL reaction, assemble on ice: a. b. c. d. 2. 3. 4. 5.

∼100 nM of recombinant Argonaute protein (GST-AGO1) (see Note 3). 4 μL of 5X cleavage buffer. 100 nM of single-stranded, phosphorylated siRNA. 1 μL of RNasin plus (40 U/μL).

Incubate the reaction at 26 ºC for 90 min. Add the target RNA (3,000–5,000 cpm) and 0.5 μg of yeast RNA. Incubate the reaction at 26 ºC for 90 min. Purify the RNA.

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Fig. 3. In vitro target RNA cleavage assay (a) luc guide siRNA-directed target RNA cleavage using recombinant Drosophila AGO1. The 5′ -end radiolabeled RNA target, luc target RNA (also see Fig. 1), was incubated with purified GST-AGO1 pre-bound with luc guide siRNA or with GST itself as a negative control. Schematic drawings of luc target RNA and its cleaved product (cleaved RNA) are indicated on the right, which show where the RNA bands migrated on the gel. (b) miRNA-directed target RNA cleavage using immunopurified hAgo2 from HeLa cells. The 5′ -end radiolabeled RNA target, miR-21 target RNA (also see Fig. 1), was incubated with hAgo2 immunopurified with 4G8, 4G8 itself (a negative control), and immunoprecipitates with non-immune IgG (n.i.: another negative control). Schematic drawings of the RNA target and its cleaved product (cleaved RNA) are indicated on the right, which show where they migrated on the gel. 6. Separate in a 6% acrylamide denaturing gel. 7. Expose the gel to an imaging plate and visualize the signals using the BAS-2500 system (Fig. 3a).

3.4.2. Slicer Assay Using Immunopurified hAgo2 from HeLa Cells 1. For a 30-μL reaction, assemble on ice: a. b. c. d. e. f.

Immunopurified hAgo2 on beads. 6 μL of 5X cleavage buffer. 1 μL of RNasin plus (40 U/μL). 1 μL of yeast RNA (0.5 mg/mL). 1 μL of miR-21 target RNA (3,000–5,000 cpm/μL). 21 μL of H2 O.

2. Incubate the reaction at 37 ºC for 90 min.

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3. Purify the RNA using ISOGEN-LS. 4. Separate in a 6% acrylamide denaturing gel. 5. Expose the gel to an imaging plate and visualize the signals using the BAS-2500 system (Fig. 3b).

4. Notes 1. In our case, GST-AGO1 was copurified with heat-shock proteins, DnaK (Hsp70) and GroEL (29), both known to be chaperonins. It is known that many newly synthesized bacterial proteins avoid aggregation by folding with the aid of chaperonins; thus, the AGO1 protein may be fortuitously solubilized by DnaK and GroEL. Expression of recombinant hAgo2 in E. coli was also successfully done through the coexpression of human HSP90 (24,33). Interestingly, a potential connection between heat-shock proteins and the RNAi pathway has been reported: (a) A recent genome-wide screen for identifying components of the RNAi pathway in Drosophila cells revealed seven genes that affect the RNAi response, two were the heat-shock proteins Hsc70-3 and Hsc70-4 (34); and (b) human HSP90 was shown to be associated with hAgo2 in coimmunoprecipitation studies (23) (also see Note 2). These findings suggest the possibility that some heat-shock proteins may not only be chaperonins for Argonaute proteins but also are critical regulators for Argonaute functions. 2. We intentionally purified hAgo2 under harsh conditions with a salt concentration of 1 M of NaCl, where most of the proteins associated with hAgo2 in cells would ideally be stripped away from hAgo2. Actually, as indicated in Fig. 2a, under such conditions we could immunoprecipitate a good amount of hAgo2 but did not observe any clear protein bands specific in the 4G8-IP lane. Even a band that most likely would correspond to HSP90 was not detected. It could be argued that 1 M of NaCl could lead to the dissociation of small RNAs from hAgo2; however, Northern blotting analysis clearly revealed that this was not the case, as we could still detect miR-21 association with hAgo2 immunopurified from HeLa cells under the harsh conditions (data not shown). Indeed, it has been noted that the potassium chloride concentration in the wash steps of the affinity column purification could be increased up to 2.5 M without loss of RISC activity (36). Thus, it is critical to selectively adjust immunoprecipitation conditions (salt and/or detergent concentrations, incubation time for binding, and so on) according to the specific aim. 3. We normally add Mg2+ to buffers at a final concentration of 2–5 mM when Argonautes are purified from cells. It has been reported that the Slicer activity of Argonaute proteins is Mg2+ -dependent and that Argonautes use an active site Asp-Asp-His (DDH) motif in the PIWI domain for metal ion coordination (35). Through our experiments, we now understand that Mg2+ may also be needed for stabilizing Argonaute proteins. In other words, in vitro target RNA cleavage assay does not go well when Argonautes purified in buffers lacking Mg2+ are used.

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Acknowledgments K. M. is a research fellow supported by the Japan Society for the Promotion of Science (JSPS). This work was supported by grants to M. C. S. and H. S. from the Ministry of Education, Culture, Sports, Science and Technology of Japan, the JSPS, and the New Energy and Industrial Technology Development Organization. M.C.S. is also supported by CREST from Japan Science and Technology Agency.

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4 Identifying siRNA-Induced Off-Targets by Microarray Analysis Emily Anderson, Queta Boese, Anastasia Khvorova, and Jon Karpilow

Summary RNA interference (RNAi) is an endogenous gene regulatory pathway that the research community has adopted to facilitate the creation of a functional map of the human genome. To achieve this, small interfering RNAs (siRNAs), short synthetic duplexes having complete homology to the intended target, are introduced into cells to silence gene expression via a posttranscriptional cleavage mechanism. While siRNAs can be designed to effectively knock down any target gene, recent studies have shown that these small molecules frequently trigger off-target effects. These unintended events can have a significant impact on experimental outcomes and subsequent data interpretation. As RNAi is envisioned to play a central role in developing a functional map of the human genome, the development of reliable protocols for identifying off-targeted genes is essential. This chapter focuses on the underlying features of siRNA-mediated off-targeting and the stateof-the-art methodology used to identify off-targeted genes via microarray-based gene expression analysis. Future adoption of standards in this field will allow a clean distinction between sequence-specific off-target gene regulation and other forms of gene modulation resulting from delivery effects and other events unrelated to the RNAi pathway.

Key Words: siRNA; RNAi; microarray; off-target effect; transfection.

1. Introduction 1.1. miRNA Targeting and siRNA Off-Targeting: The Caveats of Adopting Biological Pathways Detailed studies of the endogenous substrates of the RNAi pathway (miRNA) (Fig. 1) have provided keen insights into the source of siRNA From: Methods in Molecular Biology, vol. 442: RNAi: Design and Application Edited by: S. Barik © Humana Press, Totowa, NJ

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Fig. 1. The endogenous RNAi pathway depicting the key steps associated with the processing of and silencing by miRNAs and synthetic siRNAs. In the endogenous pathway: (1) miRNAs are transcribed forming complex stem-loop pri-miRNA structures; (2) Drosha processes these primary transcripts into pre-miRNAs in the nucleus; (3) pre-miRNAs are transported to the cytoplasm via Exportin 5; (4) Dicer further processes the pre-miRNAs into double-stranded miRNAs with characteristic mismatches and bulges; (5) RISC loads the active strand to form an activated complex and (6a) effect translational repression in cases of partial identity within the 3′ -UTR of a given target or (6b) mRNA cleavage in cases of complete identity. mRNA regulation is also mediated by the introduction of long dsRNAs (a1) or siRNAs (a2) directly into the cytoplasm. In the case of long dsRNAs, these molecules are processed by DICER (b) into siRNAs that interact with RISC (c) and effect RNAi silencing via degradation of the mRNA target (d).

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off-target effects. After undergoing a complex process of maturation (pri-miRNA → pre-miRNA → mature miRNA) that involves both nuclear and cytoplasmic RNase III endonucleases (Drosha and Dicer, respectively), the guide strand of miRNAs mediates translation attenuation of prospective targets through annealing of the miRNA seed region (positions 2–7) to complementary sites in the 3′ UTR of the target gene (1,2). These seemingly simple interactions allow modest (∼2–4-fold) changes in the levels of gene expression and lead to the orchestrated suppression of dozens to hundreds of genes by a single miRNA. While the primary mode of action of siRNAs is distinctly different from that of miRNAs (i.e., target cleavage vs. translation attenuation), there are several overlapping attributes that tie miRNA gene targeting with siRNA off-targeting. Like miRNAs, siRNA off-targeting induces modest changes in the expression of dozens to hundreds of messages. Moreover, studies by a number of groups have demonstrated a strong association between a transcript being an off-target and the presence of one or more siRNA seed complements in the 3′ UTR of off-targeted genes. Given the parallels among the number of targets, the level of gene modulation, and the apparent importance of 3′ UTR target sites for both molecules, the overall sentiment by the research community is that off-targeting by siRNA results from the promiscuous entry of these molecules into the miRNA pathway. 1.2. Techniques for Identifying Off-Targets The mounting evidence that suggests the presence of off-targets has been derived from multiple sources. In an investigation of 10 separate siRNAs targeting the MEN1 gene, Scacheri and colleagues used Western blot analysis to characterize the unexpected variation in the levels p21 and p53, two genes considered to be indicators of overall changes in cellular physiology (3). As each siRNA altered expression of these proteins by differing amounts, the authors predicted that the siRNA affected the expression of targets other than MEN1 and that these unintended targets disproportionately impacted p21 and p53 expression. In a different approach, Fedorov and coworkers employed a phenotypic assay that monitored cell viability to distinguish differences in siRNAs targeting a nonessential housekeeping gene, PPIB (4). Akin to Scacheri et al.‘s findings, each duplex induced distinctly different levels of RNAidependent toxicity, thus lending further support to the notion that siRNAs modulate expression of genes other than the intended target. While these and similar approaches provided evidence of siRNA off-target activity, the most definitive data that quantified off-targeted genes were first reported by Jackson et al. (5). Using microarray-based gene expression

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profiling, the group monitored the genome-wide effects resulting from siRNA transfection and identified unique off-target signatures for multiple duplexes. Subsequent applications of microarray technologies have been responsible for the identification of (1) strong similarities between siRNA off-targeting and miRNA targeting mechanisms (6–8), (2) the importance of the 3′ UTR transcriptome seed complement frequency (SCF) in defining the extent of off-target signatures (9), and (3) strand-specific modification patterns that greatly enhance siRNA specificity (10). Thus, gene expression technologies have served to reveal the global effects of siRNAs and focus efforts toward improving and enhancing siRNA specificity. 1.3. Application of Microarrays to Off-Target Analysis Successful application of microarray-based technologies to off-target studies requires attention to three principal phases: pre-experimental assay optimization, global gene expression profiling, and post-experimental data analysis (Fig. 2). To facilitate the generation of reliable data sets, rigorous optimization of cell lines, plating conditions, delivery methods, and siRNA concentrations must be performed to minimize the confounding effects that can result from small alterations in experimental protocols. Once optimal conditions for potent silencing (>75% reduction of mRNA) without adverse viability effects (100 ng/μL). Store samples at –80 ºC. 3. Assess the quality of the RNA preparation using a 2100 Bioanalyzer (Agilent) with the RNA 6000 Nano LabChip. Acceptable samples should have an RNA Integrity Number (RIN) of at least 9 and can be stored at –80 ºC.

3.4. RNA Amplification and Labeling 1. Amplify total RNA and label using the Low Input RNA Fluorescent Linear Amplification Kit (Agilent, Cat# 5184-3523) according to the manufacturer’s protocol (http://www.chem.agilent.com/). Cy5™ CTP and Cy3™ CTP are obtained from Perkin Elmer (Wellesley, MA). Typically, 650 ng of total RNA is used per reaction. Experimental samples are labeled with Cy5™ while mock (lipid-treated) samples are labeled separately with Cy5™ and Cy3™. Untransfected samples are labeled with Cy3™. Remove excess dye from the labeled samples with Qiagen RNeasy Mini columns, implementing two 1-min. “dry” spins to remove excess ethanol (with a 180 degree rotation of the column in between each spin) before sample elution. 2. Quantify yields of labeled RNA: Yields of labeled samples are quantified with a NanoDrop instrument (NanoDrop). Typical yields are 300–500 ng/μL starting with 650 ng of total RNA and should be uniformly colored depending upon the fluorophore (pink for Cy3™, blue for Cy5™). Assess dye incorporation by measuring the respective dye wavelengths (Cy3™: 550 nm, Cy5™: 650 nm) with the “microarray” protocol on the NanoDrop instrument.

3.5. Hybridization and Scanning 1. Perform hybridizations to Human 1A (V2) microarray (Agilent, Cat# G4110B) according to the manufacturer’s instructions using 750 ng of both Cy5™ and Cy3™ labeled sample per array. Hybridizations include: a. Each siRNA-transfected sample (Cy5™) is hybridized along with the mocktransfected (Cy3™-labeled) sample as a reference. b. A control array for gene expression effects of transfection is assembled by using mock-transfected (Cy5™) and untransfected (Cy3™) samples. c. A control array for assessing signal/expression levels in the absence of siRNA treatment as well as for determining dye-biased targets is assembled with mock-transfected (Cy5) and mock-transfected (Cy3) (a self-self array).

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2. After a 17–20-hour hybridization at 60 ºC, wash arrays for 1 minute each in 6X and then 0.06X SSPE containing 0.025% N-lauroylsarcosine. Subsequently, submerge the arrays in acetonitrile and wash in nonaqueous drying and stabilization buffer (Agilent, Cat# 5185-5979). 3. Scan arrays using Agilent’s model G2505B Microarray scanner and process the raw .tiff images according to the default protocols in Feature Extraction v 9.1 (Agilent).

3.6. Data Analysis 1. Data analysis is initiated in Microsoft Excel. Construct a flat text file of feature identifier columns (Feature Number, Row, Column, Control Type, Gene Name, Systematic Name, and Description if available) and select data columns for each sample (Log Ratio, PValue LogRatio, gProcessedSignal, rProcessedSignal, gIsWellAboveBG, rIsWellAboveBG). Record the processed data for saturated signals from the mock-mock array in separate columns [saturated green (gIsSaturated) and red signals (rIsSaturated), respectively] along with a column for the log of combined red and green signals (gProcessedSignal + rProcessedSignal) computed from the mock-mock array. 2. Conduct further analysis in Spotfire DecisionSite using the Functional Genomics Module. First, filter the data by control and low-signal/low-expression probes to eliminate sequences for which it would be difficult to accurately determine a twofold reduction in signal (Log Ratio –0.3). This is achieved by filtering probes for which the log (combined signal) from the mock-mock array is below 2.8, or approximately 630 intensity units. Assess the absolute range of processed signals among all of the arrays in a batch to make sure that each hybridization was successful. Remove probes with signals that saturate in either channel in the mock-mock array. Typically, by employing these procedures, the final probe set available for analysis is reduced from 21,073 to approximately 13,000–15,000 probes. 3. Remove probes with signals that do not change more than twofold in any sample (–0.3 < Log Ratio < 0.3). Plot the signals for the remaining probes on a heatmap after unsupervised hierarchical clustering (unweighted average, Euclidean distance, average value). Further comparison of siRNAtreated sample columns with mock-mock and mock-untransfected hybridizations will allow probes that appear to be differentially expressed in either of these control arrays or those that are inherently noisy (for instance, Log Ratio vs. mock of 90% mRNA knockdown, 100 nM) targeting a nonessential gene (GAPDH, PPIB). In cases where cells are refractory to lipid-mediated delivery of siRNA, electroporation and DNA-based RNAi are often evaluated. Electroporation induces fewer nonspecific gene expression changes than lipid-mediated transfection (11), yet to minimize cytotoxicity this method still requires extensive optimization around parameters such as siRNA concentration, cell density, buffers, voltage, pulse

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Fig. 4. Effects of lipid-based transfection on gene expression profiles. HeLa cells were transfected with four different lipid-based transfection in the presence and absence of a control siRNA (100 nM). As shown by the heatmaps for each treatment (normalized to mock or untransfected), lipid-based transfection induce both common and unique (boxes) changes in gene expression.

length and number, and interpulse interval length. Several manufacturers have developed specific buffers and preprogrammed protocols for cell types (e.g., amaxa Nucleofector® II), while alternative platforms provide extended flexibility by allowing for tailored protocols (e.g., Bio-Rad Gene Pulser XL). Viral delivery systems offer another alternative for siRNA (or small hairpin RNA, shRNA) delivery and offer the added benefit of long-term silencing. Still, it is important to consider the potential shortcomings of these systems. First, work by Grimm and associates at Stanford (14) demonstrated that expression of shRNA from a viral construct can interfere with the processing of native RNAi substrates, miRNAs. Given this finding, it may be difficult to distinguish off-targets from changes in gene expression that result from perturbations in the endogenous miRNA processing mechanism. Second, because DNA-based RNAi provides long-term silencing, the profile changes observed in transduced cells will include both off-target effects as well as downstream effects resulting from silencing of the target gene. Differentiation between these two classes can be achieved

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by simultaneously performing control studies with a second (equally functional) shRNA targeting the same gene, but this requires obvious additional costs and data analysis to correctly identify sequence-specific off-targets. Last, while there are multiple, commercially available custom viral-based silencing services (e.g., OpenBiosystems, Sigma-Aldrich), very little is known about how these molecules are processed. Studies by Vermeulen et al. demonstrated that Dicer processing of long siRNA (e.g., >23-mer duplexes) can often lead to multiple products (15). Given that the number and identity of most off-targets are determined by the 5′ terminal region of the guide strand (e.g., the seed region), the diverse population of molecules created by successive Drosha and Dicer processing of expressed hairpins could conceivably generate an expanded number of off-targeted genes. 6. siRNA attributes: Researchers planning to perform microarray-based off-target studies should also be conscious of the attributes associated with siRNA that contribute to the size, makeup, and intensity of the off-target signature. Studies by several groups have now demonstrated that siRNA off-targets are concentrationdependent (5), with off-target (and on-target) gene knockdown generally diminishing between 1–10 nM when lipid-mediated delivery technologies are employed. For these reasons, we typically perform off-target analysis at concentrations between 25–100 nM, which allows for clear identification of genes that are downregulated within 24 h after transfection. In addition to concentration, strand selection and nucleotide content play a significant role in the off-target signature. Off-target events can be induced by either strand of the siRNA duplex and both bioinformatic strategies and chemical modifications can be used to influence the contribution of either strand to the overall signature. Most siRNA rational design algorithms (which select functional sequences on the basis of thermodynamic and recognized nucleotide positional preferences) include differential end stability as a key element in siRNA selection to ensure RISC entry of the antisense strand. This process, which selects for disproportionate levels of stability at the duplex termini, can greatly minimize the sense strand off-target profiles and greatly simplify subsequent analysis. In addition, chemical modification patterns can also be incorporated into the sense and/or antisense strand to alter the off-target signature of each strand, respectively (10).

Specifically, modification patterns have now been identified that can (a) interfere with a strand’s ability to enter RISC or (b) reduce the efficiency at which nonidentical pairings are permitted (10). While these techniques can help simplify subsequent off-target analysis, it is critical to note that in some instances, chemical modifications (as well as base-pair mismatches) can significantly alter off-target profiles. For example, in some cases, elimination of sense strand competition by chemical modification enhances the functionality of the antisense strand and amplifies the associated signature. In contrast, the addition of mismatches within the seed region (positions ∼2–7) has been shown to shift

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off-target profiles, further supporting bioinformatic studies that suggest siRNA off-targeting is similar to miRNA targeting (7). Recent findings by our group have also shown that the number of off-targets generated by siRNA strongly correlates with the frequency at which the seed complement appears in the 3′ UTR transcriptome (referred to as the Seed Complement Frequency, SCF). siRNA with low SCF generally induce modest off-target profiles, while duplexes with medium-to-high SCF may exhibit more robust signatures. As the observed association has direct consequences on the rate at which false positives are generated in siRNA-mediated phenotypic screens, performing a preliminary bioinformatic analysis of the SCF for any siRNA can provide insights into the potential number and identity of offtargeted genes and aid in the selection of a duplex for future studies. While concentration, strand selection, and seed frequency represent properties that can alter the off-target signature in an RNAi-dependent fashion, the length, structure, and sequence content of an siRNA can impact expression profiles through RNAi-independent mechanisms (see Note 6). Long siRNA (>23-mers) can induce a broad collection of IFN-response genes and toxicity at concentrations as low as 10 nM (25). In addition, particular sequence motifs and terminal end structures have likewise been shown to activate the innate immune response (26,27). Given these observations, we consistently utilize siRNA that are less than 23 bp in length and have been rationally designed to (a) bias antisense strand entry into RISC, (b) eliminate known sequences that induce the innate immune response, and (c) utilize seeds regions that have low SCFs.

References 1. Rehwinkel, J., Behm-Ansmant, I., Gatfield, D., and Izaurralde, E. (2005). A crucial role for GW182 and the DCP1:DCP2 decapping complex in miRNA-mediated gene silencing. RNA 11, 1640–1647. 2. Rossi, J. J. (2005). RNAi and the P-body connection. Nat. Cell Biol. 7, 643–644. 3. Scacheri, P. C., Rozenblatt-Rosen, O., Caplen, N. J., et al. (2004). Short interfering RNAs can induce unexpected and divergent changes in the levels of untargeted proteins in mammalian cells. Proc. Natl. Acad. Sci. USA 101, 1892–1897. 4. Fedorov, Y., Anderson, E. M., Birmingham, A., et al. (2006). Off-target effects by siRNA can induce toxic phenotype. RNA 12, 1188–1196. 5. Jackson, A. L., Bartz, S. R., Schelter, J., et al. (2003). Expression profiling reveals off-target gene regulation by RNAi. Nat. Biotechnol. 21, 635–637. 6. Lin, X., Ruan, X., Anderson, M. G., et al. (2005). siRNA-mediated off-target gene silencing triggered by a 7 nt complementation. Nucleic Acids Res. 33, 4527–4535.

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7. Jackson, A. L., Burchard, J., Schelter, J., et al. (2006). Widespread siRNA “offtarget” transcript silencing mediated by seed region sequence complementarity. RNA 12, 1179–1187. 8. Birmingham, A., Anderson, E. M., Reynolds, A., et al. (2006). 3′ UTR seed matches, but not overall identity, are associated with RNAi off-targets. Nat. Meth. 3, 199–204. 9. Anderson, E. M., Birmingham, A., Baskerville, S., et al. (2008). Experimental Validation of the Importance of Seed Frequency to siRNA Specificity. RNA (in press). 10. Jackson, A. L., Burchard, J., Leake, D., et al. (2006). Position-specific chemical modification of siRNAs reduces “off-target” transcript silencing. RNA 12, 1197–1205. 11. Fedorov, Y., King, A., Anderson. E., et al. (2005). Different delivery methods— Different expression profiles. Nat. Meth. 2, 241. 12. Spagnou, S., Miller, A., and Keller, M. (2004). Lipidic carriers of siRNA: Differences in the formulation, cellular uptake, and delivery with plasmid DNA. Biochemistry 43, 13348–13356. 13. Lv, H., Zhang, S., Wang, B., Cui, S., and Yan, J. (2006). Toxicity of cationic lipids and cationic polymers in gene delivery. J. Control Release 114, 100–109. 14. Grimm, D., Streetz, K. L., Jopling, C. L., et al. (2006). Fatality in mice due to oversaturation of cellular microRNA/short hairpin RNA pathways. Nature 441, 537–541. 15. Vermeulen, A., Behlen, L., Reynolds, A., et al. (2005). The contributions of dsRNA structure to Dicer specificity and efficiency. RNA 11, 1–9. 16. Harborth, J., Elbashir, S. M., Vandenburgh, K., et al. (2003). Sequence, chemical, and structural variation of small interfering RNAs and short hairpin RNAs and the effect on mammalian gene silencing. Nucleic Acid Drug Dev. 13, 83–105. 17. Khvorova, A., Reynolds A., and Jayasena, S. (2003). Functional siRNAs and miRNAs exhibit strand bias. Cell 115, 209–216. 18. Chalk, A. M., Wahlestedt, C., and Sonnhammer, E. L. (2004). Improved and automated prediction of effective siRNA. Biochem. Biophys. Res. Commun. 319, 264–274. 19. Ding, Y., Chan, C. Y., and Lawrence, C. E. (2004). Sfold web server for statistical folding and rational design of nucleic acids. Nucleic Acids Res. 32(Suppl_2), W135–W141. 20. Henschel, A., Buchholz, F., and Habermann, B. (2004). DEQOR: A web-based tool for the design and quality control of siRNAs. Nucleic Acids Res. 32(Suppl_2), W113–W120. 21. Naito, Y., Yamada, T., Ui-Tei, K., Morishita, S., and Saigo, K. (2004). siDirect: Highly effective, target-specific siRNA design software for mammalian RNA interference. Nucleic Acids Res. 32(Suppl_2), W124–W129. 22. Pancoska, P., Moravek, Z., and Moll, U. M. (2004). Efficient RNA interference depends on global context of the target sequence: Quantitative analysis of silencing efficiency using Eulerian graph representation of siRNA. Nucleic Acids Res. 32, 1469–1479.

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23. Reynolds, A., Leake, D., Boese, Q., Scaringe, S., Marshall, W. S., and Khvorova, A. (2004). Rational siRNA design for RNA interference. Nat. Biotechnol. 22, 326–330. 24. Schubert, S., Grunweller, A., Erdmann, V. A., and Kurreck, J. (2005). Local RNA target structure influences siRNA efficacy: Systematic analysis of intentionally designed binding regions. J. Mol. Biol. 348, 883–893. 25. Reynolds, A., Anderson, E. M., Vermeulen, A., et al. (2006). Induction of the interferon response by siRNA is cell type—and duplex length—dependent. RNA 12, 1–6. 26. Judge, A. D., Bola, G., Lee, A. C., and MacLachlan, I. (2006). Design of noninflammatory synthetic siRNA mediating potent gene silencing in vivo. Mol. Ther. 13, 494–505. 27. Marques, J. T., Devosse, T., Wang, D., et al. (2006). A structural basis for discriminating between self and nonself double-stranded RNAs in mammalian cells. Nat. Biotechnol. 24, 559–565. 28. Bammler, T., Beyer, R. P., Bhattacharya, S., et al. (2005). Standardizing global gene expression analysis between laboratories and across platforms. Nat. Meth. 2, 351–356. 29. Irizarry, R. A., Warren, D., Spencer, F., et al. (2005). Multiple-laboratory comparison of microarray platforms. Nat. Meth. 2, 345–350. 30. Larkin, J. E., Frank, B. C., Gavras, H., Sultana, R., and Quackenbush, J. (2005). Independence and reproducibility across microarray platforms. Nat. Meth. 2, 337–344. 31. MAQC Consortium: Shi, L., Reid, L. H., Jones, W. D., et al. (2006). The MicroArray Quality Control (MAQC) project shows inter- and intraplatform reproducibility of gene expression measurements. Nat. Biotechnol. 24, 1151–1161. 32. Patterson, T. A., Lobenhofer, E. K., Fulmer-Smentek, S. B., et al. (2006). Performance comparison of one-color and two-color platforms within the MicroArray Quality Control (MAQC) project. Nat. Biotechnol. 24, 1140–1150. 33. Dobbin, K., and Simon, R. (2002). Comparison of microarray designs for class comparison and class discovery. Bioinformatics 18, 1438–1445. 34. Dobbin, K., Shih, J. H., and Simon, R. (2003). Questions and answers on design of dual-label microarrays for identifying differentially expressed genes. J. Natl. Cancer Inst. 95, 1362–1369.

II Application of RNAi in Diverse Organisms

5 Hydrodynamic Delivery of siRNA in a Mouse Model of Sepsis Doreen E. Wesche-Soldato, Joanne Lomas-Neira, Mario Perl, Chun-Shiang Chung, and Alfred Ayala

Summary The use of siRNA in vitro as well as in animal models has become more widespread in recent years, leading to further questions as to the best mode of delivery that will achieve optimal knockdown. While the exact mechanism of siRNA uptake at a cellular level has yet to be fully elucidated, various delivery techniques are being researched, including the use of viral vectors of shRNA, liposome encapsulations, and hydrodynamic delivery of naked siRNA. We describe the use of hydrodynamic administration as a technique to deliver, in vivo, naked siRNA constructs into experimental animals as a method of transient gene knockdown. This method may prove useful in situations where knockout animals do not exist, or to determine the effect of gene knockdown at specific time points during an experiment.

Key Words: siRNA; hydrodynamics; sepsis; mice. 1. Introduction Hydrodynamic infusion was first described with plasmid DNA (1) in which plasmid DNA containing cDNA of luciferase and -galactosidase as reporter genes was given as a large-volume, rapid tail vein injection. Among the organs expressing the transgene, including the lung, spleen, heart, kidney, and liver, the liver showed the highest level of gene expression (see Note 1). This study also demonstrated that volume and rate of injection were critical in the uptake of naked constructs, in that the plasmid had to be in a large fluid volume (i.e., equivalent of about one time the animal’s blood volume) rapidly injected. From: Methods in Molecular Biology, vol. 442: RNAi: Design and Application Edited by: S. Barik © Humana Press, Totowa, NJ

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One of the first studies that utilized this technique for siRNA showed that approximately 88% of hepatocytes take up siRNA after a hydrodynamic injection of 50 μg of Cy5-tagged siRNA in 2 mL (2). It should also be noted that fluorochrome-tagged siRNA is said to be not functional but can be useful for determining uptake of siRNA in specific cell types. Our own localization studies using GFP siRNA in GFP transgenic mice showed, in accordance with Liu et al., that siRNA was taken up in all major organs (3). In contrast to what has been suggested in vitro and with viral vectors (4–6), our lab and others have found no evidence of naked siRNA stimulating the interferon pathway in vivo (3,7,8). This latter point is not trivial, as the administration of any agent that can induce inflammation, can be a confounding variable in the examination of a given experimental pathology. 2. Materials 2.1. siRNA 1. Custom siRNA (19–21-mer duplexes) (Dharmacon, Lafayette, CO) 2′ deprotected, duplexed, desalted (ready to use). Predesigned siRNAs are also available using siGENOME™ (Dharmacon). 2. Control siRNA: GFP siRNA (Dharmacon) can be used, or a scrambled version of the experimental siRNA can be used. Scrambled sequences must undergo a BLAST search to rule out specificity against other possible sequences in the mouse genome, i.e., off-target effects. 3. Ice-cold sterile PBS.

2.2. Animal Groups and Injection Materials 1. The strain of mouse to be used, divided into at least three groups. Group 1: Saline control; group 2: nonsense siRNA control (GFP or scrambled siRNA); and group 3: siRNA-treated group. 2. Alcohol prep pads. 3. 25-gauge needles and 3-mL (cc) syringes. 4. A heat source: a heat lamp or a 55 °C water bath and gauze. 5. Rodent restraint.

2.3. Materials for Cecal Ligation and Puncture (CLP) 1. 2. 3. 4. 5. 6.

Anesthesia setup and isoflurane (Abbott Laboratories, North Chicago, IL). Surgical instruments including forceps, surgical scissors, and a needle holder. Lidocaine (Abbott Laboratories). Lactated Ringer’s solution (Baxter Healthcare Corp., Deerfield, IL). Sterile surgical ties. Electric razor to shave animal’s abdomen fur (Wahl).

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7. Betadine solution (Purdue Products, Stamford, CT). 8. 6-0 Ethilon suture material (ETHICON, Inc., Somerville, NJ).

3. Methods The development of clinical as well as pharmaceutical interventions has proven to be a difficult task in treating sepsis. As human and animal sepsis is a complex condition, the timing of the therapy is critical (see Note 2). That said, as it is not known beforehand if an individual will become septic, possible treatments must be able to be given after the onset of sepsis, or as a “posttreatment.” In this method, hydrodynamic delivery of siRNA is given 30 min post-CLP and as late as 12 h post-CLP. The idea of whether the injection can be delivered while the animal is under anesthesia is still of some controversy. While the protocol outlined here was performed in our laboratory without anesthesia, other labs have employed a light dosage of either isoflurane (9) or pentobarbital anesthesia (10) to carry out the hydrodynamic injection without morbidity or mortality (see Note 3). 3.1. Cecal Ligation and Puncture 1. The surgical procedure to generate sepsis, as previously described (11), can be carried out in any number of animal strains (i.e., outbred, inbred, knockout, etc.). We have commonly used C3H/HeN, Balb/C, or C57BL6 male mice. 2. The mice are lightly anesthetized using isoflurane. The abdomen is shaved and scrubbed with Betadine. 3. A midline incision (1.5–2 cm) is made below the diaphragm first in the skin and then in the muscle layer. 4. The cecum is isolated, ligated, and punctured twice with a 22-gauge needle. The cecum is then gently compressed to extrude a small amount of cecal material from each of the punctures. 5. The cecum is returned to the abdomen, and the muscle and skin incisions are then closed with 6-0 Ethilon suture material. Before suturing the skin, 2–3 drops of Lidocaine are administered to the wound for analgesia. 6. The mice are subsequently resuscitated with 1.0 mL of Lactated Ringer’s solution subcutaneously. Sham controls are subjected to the same surgical laparotomy and cecal isolation, but the cecum is neither ligated nor punctured.

3.2. Hydrodynamic Delivery of siRNA 1. Prior to use, the siRNA must be reconstituted and aliquoted. By following the amount provided in the tube (in μg), add enough sterile PBS to yield a concentration of 5 μg/μL, or 50 μg/10 μL. Separate the reconstituted siRNA into 10-μL aliquots, and store in –20 °C. If siRNA is tagged with a fluorochrome, be sure to protect from light.

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2. For each mouse to receive a hydrodynamic injection, add 1.5 mL of sterile PBS to an Eppendorf tube. Thaw a 10-μL aliquot of siRNA per mouse and add to the 1.5 mL of PBS. This will equate to 50 μg of siRNA per mouse. 3. Draw up the volume of siRNA into a 3-cc syringe capped with a 25-gauge needle. Make sure to purge the syringe and needle of air bubbles. 4. Restrain the mouse in the rodent restraint in such a way that the tail is completely exposed. For this aspect, short-term exposure to a volatile anesthetic like isoflurane can be utilized during restraint of the animal, but the anesthetic should be taken off just prior to the actual tail vein injection. 5. Clean the tail with an alcohol prep and place the heat source over the tail in order to dilate the veins, making them more visible. Three vascular structures will become visible, one in the middle and one on each side. Choose one of the “side” tail veins and proceed to inject the volume of siRNA within 5 sec. If the needle is placed correctly in the vein, the volume will inject in very easily. Upon completion of the injection, remove the needle and immediately hold a piece of gauze over the injection site until any bleeding has stopped and then return the mouse to its cage.

4. Notes 1. One way to determine the distribution of siRNA would be to inject GFP siRNA into GFP transgenic mice C57BL/6-TgN (ACTbEGFP)/Osb (Jackson Labs). In these mice, GFP constitutive transcription is under the control of a chicken betaactin promoter and a cytomegalovirus enhancer, making all of the tissues, with the exception of erythrocytes and hair, appear green under ultraviolet excitation light (12). At the desired time point, mice are killed and tissues of interest are harvested and prepared for frozen sectioning. The tissues are then viewed using a fluorescent microscope and compared to that of a GFP mouse receiving a hydrodynamic injection of only PBS as a control. Our own localization studies using GFP siRNA in GFP transgenic mice showed that siRNA was taken up in all major organs (3). Localization of siRNA can also be determined using a fluorochrome-tagged siRNA, such as FITC or rhodamine (2). The siRNA manufacturer can add these tags to your custom siRNA. As with the GFP mice, wild-type mouse tissues that received a hydrodynamic injection of fluorochrome-tagged siRNA can also be viewed under fluorescence microscopy to look for the presence of the siRNA in the tissues. This approach is potentially also amenable to dual or three-color flow cytometric analysis, which might also allow potential assessment of cellular as well as tissue-specific targeting. 2. To assess the extent and duration of siRNA silencing in vivo at the mRNA level, RT-PCR or quantitative RT-PCR can be done on the tissues of interest using primers for the gene you are attempting to silence. For protocols concerning RNA isolation for primer production, RT-PCR, or quantitative RT-PCR, the reader is referred to the Current Protocols in Immunology series (John Wiley & Sons, Inc.).

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To assess siRNA silencing at the protein level, a Western blot can be performed using protein from the tissues of interest. Assuming one has a relatively specific poly- and/or monoclonal antibody, which can be utilized for Western blotting, the reader is again referred to Current Protocols in Immunology for details on tissue/cell lysis, gel electrophoresis, protein transfer, and detection. The extent of siRNA-induced silencing on a cellular level can also be determined by flow cytometry if antibodies for the siRNA target gene product of interest and given cell phenotype exist as direct fluorochrome conjugates. This may also be possible by immuno-histochemical or whole-tissue section approaches but requires more controls and blinded assessment or a method for clearly quantitating image probe intensity, as siRNA treatment typically produces gene knockdown (partial suppression) and not a complete knockout. 3. As mentioned in the introduction to this chapter, inflammation may be of some concern in that the in vivo infusion of siRNA (being a double-stranded RNA) might induce an inflammatory response through the activation of the interferon or the classic NF-B pathway (5). We have shown, however, that animals treated solely with siRNA did not show an increase in plasma IFN- levels as did animals treated with poly I:C (3). IFN- is well documented to rise in response to TLR3, PKR, and/or IRF-STAT1 activation by double-stranded RNA (13,14). This would suggest that hydrodynamic delivery of siRNA at this concentration differs distinctly from alternative forms of double- or single-stranded RNAs typically derived from viruses or poly (I:C), as a stimulant of interferon activation. In addition, naked siRNA constructs given locally in the form of an intratracheal instillation did not induce local tissue inflammation (7). 4. Off-target effects can occur if your siRNA sequence has homology to mRNA transcripts other than your gene of interest (15). In this case, transcripts that are not related to your gene of interest may be silenced as well. Off-target effects can be significantly reduced by careful selection of your target sequence, thorough sequence validation, and adequate experimental controls. To optimize gene silencing, it is highly recommended that you test more than one siRNA target sequence per gene (particularly if your target gene has not been published or tested by the company that produces the siRNA). If the results are consistent from at least two different siRNA target sequences, it is more likely that any biological effects observed in your experiments are specifically due to loss of the gene of interest. 5. Due to the nature of the hydrodynamic injection, there are obvious limitations in that an injection of this volume and rate cannot be given clinically. However, the primary objective of such an approach is, first, to document the significance, i.e., contribution, a given gene makes to the pathology seen in a given experimental model. Second, as is the case in our studies of sepsis, it is also possible to document that siRNA can be used in a posttreatment scenario (3). In those studies we also found that hydrodynamic fluid delivery had no marked deleterious effects when administered at 30 min, 1 h, or 12 h post-CLP. If anything, there was a transient survival benefit at 36 h post-CLP, which was subsequently lost

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Wesche-Soldato et al. (3). It is also important to appreciate that all arms of a study such as this should receive hydrodynamic fluid administration, so that the differences seen remain independent of those effects simply attributable to large volume fluid administration alone. With respect to the duration of target gene suppression, we found that the suppression of both message and protein production in the liver lasts up to 10 days in our model (3), but we have not explored the nature of this persistence. At present, we can only speculate that cells in the liver under in vivo conditions, which do not divide at the frequency of cell lines in culture (to which comparisons are made), lead to less dilution of the functional siRNA levels present and thus may allow the silencing effect to persist much longer (2). While this is only a prolonged but not permanent suppression, toxicity has not been a concern thus far. This, unfortunately, is not the case with vectored short hairpin (sh) RNA constructs. Recent studies suggest cumulative toxicity may be observed with such vectored systems over time (5). In this case, however, prolonged suppression allows for further time points to be studied in which to analyze the effect of gene target knockdown. The duration of knockdown is most likely also cell- or tissue type-specific, so this needs to be redetermined based on the tissue of interest. In conclusion, hydrodynamic delivery of naked siRNA constructs, while not necessarily a clinically significant modality, represents a potentially valuable tool for the in vivo transient silencing of select target genes in any number of experimental settings in mice. Here we have provided a basic protocol for this approach and have attempted to outline some of the controls as well as concerns the investigator should keep in mind. Finally, as more is understood about the mechanism of RNAi action, its delivery, and the uptake of siRNA into cells, it is likely this method will be modified or an alternative developed for better use in the clinical setting.

References 1. Liu, F., Song, Y. K., and Liu, D. (1999). Hydrodynamics-based transfection in animals by systemic administration of plasmid DNA. Gene Ther. 6, 1258–1266. 2. Song, E., Lee, S. K., Wang, J., et al. (2003). RNA interference targeting Fas protects mice from fulminant hepatitis. Nat. Med. 9, 347–351. 3. Wesche-Soldato, D. E., Chung, C. S., Lomas-Neira, J., Doughty, L. A., Gregory, S. H., and Ayala, A. (2005). In vivo delivery of caspase 8 or Fas siRNA improves the survival of septic mice. Blood 106, 2295–2301. 4. Sledz, C. A., Holko, M., de Veer, M. J., Silverman, R. H., and Williams, B. R. (2003). Activation of the interferon system by short-interfering RNAs. Nat. Cell Biol. 5, 834–839. 5. Bridge, A. J., Pebernard, S., Ducraux, A., Nicoulaz, A.-L., and Iggo, R. (2003). Induction of an interferon response by RNAi vectors in mammalian cells. Nat. Genet. 34, 263–264.

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6. Kariko, K., Bhuyan, P., Capodici, J., and Weissman, D. (2004). Small interfering RNAs mediate sequence-independent gene suppression and induce immune activation by signaling through Toll-like receptor 3. J. Immunol. 172, 6545–6549. 7. Perl, M., Chung, C. S., Lomas-Neira, J., et al. (2005). Silencing of Fas- but not caspase-8 in lung epithelial cells ameliorates experimental acute lung injury. Am. J. Pathol. 167, 1545–1559. 8. Heidel, J. D., Siwen, H., Liu, X. F., Triche, T. J., and Davis, M. E. (2004). Lack of IFN response in animals to naked siRNAs. Nat. Biotechnol. 22, 1579–1582. 9. Sebestyen, M. G., Budker, V. G., Budker, T., et al. (2006). Mechanism of plasmid delivery by hydrodynamic tail vein injection. I. Hepatocyte uptake of various molecules. J. Gene Med. 8, 852–873. 10. Feng, D. M., He, C. X., Miao, C. Y., et al. (2004). Conditions affecting hydrodynamics-based gene delivery into mouse liver in vivo. Clin. Exp. Pharmacol. Physiol. 31, 850–855. 11. Ayala, A., Herdon, C. D., Lehman, D. L., Ayala, C. A., and Chaudry, I. H. (1996). Differential induction of apoptosis in lymphoid tissues during sepsis: Variation in onset, frequency, and the nature of the mediators. Blood 87, 4261–4275. 12. Okabe, M., Ikawa, M., Kominami, I., Nakanishi, T., and Nishimune, Y. (1997). “Green mice” as a source of ubiquitous green cells. FEBS Lett. 407, 313–319. 13. Levy, D. E. (2002). Whence interferon? Variety in the production of interferon in response to viral infection. J. Exp. Med. 195, F15–F18. 14. Asselin-Paturel, C., Boonstra, A., Dalod, M., et al. (2001). Mouse type I IFNproducing cells are immature APCs with plasmacytoid morphology. Nat. Immun. 2, 1144–1150. 15. Jackson, A. L., Bartz, S. R., Schelter, J., et al. (2003). Expression profiling reveals off-target gene regulation by RNAi. Nat. Biotechnol. 21, 635–637.

6 Nasal Delivery of siRNA Vira Bitko and Sailen Barik

Summary The intranasal administration of siRNA has opened new vistas in drug delivery and respiratory therapy. In this strategy, synthetic siRNA with or without chemical modifications can be applied intranasally. Various delivery vehicles have been tested and optimized. With a few exceptions, all promote significant uptake of siRNA into the lung tissue and offer protection against respiratory viruses such as respiratory syncytial virus (RSV), parainfluenza virus (PIV), and influenza virus. No major adverse immune reaction has been encountered. Nasally applied siRNA remains within the lung and does not have systemic access, as judged by its absence in other major organs such as the lung, liver, heart, and kidney. We provide techniques for using the nose as a specific route for siRNA delivery into the lung of laboratory animals, which has enormous potential for clinical applications.

Key Words: siRNA; intranasal; RNAi; antiviral; RSV; parainfluenza; influenza.

1. Introduction Small interfering siRNAs (siRNAs) have emerged as a promising tool to downregulate gene expression in worms, plants, animals, and humans (1–8) with great potential to serve both as a research tool and as a therapeutic modality for targeted knockdown of disease-causing genes. Several studies have demonstrated the possibility of using siRNA to treat viral infections (7–17), inflammatory diseases (18,19), cancer (20,21), and genetic (22,23) and ocular (16,24–27) diseases. The success of siRNA in RNAi is due to its complementarity to its target mRNA. RNAi includes several steps (see Fig. 1 in Chapter 4) (28). In brief, the antisense strand of the siRNA, as part of From: Methods in Molecular Biology, vol. 442: RNAi: Design and Application Edited by: S. Barik © Humana Press, Totowa, NJ

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Fig. 1. Nasal instillation in mice. From left: gloved hand, anesthetized white BALB/c mouse, pipette tip being used for instillation. Note that the proper angle of the tip relative to the nostril is important to prevent spillage of solution and nosebleed.

the RNA-induced silencing complex (RISC), serves as the guide strand to engage the complementary mRNA that is then cleaved by the endonuclease component of RISC, namely Argonaute 2, resulting in gene silencing. Perhaps the most promising area of siRNA application is in vivo, which allows a close examination of gene function in living organisms and combating of viral infections. Application of RNAi in vivo is, however, still in its early stages, specific delivery being a significant problem. The two major routes of siRNA delivery are systematic and local [e.g., ocular (25–27), cerebral (29,30), peritoneal (31), and nasal (7,8,10,32)]. The nasal route presents an interesting regimen since it allows noninvasive means of delivery of siRNA to lung cells, which is experimentally and therapeutically useful. It can also be useful in treating such debilitating respiratory tract disorders as chronic obstructive pulmonary disease (COPD), cystic fibrosis, asthma, as well as many viral infections of the lung.

2. Materials We have successfully used the reagents described below, but various equivalents are available commercially that can be optimized. We also list a number of popular Web sites that offer useful advice on various aspects of RNAi (Table 1).

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Table 1 Useful Web Sites URL www.ambion.com

www.dharmacon.com

www.invitrogen.com

www.mirusbio.com

Content Predesigned and custom siRNA, siRNA design algorithm, transfection reagents for in vivo and in vitro siRNA transfection Custom siRNA design at the siDesign™ Center, large selection of predesigned siRNA targeting most human, mouse, and rat genes, transfection reagents Custom siRNA using the BLOCK-iT™ RNAi Designer algorithm, Stealth™ Select RNAi (predesigned sequences to the majority of the human, rat, and mouse genes) Transfection reagents for in vivo and in vitro siRNA delivery, reagents for siRNA localization and tracking, fluorescent delivery controls

2.1. RNA Work 1. DEPC-treated water (Sigma-Aldrich, St. Louis, MO). 2. RNase-free ART® aerosol resistant pipette tips (Molecular BioProducts, San Diego, CA). 3. RNase-free microfuge tubes (Ambion, Austin, TX). 4. 9.5 M of ammonium acetate. 5. 95% and 100% Ethanol. 6. 2′ -Deprotection buffer: 100 mM acetic acid-TEMED, pH 3.8 (Dharmacon, Lafayette, CO). 7. siRNA buffer: 20 mM KCl, 6 mM HEPES-KOH, pH 7.5, 0.2 mM MgCl2 .

2.2. Animals and Anesthesia 1. BALB/c mice, 8–10 weeks old, weighing 16-20 g (Charles River Laboratories, Wilmington, MA). 2. 5 mg/mL of sodium pentobarbital (Nembutal™). 3. 25-gauge single-use needles (VWR, Westchester, PA). 4. 1-cc single-use syringes with BD Luer-Lok tip (VWR).

2.3. siRNA Transfection 1. TransIT-TKO siRNA transfection reagent (Mirus Bio Corporation, Madison, WI). 2. Opti-MEM I Reduced Serum Medium (Gibco™, Invitrogen Corporation, Carlsbad, CA). 3. RNase-free gel-loading microcapillary tips (VWR). 4. RNase-free microfuge tubes (Ambion).

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3. Methods 3.1. siRNA 2′ -Deprotection, Annealing, and Desalting 1. Add 200 μL of 2′ -deprotection buffer to tube containing 0.1 μmol of 2′ -ACE protected, single-stranded RNA. 2. Combine the volumes of complementary strands of RNA, vortex for 10 sec, and centrifuge for 30 sec. 3. Heat the mixture at 60 °C for 45 min. 4. Remove from heat and centrifuge briefly, 5–10 sec. 5. Allow solution to cool to room temperature over 30 min. 6. Add 40 μL of 9.5 M ammonium acetate and 1.5 mL of 100% ethanol to the 400 μL of siRNA duplex solution, and vortex. 7. Place the tubes at –20 °C for >16 h or at –70 °C for 2 h. 8. Centrifuge at 14,000 x g for 30 min at 4 °C. 9. Carefully remove the supernatant away from the pellet. 10. Rinse the pellet with 200 μL of ice-cold 95% ethanol. 11. Dry under vacuum using Speed-Vac. 12. The dry pellet can be stored at –20 °C until use or resuspended in an appropriately buffered solution (20 mM of KCl, 6 mM of HEPES-KOH, pH 7.5, 0.2 mM of MgCl2 ).

3.2. siRNA–Vehicle Complex Formation Perform siRNA complex formation immediately before nasal administration. Determine the optimal siRNA amount by titrating from 3 to 15 nmol per mouse (see Note 2). 1. In a sterile, RNase-free plastic tube, add 35 μL of Opti-MEM reduced-serum medium. 2. Add 5 μL of the TransIT-TKO transfection reagent into the tube containing the Opti-MEM medium. 3. Mix thoroughly by vortexing for 10 sec. 4. Incubate at room temperature for 10 min. 5. Add desired amount of siRNA in 1 μL of siRNA buffer to the diluted TransITTKO reagent. 6. Carefully mix by gentle pipetting. 7. Incubate at room temperature for 20 min.

3.3. Animal Anesthesia Prior to nasal administration of siRNA, the mouse must be anesthetized to minimize any pain or discomfort. Nembutal is administered by intraperitoneal (IP) injection. The recommended drug dosage for mice is 50 mg/kg (see Note 1). 1. Gently lift the mouse by the tail and place it on a cage lid. 2. Grip the loose skin of the neck to immobilize the head of the animal.

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3. With the head immobilized, extend the tail to draw the skin tight over the abdomen by gripping the tail with your little finger. 4. The animal should be held in a head-down position. 5. Disinfect injection site. 6. Insert the hypodermic needle into the lower right or left quadrant of the abdomen. 7. Place animal back into the cage and wait until anesthesia takes effect. 8. The animal is ready for siRNA administration when no voluntary movement is observed.

3.4. siRNA Administration 1. Place anesthetized animal on a lab towel facing up (Fig. 1). 2. With the mouse’s head immobilized, insert microcapillary tip containing siRNA/transfection reagent complexes into the nostril. 3. Instill solution slowly over a 2–3-min period, allowing the mouse to breathe the liquid in. 4. Place animal back into the cage and monitor for at least 45 min to avoid depression of cardiac and/or respiratory functions. 5. Test for the desired RNAi effect at appropriate intervals. For antiviral studies, instill virus through the nostril as well (see Notes 3 and 4). For human RSV, which does not infect mice well, use 107 –108 virus particles per animal, and measure standard lung titer assay and/or clinical symptoms (such as body weight, respiration rate, etc.).

4. Notes 1. We have described a simple method of nasal delivery of siRNA in the mouse model, but it can be successfully scaled up or down to use in other laboratory animals. 2. To diminish any possible toxic effect of delivery reagents, siRNA can be introduced “naked,” i.e., without transfection reagent, which exhibits about 70–80% activity of the reagent-complexed siRNA. It is best not to use polyethyleneimine (PEI). Although PEI is often used to form complexes with nucleic acids, mice apparently do not tolerate PEI through the nose. In our attempts, essentially all mice died within minutes of inhalation of the PEI–siRNA complex. 3. Administration of excessive liquid will “drown” the mouse and cause death. Try to keep the total volume under 45 μL in routine application, although up to 100 μL may be tolerated (see Chapter 11). 4. The procedure also works in an aerosolized application. We have used a small homemade enclosure in which the anesthetized mouse is placed and the siRNA complex (made as in Section 3.2) is sprayed in using a handheld nebulizer (the common type used as an inhaler by asthmatics). In this method, a larger amount of siRNA is needed because most of the mist is wasted and a small fraction is inhaled by the animal. To optimize cost versus benefit, it is recommended that various amounts of mist and duration of exposure are tried for a given enclosure

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References 1. Fire, A., Xu, S., Montgomery, M. K., Kostas, S. A., Driver, S. E., and Mello, C. C. (1998). Potent and specific genetic interference by double-stranded RNA in Caenorhabditis elegans. Nature 391, 806–811. 2. Elbashir, S. M., Martinez, J., Patkaniowska, A., Lendeckel, W., and Tuschl, T. (2001). Functional anatomy of siRNAs for mediating efficient RNAi in Drosophila melanogaster embryo lysate. EMBO J. 20, 6877–6888. 3. Hammond, S. M., Bernstein, E., Beach, D., and Hannon, G. J. (2000). An RNAdirected nuclease mediates post-transcriptional gene silencing in Drosophila cells. Nature 404, 293–296. 4. Nykanen, A., Haley, B., and Zamore, P. D. (2001). ATP requirements and small interfering RNA structure in the RNA interference pathway. Cell 107, 309–321. 5. Billy, E., Brondani, V., Zhang, H., Muller, U., and Filipowicz, W. (2001). Specific interference with gene expression induced by long, double-stranded RNA in mouse embryonal teratocarcinoma cell lines. Proc. Natl. Acad. Sci. USA 98, 14428–14433. 6. Barik, S. (2004). Control of nonsegmented negative-strand RNA virus replication by siRNA. Virus Res. 102, 27–35. 7. Barik, S. (2005). Silence of the transcripts: RNA interference in medicine. J. Mol. Med. 83, 764–773. 8. Bitko, V., and Barik, S. (2007). Intranasal antisense therapy: Preclinical models with a clinical future? Curr. Opin. Mol. Ther. 9, 119–125. 9. Bitko, V., and Barik, S. (2001). Phenotypic silencing of cytoplasmic genes using sequence-specific double-stranded short interfering RNA and its application in the reverse genetics of wild type negative-strand RNA viruses. BMC Microbiol. 1, 34. 10. Bitko, V., Musiyenko, A., Shulyayeva, O., and Barik, S. (2004). Inhibition of respiratory viruses by nasally administered siRNA. Nat. Med. 11, 50–55. 11. Shin, D., Kim, S. I., Kim, M., and Park, M. (2006). Efficient inhibition of hepatitis B virus replication by small interfering RNAs targeted to the viral X gene in mice. Virus Res. 119, 146–153. 12. Zender, L., Hutker, S., Liedtke, C., et al. (2003). Caspase 8 small interfering RNA prevents acute liver failure in mice. Proc. Natl. Acad. Sci. USA 100, 7797–7802. 13. Morrissey, D. V., Lockridge, J. A., Shaw, L., et al. (2005). Potent and persistent in vivo anti-HBV activity of chemically modified siRNAs. Nat. Biotechnol. 23, 1002–1007. 14. Song, E., Lee, S. K., Wang, J., et al. (2003). RNA interference targeting Fas protects mice from fulminant hepatitis. Nat. Med. 9, 347–351.

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15. Palliser, D., Chowdhury, D., Wang, Q. Y., et al. (2006). An siRNA-based microbicide protects mice from lethal herpes simplex virus 2 infection. Nature 439, 89–94. 16. Bitko, V., Musiyenko, A., and Barik, S. (2007). Viral infection of the lungs through the eye. J. Virol. 81, 783–790. 17. Banerjea, A., Li, M. J., Bauer, G., Remling, L., Lee, N. S., Rossi, J., and Akkina, R. (2003). Inhibition of HIV-1 by lentiviral vector-transduced siRNAs in T lymphocytes differentiated in SCID-hu mice and CD34+ progenitor cell-derived macrophages. Mol. Ther. 8, 62–71. 18. Nigo, Y. I., Yamashita, M., Hirahara, K., et al. (2006). Regulation of allergic airway inflammation through Toll-like receptor 4-mediated modification of mast cell function. Proc. Natl. Acad. Sci. USA 103, 2286–2291. 19. Khoury, M., Louis-Plence, P., Escriou, V., et al. (2006). Efficient new cationic liposome formulation for systemic delivery of small interfering RNA silencing tumor necrosis factor alpha in experimental arthritis. Arthritis Rheum. 54, 1867–1877. 20. Landen, C. N., Merritt, W. M., Mangala, L. S., et al. (2006). Intraperitoneal delivery of liposomal siRNA for therapy of advanced ovarian cancer. Cancer Biol. Ther. 5, 1708–1713. 21. Zhang, Y. A., Nemunaitis, J., Samuel, S. K., Chen, P., Shen, Y., and Tong, A. W. (2006). Antitumor activity of an oncolytic adenovirus-delivered oncogene small interfering RNA. Cancer Res. 66, 9736–9743. 22. Rodriguez-Lebron, E., Denovan-Wright, E. M., Nash, K., Lewin, A. S., and Mandel, R. J. (2005). Intrastriatal rAAV-mediated delivery of anti-huntington shRNAs induces partial reversal of disease progression in R6/1 Huntington’s disease transgenic mice. Mol. Ther. 12, 618–633. 23. Wang, Y. L., Liu, W., Wada, E., Murata, M., Wada, K., and Kanazawa, I. (2005). Clinico-pathological rescue of a model mouse of Huntington’s disease by siRNA. Neurosci. Res. 53, 241–249. 24. Nakamura, H., Siddiqui, S. S., Shen, X., et al. (2004). RNA interference targeting transforming growth factor-beta type II receptor suppresses ocular inflammation and fibrosis. Mol. Vis. 10, 703–711. 25. Tang, W., Yang, X., Maguire, A. M., Bennett, J., and Tolentino, M. J. (2003). Small interfering RNA (siRNA) targeting VEGF effectively inhibits ocular neovascularization in a mouse model. Mol. Vis. 9, 210–216. 26. Reich, S. J., Fosnot, J., Kuroki, A., et al. (2003). Small interfering RNA (siRNA) targeting VEGF effectively inhibits ocular neovascularization in a mouse model. Mol. Vis. 9, 210–216. 27. Shen, J., Samul, R., Silva, R. L., et al. (2006). Suppression of ocular neovascularization with siRNA targeting VEGF receptor 1. Gene Ther. 13, 225–234. 28. Rana, T. M. (2007). Illuminating the silence: Understanding the structure and function of small RNAs. Nat. Rev. Mol. Cell. Biol. 8, 23–36. 29. Dorn, G., Patel, S., Wotherspoon, G., et al. (2004). siRNA relieves chronic neuropathic pain. Nucleic Acids Res. 32, e49.

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30. Hassani, Z., Lemkine, G. F., Erbacher, P., et al. (2005). Lipid-mediated siRNA delivery down-regulates exogenous gene expression in the mouse brain at picomolar levels. J. Gene Med. 7, 198–207. 31. Sioud, M., and Sorensen, D. R. (2003). Cationic liposome-mediated delivery of siRNAs in adult mice. Biochem. Biophys. Res. Commun. 312, 1220–1225. 32. Massaro, D., Massaro, G. D., and Clerch, L. B. (2004). Noninvasive delivery of small inhibitory RNA and other reagents to pulmonary alveoli in mice. Am. J. Physiol. Lung. Cell Mol. Physiol. 287, L1066–L1070.

7 RNA Interference as a Genetic Tool in Trypanosomes Vivian Bellofatto and Jennifer B. Palenchar

Summary RNA interference (RNAi) is a cellular mechanism that is often exploited as a technique for quelling the expression of a specific gene. RNAi studies are carried out in vivo, making this a powerful means for the study of protein function in situ. Several trypanosomatids, including those organisms responsible for human and animal diseases, naturally possess the machinery necessary for RNAi manipulations. This allows for the use of RNAi in unraveling many of the pressing questions regarding the parasite’s unique biology. The completion of the Trypanosoma brucei genome sequence, coupled with several powerful genetic tools, has resulted in widespread utilization of RNAi in this organism. The key steps for RNAi-based reduction of gene expression, including parasite cell culture, DNA transfection, RNAi expression, and experimental execution, are discussed with a focus on procyclic forms of Trypanosoma brucei.

Key Words: RNAi; RNA eukaryotic gene expression.

interference;

trypanosomes;

parasitic

protozoa;

1. Introduction Endogenous RNA interference (RNAi) machineries, which normally regulate gene expression on many levels, can be manipulated at will to alter specific protein levels inside cells. In contrast to more traditional molecular genetic techniques, the timely and laborious process of producing genetic knockouts at the DNA level can be circumvented, thus facilitating a speedy initial study of gene function. Using RNAi, the essential nature of a gene can be assessed and the biological effects of the ablation of a nonessential gene can be deduced. From: Methods in Molecular Biology, vol. 442: RNAi: Design and Application Edited by: S. Barik © Humana Press, Totowa, NJ

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Fig. 1. Important vectors for dsRNA expression in T. brucei. (a)–(c). A simple schematic of the vectors used to generate stably transfected cell lines. Both the pZJM (23) and p2T7-177 (17) are tetracycline-regulated and contain opposing T7 promoters to generate dsRNA fragments that trigger RNAi. pZJM is targeted to the transcriptionally silent rDNA spacer region of the genome, whereas p2T7-177 is targeted to the minichromosome 177 base-pair repeat region. pZJM has an upstream T7 RNAPdriven phleomycin resistance gene, and p2T7-177 has an upstream rRNA promoterdriven phleomycin-resistance gene; these genes are necessary to maintain the integrated plasmid in the parasite genome. In parts (a) and (b), the short underlines indicate the location of dual T7 terminators (15,17). In part (c), the stem-loop vector (20) contains a procyclin (PARP) promoter, is tetracycline-regulated, and is targeted to the rDNA region of the genome as well. The vector is unique in that dsRNA is generated through a stem-loop structure.

The phenomenon of RNAi (see Fig. 1 in Chapter 4) was first observed in 1990 when Jorgensen and colleagues attempted to deepen the color of petunias through the introduction and overexpression of a chimeric petunia chalcone synthase gene. Instead, they observed “co-suppression” resulting in white or variegated flowers (1). Several years later, Mello, Fire and colleagues injected sense and antisense strands of RNA into C. elegans (2). In these experiments, a few double-stranded RNA (dsRNA) molecules per cell were sufficient to silence the homologous gene. This work garnered the pair the 2006 Nobel Prize

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in Medicine or Physiology. RNAi has been found in organisms as diverse as humans, flies, fungi, plants, and parasitic protozoa. The expression of dsRNA triggers the destruction of the complementary mRNA. This dsRNA is a substrate for the ribonuclease Dicer (3). Dicer cleaves the dsRNA into ∼20–25 nucleotide small interfering (si) RNAs. siRNAs enter the RNA induced silencing complex (RISC), where they are unwound. The antisense strand (or guide strand) targets RISC to the complementary mRNA, which is subsequently degraded through RISC nuclease activity. RNAi in trypanosomes was discovered by the Ullu and Tschudi laboratory (4) in 1998 during a study designed to address mRNA splicing in the African trypanosome, Trypanosoma brucei. In this seminal work, they observed that dsRNA, derived from the sequences that encode the alpha-tubulin genes, caused cells to round up or become “fat.” Notably, this was one of the earliest discoveries of RNAi activity in eukaryotic cells, after the initial work on plants and worms (2,5). Several trypanosomatids possess endogenous RNAi machinery, including T. brucei spp. and T. congolense (6). RNAi is notably absent from T. cruzi and Leishmania spp. It is unknown if RNAi functions in either in Leptomonas spp. or Crithidia spp. The combination of available genetic tools, a completed genome sequence, and many pressing trypanosome biology questions make the utilization of RNAi in T. brucei highly relevant. The utility of RNAi in this parasite is highlighted by a recent, chromosome-wide analysis of gene function by Subramaniam et al. (7). In addition to the importance of T. brucei as an infectious agent, this organism as well as the closely related Leishmania spp. are excellent model organisms for the study of intracellular processes in eukaryotic cells. T. brucei is a single-cell organism that grows rapidly, dividing every ∼10 h, axenically in synthetic media. It possesses simplified pathways for many intracellular processes, including having few cis-introns in its mRNA and few, if any, discreet promoters for RNA polymerase (RNAP) II-dependent mRNA coding genes. Unlike many other wellstudied eukaryotes, it appears to contain few transcription factors, and possibly no transcriptional activators. Equally useful is its overexpression of several molecular traits that are relatively elusive in other organisms. For example, in other eukaryotes, hypermethylated RNA cap structures (designated cap 1 and 2; cap 0 is the me7 GpppG-monomethylated and more common cap structure) are challenging to study due to their low abundance. In T. brucei, a hypermethylated cap, designated cap 4, is highly abundant, present on the 5′ end of every mRNA. Indeed, RNAi is being utilized to examine cap 4 biology (8). The phenomenon of RNA trans-splicing, the process in which a spliced leader (SL) RNA is added upstream of a translatable region of RNA to produce a stable mRNA, occurs to a different extent in a wide variety of eukaryotes, from

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trypanosomes to simple chordates (reviewed in 9). Trypanosomes utilize transsplicing for the production of every functional mRNA. Most of our knowledge of SL RNA production and function in trans-splicing comes from trypanosome studies. Here, too, RNAi is proving an invaluable tool in understanding SL RNA biogenesis and function (10–12). In the T. brucei system, adequate genetics are available, including homologous recombination upon DNA transfection, cell lines with small moleculeresponsive gene expression (under tetracycline control), and bacteriophage T7 RNAP-dependent RNAi expression, allowing for the facile expression of dsRNA for posttranscriptional downregulation of endogenous mRNAs. Here we describe the current RNAi methodology for T. brucei, the ways in which it has been employed, and the important considerations that must be adhered to in data interpretation (see Notes 1–9). It is required that the researcher is in a Trypanosome laboratory that is already familiar with the growth and the basic genetics of the parasite. Still, in order for the reader to understand the rationale behind the experiments, we will also provide a theoretical background of the major items in Sections 2 and 3. 2. Materials 1. Procyclic (tsetse midgut form) trypanosomes, specifically wild-type T. brucei strain Lister 427 and its derivatives. This protocol is written for Strain 29-13, which expresses T7 RNA polymerase as well as tetracycline repressor (TetR). The specifics of this strain are as follows: The T7 RNAP, a single subunit RNA polymerase, and one copy of the TETR are encoded by the plasmid pLEW13 that is maintained as an integrated molecule in the -tubulin locus in T. brucei by G418 selection. These two genes are maintained by drug selection, as they are coupled to genes that encode resistance to G418 (to maintain the T7 RNAP) and hygromycin (HYG) (to maintain the TETR). The transcriptional abundance of all three foreign genes [T7 RNAP, NEOR (encoding G418R), and TETR] is roughly equal to that of a single -tubulin transcript. A 3′ untranslated region (3′ UTR) from the T. brucei aldolase gene helps destabilize the T7 RNAP mRNA and thus indirectly helps manage the amount of TETR made in the cells. To better control the level of tetracycline-inducible regulation of introduced constructs, the Cross group (13) added a second TETR gene, under the control of a modified T7 RNAP-dependent promoter, into the “rnp1” locus. The TETR contains a 3′ UTR from actin, which ensures a short half-life to the repressor. This additional DNA is maintained under HYG selection. This HYGR, G418R strain, named 29-13, is to date the optimum strain for RNAi experiments in procyclic T. brucei. 2. Parasite growth medium, SDM-79. It is a semidefined medium, supplemented with 10% heat-inactivated fetal calf serum and 7.5 mg/L of hemin (14). 3. Cytomix: 2 mM EGTA (pH 7.6), 120 mM KCl, 0.15 mM CaCl2 , 10 mM K2 HPO4 /KH2 PO4 (pH 7.6), 25 mM HEPES (pH 7.6), 5 mM MgCl2 , 0.5% glucose,

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100 μg/mL BSA, and 1 mM hypoxanthine. The pH is adjusted to 7.6 (KOH) and sterile-filtered. 4. RIPA buffer: 300 mM NaCl, 1% NP-40, 0.25% Na-Deoxycholate, 50 mM TrisHCl (pH 7.4), and 1 mM EDTA. 5. Drugs for selection: Drug Stock solution Final concentration in media Hygromycin 50 mg/mL in PBS 50 μg/mL G418 50 mg/mL 15 μg /mL Phleomycin 10 mg/mL 2.5 μg /mL Tetracycline 1 mg/mL 1 μg/mL All stocks are sterile-filtered and stored at –20 ºC. Upon thawing, phleomycin is stable for several months at 4 ºC, and G418 and hygromycin are stable for up to one year in aqueous solution at this temperature. 6. pZJM vector (Fig. 1): This is the original highly utilized vector for expressing heritable and controllable dsRNA (15). It utilizes two opposing T7 RNAP promoters under the control of adjacent, cis-acting tetracycline operators. An advantage of the pZJM vector is that only a single-step PCR is required to position the dsDNA that encodes the two complementary RNAs between the opposing T7 RNAP promoters. The RNAs anneal inside the nucleus to form a dsRNA that enters the siRNA pathway. Linearized pZJM integrates into one of the rRNA spacer regions in the T. brucei genome and is maintained using phleomycin selection. Limiting dilution in selection medium allows for the selection of clonal transfectants. 7. p2T7-177 vector (Fig. 1), a derivative of p2T7 (16): It was originally designed for integration into minichromosomes, which are generally transcriptionally silent chromosomes in T. brucei (17). 8. Standard cloning protocols and reagents such as restriction enzymes, ligases, phenol/chloroform, LB-agar plates with selective antibiotics, etc.

3. Methods 3.1. T. brucei Cell Culture 1. Store parasites at –80 ºC for medium-term storage (a few weeks) and in liquid nitrogen for longer periods. An upright –80 ºC freezer is not optimal, as temperature fluctuations decrease cell viability over time. 2. When needed, revive parasites by rapid thawing at room temperature. Add 1 mL of thawed cell stock slowly to 10 mL of prewarmed (27 ºC) complete SDM-79 media and incubate at 27 ºC for 24 h before challenging with G418 and hygromycin. Grow cells for several days and maintain between 2 × 106 /mL and 1 × 107 /mL before using in transfection.

3.2. Choice and Design of dsRNA Sequence The specific knockdown of a single gene product requires production of dsRNA that is complementary to a unique region of the gene’s mRNA. Off-target results

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can occur if the dsRNA is close in sequence to nontarget mRNAs. Similarly, if common sequences are used, multiple mRNA can be destroyed simultaneously. For example, using a dsRNA that is complementary to the SL will potentially destroy all T. brucei mRNAs. Nonspecific results may be avoided by taking care in designing a dsRNA that only complements the target mRNA. This is best determined by “BLASTing” the dsRNA sequence against the trypanosome genome (http://www.genedb.org). Either a region within the 5′ untranslated region, the open reading frame, or the 3′ untranslated region may be suitable. In all cases, it is optimal to choose a dsRNA sequence that is complementary to ∼300–500 contiguous bases of the mRNA. The RNAit program, found at the Trypanosomatid Functional Analysis Network (http://trypanofan.path.cam.ac.uk/software/RNAit.html), is useful for choosing the unique region of the target gene that will be recognized by the dsRNA. The RNAit program allows the user to circumvent the tedious manual BLAST searching necessary to ensure the suitability of their target sequence. The basic steps are 1. Enter the target sequence. 2. Generate PCR primers based upon melting temperature and length. 3. BLAST the resultant hypothetical PCR product against trypanosome genomic DNA sequence. 4. Parse the results to see whether or not the sequence is suitable for RNAi.

3.3. Construction of RNAi Clones Although RNAi in trypanosomes was originally observed through the transient transfection of synthetic dsRNA (4), stable DNA transfection, which allows for heritable expression of dsRNA, the selection of clonal cell lines, and conditional RNAi expression, is more commonly used. 1. Choose from vectors pZJM or p2T7-177 (see their description in Sections 2 and 4). 2. Clone the sequence designed in Section 2.2 in one of the above vectors, grow it as plasmid DNA in E. coli, and then linearize by restriction at a single unique site. Linearization facilitates homologous recombination-directed integration into a T. brucei chromosome upon DNA transfection.

3.4. Introduction of RNAi Clones 1. Linearize the pZJM and p2T7-177 clones with the restriction enzyme NotI. Digest approximately 20 μg of DNA with NotI and then purify using a phenol/chloroform extraction step. Ethanol-precipitate the linearized DNA and then resuspend in ∼40 μL of sterile deionized water. Determine DNA concentration by absorbance at 260 nm.

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2. Grow trypanosomes (see Section 3.1) to log phase (∼6 × 106 cells/mL) and plan to use approximately 1–2 × 107 cells per transfection. 3. Harvest the cells from culture by centrifugation at 4 o C, 10 min, at 700 x g, and wash with ZFM (22) or cytomix at 4 o C. 4. Resuspend cells in ZFM or cytomix (0.5 mL per transfection) and transfer to a prechilled 4-mm electroporation cuvette containing 5–10 μg of DNA that has been linearized and purified (in step 1 above). Immediately subject parasites to electroporation using a BTX600 model electroporator set to a charge of 1.6 kV, 2.5 kV/resistance, and R2 resistance, one pulse. The time constant value after the pulse will be ∼0.35. (Using a Bio-Rad Gene Pulser, the settings are 1500V, 25 μF, and 2 pulses.) As a negative control, a transfection containing no DNA is also performed. 5. Immediately transfer cells to 10 mL of prewarmed (27 o C) culture medium and incubate for ∼16 h before drug selection is initiated in the next step. 6. For selection, add G418, hygromycin, and phleomycin (for procyclic T. brucei strain 29-13 transfected with a phleomycin-resistance dsRNA-synthesizing vector, such as pZJM). The selection process takes approximately one week, with a noticeable difference between the control (no DNA) and the transformed cells at this point. 7. Isolation of clonal cell lines: This is commonly done through limited dilution that may be achieved in several ways (http//tryps.rockefeller.edu). During routine cell culture, SDM-79 media contains 10% fetal calf serum. This amount is increased to 15% during generation of clonal cell lines to increase cloning efficiency. In one approach, cells are diluted to 5 cells/mL and 100-μL aliquots are distributed into a 96-well plate and incubated at 27 o C with CO2 . In ∼2 weeks, a noticeable increase in cell density occurs in many of the wells. In a second approach, wild-type Lister 427 parasites are added as “feeder” cells (∼0.5–1 × 103 feeder cells/mL). Four clonal cell lines should be isolated and used in each RNAi induction experiment. In the dilution procedure, note that the cells can be successfully diluted when the culture reaches a density of ∼4 × 106 cells/mL. Dilute newly transfected cells relatively slowly (1:1), always into prewarmed media, and eventually transfer to a 25-cm2 T-flask when the volume reaches ∼3 mL. Maintain the clonal cell lines at 3–6 × 106 cells/mL. Overdilution of the trypanosomes results in reduced growth or cell death. 8. Storage of clonal cells: Upon production of a stably transfected cell line, the parasites should be stored long-term in liquid nitrogen as follows. Grow parasites to a density of ∼8 × 106 cells/mL, then pellet at 700 x g, 10 min, 4 o C. Remove the supernatant immediately and resuspend the parasite pellet in complete SDM-79 medium containing 10% glycerol (no drugs). Each 10 mL of starting culture is resuspended in 1 mL of the glycerol-containing medium. These cells are slowly frozen at –80 o C for three days and then stored in liquid nitrogen.

To use the frozen cells, thaw as described in Section 3.1. Selection drugs can be added 16–24 h after thawing.

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3.5. dsRNA Expression to Knock Down a Specific Protein Induction of RNAi is accomplished using a regulated tetracycline-controlled promoter. Tetracycline releases the TETR from the operator region on the vector DNA, thus allowing dsRNA production. As stated above, four clonal cell lines should be used for each induction. 1. On day 0, split the starting culture that is in log phase into two T-flasks. Add tetracycline (or doxycycline) to a final concentration of 1 μg/mL to only one of them, the second flask serving as a noninduced control. 2. Count parasites daily at approximately the same time each day by placing 10 μL of culture onto a Neubauer hemocytometer. At least 100 parasites are counted to obtain meaningful values. Only live cells should be counted; Trypan Blue may be used to differentiate between live and dead cells when counting. In addition, it is crucial to observe cell morphology, as this may result in the development of interesting hypotheses regarding trypanosome biology. Alternatively, cell densities can be determined using a Coulter counter model Z1 (Coulter Electronics). The total number of live cells per mL is calculated and plotted as a function of time since tetracycline induction. Sample data from our laboratories are shown in Fig. 2. From the beginning of the experiment, both the RNAi-induced and control cells are monitored by microscopy for cell motility and morphology. When either the induced or noninduced culture density reaches ∼8 × 106 cells/mL, it should be diluted to 2 × 106 cells/mL using a prewarmed (27 o C) medium containing the selection drugs (G418, hygromycin, and phleomycin) and tetracycline. This is best done by pelleting the culture and resuspending cells in fresh medium containing freshly added drugs. The length of time required to observe a growth phenotype of an essential gene is variable depending upon target protein stability and protein

Fig. 2. Growth curve of a procyclic cell line stably transfected with an inducible RNAi construct that ablates an essential gene. At day 0, tetracycline was added to initiate dsRNA production. By day 4, growth of the RNAi-induced cell culture deviates from that of the noninduced control.

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concentration. Growth deviations from the noninduced control are generally seen within 1 to 4 days. 3. Follow the induced and noninduced paired cultures daily for 12 days, diluting as necessary to maintain cell densities between 2–8 × 106 cells/mL. In some cases, cells harboring an RNAi construct that knocks down an essential protein will begin to recover and resume logarithmic growth rates around day 10. This observation is likely the result of an epigenetic change in genotype, resulting in a loss of RNA interference of the target mRNA. Thus, a gene product is considered essential to cell viability if cells halt log-phase growth within five days of RNAi induction. Clearly, analysis of a gene that produces an essential, albeit long-lived and highly abundant, protein may show a phenotypic effect as a result of protein dilution in the population; therefore, growth arrest may be delayed. 4. To monitor protein levels throughout the experiment, remove approximately 5 × 106 cells daily from both induced and noninduced cultures (the volume necessary to achieve this number of cells can be determined from the daily cell count). Centrifuge cells at 1400 x g, 10 min, 4 o C, and discard the supernatant. Resuspend the pellet in 5 mL of chilled (4 o C) PBS and recentrifuge. Finally, resuspend the pellet in 50 μL of RIPA buffer containing the protease inhibitors PMSF, pepstatin, and leupeptin. Vortex briefly and incubate on ice for 10 min, and centrifuge again at 14,000 x g, 10 min, 4 o C. Use the supernatant (containing cell extract from 1 × 105 cells/μL) for Western blot analysis. As a general rule, to accurately monitor and compare protein levels between samples, it is necessary that the extract from the same number of cells be loaded into each well of the SDS-PAGE. Using 20 μL of extract for the Western blot analysis is sufficient to visualize protein levels using specific antibodies and the ECL system of detection. Positive controls and analysis of mRNA are described later (see Section 4).

4. Notes 1. Potential artifacts: The experimental manipulations of DNA transfection, plasmid DNA integration, drug challenge, and dsRNA expression are stressful for the parasites and many elicit biological responses that are artifactual. Thus, it is crucial that interesting results obtained with RNAi assays be followed up with additional genetic and biochemical experiments. Additionally, off-target effects must also be considered in all RNAi experiments. The ablation of nontarget mRNA may cause phenotypic changes that are unrelated to the gene of interest. 2. Target stability: RNAi knockdown of target mRNA and protein levels is a function of mRNA synthesis, translation, and turnover rates. Knockdown of a rapidly transcribed and long-lived mRNA is therefore more difficult than the destruction of a low-level and unstable mRNA. 3. Reversion: Researchers often analyze RNAi-induced cells for the reversion of the observed phenotype. Tetracycline is removed, leading to the shutoff of dsRNA, allowing the return of target mRNA and encoded protein. For example, this

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5.

6.

7.

8.

Bellofatto and Palenchar experimental procedure has been used to follow the reacquisition of protein in protein localization studies (19). It is important to ensure that the regrowth of the transfected cell line is not due to the emergence of revertant cells; this is controlled for by the readdition of tetracycline and observation of the original ablation phenotype. Multiple clones: It has been observed that in some cases only one of several clonal cell lines that are maintained in drug selection (i.e., phleomycin for pZJM and p2T7-177 integrants) actually shows ablation of the target mRNA and encoded protein. Thus, it is important to analyze several clones for both mRNA and protein levels to maximize the chance of recovering a desired cell line. Positive controls: A useful positive control is the knockdown of alpha-tubulin, as described initially by the Ullu and Tschudi laboratory (20). Either the pLEWFAT construct, which contains a dsRNA that complements the 5‘UTR of the alphatubulin mRNA, or a derivative of the pZJM that produces a dsRNA that is complementary to 650 bp of the coding region of alpha-tubulin, has been used (20,23). For this control, the 650-bp region is cloned into the vector backbone that is being used for the experiments. A stable cell line containing the alphatubulin dsRNA should be generated in parallel with the gene under study. Prior to cloning out the experimental cell lines, the nonclonal tubulin targeted parasites are induced with tetracycline to decrease tubulin protein levels. Cells should appear “fat” by microscopy after 24–48 h of culture. This observation confirms that the transfection and induction steps of the experiment were successful. Other assays: Analysis of specific mRNA and/or protein shutoff is performed using Northern blotting, to assess mRNA levels, or Western blotting, to assess protein levels. For RNA detection, use total RNA, which is ∼10% mRNA, following a standard Northern protocol and probing with 32 P-labeled DNA used for the transfection step. It is possible to detect both the loss of the target mRNA and the presence of the dsRNA after RNAi induction. It is preferable to assess protein levels, assuming a specific antibody is available, as discussed above. Background expression: Inserting the T7 RNAP-driven, tetracycline-inducible dsRNA-producing vector into this locus results in less “background”-level expression of the dsRNA. Low-level “background” expression can be harmful to the experimental outcome, as it can result in partial inhibition of an essential gene and thus uncontrolled genetic changes during transfected cell selection. To obviate this problem, a more highly controlled ectopic dsRNA, such as that obtainable by cloning into transcriptionally silent minichromosomes, is preferable. In addition, currently available derivatives of the p2T7 vector allow for easy cloning of PCR products (the p2T7TAblue plasmid) (18). Several vectors are now available for the construction of stably transfected cell lines in which the expression of dsRNA is under the control of tetracycline. Vectors for stem-loop or hairpin dsRNA production: These constructs were used in the early days of trypanosome RNAi experimentation (19,20). Although constructing the hairpin vectors requires two cloning steps, expression of the hairpin RNA from these vectors is tightly regulated and highly expressed upon

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induction (21). These vectors produce a single RNA molecule that folds back on itself to form a stem-loop structure that then enters the siRNA pathway. 9. RNAi protocols for analysis of gene expression in bloodstream forms of T. brucei can be found at http://tryps.rockefeller.edu/ and at http://trypanofan. path.cam.ac.uk.

Acknowledgments We thank Chris Utter for critical reading of the manuscript. This work was supported by grants AI29478 and AI53835 to V. B., who is also a recipient of a Burroughs Wellcome Fund New Investigator Award in Molecular Parasitology. J. B. P. is supported by the Department of Chemistry at Villanova University.

References 1. Napoli, C., Lemieux, C., and Jorgensen, R. (1990). Introduction of a chimeric chalcone synthase gene into petunia results in reversible co-suppression of homologous genes in trans. Plant Cell 2, 279–289. 2. Fire, A., Xu, S., Montgomery, M. K., Kostas, S. A., Driver, S. E., and Mello, C. C. (1998). Potent and specific genetic interference by double-stranded RNA in Caenorhabditis elegans. Nature 391, 806–811. 3. Hutvagner, G., and Zamore, P. D. (2002). RNAi: Nature abhors a double-strand. Curr. Opin. Genet. Dev. 12, 225–232. 4. Ngo, H., Tschudi, C., Gull, K., and Ullu, E. (1998). Double-stranded RNA induces mRNA degradation in Trypanosoma brucei. Proc. Natl. Acad. Sci. USA 95, 14687–14692. 5. Metzlaff, M., O’Dell, M., Cluster, P. D., and Flavell, R. B. (1997). RNAmediated RNA degradation and chalcone synthase A silencing in petunia. Cell 88, 845–854. 6. Inoue, N., Otsu, K., Ferraro, D. M., and Donelson, J. E. (2002). Tetracyclineregulated RNA interference in Trypanosoma congolense. Mol. Biochem. Parasitol. 120, 309–313. 7. Subramaniam, C., Veazey, P., Redmond, S., et al. (2006). Chromosome-wide analysis of gene function by RNA interference in the African trypanosome. Eukaryot. Cell 5, 1539–1549. 8. Li, H., and Tschudi, C. (2005). Novel and essential subunits in the 300-kilodalton nuclear cap binding complex of Trypanosoma brucei. Mol. Cell. Biol. 25, 2216–2226. 9. Palenchar, J. B., and Bellofatto, V. (2006). Gene transcription in trypanosomes. Mol. Biochem. Parasitol. 146, 135–141. 10. Biton, M., Mandelboim, M., Arvatz, G., and Michaeli, S. (2006). RNAi interference of XPO1 and Sm genes and their effect on the spliced leader RNA in Trypanosoma brucei. Mol. Biochem. Parasitol. 150, 132–143.

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11. Palenchar, J. B., Liu, W., Palenchar, P. M., and Bellofatto, V. (2006). A divergent transcription factor TFIIB in trypanosomes is required for RNA polymerase II-dependent spliced leader RNA transcription and cell viability. Eukaryot. Cell 5, 293–300. 12. Zeiner, G. M., Foldynova, S., Sturm, N. R., Lukes, J., and Campbell, D. A. (2004). SmD1 is required for spliced leader RNA biogenesis. Eukaryot. Cell 3, 241–244. 13. Wirtz, E., Leal, S., Ochatt, C., and Cross, G. A. (1999). A tightly regulated inducible expression system for conditional gene knock-outs and dominant-negative genetics in Trypanosoma brucei. Mol. Biochem. Parasitol. 99, 89–101. 14. Cross, G. A., and Manning, J. C. (1973). Cultivation of Trypanosoma brucei spp. in semi-defined and defined media. Parasitology 67, 315–331. 15. Wang, Z., and Englund, P. T. (2001). RNA interference of a trypanosome topoisomerase II causes progressive loss of mitochondrial DNA. EMBO J. 20, 4674–4683. 16. LaCount, D. J., Bruse, S., Hill, K. L., and Donelson, J. E. (2000). Double-stranded RNA interference in Trypanosoma brucei using head-to-head promoters. Mol. Biochem. Parasitol. 111, 67–76. 17. Wickstead, B., Ersfeld, K., and Gull, K. (2002). Targeting of a tetracyclineinducible expression system to the transcriptionally silent minichromosomes of Trypanosoma brucei. Mol. Biochem. Parasitol. 125, 211–216. 18. Alibu, V. P., Storm, L., Haile, S., Clayton, C., and Horn, D. (2005). A doubly inducible system for RNA interference and rapid RNAi plasmid construction in Trypanosoma brucei. Mol. Biochem. Parasitol. 139, 75–82. 19. Bastin, P., Ellis, K., Kohl, L., and Gull, K. (2000). Flagellum ontogeny in trypanosomes studied via an inherited and regulated RNA interference system. J. Cell Sci. 113, 3321–3328. 20. Shi, H., Djikeng, A., Mark, T., Wirtz, E., Tschudi, C., and Ullu, E. (2000). Genetic interference in Trypanosoma brucei by heritable and inducible double-stranded RNA. RNA 6, 1069–1076. 21. Djikeng, A., Shen, S., Tschudi, C., and Ullu, E. (2004). Analysis of gene function in Trypanosoma brucei using RNA interference. Meth. Mol. Biol. 265, 73–83. 22. Bellofatto, V., and Cross, G. A. M. (1989). Expression of a bacterial gene in a trypanosomatid protozoan. Science 244, 1167–1169. 23. Wang, Z., Morris, J. C., Drew, M. E., and Englund, P. T. (2000). Inhibition of Trypanosoma brucei gene expression by RNA interference using an integratable vector with opposing T7 promoters. J. Biol. Chem. 275, 40174–40179.

8 Lentivirus-Mediated RNA Interference in Mammalian Neurons Scott Q. Harper and Pedro Gonzalez-Alegre

Summary The ability to manipulate RNAi in cultured mammalian cells has provided scientists with a very powerful tool to influence gene expression. Neurons represent a cell type that initially displayed resistance to transduction by siRNAs or shRNA, when attempting to silence expression of endogenous genes. However, the development of lentiviral systems with that goal has facilitated the exogenous manipulation of RNAi in these postmitotic cells. Lentiviral-mediated RNAi experiments in cultured mammalian neurons can be designed to address a wide variety of biological questions or to test potential therapeutic hairpins before moving to treatment trials in vivo. We provide a practical approach to accomplish siRNA-mediated silencing of the disease-linked protein torsinA in primary neuronal cultures through the generation of lentiviral vectors expressing shRNAs.

Key Words: PNC (primary neuronal cultures); shRNA; torsinA; RNAi; silencing; lentivirus; FIV (feline immunodeficiency virus). 1. Introduction TorsinA is an endoplasmic reticulum resident glycoprotein that, when mutated, leads to the dominantly inherited disease DYT1 dystonia (1). While the function of this AAA protein (ATPases Associated to diverse cellular Activities) remains unknown, the mutant protein acts through a dominant negative effect by recruiting wild-type torsinA from the endoplasmic reticulum to the nuclear envelope (2–4), thus leading to torsinA loss of function (5). As is true for many other neurological disease-linked proteins, torsinA is a widely expressed protein in both in neural and nonneural tissues (6). However, the resulting phenotype of the mutation is restricted to dysfunction of a specific subset of From: Methods in Molecular Biology, vol. 442: RNAi: Design and Application Edited by: S. Barik © Humana Press, Totowa, NJ

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neurons. Therefore, establishing neuronal models of torsinA insufficiency may help us determine the function of this protein and uncover the pathobiology underlying DYT1 dystonia. Furthermore, specifically silencing expression of mutated torsinA has been proposed as a promising therapeutic strategy for this incurable disease (7,8). Primary neurons derived from the embryonic mouse brain are widely used in the investigation of different aspects of neuronal function, including development and differentiation, synaptic transmission, or excitotoxicity, among others (9). Using RNAi in this cellular model greatly expands the repertoire of biological questions that can be addressed in this system. The development of lentiviral systems to mediate RNAi in primary neurons has facilitated these types of studies (8,10,11). Here we will describe the protocol we have developed to silence torsinA expression in primary neuronal cultures derived from wildtype embryonic mice. This protocol can easily be adapted to the study of many other proteins of interest.

2. Materials 2.1. Generation of U6shRNA 1. 2. 3. 4. 5. 6. 7. 8. 9. 10. 11. 12. 13. 14. 15.

pAd5mU6 (template plasmid for PCR amplification) (see Note 1). Oligonucleotide primers. Platinum PCR SuperMix High Fidelity 1.1X (Invitrogen, Carlsbad, CA). pCR2.1 vector in TOPO-TA Cloning kit (Invitrogen). Quantum Prep Freeze ‘N Squeeze DNA Gel Extraction Spin Columns (Bio-Rad, Hercules, CA). E. coli strains DH5, TOP10 (Invitrogen). Ampicillin. 40 mg/mL X-gal in dimethylformamide. Luria–Bertani (LB) broth. 37 ºC shaking incubator. Quantum Prep Plasmid Miniprep Kit (Bio-Rad). Restriction enzyme EcoRI (New England Biolabs). Agarose and ethidium bromide. DNA sequencing capability. Spectrophotometer.

2.2. Generation of FIV.shRNA.eGFP 1. 2. 2. 4.

Restriction enzymes EcoRI and MfeI (New England Biolabs). CIAP (calf intestinal alkaline phosphatase) (New England Biolabs). FIV shuttle (pVETL CMV-eGFP) (10) packaging and envelope plasmids (12). Rapid DNA ligation kit (Roche, Indianapolis, IN).

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5. HEK393 cells. 6. Tissue culture incubator at 37 ºC with 5% CO2 . 7. HEPES-buffered saline (HBS): 5.0 g HEPES, 8.0 g NaCl, 0.37 g KCl, 0.188 g Na2 HPO4 .7H2 O, and 1.0 g glucose. Bring to 1 L in ddH2 O, adjust pH to 7.1 with concentrated NaOH, filter-sterilize, and store at 4 ºC. 8. 2.5 M of CaCl2 . 9. DMEM (Dulbecco’s modified Eagle’s medium): DMEM-10: DMEM (Gibco BRL) with 10% fetal calf serum, 5 mL of Penicillin/Streptomycin (Pen/Strep) (Gibco BRL). DMEM-2: DMEM with 2% fetal bovine serum (FBS) and 5 mL of Pen/Strep. 10. Lactose buffer: phosphate-buffered saline (PBS), pH 7.4 (Sigma P-3813), with 40 mg/mL of lactose, filter-sterilized. 11. Sorvall Centrifuge RC 26 Plus, with SLA 1500 rotor. 12. Centrifugation bottles (250-mL capacity).

2.2.1. Titer by Transgene Expression 1. 2. 3. 4. 5. 6.

HT-1080 cells (ATCCCRL-121) maintained in exponential growth in DMEM-10. Incubate at 37 ºC with 5% CO2 . Six-well tissue culture dishes. DMEM-2 and DMEM-10. Polybrene stock: 8 mg/mL in ddH2 O, filter-sterilized. DMEM-2/polybrene: On the day of use, dilute the polybrene stock 1:2,000 in DMEM-2 for a final polybrene concentration of 4 μg/mL. 7. Dilution tubes (3.5-mL polystyrene sterile tubes).

2.3. Primary Neuronal Cultures 1. Ketamine/xylazine (mix 10 mL at a time and store at room temperature): 1 mL of 100 mg/mL ketamine, 0.1 mL of 100 mg/mL xylazine, 8.9 mL of sterile PBS (Gibco BRL). 2. Poly-L-lysine (0.1 mg/mL): Dilute 5 mg of poly-L-lysine (Sigma) in 50 mL of sterile distilled water. Store working stock at 4 °C. 3. HBSS (Hank’s Balanced Salt Solution) dissecting medium pH 7.2: 500 mL of HBSS without calcium or magnesium (Invitrogen), 5 mL of Pen/Strep (Gibco BRL), 0.5 mL of 1 M HEPES. Store at 4 °C. 4. 10X (10 mg/mL) DNase solution: 10 mL of HBSS dissecting media, 100 mg of DNase (Sigma). Sterilize with 10-mL syringe filter. Store 1-mL aliquots at –20 °C. 5. 10X 2.5% Trypsin (Invitrogen). Aliquot 10 mL into 15-mL tubes. Store at –20 °C until thawed; then store at 4 °C. 6. Wash medium: 500 mL of DMEM, 10% FBS, 1% Pen/Strep. Store at 4 °C. 7. Neurobasal/FBS plating media: 220 mL of Neurobasal media (Gibco BRL), 2.5 mL of Pen/Strep, 2.5 mL of 200 mM glutamine stock (Sigma), 25 mL of FBS. Store at 4 °C.

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8. Neurobasal/B27 maintenance media: 500 mL of Neurobasal media, 5 mL of Pen/Strep, 5 mL of 200 mM glutamine stock, 10 mL 2% B27 (50X) (Gibco BRL). Store at 4 °C. 9. 10 μM of Ara-C. 10. Surgical material: fine and tissue forceps, small straight scissors, sterile scalpel blade, metal dissecting pan, dissecting microscope. 11. 10- and 6-cm sterile Petri dishes. 12. Hemocytometer.

2.4. Transduction of Primary Neuronal Cultures 1. 2. 3. 4. 5. 6.

FIV.eGFP.mU6shRNA. Serum-free OptiMEM (Gibco BRL). Neurobasal/B27 maintenance media (see above). Tissue culture (TC) hood. Hemocytometer. Inverted fluorescence scope (Olympus).

2.5. Lysis for Protein Analysis 1. Modified Laemmli buffer for cell lysis (2X) (500 mL): 50 mL of 1 M Tris HCI, pH 6.8, 20 g of sodium dodecyl sulfate (SDS), 1.0 g of bromophenol blue, 100 mL of glycerol, milliQ H2 O to total of 500 mL. Keep at room temperature. To harvest the protein lysates, add 1:2 dilution of 2X modified Laemmli buffer 2X SDS, 1:10 dilution of DTT (add before use), and dH2 O to final volume. 2. Dithiothreitol (DTT) (1 M): Dissolve 3.09 g of DTT in 20 mL of 0.01 M N -acetate (pH 5.2). Sterilize by filtration and store at –20 °C. 3. Teflon cell scrapers (Fisher). 4. Staining solution (500 mL): 50 mL of acetic acid, 225 mL of H2 O, 225 mL of methanol, 1.25 mL of coomassie brilliant blue (Amresco). 5. Destaining solution (6 L): 600 mL of acetic acid, 2.7 L of H2 O, 2.7 L of methanol. 6. Whatman filter paper.

2.6. SDS-PAGE (12%) 1. Mini-PROTEAN 3 electrophoresis system (Bio-Rad). 2. 10% Ammonium persulfate (APS): 0.1 g of APS (Pierce) in 1 mL of milliQ H2 O. 3. Separating gel: 3.3 mL of dH2 O, 4 mL of 30% acrylamide/bis solution (37.5:1 with 12% total concentration) (Amresco), 2.5 mL of 1.5 M Tris-HCl (pH 8.8), 100 μL of 10% SDS, 100 μL of 10% APS, 0.4 μL of N,N,N,N’-Tetramethylethylenediamine (TEMED) (Amresco). 4. Stacking gel (4X): 2.1 mL of dH2 O, 0.5 mL of 30% acrylamide/bis solution (37.5:1 with 2.6% total concentration when 1X), 380 μL of 1.0 M Tris-HCl (pH 6.8), 30 μL of 10% SDS, 30 μL of 10% APS, 3 μL of TEMED.

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5. Water-saturated isobutanol: Shake equal volumes of water and isobutanol in a dark glass bottle and allow to separate. Use the top layer. 6. Running buffer (4 L, 5X): 60 g of Tris, 288 g of glycine, 20 g of SDS. First dissolve Tris and glycine in 1.8 L of H2 O, then the SDS. Add milliQ H2 O to total exactly 4 L. Shake well. Store in 4 °C. Dilute to 1X with milliQ H2 O before use. 7. Prestained molecular weight markers: Kaleidoscope markers (Bio-Rad, Hercules, CA).

2.7. Western Blotting for TorsinA 1. Mini Trans-Blot Cell tank transfer system (Bio-Rad). 2. Transfer buffer (4L, 5X): 60 g of Tris, 288 g of glycine, milliQ H2 O to total of exactly 4 L. Shake well and store at 4 °C. To make 1X buffer for use (20 L): Add 12 L of milliQ H2 O to 4 L of 5X transfer buffer and add 4 L of methanol (always add methanol last or it will precipitate the solute). For transfer, set up buffer plus 0.05% (w/v) SDS. 3. Supported nitrocellulose membrane (Millipore, Bedford, MA). 4. 3MM Chr chromatography paper (Whatman, Maidstone, UK). 5. Tris-buffered saline (TBS) (2L): 48.4 g of Tris, 160 g of NaCl, 76 mL of 1 M HCl. Add milliQ H2 O to 90% of final volume. Adjust pH to 7.6 and add milliQ H2 O to final 2-L volume. Shake well and store at 4 °C. 6. TBS with Tween (TBS-T) (20 L): 20 mL of Tween, 2 L of 10X TBS; bring volume up to 20 L with milliQ H2 O. 7. Blocking buffer: 5% (w/v) nonfat dry milk in TBS-T. 8. Secondary antibody: Goat anti-mouse conjugated to horseradish peroxidase (Jackson ImmunoRes). 9. ECL Plus Western blotting detection reagents (Amersham Biosciences, Arlington Heights, IL). 10. Bio-Max XAR film (Kodak, Rochester, NY). 11. Autoradiography cassette (Fisher Biotech). 12. Developer.

2.8. Stripping and Reprobing Blots for -Tubulin 1. Stripping buffer (500 mL): 3.5 mL of 2-mercaptoethanol, 50 mL of 20% SDS, 31.25 mL of 1 M Tris HCI (pH 6.8), 415.25 mL of milliQ H2 O. Store at room temperature. 2. TBS-T. 3. Blocking solution. 4. Primary antibody: Mouse monoclonal anti--tubulin (Sigma). 5. Secondary antibody: Goat anti-mouse conjugated to horseradish peroxidase (Jackson ImmunoRes). 6. ECL Plus Western blotting detection reagents (Amersham Biosciences). 7. Bio-Max XAR film (Kodak).

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8. Autoradiography cassette (Fisher Biotech). 9. Developer.

2.9. Immunofluorescence for TorsinA 1. 18-mm round coverslips (Fisher). 2. 38 × 77 frosted glass slides (BRL, Waban, MA). 3. 4% PFA (paraformaldehyde)/PBS: Stir 20 g of PFA (Sigma) into 250 mL of sterile milliQ H2 O. Then add ∼10 drops of 5 N NaOH solution. Heat (to ∼65 °C) and stir in hood until PFA is dissolved. Add 50 mL of 10X PBS (0.1 M) and allow it to cool to room temperature. Adjust pH to 7.4 using 1 M of HCl (∼5 mL). Fill to 500-mL total with sterile milliQ H2 O. Filter-sterilize through 500-mL filter flask. Aliquot ∼45 mL into 50-mL conical tubes and store at −20 ºC. 4. 0.05% Triton X-100/PBS: Add 500 mL of PBS to 250 μL of Triton X-100 (Fisher). 5. Blocking buffer: 5% normal goat serum (NGS) in 0.05% Triton X-100/PBS. 6. Secondary antibody: Rhodamine (TRITC)-conjugated goat anti-mouse (Jackson ImmunoRes). 7. DAPI (Sigma): Dissolve the content of a 10-mg bottle in sterile, milliQ H2 O (shake and sit for 10–15 min). Transfer solution to a 15-mL tube. Bring the volume up to 10 mL and store 1-mL aliquots at 4 °C. 8. SlowFade Anti Fade kit (Invitrogen). 9. Permount (Fisher). 10. Zeiss Axioplan fluorescence microscope (Thornwood, NY). 11. Axiocam HRm (Zeiss) digital camera.

3. Methods The approach taken in our laboratory for the generation of shRNAs is a single-step PCR amplification of a RNA polymerase III (pol III) promoter followed by the hairpin sequence and a transcriptional termination signal. We elected to use the mouse U6 (mU6) promoter to drive expression of the shRNA, but other pol III promoters can be used. Based on a plasmid template containing the mU6 promoter, we designed a forward PCR primer (Fig. 1, primer 1) that includes 22 nucleotides located 85 nucleotides upstream of the mU6 promoter in this vector, and a reverse primer (Fig. 1, primer 2) that includes the sequence encoding a pol-III termination signal (UUUUUU, therefore AAAAAA in the primer), followed by the sequence complementary to the guide or antisense strand, an 8-nt loop (primer sequence CAAGCTTC), the passenger or sense strand, and the last 21 nucleotides of the mU6 promoter, so that the final DNA product is as shown in Fig. 1. This PCR product was then cloned in a plasmid vector lacking eukaryotic promoter, resulting in a plasmid that can be used in cotransfection experiments to test its efficacy

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Fig. 1. Overview of protocol to generate shRNA-expressing plasmid vectors.

against overexpressed targets (this step is not described here, but we strongly encourage the identification of highly effective hairpins in transient cotransfection experiments before moving to primary neurons). Overall, this is a rapid strategy to quickly generate shRNA expression plasmids employing routine molecular biology techniques. Effective hairpins are then cloned into the shuttle plasmid vector that will be used to generate recombinant lentivirus. The viral vector also encodes a GFP gene as a reporter of transduction (10). The resulting lentivirus is then used to transduce cultured neurons. We determine the degree of silencing of the target by measuring levels of the target protein, but mRNA quantification through quantitative RT-PCR can also be employed. 3.1. Generation of U6shRNA 1. Set up a 50-μL PCR reaction as follows: 45 μL of Platinum® PCR SuperMix High Fidelity, 2 μL of forward primer at 50 ng/μL, 2 μL of reverse primer at 50 ng/μL, and 1 μL of pAd5mU6 template plasmid at 10 ng/μL. 2. Perform the following amplification cycles: Initial denaturation for 3 min at 94 °C (1 cycle); denaturation for 30 sec at 94 °C, annealing for 30 sec at 50 °C, extension for 30 sec at 72 °C (30 cycles); final extension for 7 min at 72 °C (1 cycle). 3. Prepare a 1% agarose gel using standard procedures with a well capacity of 50 μL. 4. Load the resulting amplification reaction and verify by agarose gel electrophoresis using appropriate standards that a single band of 434 bp was produced. 5. Extract the PCR product from the agarose gel using Quantum Prep Freeze ‘N Squeeze DNA Gel Extraction Spin Columns. Following the manufacturer’s protocol, identify the bands of interest, and excise them using a razor blade, cutting off excess agarose. Insert the DNA-containing agarose piece into the

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Harper and Gonzalez-Alegre filter cup of the spin column, and then place the filter cup into the dolphin tube. Place in freezer (−20 ºC) for 5 min, then spin in tabletop microcentrifuge at 13,000 rpm, room temperature for 3 min. Collect the purified DNA from the tube and discard the column with the remaining agarose. The purified product obtained by this method can be directly used for cloning into the pCR2.1 vector as described here (go to next step). However, if problems are encountered with this cloning step, the PCR product can be precipitated as follows: Dilute Freeze ‘N Squeeze product in 3 M Na-acetate 1/10th vol, 100% EtOH 2X vol, and 1–2 μg of glycogen. Freeze in liquid N2 until solid. Thaw and spin at 14,000 rpm for 8 min. Remove EtOH, wash pellet with 70% EtOH, and air-dry for 2 min. Resuspend pellet in 5 μL of water. Alternatively, a small amount of the PCR product can be electrophoresed for confirmation of the appropriate size, and the remainder of the PCR product ligated directly into the cloning vector, thus eliminating the DNA purification step. In this case, it is essential that the template plasmid used in the PCR reaction contains a different bacterial resistance gene than the TOPO vector. However, components of the PCR present in the nonpurified PCR product may interfere with the cloning step to various extents. Following the manufacturer’s protocol, add 4 μL of the isolated PCR product directly to a tube containing 1 μL of pCR2.1 TOPO vector and 1 μL of salt solution (1.2 M NaCl; 0.06 M MgCl2 ; provided with TOPO kit). Incubate 5 min at room temperature. Transform 3 μL of TOPO cloning reaction into chemically competent E. coli (provided with the cloning kit) using standard procedures. Spread 50 μL of 40 mg/mL X-gal onto an ampicillin-selective LB plate, prewarm it to 37 ºC, and then spread 1/10th vol of the transformed bacteria. Centrifuge the remaining transformation at a low speed (e.g., 2,000 rpm for 1 min) to pellet bacteria. Remove supernatant and resuspend pellet in 100 μL of LB media. Spread remaining cells on a second plate. Incubate plates overnight at 37 °C. Next day, pick 5–10 white colonies from the plate into a culture tube containing 5 mL of LB medium and 100 μg/mL of ampicillin. Grow overnight in shaking incubator at 37 °C (see Note 2). Isolate plasmid DNA using Quantum Prep Plasmid Miniprep Kit following the manufacturer’s instructions. Any other commercial or custom method for DNA isolation can be used as well. Digest 5 μL of miniprep DNA with EcoRI as follows: 5 μL of miniprep DNA, 3 μL of 10X EcoRI buffer, 1.5 μL of 2 mg/mL bovine serum albumin (BSA) (20X), 0.5 μL of EcoR1 (at 20 units/μL), and 20 μL of water. Incubate at 37 °C for 2 h. Run 10 μL of the digestion product in 1% agarose gel to confirm the presence of a single band of 451 nucleotides by visualizing it using a UV trans-illuminator. Plasmid vectors with an insert of the appropriate size can be sent for sequencing using M13 forward and reverse primers.

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15. This vector can now be used for screening in cotransfection experiments with overexpressed targets in cultured cells (see Note 3).

3.2. Generation of FIV.eGFP.U6shRNA The protocol described here for the generation of lentiviral vectors encoding shRNAs is adapted from the Gene Transfer Vector Core at the University of Iowa (http://www.uiowa.edu/∼gene/) (10,13). This method generates recombinant feline immunodeficiency virus (FIV) pseudotyped with the vesicular stomatitis virus envelope glycoprotein (VSV-G) (10). FIV particles are generated using a triple plasmid system based on the cotransfection of 293T cells with three plasmids (shuttle, packaging, and envelope), followed by harvesting of particle-containing culture medium and concentration of particles (13). However, other lentiviral vectors are available and can be used, such as those based on the human immunodeficiency virus. Many scientists do not generate their own recombinant lentiviral vectors but purchase them from different academic or industry-based facilities. At this point, the plasmid vector encoding your effective pol III-shRNA can be sent to those centers for the generation of shuttle vectors. 3.2.1. Generation of Shuttle Plasmid 1. Digest 3 μL of pVETL CMVeGFP plasmid vector with MfeI for 1 h in a 37 ºC water bath. After 1 h, add 3 μL of CIAP buffer and 1 μL of CIAP and incubate for an additional hour in a 37 ºC water bath. Run in 1% agarose gel, and purify the digested vector using Quantum Prep Freeze ‘N Squeeze DNA Gel Extraction Spin Columns as above. Precipitate DNA with Na acetate as described earlier. 2. Excise mU6shRNA from the pCR2.1 vector using EcoRI as above (using 10 μL of the plasmid vector here). Run in 1% agarose gel and purify the digested insert (band of 451 nucleotides) using Quantum Prep Freeze ‘N Squeeze DNA Gel Extraction Spin Columns. 3. EcoRI and MfeI are compatible ends. Ligate mU6shRNA into pVETL CMVeGFP using the Rapid DNA ligation kit. Prepare the ligation reaction as follows: 5 μL of insert (mU6shRNA), 1 μL of vector (pVETL CMVeGFP), 2 μL of 5X DNA dilution buffer, and 2 μL of water, mixing well. Add 10 μL of 2X ligation buffer and 1 μL of T4 DNA Ligase, mixing well. Incubate for 5 min at room temperature. Rapidly transform competent cells using 10 μL of ligation reaction as above and grow in ampicillin-containing LB plates as above (except no need to spread X-gal). 4. Select 10 colonies per plate, grow in 5 mL of LB/ampicillin media overnight, and isolate plasmid DNA using Quantum Prep Plasmid Miniprep Kit as above. 5. Perform a diagnostic reaction to confirm ligation with NotI and HincII. If there is no insertion of the U6shRNA, the resulting band will be 544 bp. If there is insertion, the resulting band should be 995 bp.

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3.2.2. Virus Production 1. Seed 293T cells into 18 150-mm-diameter flat-bottom tissue culture dishes at a density of 107 cells per dish. 2. The next day, add 34 mL of room-temperature HBS to two 50-mL conical tubes. 3. Add 225 μg of the packaging plasmid, 337.5 μg of the vector plasmid, and 112.5 μg of the envelope plasmid to each HBS-containing tube and vortex well. 4. Slowly add 1.7 mL of room-temperature 2.5 M CaCl2 to each tube while slowly vortexing or shaking the HBS–plasmid mixture. 5. Let the solution stand for 25 min to allow precipitate formation. The solution should appear slightly translucent or cloudy. 6. Add both tubes of precipitate directly to 200 mL of DMEM. Briefly mix. 7. Aspirate off the medium from the cells (nine plates at a time). 8. Gently pipette the transfection solution onto the cells (15 mL per dish), and return the cells to the incubator. 9. Four to six hours after transfection, aspirate off the medium and provide 15 mL of fresh DMEM-10 per dish. 10. Collect the medium (containing vector particles) at 24, 36, and 72 h, each time replacing this medium with fresh DMEM-10. At each collection, filter the medium through a 0.45-μm filter (Nalgene PES, low-protein-binding 500-mL bottle-top filter) and store short-term at 4 ºC or long-term in 50-mL aliquots at −80 ºC. 11. Just before intended use, concentrate the particles by centrifuging the collected medium at 4 ºC for 16 h at 7,400g (7,000 rpm in the SLA 1500 rotor, Sorvall Centrifuge, 275-mL capacity tubes). Carefully pour off the supernatants and resuspend particles in lactose buffer. We typically resuspend the particles produced from an 18-plate transfection into a total volume of 3 mL.

3.2.3. Determining Titer by Limiting Dilution and Assay of Transgene (GFP) Expression 1. One day before transduction, seed a six-well flat-bottom plate with 2 × 106 HT-1080 cells per well in DMEM-10. 2. For transduction, make a 10-fold dilution series of concentrated FIV as follows: Add 1.584 mL of DMEM-2/polybrene in the first tube and 1.35 mL in tubes 2 through 6. Add 15 μL of virus to the first tube and vortex. Transfer 150 μL from the first to the second tube and vortex, and so on for the remaining tubes. 3. Remove culture media from wells. Add 1 mL of each dilution to separate wells. For wells 1 through 6, the dilution factors will thus be 103 , 104 , 105 , 106 , 107 , and 108 , respectively. Return the cells to the incubator. 4. Incubate the HT-1080 cells for 72 h and then feed with 1 mL of DMEM-10. 5. Incubate further for 24 h. Rinse monolayers with PBS. 6. Using an inverted fluorescent microscope, visualize and count the number of GFP-expressing cells in each well. The first two wells will often have too many

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positive cells to count. Doublets or small clusters of cells are counted as one, as they likely originated by division of a single transduced cell. 7. For each well, multiply the total blue cell count by the dilution volume (1 mL) and by the dilution factor. Determine the mean of all the wells. This number represents the transducing units per milliliter (TU/mL) of concentrated virus. By using this method, our concentrated FIVeGFP preparations typically contain 108 –109 TU/mL.

3.3. Primary Neuronal Cultures 1. Add poly-D-lysine to 12-well dishes, and leave for at least 30 min before aspiration at step 11 (if experiments are designed for immunofluorescence analysis, place a round coverslip before adding the poly-D-lysine). 2. Sacrifice pregnant mouse at embryonic day 16 by injecting with ketamine/xylazine (150 μg/g of ketamine, 20 μg/g of xylazine). 3. Place mouse on paper towels lining the dissection pan and soak abdomen with 70% EtOH. Rinse instruments with 70% EtOH. 4. Make incision by grabbing skin with tissue forceps and cutting laterally along lower abdomen with large straight scissors to expose uterine horns. Rinse instruments with 70% EtOH. 5. Remove uterus by grabbing with tissue forceps and gently pulling upward while cutting away connective tissue and fat with scissors. Rinse uterus with 70% EtOH and place in 10-cm dish containing ice-cold HBSS dissecting media. 6. To remove embryos, carefully tear open uterus above each embryo with tissue forceps and fine forceps. Embryos will have individual chorionic sacks to remove as well. Sever umbilical cord and transfer embryo to new 10-cm dish with ice-cold HBSS dissecting media. 7. Decapitate embryos using small scissors and transfer heads to new 10-cm dish with ice-cold HBSS dissecting media, ensuring that heads are completely immersed in subsequent steps. Place Petri dish on ice and move to dissecting microscope. 8. To remove the brain, begin by removing the skin from the skull using one pair of fine forceps to hold the head through the orbits, using the other pair of forceps to remove the skin. Puncture the skull with fine forceps in the junction between cortices and brain stem. Gently remove skull by tearing with fine forceps starting with tips close together and pulling apart. Pinch off olfactory bulbs. With closed forceps, pry brain away from head starting at its anterior end, and then pinch off at base of brain stem. Transfer brain and brain stem to new 6-cm dish of ice-cold HBSS dissecting media. 9. To isolate cortex, pry hemisphere away from brain stem and gently pinch off. Remove meninges, choroid plexus, basal ganglia, and hippocampus. Place isolated cortex into new 6-cm dish of ice-cold HBSS dissecting media. Eight to 10 brains per dish are optimal. The following steps are performed inside a TC hood. 10. Cut up dissected brain with scalpel blade into small pieces (∼2 mm). Using a 5-mL pipette, transfer tissue pieces from the 6-cm dish (cortices from 8–10 brains) to a 15-mL tube.

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11. Bring volume to 8 mL with HBSS dissecting media, and add 1 mL of 10X DNase and 1 mL of 10X trypsin. Invert tubes and incubate in a small 37 °C water bath for 12–15 min. During this incubation, aspirate poly-D-lysine and add Neurobasal/FBS plating media to plates. Store plates in 37 °C, 5% CO2 incubator until use. 12. Quick-spin 15-mL tube in centrifuge at 500 rpm to pellet tissue. Gently aspirate supernatant and wash tissue twice by adding 10 mL of room-temperature DMEM wash media, resuspending, quick-spinning, and removing supernatant. After second rinse, leave