siRNA and miRNA Gene Silencing: From Bench to Bedside (Methods in Molecular Biology, 487)

siRNA and miRNA Gene Silencing

METHODS IN MOLECULAR BIOLOGY™

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METHODS

MOLECULAR BIOLOGY™

siRNA and miRNA Gene Silencing From Bench to Bedside

Edited by

Mouldy Sioud Department of Immunology, Institute for Cancer Research, The Norwegian Radium Hospital, University of Oslo, Norway

Editor Mouldy Sioud Department of Immunology Institute for Cancer Research The Norwegian Radium Hospital University of Oslo Norway

Series Editor John M. Walker School of Life Sciences University of Hertfordshire Hatfield, Hertfordshire AL10 9AB, UK

ISBN: 978-1-60327-546-0 e-ISBN: 978-1-60327-547-7 ISSN: 1064-3745 e-ISSN: 1940-6029 DOI: 10.1007/978-1-60327-547-7 Library of Congress Control Number: 2008939423 © Humana Press, a part of Springer Science+Business Media, LLC 2009 All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Humana Press, C/o Springer Science+Business Media, LLC, 233 Spring Street, New York, NY 10013 USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. While the advice and information in this book are believed to be true and accurate at the date of going to press, neither the authors nor the editors nor the publisher can accept any legal responsibility for any errors or omissions that may be made. The publisher makes no warranty, express or implied, with respect to the material contained herein. Printed on acid-free paper 9 8 7 6 5 4 3 2 1 springer.com

Preface RNA interference (RNAi) refers to the process by which dsRNA molecules silence a target through the specific destruction of their mRNA molecules. Subsequent to the discovery that small interfering RNAs (siRNAs) mimicking the Dicer cleavage products can silence mammalian genes, RNAi has become the experimental tool of choice to suppress gene expression in a wide variety of organisms. In addition, RNAi has also become a method of choice for key steps in the development of therapeutic agents, from target discovery and validation to the analysis of the mechanisms of action of small molecules. To date, several strategies have been devised to trigger the RNAi pathway, each of which is adapted and optimized for different cell systems. Although the technology has several advantages over other methods, the specificity of gene silencing is not absolute and there is a danger of off-target effects and activation of innate immunity. Also, strategic success of therapeutic siRNAs will depend on the development of a delivery vehicle that can target pathogenic cells and from our understanding of the biogenesis of microRNAs (miRNAs). The purpose of this book is to provide readers with the recent advances in siRNA design, expression, delivery, in vivo imaging, and methods to minimize siRNA unwanted effects and use in patients. To design an effective siRNA, one must consider the base composition of the chosen site and whether the target site will be accessible. Chap. 1 critically reviews the published design guide rules and presents new statistical and clustering design strategies that are useful for selecting effective siRNA sequences. If the chosen target is an RNA virus that can mutate rapidly, one may consider to target conserved site sequences and/or to combine diverse siRNA sequences. Recent studies indicated that certain siRNA sequences can activate innate immunity resulting in the production of pro-inflammatory cytokines and type I interferons. Moreover, siRNAs can also silence the expression of unrelated genes, a phenomenon known as off-target effects that is mediated largely by limited target sequence complementary to the seed region of the siRNA guide strand. Unfortunately, the current tools for siRNA design (see Chap. 1) cannot eliminate all the potential unwanted effects of siRNAs. Chap. 2 offers valuable and detailed description of how to eliminate siRNA unwanted effects, including the activation of innate immunity and off-target effects. Also, it describes siRNA-based methods for enhancing tumor immunity. Notably, some of the main challenges in using siRNAs in vivo are the delivery, tissue targeting, and monitoring of siRNA potency in vivo. In vitro, siRNA duplexes have been delivered to target cells mainly by lipid-mediated transfection or via electroporation. However, these methods are not broadly applicable in patients. Approaches to improve in vivo delivery of siRNAs are currently being pursued using nanoparticles, new lipid formulations, and receptor-mediated targeting. Chaps. 3–8 describe new formulations and strategies with promising applications in vitro and in vivo. While Chap. 5 describes the first multimodal nanoparticles to deliver siRNAs, image siRNA v

Preface

uptake, and monitor gene silencing in tumors, Chap. 6 describes the first detailed protocol for siRNA magnetofection that is applicable in vitro and in vivo. Being RNA, siRNAs are prone to nuclease-mediated degradation in serum and the cytosol, which has a negative impact on their use in cells and patients. Chemical modifications of ribose (e.g., locked nucleic acids, 2¢-deoxy, 2¢-fluoro, 2¢-O-methyl) can enhance nuclease resistance without interfering with siRNA silencing potency. Chap. 9 describes the development of nuclease-resistant siRNAs with the potential to progress into a new class of therapeutic drugs. Chap. 10 describes new vectors for RNAi in which a synthetic siRNA/miRNA is expressed from a synthetic stem-loop precursor based on the miRNA 155 and miRNA 30 precursors. These new vectors offer several advantages over traditional RNAi vectors driven by RNA polymerase III promoters. These include the expression of several artificial miRNAs from a single transcript and tissue-specific expression as discussed in Chap. 3. By using siRNAs to downregulate gene expression in human cells, a number of therapeutic target genes have been validated both in vitro and in vivo. Several relevant examples are featured in Chaps. 11–15. These include oncogenes, growth factors, immune regulatory factors, urokinase plasminogen activator and its receptor, matrix metalloproteinases, hypoxia-induced factor, and telomerase. In addition to interfering with endogenous genes, siRNAs have been used to block viral replication. Nevertheless, in vitro and in vivo experiments have revealed potential problems of viral escape mutants. Chap. 16 describes the treatment of respiratory viral diseases with chemically modified new generation of siRNAs. And Chap. 17 describes the recent progress in using siRNAs as treatment for HIV-1 infection and several excellent recommendations are offered. Notably, the success of siRNAs will depend not only on the development of delivery strategies and chemical modifications, but also on our understanding of miRNA biogenesis. Naturally occurring miRNA are 19–24 nt in length cleaved from 60to 110-nt hairpin precursors that are produced from large primary transcripts. To date over 1000 miRNAs have been identified in humans. They play critical roles in developmental and physiological processes by regulating target gene expression at the post-transcriptional level. It is therefore not surprising that deregulation of miRNA expression could result in specific disease phenotypes. Recent studies have implicated miRNAs in cancer development. A group of three chapters describe the recent progress in understanding miRNA expression, function, and involvement in diseases. Chaps. 18 and 20 focuses on the recent progress in understanding the components involved in miRNA function, biogenesis, and interference with virus infection, and Chap. 19 demonstrates that intron-derived miRNA can induce RNAi not only in vitro but also in adult mice. Chap. 21 describes the design of effective miRNA sequences and their applications as anti-gene agents. The book ends by describing the first clinical trial in a patient with leukemia using a synthetic siRNA against Bcl-Abl fusion transcript (Chap. 22). It was found that the use of siRNAs in humans is safe, thus facilitating the progression of synthetic siRNA-based drugs to clinical trials. Topics covered in this volume will be of interest to researchers, teachers, students, and biotech companies interested in RNAi, gene regulation, and gene and immunotherapy. It is my hope that the readers will benefit from this collection of

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excellent chapters dealing with the recent advances of RNAi technology from the bench to bedside. Finally, I would like to thank the authors for their contributions, Anne Dybwad for excellent editorial assistance, the series editors, John Walkers, and all those involved in the production of the book. Mouldy Sioud

Contents Preface.................................................................................................................. Contributors ......................................................................................................... 1.

2. 3.

5. 6.

10. 11. 12.

Methods for Selecting Effective siRNA Sequences by Using Statistical and Clustering Techniques............................................................. Shigeru Takasaki Deciphering the Code of Innate Immunity Recognition of siRNAs ............... Mouldy Sioud Targeted Delivery of Antisense Oligonucleotides and siRNAs into Mammalian Cells ................................................................ Mouldy Sioud Local and Systemic Delivery of siRNAs for Oligonucleotide Therapy ............ Fumitaka Takesh*ta, Naomi Hokaiwado, Kimi Honma, Agnieszka Banas, and Takahiro Ochiya Imaging of siRNA Delivery and Silencing ..................................................... Anna Moore and Zdravka Medarova Recent Advances in Magnetofection and Its Potential to Deliver siRNAs in Vitro ............................................................................................ Olga Mykhaylyk, Olivier Zelphati, Edelburga Hammerschmid, Martina Anton, Joseph Rosenecker, and Christian Plank In Vitro and In Vivo Gene Silencing by TransKingdom RNAi (tkRNAi) ............................................................................................ Shuanglin Xiang, Andrew C. Keates, Johannes Fruehauf, Youxin Yang, Hongnian Guo, Thu Nguyen, and Chiang J. Li Bacterial Delivery of siRNAs: A New Approach to Solid Tumor Therapy ............................................................................................ De-Qi Xu, Ling Zhang, Dennis J. Kopecko, Lifang Gao, Yueting Shao, Baofeng Guo, and Lijing Zhao The Therapeutic Potential of LNA-Modified siRNAs: Reduction of Off-Target Effects by Chemical Modification of the siRNA Sequence ................................................................................. Kees Fluiter, Olaf R. F. Mook, and Frank Baas pSM155 and pSM30 Vectors for miRNA and shRNA Expression ................. Junzhu Wu, Akua N. Bonsra, and Guangwei Du Targeting Oncogenes with siRNAs ............................................................... Olaf Heidenreich Targeting Stromal–Cancer Cell Interactions with siRNAs ............................. Seyedhossein Aharinejad, Mouldy Sioud, Trevor Lucas, and Dietmar Abraham

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13.

Therapeutic Potential of siRNA-Mediated Targeting of Urokinase Plasminogen Activator, Its Receptor, and Matrix Metalloproteinases ...................................................................... Christopher S. Gondi and Jasti S. Rao 14. Silencing of HIF-1α by RNA Interference in Human Glioma Cells In Vitro and In Vivo ............................................................................ David L. Gillespie, Jeannette R. Flynn, Brian T. Ragel, Maria Arce-Larreta, David A. Kelly, Sheryl R. Tripp, and Randy L. Jensen 15. RNA Interference-Mediated Validation of Genes Involved in Telomere Maintenance and Evasion of Apoptosis as Cancer Therapeutic Targets ...................................................................... Marco Folini, Marzia Pennati, and Nadia Zaffaroni 16. Treating Respiratory Viral Diseases with Chemically Modified, Second Generation Intranasal siRNAs ........................................................... Sailen Barik 17. Progress in the Therapeutic Applications of siRNAs Against HIV-1 .............. Miguel Angel Martínez 18. Protein Components of the microRNA Pathway and Human Diseases .......... Marjorie P. Perron and Patrick Provost 19. Intron-Mediated RNA Interference and microRNA Biogenesis Shao-Yao Ying and Shi-Lung Lin .................................................................. 20. Emergence of a Complex Relationship Between HIV-1 and the microRNA Pathway ......................................................................... Dominique L. Ouellet, Isabelle Plante, Corinne Barat, Michel J. Tremblay, and Patrick Provost 21. Synthetic microRNA Targeting Glioma-Associated Antigen-1 Protein .......... Naotake Tsuda, Takahi Mine, Constantin G. Ioannides, and David Z. Chang 22. Therapeutic Targeting of Gene Expression by siRNAs Directed Against BCR-ABL Transcripts in a Patient with Imatinib-Resistant Chronic Myeloid Leukemia .......................................................................... Michael Koldehoff and Ahmet H. Elmaagacli Index ....................................................................................................................

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331 343 369

387 415

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Contributors DIETMAR ABRAHAM • Laboratory for Cardiovascular Research, Center for Anatomy and Cell Biology, Medical University of Vienna, Vienna, Austria SEYEDHOSSEIN AHARINEJAD • Laboratory for Cardiovascular Research, Center for Anatomy and Cell Biology, Medical University of Vienna, Vienna, Austria MARTINA ANTON • Institute of Experimental Oncology, Technische Universität München, Munich, Germany MARIA ARCE-LARRETA • Department of Neurosurgery, Huntsman Cancer Institute in the Division of Pediatric Hematology/Oncology, University of Utah School of Medicine, Salt Lake City, UT, USA FRANK BAAS • Department of Neurogenetics, AMC, Amsterdam, The Netherlands AGNIESZKA BANAS • Section for Studies on Metastasis, National Cancer Center Research Institute, Tokyo, Japan CORINNE BARAT • Research Center in Infectious Diseases, CHUL Research Center, Quebec, Canada SAILEN BARIK • Department of Microbiology and Immunology, College of Medicine, University of South Alabama, Mobile, AL, USA AKUA N. BONSRA • Department of Pharmacology and the Center for Developmental Genetics, Stony Brook University, Stony Brook, NY, USA DAVID Z. CHANG • Departments of Immunology, The University of Texas M. D. Anderson Cancer Center, Houston, TX, USA GUANGWEI DU • Department of Pharmacology and the Center for Developmental Genetics, Stony Brook University, Stony Brook, NY, USA AHMET H. ELMAAGACLI • Department of Bone Marrow Transplantation, University Hospital of Essen, Essen, Germany KEES FLUITER • Department of Neurogenetics, AMC, Amsterdam, The Netherlands JEANNETTE R. FLYNN • Center for Children, Huntsman Cancer Institute in the Division of Pediatric Hematology/Oncology, University of Utah School of Medicine, Salt Lake City, UT, USA MARCO FOLINI • Dipartimento di Oncologia Sperimentale e Laboratori, Fondazione IRCCS Istituto Nazionale dei Tumori, Milano, Italy JOHANNES FRUEHAUF • Skip Ackerman Center for Molecular Therapeutics, Division of Gastroenterology, Beth Israel Deaconess Medical Center, Harvard Medical School, Boston, MA, USA LIFANG GAO • Prostate Diseases Prevention and Treatment Research Centre, and Department of Pathophysiology, School of Basic Medicine, Jilin University, Changchun, China DAVID L. GILLESPIE • Department of Neurosurgery, Huntsman Cancer Institute in the Division of Pediatric Hematology/Oncology, University of Utah School of Medicine, Salt Lake City, UT, USA xi

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Contributors

CHRISTOPHER S. GONDI • Department of Cancer Biology and Pharmacology, University of Illinois College of Medicine at Peoria, Peoria, IL, USA BAOFENG GUO • Prostate Diseases Prevention and Treatment Research Centre, and Department of Pathophysiology, School of Basic Medicine, Jilin University, Changchun, China HONGNIAN GUO • Skip Ackerman Center for Molecular Therapeutics, Division of Gastroenterology, Beth Israel Deaconess Medical Center, Harvard Medical School, Boston, MA, USA EDELBURGA HAMMERSCHMID • Institute of Experimental Oncology, Technische Universität München, Munich, Germany OLAF HEIDENREICH • Newcastle University, Northern Institute for Cancer Research, Medical School, Newcastle upon Tyne, UK NAOMI HOKAIWADO • Section for Studies on Metastasis, National Cancer Center Research Institute, Tokyo, Japan KIMI HONMA • Koken Bioscience Institute, Tokyo, Japan CONSTANTIN G. IOANNIDES • Departments of Gynecologic Oncology, The University of Texas M. D. Anderson Cancer Center, Houston, TX, USA RANDY L. JENSEN • Department of Neurosurgery, Huntsman Cancer Institute in the Division of Pediatric Hematology/Oncology, University of Utah School of Medicine, Salt Lake City, UT, USA ANDREW C. KEATES • Skip Ackerman Center for Molecular Therapeutics, Division of Gastroenterology, Beth Israel Deaconess Medical Center, Harvard Medical School, Boston, MA, USA DAVID A. KELLY • Department of Neurosurgery, Huntsman Cancer Institute in the Division of Pediatric Hematology/Oncology, University of Utah School of Medicine, Salt Lake City, UT, USA MICHAEL KOLDEHOFF • Department of Bone Marrow Transplantation, University Hospital of Essen, Essen, Germany DENNIS J. KOPECKO • Laboratory of Enteric and Sexually Transmitted Diseases, Center for Biologics Evaluation and Research, Food and Drug Administration, Bethesda, MD, USA CHIANG J. LI • Skip Ackerman Center for Molecular Therapeutics, Division of Gastroenterology, Beth Israel Deaconess Medical Center, Harvard Medical School, Boston, MA, USA SHI-LUNG LIN • Department of Cell & Neurobiology, Keck School of Medicine, University of Southern California, Los Angeles, CA, USA TREVOR LUCAS • Laboratory for Cardiovascular Research, Center for Anatomy and Cell Biology, Medical University of Vienna, Vienna, Austria MIGUEL ANGEL MARTÍNEZ • Fundacio irsiCaixa, Universitat Autònoma de Barcelona (UAB), Spain ZDRAVKA MEDAROVA • Molecular Imaging Laboratory, MGH/MIT/HMS Athinoula A. Martinos Center for Biomedical Imaging, Department of Radiology, Massachusetts General Hospital/Harvard Medical School, Charlestown, MA, USA

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TAKAHI MINE • Departments of Gynecologic Oncology and Immunology, Kurume University, Kurume, Japan OLAF R. F. MOOK • Department of Neurogenetics, AMC, Amsterdam, The Netherlands ANNA MOORE • Molecular Imaging Laboratory, MGH/MIT/HMS Athinoula A. Martinos Center for Biomedical Imaging, Department of Radiology, Massachusetts General Hospital/Harvard Medical School, Charlestown, MA, USA OLGA MYKHAYLYK • Institute of Experimental Oncology, Technische Universität München, Munich, Germany THU NGUYEN • Skip Ackerman Center for Molecular Therapeutics, Division of Gastroenterology, Beth Israel Deaconess Medical Center, Harvard Medical School, Boston, MA, USA TAKAHIRO OCHIYA • Section for Studies on Metastasis, National Cancer Center Research Institute, Tokyo, Japan DOMINIQUE L. OUELLET • Centre de Recherche en Rhumatologie et Immunologie, CHUL Research Center, Quebec, Canada MARZIA PENNATI • Dipartimento di Oncologia Sperimentale e Laboratori, Fondazione IRCCS Istituto Nazionale dei Tumori, Milano, Italy MARJORIE P. PERRON • Centre de Recherche en Rhumatologie et Immunologie, CHUL Research Center, Quebec, Canada CHRISTIAN PLANK • Technische Universität München, Munich, Germany ISABELLE PLANTE • Centre de Recherche en Rhumatologie et Immunologie, CHUL Research Center, Quebec, Canada PATRICK PROVOST • Centre de Recherche en Rhumatologie et Immunologie, CHUL Research Center, Quebec, Canada BRIAN T. RAGEL • Department of Neurosurgery, Huntsman Cancer Institute in the Division of Pediatric Hematology/Oncology, University of Utah School of Medicine, Salt Lake City, UT, USA JASTI S. RAO • Department of Cancer Biology and Pharmacology, University of Illinois College of Medicine at Peoria, Peoria, IL, USA JOSEPH ROSENECKER • Forschungszentrum der Kinderklinik und Poliklinik Dr. von Haunersches Kinderspital, Kubus Rückgebäude, München, Germany YUETING SHAO • Prostate Diseases Prevention and Treatment Research Centre, and Department of Pathophysiology, School of Basic Medicine, Jilin University, Changchun, China MOULDY SIOUD • Department of Immunology, Institute for Cancer Research, The Norwegian Radium Hospital, University of Oslo, Norway SHIGERU TAKASAKI • RIKEN Genomic Sciences Center (GSC),Tsurumi-ku, Yokohama, Kanagawa, Japan FUMITAKA TAKEsh*tA • Section for Studies on Metastasis, National Cancer Center Research Institute, Tokyo, Japan MICHEL J. TREMBLAY • Research Center in Infectious Diseases, CHUL Research Center, Quebec, Canada

xiv

Contributors

SHERYL R. TRIPP • ARUP Laboratories, Salt Lake City, UT, USA NAOTAKE TSUDA • Departments of Gynecologic Oncology and Immunology, Kurume University, Kurume, Japan JUNZHU WU • Department of Pharmacology and the Center for Developmental Genetics, Stony Brook University, Stony Brook, NY, USA SHUANGLIN XIANG • Skip Ackerman Center for Molecular Therapeutics, Division of Gastroenterology, Beth Israel Deaconess Medical Center, Harvard Medical School, Boston, MA, USA DE-QI XU • Laboratory of Enteric and Sexually Transmitted Diseases, Center for Biologics Evaluation and Research, Food and Drug Administration, Bethesda, MD, USA YOUXIN YANG • Skip Ackerman Center for Molecular Therapeutics, Division of Gastroenterology, Beth Israel Deaconess Medical Center, Harvard Medical School, Boston, MA, USA SHAO-YAO YING • Department of Cell & Neurobiology, Keck School of Medicine, University of Southern California, Los Angeles, CA, USA NADIA ZAFFARONI • Dipartimento di Oncologia Sperimentale e Laboratori, Fondazione IRCCS Istituto Nazionale dei Tumori, Milano, Italy OLIVIER ZELPHATI • OZ Biosciences, Parc Scientifique et Technologique de Luminy, Marseille, Cedex 9, France LING ZHANG • Prostate Diseases Prevention and Treatment Research Centre, and Department of Pathophysiology, School of Basic Medicine, Jilin University, Changchun, China LIJING ZHAO • Prostate Diseases Prevention and Treatment Research Centre, and Department of Pathophysiology, School of Basic Medicine, Jilin University, Changchun, China

Chapter 1 Methods for Selecting Effective siRNA Sequences by Using Statistical and Clustering Techniques Shigeru Takasaki Abstract Short interfering RNAs (siRNAs) have been widely used for studying gene functions in mammalian cells but vary markedly in their gene-silencing efficacy. Although many design rules/guidelines for effective siRNAs based on various criteria have been reported recently, there are only a few consistencies among them. This makes it difficult to select effective siRNA sequences targeting mammalian genes. This chapter first reviews the reported siRNA design guidelines and clarifies the problems concerning the current guidelines. It then describes the recently reported new scoring methods for selecting effective siRNA sequences by using statistics and clustering techniques such as the self-organizing map (SOM) and the radial basis function (RBF) network. In the proposed three methods, individual scores are defined as a gene degradation measure based on position-specific statistical significances. The effectiveness of the methods was confirmed by evaluating effective and ineffective siRNAs for recently reported genes and comparison with other reported scoring methods. The sizes (values) of these scores are closely correlated with the degree of gene degradation, and the scores can easily be used for selecting high-potential siRNA candidates. The evaluation results indicate that the proposed new methods are useful for selecting siRNA sequences targeting mammalian mRNA sequences. Key words: siRNA design, RNA interference, gene silencing, SOM classification, statistical significance, RBF network.

1. Introduction Although RNA interference (RNAi) has been successfully used for studying gene functions in both plants and invertebrates, many practical obstacles need to be overcome before it becomes an established tool for use in mammalian systems (1–6). One of the important problems is designing effective short interfering RNA (siRNA) sequences for target genes. The siRNA responsible for M. Sioud (ed.), Methods in Molecular Biology, siRNA and miRNA Gene Silencing, vol. 487 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-60327-547-7_1

Takasaki

RNA interference varies markedly in its gene-silencing efficacy in mammalian genes, where the gene-silencing effectiveness depends very much on the target sequence positions (sites) selected from the target gene (7,8). Since different siRNAs synthesized for various positions induce different levels of gene silencing, the selection of the target sequence is critical for the effectiveness of the siRNA. We therefore need useful criteria for gene-silencing efficacy when we design siRNA sequences (9,10). Zamore et al. and Jayasena et al. showed that the 5′ end of the antisense strand might be incorporated into the RNA-induced silencing complex (RISC). Strand incorporation may depend on weaker base pairing and thus an A–T terminus may lead to more strand incorporation than a G–C terminus (11,12). Other factors reported to be related to gene-silencing efficacy are GC content, point-specific nucleotides, specific motif sequences, and secondary structures of mRNA. Several siRNA design rules/guidelines using efficacy-related factors have been reported (13 –17). Although the positional nucleotide characteristics for siRNA designs seem to be the most important factor determining effective siRNA sequences, there are few consistencies among the reported rules/guidelines (18–23). This implies that these rules/ guidelines might result in the generation of many candidates and thus make it difficult to extract a few for synthesizing siRNAs. Furthermore, there is in RNAi a risk of off-target regulation: a possibility that the siRNA will silence other genes whose sequences are similar to that of the target gene. When we use gene silencing for studying gene functions, we have to first somehow select high-potential siRNA candidate sequences and then eliminate possible off-target ones (24). This chapter first reviews the reported siRNA design guidelines and clarifies the problems concerning the reported guidelines. It then describes the recently reported new scoring methods for selecting effective siRNA sequences by using statistical and clustering techniques (25–32). In the statistical method, many effective siRNA sequences are examined in the literature (31), because it can be hypothesized that position-specific nucleotides play important roles in gene-silencing efficacy. If specific features of nucleotide frequencies appeared in many effective siRNAs, they mean the positional nucleotide characteristics for siRNA designs. The features of nucleotide frequencies at individual positions are then analyzed by using the statistical significance test. As these features can be considered as new guidelines, a measure (score) for selecting effective siRNA candidates is defined based on the positional features of specific significant nucleotides. The effectiveness of the proposed measure was confirmed by comparing the computed scores with those of the recently reported other selection methods (28,29,31).

Methods for Selecting Effective siRNA Sequences

The chapter then describes how to extract individual nucleotide features from many effective siRNA sequences by using mathematical clustering techniques – the SOM and the RBF network (see later Sects. 3.2.1 and 3.2.2) (25–27). In the SOM-based clustering method, siRNA classification from many effective siRNAs is first described. It is then shown how positional nucleotide features are extracted from the classified groups and is demonstrated how the extracted features are integrated as a measure (score). It is finally confirmed that the SOM method is effective by evaluating the relations between the scores and effective/ineffective siRNAs reported in the literature and comparing them with those of other reported scoring methods (30,33). In the RBF-network-based method, after the siRNA classification is carried out by using the RBF network (25,26), the preferred and unpreferred nucleotides for effective siRNAs at individual positions are determined by significance testing and are used to calculate a score that measures a sequence’s potential for gene degradation. The effectiveness of the proposed scoring method was then confirmed by using it to evaluate RNA sequences recently reported to effectively or ineffectively suppress the expression of various genes (see later subsection) and comparing it with other scoring methods (32,33). As a result of various evaluations, it is found there are good correlations between the sizes (values) of the proposed individual scores and the effectiveness and ineffectiveness of the recently reported siRNA sequences. The evaluation results indicate that the three methods would be useful for selecting siRNA sequences for mammalian genes.

2. RNA Interference and siRNA Sequence Selection Problem 2.1. RNA Interference

RNA interference (RNAi) is a phenomenon that silences gene expression by introducing double-stranded RNA (dsRNA) hom*ologous to the target mRNA (1). After this phenomenon was discovered in the nematode Caenorhabditis elegans, it gradually became clear that similar phenomena occur in the cells of plants, fungi, and mammals (1–6). RNAi has been reported to result from the following sequence of events (2,5,6). Long dsRNA is first cleaved into siRNA species by an RNAase III enzyme, Dicer. These siRNAs are then incorporated into an RNA-induced silencing complex (RISC), where the duplex siRNA is unwound so the antisense strand can guide RISC to the target mRNA having the complementary sequence. Finally, the target mRNA is cleaved at a single site in the center of the duplex region between the guide siRNA and the target mRNA (28). Among the events that are

Takasaki

still unclear, however, are the mechanism of the target mRNA cleavage and the mechanism by which the center of the duplex region in the RISC is identified. Furthermore, although RNAi has been widely used for studying gene functions, the effectiveness with which the genes in mammalian cells can be silenced this way depends very much on target sequence positions (sites) selected from the target gene. That is, different siRNAs synthesized from various positions induce different levels of gene silencing. This indicates that the selection of the target sequence position (site) is critical for the effectiveness of the siRNA (3–6). 2.2. siRNA Sequence Selection Problem 2.2.1. Related Works Regarding the siRNA Sequence Selection Problem

2.2.2. The Reported Guidelines for siRNA Sequence Design

To use RNAi as a biological tool for mammalian cell experiments, we first need to identify target sequences causing gene degradation. They have so far been identified by using a trail-and-error method (3,8), but siRNAs extracted from different regions of the same gene have varied remarkably in their effectiveness. The difficulty of using the trail-and-error method to select target sequences causing gene silencing increases when the coding regions are long, as they are in mammalian cells. This is because the larger the number of candidates becomes, the more difficult it is to get gene-silencing candidates. The earliest guidelines for siRNA sequence design were proposed by Elbashir et al. (4,8,40). They suggested that synthesizing siRNA duplexes of 21 nucleotides (nt) length – 19 nt base-paired sequence with 2 nt 3′ overhang at the ends – mediates efficient cleavage of the target mRNA. Their rules are summarized as follows. (1) Select the target region from the open reading frame (ORF) of a given cDNA sequence preferably 50–100 nt downstream of the start codon. Avoid 5′ or 3′ untranslated regions (UTRs) or regions close to the start codon as these may be richer in regulatory protein-binding sites. (2) Search for sequences 5′-AA(N19)UU, where N is any nucleotide, in the mRNA sequence and choose those with approximately 50% GC content. Highly G-rich sequences should be avoided because they tend to form G-quartet structures. If there are no 5′-AA(N19)UU motifs present in the target mRNA, search for 5′-AA(N21) or 5′-NA(N21), and synthesize the sense siRNA as 5′-(N19) TT and the antisense siRNA as 5′-(N′19)TT, where N′19 denotes the reverse complement sequence of N19 and T indicates 2′-deoxythymidine. (3) Blast-search the selected siRNA sequences against EST libraries or mRNA sequences of the respective organism to ensure that only a single gene is targeted.

Methods for Selecting Effective siRNA Sequences

(4) It may be advisable to synthesize several siRNA duplexes to control for the specificity of the knockdown experiments; those siRNAs duplexes that are effective for silencing should produce exactly the same phenotype. Furthermore, a nonspecific siRNA duplex may be needed as control. (5) If the siRNA does not work, first verify that the target sequence and the cell line used are derived from the same organism. Finally, make sure that the mRNA sequence used for selection of siRNA duplexes is reliable; it could contain sequencing errors, mutations, or polymorphisms. After that, many siRNA design guidelines/rules were reported as follows. Reynolds et al. analyzed 180 siRNAs systematically, targeting every other position of two 197-base regions of firefly luciferase and human cyclophilin B mRNA (90 siRNAs per gene), and reported the following eight criteria for improving siRNA selection (18). G1 (Reynolds et al.) eight criteria: (1) G/C content 30–52%, (2) at least 3 As or Ts at positions 15–19, (3) absence of internal repeats, (4) an A at position 19, (5) an A at position 3, (6) a T at position 10, (7) a base other than G or C at position 19, and (8) a base other than G at position 13. Ui-Tei et al. examined 72 siRNAs targeting six genes and reported four rules for effective siRNA designs (19). They are summarized as follows. G2 (Ui-Tei et al.) four rules: (1) A or T effective and G or C ineffective at position 19, (2) G or C effective and A or T ineffective at position 1, (3) at least 5 T or A residues from positions 13 to 19, and (4) no GC stretch more than 9 nt long. Amarzguioui and Prydz analyzed 46 siRNAs targeting four genes and reported the following six rules for effective siRNA designs based on their literature (20). G3 (Amarzguioui and Prydz) six rules: (1) G or C positive and T negative at position 1, (2) A positive at position 6, (3) T negative at position 10, (4) T positive at position 13, (5) C positive at position 16, and (6) A or T positive and G negative at position 19. Jagla et al. tested 601 siRNAs targeting one exogenous and three endogenous genes and reported four rules as follows (22). G4 (Jagla et al.): (1) A or T positive at position 19, (2) A or T positive at position 10, (3) G or C positive at position 1, and (4) more than three A/Ts between positions 13 and 19. Hsieh et al. examined 138 siRNAs targeting 22 genes and reported the following position-specific characteristics (21). G5 (Hsieh et al.): (1) T positive and G negative at position 19, (2) C or G positive and A or T negative at position 11, (3) G positive at position 16, (4) A positive at position 13, and (5) C negative at position 6.

Takasaki

The above previous works for positional characteristics in siRNA designs are summarized in Table 1.1a. Other scoring, screening, and designing methods for functional siRNAs have also been reported recently. Chalk et al. reported seven rules (“Stockholm rules”) based on thermodynamic properties. They are (1) total hairpin energy 20 kHz) sound waves are emitted from a transducer placed against the skin and the ultrasound is reflected back from the internal organs under examination. Contrast in the images obtained depends on the imaging algorithm used, backscatter, attenuation of the sound, and sound

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speed. Ultrasound imaging using diagnostic ultrasound instrumentation operating in the 7.5–15 MHz frequency range has been successfully applied to a variety of mouse models (26), yielding images with a spatial resolution of 300–500 µm. The strengths of ultrasound in cardiac, obstetric, vascular, and abdominal imaging appear most likely to extend to the mouse as a model when the technology is scaled down to achieve high resolution and a level of practicality/functionality similar to that available with present clinical ultrasound systems. Another emerging concept is that of using targeted ultrasonic contrast agents for molecular imaging of specific cell-surface receptors, especially within the vascular compartment (27). All of the above modalities, theoretically, can be applied to imaging of siRNA delivery and function as well as monitoring its time-dependent therapeutic influence on disease progression in the same animal or patient. The choice of modality depends on the spatial and temporal resolution needed, and the availability of amplification strategies (molecular imaging probes).

3. Imaging of RNA InterferenceCurrent Status 3.1. Imaging of siRNA Using Bioluminescence

The field of in vivo RNA interference imaging is still in its early stages of development. Studies that describe the imaging of RNA interference are mostly restricted to fluorescence or bioluminescence reporter imaging. A lot of the early proof-of-principle studies on in vivo RNAi utilized suppression of luciferase transgene expression by either synthetic siRNAs or small-hairpin RNAs transcribed in vivo, achieving impressive 80–90% silencing efficiencies in the liver (28). In the same study, the authors fused luciferase RNA with an endogenous gene encoding nonstructural protein 5B from viral-polymerase-encoding region of hepatitis C virus. By measuring the bioluminescence signal, they demonstrated that siRNA targeting reduced luciferase expression from HCV protein–luciferase fusion by 75%. This result suggests that it may be feasible to use in vivo imaging to monitor siRNA as a therapeutic tool and establish a method for the in vivo quantitative imaging of the silencing effect. In a similar study, Lewis et al. (29) co-injected a plasmid expressing luciferase with antiluciferase synthetic siRNA and observed significant 80–90% silencing, as measured by bioluminescence imaging in a variety of organs. This investigation, for the first time, utilized in vivo imaging to identify, on a whole-body scale, the scope of siRNA biological activity. More recently, bioluminescence imaging of luciferase expression was used to demonstrate the dose- and time-dependence of vector-based in vivo RNAi after

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hydrodynamic injection (30). Bioluminescence imaging has also been employed for the development of optimized methods for lipid-based siRNA delivery, where siRNA was complexed with polyethyleneimine-based polyplexes and cationic lipid-based lipoplexes. RNAi directed against luciferase resulted in nearly 80% inhibition with as little as 0.2 pmol siRNA (31). This inhibition was dose dependent and specific. A group from Japan utilized BLI for imaging siRNA complexed with atelocollagen. They showed that site-specific in vivo administration of an antiluciferase siRNA/atelocollagen complex reduced luciferase expression in a xenografted tumor. Furthermore, atelocollagen-mediated transfer of siRNA in vivo showed efficient inhibition of tumor growth in an orthotopic xenograft model of a human nonseminomatous germcell tumor (32). Another study from the same group utilized BLI for monitoring the effect of systemic administration of siRNA complexed with atelocollagen and directed against potential targets to bone metastasis (33). This treatment resulted in an efficient inhibition of metastatic tumor growth in bone tissues. In addition, this delivery method was found to be nontoxic and did not cause upregulation of serum cytokine levels. Though not using BLI directly, Takabatake et al. employed luciferase-directed siRNA for in vivo delivery to the kidney. Their method utilized injection via the renal artery followed by electroporation (34). Luciferase targeting by siRNA encapsulated in the interior of pegylated immunoliposomes modified with monoclonal antibody to the transferrin receptor resulted in 64–68% knockdown of the transcript (35). In a comprehensive imaging study (36), bioluminescence was applied for the noninvasive assessment of P-glycoprotein silencing. Multidrug resistance (MDR) remains a major obstacle to successful chemotherapeutic treatment of cancer and can be caused by overexpression of P-glycoprotein, the MDR1 gene product. shRNA constructs against human MDR1 mRNA synthesized in this study inhibited expression of P-glycoprotein by >90%. Furthermore, after somatic gene transfer by hydrodynamic infusion of a MDR1-Firefly luciferase (MDR1-FLuc) fusion construct into mouse liver, the effect of shRNAi delivered in vivo on P-glycoprotein-FLuc protein levels was documented with bioluminescence imaging using d-luciferin. shRNAi against MDR1 reduced the bioluminescence output of the P-glycoprotein-FLuc reporter fourfold in vivo compared with mice treated with control or scrambled shRNAi (36) (Fig. 5.1). Hu-Lieskovan et al. reported on the delivery of siRNA by a nonviral delivery system, which uses a cyclodextrin-containing polycation to bind and protect siRNA and transferrin as a targeting ligand for delivery to transferrin receptor–expressing tumor cells. Bioluminescence imaging was employed here to show the effect of siRNA targeting metastatic Ewing’s sarcoma chimeric fusion gene EWS-FLI1, found in 85% of patients with Ewing’s family of

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shRNAi

Image Min:-2.0726e+05 Max= 5.659e+06 p/s ec/cm^2/sr

Coelenterazine cp

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Color Bar Min = 2.5e +05 Max = 3.1e+06

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Image Min = -22076 Max = 9.5122e+05 p/s ec/cm^2/sr

Color Bar Min = 90000 Max = 1.56e+06

Fig. 5.1. Downregulation of P-glycoprotein in vivo: shRNAi-mediated reduction in P-glycoprotein protein levels. Representative bioluminescence images of RLuc expression with coelenterazine cp (top) and P-glycoprotein-FLuc expression with D-luciferin (bottom). Mice were hydrodynamically transfected with pMDR1-FLuc (1 µg) combined with pRLuc-N3 (1 µg; as transfection control) and treated as indicated with 10-fold excess of control (left), scrambled shRNAi (middle), or shRNAi against MDR1 (right). (Reprinted by permission from AACR: Clin Cancer Res., 11, 4487–4494 (2005)).

tumors (EFT). BLI showed that only the targeted, formulated siRNA against this gene achieves long-term tumor growth inhibition (37). More recently, bioluminescence imaging was applied together with a mathematical model of siRNA delivery and function, in order to define, in a more general context, the effects of target-specific and treatment-specific parameters on siRNAmediated gene silencing (38). This particular study demonstrated that in rapidly dividing subcutaneous tumors, the silencing effect may persist ~10 days; in nondividing hepatocytes it may last as long as 3–4 weeks. More importantly, this study represents the origin of the current belief that siRNA dilution due to cell division,

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and not intracellular siRNA half-life, governs the duration of gene silencing. As a result of this study, now it is possible to apply this mathematical model in treatment design and to predict the dosing schedule required to maintain persistent silencing of target proteins with different half-lives in rapidly dividing or nondividing cells. While many studies have explored bioluminescence imaging for the quantitative evaluation of RNAi efficiency (32, 34, 35, 39), this imaging modality lacks clinical a equivalent, therefore precluding it from application in humans. 3.2. Imaging of siRNA Using GFP Reporter

Green fluorescent protein and its modifications have been used mostly for ex vivo imaging and microscopy to monitor the efficiency of siRNA action. As such, high-pressure hydrodynamic injection of siRNA–EGFP resulted in substantial reduction of EGFP expression in the liver in a large percentage of hepatocytes at 48 h after injection, as determined by fluorescent microscopy (29). Similar questions have been addressed using GFP reporters (8, 36, 40, 41). For example, the utility of lentiviral vectors for the in vivo delivery of short-hairpin RNAs to the brain was demonstrated using an EGFP reporter (42). However, this method has been restricted to ex vivo analysis with fluorescence microscopy or flow cytometry. An in vivo GFP fluorescence study reported by Rubinson et al. involved imaging of transgenic mice in which a lentiviral vector carrying shRNA also incorporated EGFP as a reporter gene permitting the evaluation of shRNA delivery (43). Importantly the effect (94%) persisted into adulthood, due to integration of the vector into the host genome. Another in vivo imaging study involved intratumoral injection of siRNA against EGFP into EGFP-expressing B16F10 melanoma tumors followed by application of an external electric field. When the EGFP–siRNA was electrotransferred, a significant decrease in the GFP fluorescence of the tumor was observed by direct imaging of the animal (Fig. 5.2) and quantification of the fluorescence levels within 2 days following the treatment. The decrease was maximal at days 2–4 (reduction to about 30% of the physiological saline control) (44). However, this is a very artificial system, requiring transgenesis and intratumoral injection, and only applicable in a research scenario. Besides this study, direct investigation of the delivery of siRNA to target tissues is exclusively based on ex vivo experiments (45–50). These investigations provided answers to important new questions, such as the subcellular distribution of siRNAs delivered as part of various formulations, including tumor-targeted carriers (46) and their bioavailability.

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Fig. 5.2. Fluorescence analysis of tumors by in vivo stereomicroscopy in live animals. Ten days after the subcutaneous injection of 1 × 106 B16-F10 GFP cells, tumors having a mean diameter of 5–7 mm were treated. PBS + EP: intratumoral injection of 50 µl saline solution followed by application of EP. p76 + EP: intratumoral injection of 50 µl of saline solution containing 12 µg of p76 siRNA followed by application of EP. Anti-GFP − EP and anti-GFP + EP: intratumoral injection of 50 µl of saline solution containing 12 µg of egfp22 siRNA, followed (anti-GFP + EP) or not (anti-GFP − EP) by application of EP. B16-F10 GFP-derived tumors were clearly detected under the animal’s skin upon fluorescence excitation and tumor margin could be easily defined. This enabled measurement of the tumor area and fluorescence intensity over a period of 5 days after treatment. However, owing to the growth characteristics of B16F10 tumors, necrosis occurred 3 days after the beginning of the treatment, as shown by the heterogeneity of the fluorescence pattern in the central region. (a) Representative images of EGFP fluorescence in B16-F10 tumors observed by noninvasive imaging in live animals 2 days after the different treatments. (b) Pseudocolor 8-bit images showing a representative analysis over time of tumor fluorescence (fluorescence levels: 256; white: most intense; black: least intense) upon treatment with the siRNA egfp22, which allow to see the different nodules that sometimes form the tumor. In the anti-GFP + EP sample a decrease in fluorescence can be observed in the nodules that were injected with the specific siRNA on day 0 but not in the uninjected small nodule emerging close to the main tumor (arrows). (Reprinted by permission from Macmillan Publishers Ltd.: Gene Therapy, 14, 752–759 (2007)).

3.3. Imaging of siRNA Using Other Optical Imaging Approaches

Application of quantum dots for imaging siRNA delivery and silencing is so far limited to in vitro studies with the possibility to translate them to in vivo applications. One study utilized cationic liposomes to co-deliver green quantum dots and siRNA targeting the lamin a/c gene (Lmna) into murine fibroblasts, followed by flow cytometry. The authors report that in co-transfected cells gene silencing correlated directly with intracellular fluorescence and resulted in about 90% knockdown in highly fluorescent cells (51). Earlier this year the same group reported on a new system for siRNA delivery to tumor cells that consisted of a PEGylated quantum dot core as a scaffold with siRNA (to EGFP) and tumorhoming peptides attached to it. This system afforded intracellular localization of the quantum dot particle followed by endosomal escape mediated by the addition of cationic liposomes. A rather

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modest knockdown in EGFP fluorescence (about 29%) was achieved using this method (52). In another study quantum dots were encapsulated into chitosan nanoparticles labeled with the HER2 antibody targeting the HER2 receptors to deliver HER2/ neu siRNA. Targeted delivery of HER2 siRNA to HER2-overexpressing SKBR3 breast cancer cells was shown to be specific and confirmed by quantum dot microscopy (53). While quantum dots remain an attractive tool for in vitro and animal testing, where fluorescence is the most accessible and common imaging modality, concerns over their cytotoxicity and the limited tissue penetration of light should be taken into account. The studies on direct siRNA imaging so far have been limited to end-labeling of the molecule with fluorophores to monitor delivery of siRNA into cells. In the study by Chang et al., the authors propose to utilize molecular beacons (MBs) to leverage RNAi towards diagnostic and monitoring applications (54). The MBs proposed here would rely on a fluorescence resonance energy transfer (FRET) fluorophore pair to generate signals upon binding to complementary sequences. The siRNA probe utilized in this study targeted the hTR sequence of telomerase mRNA. Cancer cells express high levels of telomerase mRNA, while normal cells have very limited expression of telomerase. The probe contained a Cy3-Cy5 fluorophore pair conjugated to the 3′ terminus of the antisense and 5′ terminus of the sense strand, respectively, and a highly flexible poly(ethyleneglycol) (PEG) molecule covalently linked to the RNA strands as the loop of an siRNA-based MB. Confocal imaging demonstrated the activation of the siRNA probe in cancer cells in contrast to normal cells. The authors reported that effective gene silencing (about 80%) of telomerase by the siRNA-based probe was achieved (54). This approach could potentially provide promising new ways to evaluate mRNA expression in diseases and serve as useful reporter/ gene silencing tools for target validation in basic science and for dual imaging and therapy in clinical research. 3.4. Imaging of siRNA Using SPECT

The only study that we are aware of utilizing SPECT imaging for visualizing cellular delivery and animal distribution of siRNA was published by a group from the University of Massachusetts Medical School (55). siRNA in this study was chemically modified with an S-acetyl-N-hydroxysuccinimide (NHS) hydrazino nicotinamide (HYNIC) chelator to introduce 99mTc, a common isotope for SPECT imaging. Cellular accumulation of radiolabeled RNAs (both sense and antisense against the RIa mRNA of the Type I regulatory subunit a of cAMP-dependent protein kinase A) was tested in the ACHN kidney cancer cell line, which was selected for its high RIa mRNA expression. Cellular uptake of 99mTc-labeled small RNA was time dependent and reached 3 × 105 siRNA molecules after 24 h of incubation. There was no

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difference in cellular localization between radiolabeled sense and antisense RNA as confirmed by autoradiography. The delivery of 99mTc-labeled RNAs in vivo was tested in nude mice bearing ACHN human kidney cancer xenografts. Both sense and antisense 99mTc-labeled RNAs accumulated in the tumors at the same rate following intravenous injection. The highest tumoral accumulation was observed at 4 h after injection, with radioactivity visualized in almost all tissues. While this study produced encouraging results regarding the absolute tumoral accumulation of radiolabeled RNAs, it is not clear whether the delivery of functional RNA was achieved and whether any therapeutic effect was observed. Nevertheless, this study paved the way for further investigations on the in vivo imaging of siRNA delivery. 3.5. Multimodal Imaging of siRNA Using MRI and Optical Imaging

From multiple recent studies, it has become apparent that the most promising strategies for the cancer therapeutic applicability of RNA interference would involve their complexing/conjugation to biocompatible carriers in order to improve their accumulation in target tissues and their bioavailability. A study from a group at Harvard was the first one to utilize the multimodal approach to image siRNA delivery and silencing in tumors (24). To facilitate siRNA delivery to tumors and enable in vivo imaging of the delivery, the authors employed magnetic nanoparticles (MN), which had an optimal half-life of 8–12 h (56) and allowed visualization by magnetic resonance imaging. These nanoparticles were also labeled with Cy5.5 optical dye to enable correlative near-infrared optical imaging (NIRF). These imaging moieties were conjugated to a siRNA duplex targeting a gene of interest, as well as to myristoylated polyarginine peptides that facilitated translocation of the construct across the cell membrane into the cytoplasm. First, a model system involving animals bilaterally injected with 9L rat gliosarcoma tumors expressing GFP or RFP proteins was used. In vivo MR imaging of tumor-bearing animals followed by NIRF imaging of the same animals demonstrated that intravenous injection of MN-NIRF-siGFP resulted in successful tumoral delivery of the construct. In vivo optical imaging in the GFP channel confirmed that efficient silencing was achieved in 9L-GFP tumors (but not in 9L-RFP tumors) (Fig. 5.3). This was further confirmed by quantitative RT-PCR of gfp mRNA transcript levels. In addition to the GFP model system, we have explored a therapeutic probe targeting the antiapoptotic gene Birc5, which encodes survivin. Survivin is a member of the inhibitor of apoptosis protein (IAP) family, which is highly expressed in most cancers and associated with chemotherapy resistance, increased tumor recurrence, and shorter patient survival, making antisurvivin therapy an attractive cancer treatment strategy (57, see

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Fig. 5.3. In vivo imaging of siRNA (against GFP) delivery and silencing in tumors. (a) In vivo MRI was performed on mice bearing bilateral 9L-GFP and 9L-RFP tumors before and 24 h after MN-NIRF-siGFP administration. Following injection of the probe, there was a significant drop in T2 relaxivity associated with the tumors (p = 0.0012 for 9L-GFP, p = 0.0049 for 9L-RFP). Note that T2 relaxation times of muscle tissue remained unchanged. (b) In vivo NIRF optical imaging of the same animals as in (a) produced a high-intensity NIRF signal associated with the tumors confirmed the delivery of the MNNIRF-siGFP probe to these tissues. (c) In vivo optical imaging of animals bearing bilateral 9L-GFP and 9L-RFP tumors 48 h after i.v. probe injection showing a dramatic decrease in 9L-GFP-associated fluorescence (p = 0.0083) and no change in 9L-RFP fluorescence. To generate GFP/RFP reconstructions, GFP and RFP images were acquired separately and then merged. (Reprinted by permission from Macmillan Publishers Ltd.: Nature Medicine, 13, 372–377 (2007)).

Chap. 14). MN-NIRF-siSurvivin nanoparticles injected intravenously in animals bearing subcutaneous human colorectal carcinoma xenografts accumulated in the tumors as evidenced by both MRI and NIRF imaging (Fig. 5.4). Time course injections in tumor-bearing animals resulted in significant silencing of the Birc5 gene and increased apoptosis compared to animals injected with nanoparticles devoid of siRNA or nanoparticles bearing mismatched control siRNA. Further development of this approach

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Fig. 5.4. Application of MN-NIRF-siSurvivin in a therapeutic tumor model. (a) In vivo MRI of mice bearing subcutaneous LS174T human colorectal adenocarcinoma (arrows) showed a significant drop in T2 relaxivity on postcontrast images of the tumors (p = 0.003) indicating probe delivery. (b) A high-intensity NIRF signal on in vivo optical images associated with the tumor (arrows) following injection of MN-NIRF-siSurvivin confirmed the delivery of the probe to this tissue (left, white light; middle, NIRF; right, color-coded overlay). (c) Quantitative RT-PCR analysis of survivin expression in LS174T tumors after injection with either MN-NIRF-siSurvivin, a mismatch control, or the parental MN nanoparticle. Survivin mRNA levels in tumors from MN-NIRF-siSurvivin treated animals were reduced by 97 ± 2%, compared to MN controls (p < 0.01) and 83 ± 2%, compared to mismatch controls (p < 0.01, data are representative of three separate experiments). (Reprinted by permission from Macmillan Publishers Ltd.: Nature Medicine, 13, 372–377 (2007)).

would include tumor-specific targeting and combination with conventional chemotherapeutics. Overall, the in vivo imaging of siRNA delivery and silencing is a fast developing field (see Chap. 4). Clearly, to develop siRNAs as therapeutic agents, monitoring of the success of in vivo delivery is critical. Noninvasive molecular imaging techniques have great potential for this purpose and can potentially provide the most direct, noninvasive, and longitudinal measurements of the in vivo delivery of siRNAs as well as their direct effect on gene expression and indirect influence on biological parameters associated with silencing.

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Chapter 6 Recent Advances in Magnetofection and Its Potential to Deliver siRNAs In Vitro Olga Mykhaylyk, Olivier Zelphati, Edelburga Hammerschmid, Martina Anton, Joseph Rosenecker, and Christian Plank Abstract This chapter describes how to design and conduct experiments to deliver siRNA to adherent mammalian cells in vitro by magnetic force–assisted transfection using self-assembled complexes of small interfering RNA (siRNA) and cationic lipids or polymers that are associated with magnetic nanoparticles. These magnetic complexes are targeted to the cell surface by the application of a magnetic gradient field. In this chapter, first we describe the synthesis of magnetic nanoparticles for magnetofection and the association of siRNA with the magnetic components of the transfection complex. Second, a simple protocol is described in order to evaluate magnetic responsiveness of the magnetic siRNA transfection complexes and estimate the complex loading with magnetic nanoparticles. Third, protocols are provided for the preparation of magnetic lipoplexes and polyplexes of siRNA, magnetofection, downregulation of gene expression, and the determination of cell viability. The addition of INF-7 peptide, a fusogenic peptide, to the magnetic transfection triplexes improved gene silencing in HeLa cells. The described protocols are also valuable for screening vector compositions and novel magnetic nanoparticle preparations to optimize siRNA transfection by magnetofection in every cell type. Key words: siRNA delivery in vitro, magnetic nanoparticles, magnetic transfection vectors.

1. Introduction Magnetofection can be defined as a method for nucleic acid delivery under the influence of a magnetic field acting on nucleic acid vectors that are associated with magnetic (nano)particles. First reports on associating a vector (viral and nonviral) with magnetic particles date back to the year 2000 (1,2). Several research

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groups have developed such methods independently (see, for example refs. 3–6). In the meantime, the term “magnetofection” is widely used in the scientific literature. We have mostly used electrostatic interactions to associate vectors with magnetic particles (7); other groups have used biotin–streptavidin or antigen– antibody interactions (5,6). We have demonstrated the suitability of magnetofection for magnetically localized gene delivery in vivo (8). However, major improvements are still required to make the method efficient enough to be widely used in in vivo applications. In contrast, magnetofection is well established and widely used for in vitro applications and has been shown to potentiate viral as well as nonviral nucleic acid delivery (8,9). Vectors associated with magnetic nanoparticles are added to cell culture supernatants. Cell culture plates are subsequently placed on magnetic plates which consist of an array of suitably positioned permanent magnets. Thus, the diffusion limitation to delivery is overcome, transfection/transduction is synchronized and greatly accelerated, and the vector dose requirement for efficient transfection/ transduction is considerably reduced. These features together constitute a substantial improvement of transfection/transduction efficiency. The mechanism of magnetofection is probably the same as for standard transfection/transduction concerning vector uptake into cells (10). Magnetic nanoparticles are co-internalized with vectors into cells and are biodegradable over long time periods. Importantly, magnetofection is applicable for both viral and nonviral vectors and among the latter for “large” nucleic acids (e.g., plasmids) or small constructs (e.g., synthetic siRNA and antisense oligonucleotides (11–13)) in a naked form or packaged as lipoplexes and polyplexes (7). The potential of magnetofection to efficiently deliver siRNA in vitro has been emphasized by several recent scientific publications. Gene silencing in primary cells are notoriously challenging due to the limited efficiency of most transfection reagents. Magnetofection has been shown to be a very effective way of transfecting siRNA in human primary endothelial cells derived from umbilical vein and from human cord blood (14,15). It contributed to demonstrating the critical role of a transcription factor in angiogenesis (15) and of a selective estrogen receptor modulator in atherosclerosis (14). Primary human gastric myofibroblasts are also difficult to transfect. Mc Caig and colleagues have successfully used magnetofection to deliver siRNA in these primary myofibroblast (15). Another striking example of magnetofection potential for siRNA delivery in primary cells has been reported by Uchida and colleagues. Indeed, successful siRNA delivery was achieved in primary rat embryonic DRG neurons (16). Efficiency of siRNA delivery mediated by magnetofection has also been reported for cell lines such as COS7 (monkey kidney) and 3Y1 (rat fibroblast), which allowed to demonstrate the

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novel role of phospho-b-catenin in microtubule growth (17). Other successful applications of siRNA delivery with magnetofection have been reported in Vero E6 cells (monkey kidney) (18) and in the human microvascular endothelial cell line-1 (HMEC1) (19). Magnetofection has also been reported to be effective for siRNA transfection in suspension cells such as MOLT-4 and Jurkat (human T cell leukemia), which permitted to show the implication of RCAS1 (a receptor-binding cancer antigen) in T cell apoptosis induced by HIV infection (20). The review by Bonetta et al. on “evaluating gene delivery methods” highlights the potential of magnetofection as a new tool to improve and target a broader range of cells and applications (21). In the same way, the benefits of using magnetofection, and particularly SilenceMag (magnetofection-based reagent optimized for siRNA delivery), to concentrate and promote efficient siRNA transfection were assessed (22). All that is required to practice magnetofection are suitable magnetic nano- or microparticles and appropriate magnetic devices. In the meantime, these tools along with standardized application protocols for various vector types and cell culture formats are commercially available (OZ Biosciences, Marseille, France, http://www.ozbiosciences.com; Chemicell, Berlin, Germany, http://www.chemicell.com). The commercially available magnet array for magnetofection produces high-gradient magnetic fields (70–250 mT and a field gradient of 50–130 T m−1) in the vicinity of the cells and sediments the full-applied vector dose on the cells to be transfected within minutes. The development of new magnetic nanoparticles is expected to lead to further improvements of the technique (12). Therefore, we focus here on a 96-well format screening procedure for magnetic nanoparticles to be used in nonviral magnetofection. We provide the protocols that comprise every step from magnetic nanoparticle synthesis and characterization to their use in siRNA magnetofection and instructions for data evaluation. We illustrate these protocols with magnetofection results obtained in adherent cancer cell lines stably transfected with GFP- and luciferase reporter genes. We describe magnetite nanoparticles, which differ in their coating material and which are efficient in siRNA delivery to adherent cells in vitro by magnetofection and can be associated with a nucleic acid, or with a nucleic acid and an enhancer or vector (viral or nonviral lipoplex or polyplex (23), which are stable enough to be stored over extended periods and are biocompatible enough for application in living cells). The compounds used here as components of the shell for the particles are known to be either useful in particle stabilization and/or in gene or drug delivery, and were found in our own experiments to be among the best for nucleic acid delivery (24).

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Subsequently, we describe how the binding of siRNA to the magnetic nanoparticles, either alone or in combination with a third agent that enhances transfection (known as an enhancer), can be characterized using radioactively labeled siRNA prepared according to the modified Terebesi procedure (25). This can be used to determine suitable ratios and mixing orders of magnetic nanoparticles, nucleic acids, and third components to choose formulations that are potentially useful for siRNA magnetofection. This protocol uses as enhancers either the commercially available transfection reagent Metafectene (4 µl µg−1 DNA) or the branched polyethylene imine (25 kDa) (PEI-25Br, nitrogen-toDNA phosphate ratio N/P = 10). Other enhancers (26) known to be efficient in the transfection of a particular cell line can, however, also be combined with magnetic nanoparticles to construct magnetic vectors. In principle, any nucleic acid can be delivered using magnetofection. For the screening purposes presented here, it is most useful to use reporter genes such as GFP and luciferase reporters. Both reporter gene systems allow rapid and sensitive result evaluation in cell lysates and even in living cells. In general, the luciferase reporter gene assay provides higher sensitivity and accuracy than the enhanced GFP (eGFP) assay. The advantage of using the GFP reporter gene is that the percentage of transfected cells can be easily determined. Vectors are prepared in serum- and supplement-free medium and transferred to the cells in triplicates in a volume of 50 µl per well. We recommend performing serial dilutions of a given vector composition such that the highest siRNA dose transferred to the cell culture plate is 200 ng per well. Accordingly, the highest vector dose has to be prepared at a final siRNA concentration of 4 µg ml−1 in the complex. According to our results, optimum nanomaterial-to-siRNA ratio for magnetic vectors comprising enhancers described in this protocol is between 0.5 and 1 in terms of ironto-siRNA wt/wt ratio. This has been tested in HeLa cells, H441 cells, and M1 cells. For magnetic duplexes comprising PEI-Mag2 nanomaterial and DNA, the optimal ratio is 1:1. Furthermore, we describe a simple method to evaluate the magnetic responsiveness of the magnetic siRNA transfection complexes and to estimate the complex loading with core-shell magnetic nanoparticles. This can be used to determine whether the magnetic properties of the vectors are sufficient to sediment the vector at the cell surface in the applied magnetic fields. Data on magnetic responsiveness of the complexes can be also used to estimate the potential to target the complexes magnetically when applied in vivo. Cell culture and transfection procedures to adherent cell lines are described. Also, preparation of magnetic vectors and the magnetofection procedure are described.

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siRNA transfection is known to provoke dose-dependent toxicity. Therefore, presenting results in terms of the absolute units of reporter gene expression normalized per weight of total protein in the examined cell lysate makes it possible to distinguish between gene downregulation and toxicity effects and is, therefore, especially important in siRNA transfection experiments. In fact, it has been shown that relatively low toxicity of the magnetic siRNA is one of the essential advantages of magnetofection (10). In this respect, the quantification of complex internalization, evaluation of siRNA transfection efficiency in living cells and in cell lysates, and the MTT-based cell viability test are described. INF-7 endosomolytic peptide derived from the N-terminal sequence of influenza virus hemagglutinin was shown to improve gene delivery with polyplexes (27–30). Recently the potential of the INF-7 peptide to improve the silencing efficiency of siRNA targeting the epidermal growth factor receptor and the K-ras oncogenes in complexes with Lipofectamine2000 was demonstrated in human epidermis carcinoma cells (31). We show an example INF7-mediated improvement of siRNA magnetofection in HeLa cells stably transduced with the GFP gene.

2. Materials 2.1. Synthesis of Magnetic Nanoparticles Suitable as Components of Magnetofection Complexes

1. Iron(II) chloride tetrahydrate (Sigma-Aldrich). 2. Iron(III) chloride hexahydrate (Sigma-Aldrich). 3. Argon 5.0 (Sauerstoffwerk Friedrichshafen GmbH). 4. 10% Hydroxylamine hydrochloride solution in water (see Note 1). 5. PEI-Mag2 precipitation/coating solution: 5 g polyethylenimine 25 kDa, branched (PEI-25Br; Sigma-Aldrich, cat. no. 40,872-7) plus 25 ml 28–30% ammonium hydroxide solution (Sigma-Aldrich, cat. no. 320145) plus 2.5 ml lithium 3-[2-(perfluoroalkyl) ethylthio]propionate (Zonyl FSA; Sigma-Aldrich) filled up with water to a total volume 100 ml, degassed with argon/helium. 6. PL-Mag1 precipitation/coating solution 1: 4 g Pluronic F-127 (Sigma-Aldrich) filled up with water to a total volume 50 ml, degassed with argon/helium. PL-Mag1 precipitation/coating solution 2: 30 ml 28–30% ammonium hydroxide solution plus 15 ml ammonium bis[2-(perfluoroalkyl) ethyl] phosphate solution (Zonyl FSE; Sigma-Aldrich, cat. no. 421391) filled up with water to a total volume 50 ml, degassed with argon/helium.

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7. PalD1-Mag1 precipitation/coating solution: 4 g palmitoyl dextran PalD1 (see Note 2) plus 30 ml 28–30% ammonium hydroxide solution filled up with water to a total volume 100 ml, degassed with argon/helium. 2.2. Determination of Magnetic Nanoparticle Concentration in Terms of Dry Weight and Iron Content

1. Ammonium acetate buffer for iron determination: Dissolve 25 g ammonium acetate (Sigma) in 10 ml water, add 70 ml glacial acetic acid, and adjust volume to 100 ml with water. 2. 10% Hydroxylamine hydrochloride (Sigma-Aldrich) in water. 3. 0.1% Phenanthroline solution: Dissolve 100 mg 1,10-phenanthroline monohydrate (Sigma, cat. no. 77500) in 100 ml water; add 2 drops concentrated hydrochloric acid (Fluka, cat. no. 84415). If necessary, warm up to obtain a clear solution. 4. Iron stock solution: Dissolve 392.8 mg ammonium iron(II) sulfate hexahydrate (Sigma, cat. no. F3754) in a mixture of 2 ml concentrated sulfuric acid and 10 ml water; add 0.05 N KMnO4 dropwise until a pink color persists and adjust the volume to 100 ml with water. 5. Standard iron solution (make fresh as required): Dilute iron stock solution 25 to 1 with water just before calibration measurements. 6. 0.05 N KMnO4 solution: Dissolve 0.790 g KMnO4 in 100 ml water.

2.3. Radiolabeling (Iodination) of siRNA

1. siRNA solution: 5 nmol (78.3 µg) GFP-22 siRNA (Qiagen) is reconstituted with 39.1 µl of the siRNA suspension buffer at 2 µg siRNA/µl and stored in aliquots at −20°C. 2. Sodium 125iodide in NaOH: Activity 1 mCi in 10 µl (Amersham Biosciences, cat. no. 1MS30). Caution: Radioactive material! Store at ambient temperature 15–20°C. Retains its iodination efficiency over 2 months storage. 3. 250 µM potassium iodide in water. Prepare on the day of DNA labeling from 25 mM potassium iodide. 4. 1 M sodium hydroxide in water. 5. 30 mM Thallium trichloride tetrahydrate (Sigma-Aldrich) solution in water. To obtain a clear solution, heat the tube to 70°C using a water bath. The solution is stable and can be stored at least for a year. 6. 1 M sodium sulfite in water. Prepare on the day of siRNA labeling. 1 M ammonium acetate buffer, pH 7. 7. Disposable Sephadex G25 PD-10 desalting columns (GE Healthcare).

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1. Suspension of the magnetic nanoparticles: Dilute stock suspension of magnetic nanoparticles in water at a concentration of 288 µg iron ml−1. Prepare just before the experiment. 2. Metafectene (Biontex Laboratories). 3. Polyethylenimine 25 kDa, branched (PEI-25Br; SigmaAldrich). 4.

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I-labeled siRNA solution: 4.8 µg ml−1 total siRNA (GFP22 siRNA, Qiagen) comprising 2 × 105 CPM ml−1 125I-labeled siRNA (from Sect. 3.3) in RPMI medium without supplements, or whatever solvent of interest.

1. NCI-H441 human pulmonary epithelial (H441) cells derived from papillary carcinoma of the lungs (ATCC). 2. NCI-H441 cells stably expressing eGFP (H441-GFP cells). 3. NCI-H441 cells stably expressing luciferase (H441-Luci cells). 4. HeLa human cervical epithelial adenocarcinoma cells (ATCC). 5. HeLa cells stably expressing eGFP (HeLa-GFP cells). 6. HeLa cells stably expressing luciferase (HeLa-Luci cells). 7. NCI-3T3 mouse fibroblasts stably expressing GFP (3T3-GFP cells). 8. H441 culture medium: Modified RPMI 1640 medium with 2 mM L-glutamine, 10 mM HEPES, 1 mM sodium pyruvate, 4.5 g l-1 glucose, 1.5 g l−1 sodium bicarbonate supplemented with 10% heat-inactivated FCS, 100 U ml−1 penicillin, 100 µg ml−1 streptomycin, and 2 mM L-glutamine. Split the cells 1 to 4–5 when they are about 80–90% confluent. 9. HeLa culture medium: DMEM supplemented with 2 mM L-glutamine, 1 mM sodium pyruvate supplemented with 10% heat-inactivated FCS, 100 U ml−1 penicillin, and 100 µg ml−1. Split the cells 1 to 5–7 when they are about 80–90% confluent. 10. 3T3 culture medium: DMEM medium supplemented with 10% heat-inactivated FCS, 100 U ml−1 penicillin, and 100 µg ml−1. 11. Trypsin/EDTA solution, 0.25%/0.02% (wt/vol) (Biochrom). 12. PBS-Dulbecco’s w\o Ca2+, Mg2+ solution (Biochrom).

2.6. Preparation of Magnetic Nanoparticle–siRNA Transfection Complexes

1. Magnetic nanoparticles are synthesized according to the procedure described in Sect. 3.1. The commercially available magnetic nanoparticles are suspended in water at 36 µg ml−1 just before use (the concentration refers to the iron content). This will result in a magnetic nanoparticle

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iron-to-siRNA ratio of 0.5:1 (wt/wt) according to the protocol described in Sect. 3.6 (see Note 3 and 4). 2. Metafectene as an enhancer: mix 5.8 µl Metafectene with 34.2 µl water in a tube for each transfection complex to be tested (resulting finally in a Metafectene-to-siRNA vol/vol ratio of 4:1 when used according to the protocol described in Sect. 3.6. 3. PEI as an enhancer: Prepare a solution of PEI-25Br in water at 45.36 µg ml−1 (resulting finally in an N/P ratio of 10 when the protocol described in Sect. 3.6 is used. 4. siRNA stock solution (100x): reconstitute 5 nmol siRNA, e.g., GFP-22 siRNA (Qiagen) or luciferase GL3 siRNA (Qiagen), at 480 µg siRNA ml−1 with 162.9 µl or 154.4 ml of the siRNA suspension buffer, respectively, and store in aliquots at −20°C. 5. siRNA solution: prepare by 1 to 100 dilution of the 100x siRNA stock solution with serum- and supplement-free medium (e.g., RPMI 1640). 2.7. Evaluation of Magnetic Responsiveness of the siRNAMagnetic Nanoparticle Complexes

1. Reagents and solutions are as in Sect. 2.6.

2.8. Magnetofection

1. Cells are plated 24 h prior to transfection according to Sect. 2.5.

2. 8 Ne-Fe-B permanent magnets 18.0 × 16.0 × 4.0 mm (IBS Magnets).

2. Magnetic transfection complexes and appropriate controls including complexes of nonsilencing siRNA(s) if necessary are prepared just before magnetofection (Sect. 3.6). 3. 96-Magnets magnetic plate (magnetic plate; OZ Biosciences). 2.9. Evaluation of Transfection Complex Association with Cells and Internalization into Cells

1. GFP-22 siRNA labeled with AlexaFluor 555 (GFP-22 siRNA, Qiagen). 2. FACS buffer: PBS supplemented with 1% FCS. 3. Hoechst 33342 stock solution: Hoechst 33342, trihydrochloride trihydrate (Invitrogen) 1 mg ml−1 water. Store in the dark at 4°C. 4. YOYO-1 iodide (491/509) stock: 1 mM solution in DMSO (Invitrogen). Store in aliquots in the dark at −20°C.

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I-labeled siRNA solution: 4.8 µg ml−1 total siRNA (GFP22 siRNA, Qiagen) comprising 1 × 106 CPM ml−1 125I-labeled siRNA (from Sect. 3.3) in RPMI medium (without supplements, or whatever solvent of interest). This should be prepared fresh for each experiment. Other reagents are as described in Sect. 2.6.

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2.11. Quantification of GFP Reporter Gene Downregulation in Cell Lysate and in Living Cells

1. Lysis buffer: 0.1% Triton X-100 in 250 mM Tris pH 7.8.

2.12. Quantification of Luciferase Reporter Gene Downregulation in Cell Lysates

1. Luciferin buffer: 35 µM d-luciferin (Roche Diagnostics), 60 mM DTT (Sigma-Aldrich, cat. no. D9779), 10 mM magnesium sulfate, 1 mM ATP, in 25 mM glycyl–glycine–NaOH buffer, pH 7.8.

2. GFP stock solution: 500 ng GFP per µl PBS. Store in small portions at −70°C. 3. BioRad protein assay reagent (BioRad). 4. BSA stock solution: 1.5 mg ml−1 BSA (Sigma) in PBS. Store at 4°C.

2. Luciferase standard stock: 0.1 mg luciferase (Roche Diagnostics) per ml and 1 mg BSA (Sigma) per ml in 0.5 M Tris– acetate buffer, pH 7.5. Store in aliquots at −70°C. 2.13. MTT-Based Test for Toxicity of the Transfection Complexes

2.14. Improvement of Reporter Gene Downregulation by Magnetic siRNA Vectors Modified with INF-7 Fusogenic Peptide

1. MTT solubilization solution: 10% Triton X-100 in 0.1 N hydrochloric acid in anhydrous isopropanol (solution can be stored at room temperature, 15–25°C). 2. MTT solution: 1 mg thiazolyl blue tetrazolium bromide (MTT; Sigma) per ml and 5 mg ml−1 glucose in PBS-Dulbecco’s solution (solution must be stored at −20°C). 1. INF-7 stock: 10 mg ml−1 in 20 mM HEPES, pH 7.4. Store in aliquots at −70°C. 2. INF-7 solution: 2.33 µg/10 µl in 20 mM HEPES, pH 7.4. Prepare before starting the experiment and keep at 4°C. 3. Other reagents are described in Sect. 2.6.

3. Methods 3.1. Synthesis of Magnetic Nanoparticles Suitable as Components of Magnetofection Complexes

1. Dissolve 0.05 mol (13.52 g) of iron(III) chloride hexahydrate and 0.025 mol (4.97 g) iron(II) chloride tetrahydrate in 300 ml water and filter using a 0.2 µm filtering flask or bottle-top filter (whenever possible, use fresh reagents); transfer the solution to a 500 ml round-bottom flask (make fresh as required). Remove dissolved oxygen by continuous argon or helium bubbling through the solution for ~10 min, (for PL-Mag1 nanomaterial add PL-Mag1 precipitation/ coating solution 1); cool the solution to 2–4°C and continue bubbling argon/helium through the solution. 2. To obtain a primary precipitate, rapidly add precipitation/ coating solution (for PL-Mag1 nanomaterial add PL-Mag1 precipitation/coating solution 2), heat the material to 90°C over a 15 min interval, and stir at this temperature for the

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next 120 min. Cool the mixture down to 25°C (no more inert gas bubbling is needed), and then incubate for 2 h with continuous stirring. 3. Sonicate the product for 10 min using a resonance frequency of ~20 kHz, 75 mW, 60 s sonication/30 s break interval. Dialyze against water over 2 d using Spectra/Por 6 50 kDa cut-off dialysis membrane to neutralize the suspension and to remove excess unbound stabilizer. Sterilize the suspension using 60Co gamma irradiation, dosage 25 kGy (32). The particle types described here can be stored for at least 1 year without losing their properties that are required for magnetofection (Table 6.1, see Note 5). 3.2. Determination of Magnetic Nanoparticle Concentration in Suspension in Terms of Dry Weight, Iron Content, and Iron Concentration per Dry Weight of Magnetic Nanoparticles

1. To determine the magnetic nanoparticles concentration in suspension in terms of iron content, take 20 µl aliquots of the magnetic nanoparticle suspension, add 200 µl concentrated hydrochloric acid and 50 µl water. Wait until the magnetic nanoparticles are completely dissolved, then adjust the volume to 5 ml with water. 2. Transfer 20 µl of the resultant solution to a microcentrifuge tube, add 20 µl concentrated hydrochloric acid, 20 µl hydroxylamine hydrochloride solution, 200 µl ammonium

Table 6.1 Characteristics of the magnetic nanoparticles synthesized according to the Sect. 3.1 (see Note 6) Nanoparticles Parameter

PEI-Mag2

PalD1-Mag1 PL-Mag1

Mean magnetite crystallite size (nm)

8.5

10.6

Mean hydrated particle diameter D (nm)

63 ± 36

55 ± 10

101 ± 20

Iron content (g Fe/g dry weight)

0.56

0.526

0.47

1.21 × 10−18

2.35 × 10−18

Average iron weight per par- 1.4 × 10−18 ticleP Fepart (g Fe/particle) Saturation magnetization of the “core” Ms (A·m2/ kgFe)

109

Effective magnetic moment of the insulated particle meff (A·m2)

5.5 × 10−20

4.3 × 10−20

9.1 × 10−20

ξ-Potential in water (mV)

+55.4 ± 1.6

−15.6 ± 1.6

−13.3 ± 1.6

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acetate buffer, 80 µl 1,10-phenanthroline solution, and 860 µl water. Mix well and allow to stand for 20 min. 3. Also prepare a blank sample by mixing 20 µl concentrated hydrochloric acid, 20 µl hydroxylamine hydrochloride solution, 200 µl ammonium acetate buffer, 80 µl 1,10-phenanthroline solution, and 880 µl water (see Note 6). 4. Measure the absorbance of the samples from Step 2 at 510 nm against the blank (Step 3) using a spectrophotometer, for example, a Beckman DU 640 spectrophotometer. 5. To construct a calibration curve for the determination of the iron concentration, add increasing amounts of iron standard solution to microcentrifuge tubes (e.g., 50, 70, and 90 up to 150 µl) and adjust the volume to 150 µl with water. Use 150 µl water instead of the iron solution to prepare a blank sample. To each tube, add 20 µl concentrated hydrochloric acid, 20 µl 10% hydroxylamine hydrochloride solution, 200 µl ammonium acetate buffer, 80 µl 0.1% 1,10-phenanthroline solution, and 730 µl water. Mix well and allow to stand for 20 min. Measure the absorbance at 510 nm against the blank. Plot the absorbance at 510 nm as a function of the iron concentration in the standard samples. Use linear regression as an approximation function for calculating the iron concentration in the magnetic nanoparticle samples. 6. To determine iron concentration per dry weight of magnetic nanoparticles, freeze-dry under high vacuum as follows: transfer 1 ml aliquots of magnetic nanoparticle suspensions into pre-weighed glass vials, freeze samples (at −80°C or in liquid nitrogen), and dry overnight under high vacuum (suitably using a lyophilizer). Weigh the vials again to calculate the dry weight. Add 1 ml concentrated hydrochloric acid. Wait until the magnetic nanoparticles are completely dissolved. Transfer 20 µl of the resultant solution to a microcentrifuge tube and determine iron content following the protocol described in Steps 3 and 5. Calculate iron concentration per dry weight of magnetic nanoparticles (see Note 7). Examples of the results are given in Table 6.1 (see Note 8). 3.3. Radiolabeling (Iodination) of siRNA (see Note 9)

1. In vial 1 (ideally a conical screw cap microcentrifuge tube) prepare a mixture of 15 µl siRNA solution (2 µg siRNA/µl) and 15 µl 0.1 M ammonium acetate buffer, pH 5. In vial 2, prepare a mixture of 15 µl 250 µM potassium iodide, 2 µl sodium 125I (0.2 mCi), and 10 µl 0.1 M sodium hydroxide. 2. Add 15 µl 30 mM thallium trichloride solution to vial 2 (see Note 10), quickly mix, and immediately transfer the content of vial 2 to vial 1, incubate the vial at 60°C for 45 min, then cool on ice.

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3. Add 7.5 µl 0.1 M sodium sulfite, then 30 µl 1 M ammonium acetate buffer, pH 7, incubate for 60 min at 60°C, then cool on ice. 4. During the incubation time of the previous step, equilibrate a Sephadex G25 PD10 desalting column with water according to the instructions of the manufacturer. Apply the reaction mixture to the column and let it penetrate the column bed. Position a rack with 20 aligned microcentrifuge tubes under the column for fraction collection. Add 2 × 5 ml water for elution and collect 11 drops each (= 400–500 µl) in the microcentrifuge tubes aligned in the rack. 5. Using a handheld radiation monitor, determine the early eluting product fractions with the highest radioactivity (it is expected that the fractions will be among fractions 6 to 9) (see Fig. 6.1a). 6. Transfer a 10 µl aliquot of the product fraction to a scintillation vial and determine the radioactivity (CPM) using a gamma counter (e.g., Wallac 1480 Wizard 3″ automatic gamma counter). In another aliquot of the product fraction from Step 4, determine the DNA concentration by measuring the absorbance D at 260 nm (see Fig. 6.1a) and using the following formula: siRNA concentration (µg/ml) = (D260) × (dilution factor) × (50 µg siRNA/ml). 3.4. Testing the siRNA Binding Capacity of the Magnetic Nanoparticles

This procedure can be accomplished within 2 h. 1. For use as a transfection enhancer, mix 20.2 µl Metafectene and 119.8 µl water, or prepare a solution of 45.4 µg PEI per ml water (N/P = 10 (see Note 11)). This should be prepared fresh before the experiment. In general, any other transfection reagent can be tested as an enhancer instead. 2. In a 96-well round bottom plate (Techno Plastic Products) add 20 µl of magnetic nanoparticle suspension (from Step 1) into well A1 (corresponding to 5.76 µg iron of magnetic nanoparticles). Add 10 µl water into each well from A2 to A6. 3. Transfer 10 µl from A1 into A2, mix, from A2 into A3, etc., down to A5. Discard excess 10 µl from A5. The A6 well is a reference. 4. Add 20 µl of enhancer dilution to each well from A1 to A6; mix well with a pipette. To measure DNA association with magnetic nanoparticles in the absence of an enhancer, add 20 µl water to each well. 5. Add 150 µl 125I-labeled siRNA solution (4.8 µg ml−1 in RPMI medium without supplements, or whatever solvent

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Fig. 6.1. Radiolabeling (iodination) of siRNA and testing siRNA association with magnetic nanoparticles in transfection complexes. (a) siRNA concentration ( ) and 125I-radioactivity in 10 µl aliquots () measured in the fractions after radiolabeled siRNA purification on Sephadex G25 PD-10 disposable columns. (b) siRNA associated and magnetically sedimented with PEI-Mag2 and PalD1-Mag1 magnetic nanoparticles in duplexes (duplex means magnetic nanoparticle plus siRNA only, without enhancer) and triplexes in the presence of PEI-25Br (nitrogen-to-siRNA phosphate ratio N/P = 10) or Metafectene (4 µl Mf/1 µg siRNA) plotted against magnetic nanoparticle concentration (in terms of iron-to-siRNA weight ratio, starting siRNA concentration of 4 µg ml−1). Plots assignment as shown in the figure. A range of iron-to-siRNA ratios (w/w) from 0.25 to 4, according to Sect. 3.4, has been examined with magnetic nanoparticles having a highly positive ξ-potential (PEI-Mag2) and with other particles having a negative ξ-potential (PalD1-Mag1). In the presence of both transfection reagents (4 µl Metafectene-Pro per µg DNA or PEI-25Br at N/P = 10), both magnetic particle types efficiently form triplexes, showing a potential suitability for magnetofection. Positively charged PEI-Mag2 particles also form duplexes with siRNA.

of interest) comprising 2 × 105 CPM ml−1 125I-labeled siRNA (from Sect. 3.3) to each well from A1 to A6; mix well with a pipette. 6. Incubate for 15 min to allow siRNA binding to magnetic nanoparticles.

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7. To sediment magnetic nanoparticles/siRNA transfection complexes, place the plate on the 96-Magnets magnetic plate (magnetic plate; OZ Biosciences) for 30 min. 8. Carefully sample 100 µl supernatant from each well using a pipette. Transfer each sample together with the pipette tip into scintillation vial. Take care to avoid disturbing magnetically sedimented complexes. 9. Measure the radioactivity (CPM) in every vial using the gamma counter. 10. Calculate magnetic sedimentation of the siRNA associated with magnetic nanoparticles (%; see Fig. 6.1b) as follows: Magnetically sedimented siRNA (%) = [1 − CPMsample /CPMref ] ¥ 100, where CPMref is the measured radioactivity from well A6 if the assay is carried out to the above protocol. 3.5. Cell Culture and Plating of Adherent Cells for Transfection

1. Culture H441 cells (human adenocarcinoma bronchial epithelial cells) at 37°C in a 5% CO2 atmosphere. Split cells at a ratio of 1:4 to 1:5 every 4–5 days before reaching 100% confluence. Seed plates 24 h before transfection (see Note 12). H441 cells are used as an example, but other cell lines could equally be used. 2. For plating, wash the cells with PBS, aspirate supernatant, and add 2 ml trypsin–EDTA (0.25%) solution per 75 cm2 cultivation flask. Shake gently so that the solution can cover the area of the cells, and then take out all of trypsin with a Pasteur pipette and incubate the flask at 37°C for 2–3 min. Observe the cells under microscope and when the cells are detached immediately add 10 ml H441 culture medium to arrest trypsin action. 3. Count the cells using a microscope counting chamber (hemocytometer) and resuspend in H441 culture medium at a density of 1.67 × 105 cells per ml, before transferring to a reagent reservoir. 4. Transfer 150 µl of the cell suspension per well to the 96-well flat bottom plate (Techno Plastic Products, cat. no. 92096) or to a clear bottom black-walled plate, 96-well (Greiner BioOne) using a multichannel pipette (see Note 13). 25,000 cells per well provide a confluence of 50% before magnetofection 24 h later. 5. Store the plate in a cell culture incubator at 37°C in a 5% CO2 atmosphere until transfection, usually 24 h later. The cells should be approximately 50% confluent at the time of transfection. 6. Adherent cells that divide more rapidly than H441 cells (NIH-3T3 or HeLa cells) would be plated at a density of 5,000–10,000 cells per well.

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1. To test the transfection complexes, add 20 µl of a suspension of magnetic nanoparticles to be tested (from Step 1 in Sect. 2.6) into wells A4, A7, and A10 and E1, E4, E7, and E10, respectively, of flat-bottom 96-well plates. 2. Add 40 µl each of the enhancer solution (from Step 2 or 3 in Sect. 2.6) to the same wells and mix using a pipette. 3. Add 300 µl each of siRNA solution from Step 4 in Sect. 2.6 (4.8 µg ml−1 in serum- and supplement-free medium (e.g., RPMI 1640), which delivers 1.44 µg siRNA per well) to the same wells and mix well using a pipette. This would result in a final volume of 360 µl in wells A4, A7, and A10 (see Note 14). 4. For the untransfected control setup, add 300 µl serum- and supplement-free medium and 60 µl water to well A1. For other controls and references (e.g., magnetic nanoparticle–siRNA duplexes without enhancer, enhancer–siRNA complexes without magnetic nanoparticles), substitute the omitted component(s) with medium and/or water, respectively. 5. Incubate for 15 min at room temperature. 6. During the incubation time, fill the remaining wells of columns 1, 4, 7, and 10 with 180 µl each of serum- and supplement-free medium (RPMI 1640). 7. Prepare a 1:1 dilution series when the 15 min incubation time has expired as follows: Transfer 180 µl each from A1, A4, A7, and A10 to B1, B4, B7 and B10, respectively, using a multichannel pipette, mix, transfer 180 µl from the respective wells in row B to row C and so on down to row D. And 180 µl each from E1, E4, E7, and E10 to F1, F4, F7, and F10, respectively, using a multichannel pipette, mix, transfer 180 µl from the respective wells in row F to row G and so on down to row H (see Note 15). The characteristics for selected siRNA transfection complexes prepared according to the protocol described in Sect. 3.6 are given in Table 6.2.

3.7. Evaluation of Magnetic Responsiveness of the siRNA– Magnetic Nanoparticle Complexes: Complex “Loading” with Magnetic Nanoparticles

To evaluate magnetically induced velocity (magnetic responsiveness) of a magnetic siRNA complex in a gradient magnetic field and estimate complex loading with magnetic nanomaterial: 1. Prepare magnetic nanoparticle–siRNA transfection complexes: add 60 µl of a suspension of magnetic nanoparticles into a tube; add 120 µl of the enhancer solution (from Step 2 or 3 in Sect. 2.6) to the same tube and mix using a pipette; add 900 µl of the siRNA solution from Step 4 in Sect. 2.6 and mix well using a pipette. This results in a final volume of 1080 µl. For magnetic nanoparticle–siRNA duplexes with-

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Table 6.2 Characteristics of selected siRNA transfection complexes Iron-tosiRNA w/w z-Potential ratio (mV)

Mean hydrated diameter D (nm)

Efficient velocity in magnetic fieldsa uz (µm/s)

Average magnetic moment of the complex Ma (10−16 A·m2)

PEI/siRNA

–

+15.2 ± 1.8

413 ± 190

–

Mf/siRNA

–

+36.1 ± 9.7

283 ± 133

–

PEI-Mag2/ siRNA

1:1

−14.0 ± 0.8

685 ± 242

1.2

17.2

20483

PEI-Mag2/ siRNA

2:1

−10.1 ± 1.2

736

1.96

30.2

35946

PEI-Mag2/ PEI/siRNA

0.5:1

+2.0 ± 4

394 ± 70

1.19

9.5

11290

PalD1-Mag1/ PEI/siRNA

0.5:1

+7.2 ± 1.5

370 ± 115

1.49

11.6

15895

PEI-Mag2/ Mf/siRNA

0.5:1

+36.4 ± 3.8

210 ± 86

0.86

3.8

4500

PalD1-Mag1/ Mf /siRNA

0.5:1

+12 ± 6.3

326 ± 175

0.72

30.2

12181

Complex

Number of magnetic particles in a complexN=M/ meff

Duplexes

Triplexes

For a magnetic field configuration as shown in Fig. 6.2.

out enhancer or enhancer–siRNA complexes without magnetic nanoparticles, substitute the omitted component(s) with medium and/or water, respectively. Incubate for ~15 min at RT. Use half of the volume to measure the mean hydrated diameter D of the complex by photon correlation spectroscopy using, for example, a Malvern Zetasizer 3000 (UK). 2. Put 500 µl of the complex suspension into an optical cuvette. Position two sets of 4 quadrangular Ne-Fe-B permanent magnets (18.0 × 16.0 × 4.0 mm) cantered beside a measuring window of the optical cuvette in a spectrophotometer (for example, Beckman DU 640 Spectrophotometer); put the cuvette with the suspension into the sample holder, and immediately start to measure the time course of the turbidity

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(optical density) at 360 nm in a “kinetic mode.” The set-up for the measurements is shown in Fig. 6.2a. The configuration of the magnetic field in the measuring window is shown in Fig. 6.2b. Examples of the experimental results for the kinetics of the magnetic sedimentation (responsiveness) for the selected complexes are given in Fig. 6.2c. 3. For evaluation of the efficient velocity υz of the complexes under a gradient magnetic field and further calculation of the average magnetic moment M of the magnetic complex(s) and estimation of the number of magnetic nanoparticles N associated with the complex see Note 16. Examples of results are shown in Table 6.2. 3.8. Magnetofection

Timing: ~30–40 min plus 48–72 h to allow reporter gene downregulation. Magnetofection should be carried out under sterile conditions. 1. Check the plates prepared for transfection according to Sect. 3.5 under the microscope for cell state and confluence. Cell confluence of ~40–50% before transfection is preferable for H441 cells. 2. Cells that divide more rapidly than H441 cells (NIH-3T3 or HeLa cells) can be transfected at lower confluence of ~30– 40%. Aspirate the medium from the wells, and add 150 µl fresh cultivation medium per well. 3. Transfer 50 µl each of the transfection complexes prepared in Sect. 3.6 into the culture plates with the seeded cells as follows: Using a multichannel pipette, mix the dilutions of transfection complex prepared in column 1 of the complex preparation plate (from Sect. 3.6) by pipetting up and down, then transfer 50 µl to the wells of columns 1, 2, and 3 (to test each composition and dilution of transfection complex in triplicate) of the cell culture plate (from Step 1). Transfer 50 µl from each well of column 4 of the complex preparation plate to columns 4, 5, and 6 of the cell culture plate. Transfer 50 µl from each well of column 7 of the complex preparation plate to columns 7, 8, and 9 of the cell culture plate. Transfer 50 µl from each well of column 10 of the complex preparation plate to columns 10, 11, and 12 of the cell culture plate. This results in delivery of 200, 100, 50, and 25 ng siRNA per well in rows A (E), B (F), C (G) and D (H), respectively. 4. Place the cell culture plate on a 96-magnet magnetic plate for 15–30 min to create at the cell layer location a permanent magnetic field with a field strength and gradient of 70–250 mT and 50–130 T m−1, respectively.

Fig. 6.2. Evaluation of magnetic responsiveness of the magnetic siRNA transfection complexes in the applied magnetic fields. (a) Setup for measurements of the magnetic responsiveness of the complexes: Gradient permanent magnetic field is created by two sets of four quadrangular Ne-Fe-B permanent magnets 17 × 4 mm positioned at a distance of 20 mm

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5. Remove the magnetic plates after 20–30 min exposure of the cells to the magnetic field (see Note 17) and incubate the plate containing the transfected cells in a cell culture incubator at 37°C in a 5% CO2 atmosphere until results are evaluated (usually 48–72 h post-transfection). 3.9. Evaluation of the Transfection Complex Association with Cells and Internalization into Cells

1. To evaluate siRNA transfection efficiency and transfection complex association with cells and internalization into cells, prepare the transfection complexes with GFP-siRNA labeled with AlexaFluor 555 according to Sect. 3.6 and perform transfection of the cells according to Sect. 3.8. 2. After incubation at 37°C in a 5% CO2 atmosphere, examine the plate using a fluorescence microscope at 490/509 nm (green fluorescence) to visualize the cells expressing the eGFP reporter gene and at 510/650 nm (red fluorescence) to visualize localization of the siRNA-Alexa555 complexes (see Fig. 6.3a and b). 3. To allow visualization of internalized complexes with respect to the cell nuclei, add 1 µl per well of the cell-permeable nuclear counterstain Hoechst 33342 (1 mg ml−1 stock solution in water). Incubate for 15–20 min. Examples of the results are shown in Fig. 6.3. 4. To analyze cell/association and internalization using a FACS Vantage microflow cytometer, wash the adherent cells prepared in Step 1 with 150 µl PBS per well; aspirate the supernatant with a Pasteur pipette; add 10 µl Trypsin–EDTA (0.25%) solution per well, and incubate the flask at 37°C for 2–3 min. 5. Observe the cells under a microscope. When the cells are detached, immediately add 200 µl cell culture medium to arrest trypsin activity. 6. Combine cells from triplicate wells of cell culture plates in a fluorescence-activated cell sorting (FACS) tube. 7. Centrifuge at 300g (1,200 r.p.m. on a Heraeus Megafuge 2.0) for 5 min; remove supernatants carefully, and add 1 ml PBS supplemented with 1% FCS (FACS buffer).

Fig. 6.2. (continued) and centered around the measuring window of an optical cuvette in a Beckman DU 640 Spectrophotometer. (b) The magnetic field and the gradient of the field applied: The magnetic field is rather uniform in a measuring window Z = ±2 mm, X(Y) = ±5 mm in the X–Y plane parallel to the surface of the magnets and measuring light beam with an average magnetic field 〈Bz〉value in the direction of the particles moving perpendicular to the measuring beam of 0.213±0.017 T with an uniform magnetic field gradient ∂B/∂z in direction of the complexes movement of 4 ± 2 T/m. (c) Time course of the normalized turbidity of the magnetic complex suspensions (optical density at 360 nm normalized to the initial one D/D0) upon application of the gradient permanent magnetic field versus time for siRNA duplexes with PEI-Mag2 nanoparticles PEI-Mag2/siRNA at iron-to-siRNA wt/wt ratio of 1:1 and 2:1, and siRNA triplexes with PEI-Mag2 and PalD1-Mag1 nanoparticles and PEI-25Br (PEI) or Metafectene (Mf) as enhancers at iron-to-siRNA wt/wt ratio of 0.5:1 prepared as described in Sect. 3.6. Plots assignment as shown in the figure.

Fig. 6.3. Enhanced GFP (eGFP) expression and cell association/internalization of transfection complexes comprising magnetic nanoparticles with human cervical carcinoma cells and human pulmonary epithelial cells stably transfected with eGFP protein (HeLa-GFP and H441-GFP cells) detected by microscopy. HeLa-GFP and H441-GFP cells were incubated for 30 min at the magnetic plate with PEI-Mag2/PEI/GFP-siRNA-Alexa555 triplexes at siRNA concentration of 100 ng/10,000 cells/0.33 cm2; iron-to-siRNA wt/wt ratio of 0.5, PEI/siRNA ratio of N/P = 10, and observed after 1.5 h and/or 48 h with a fluorescence microscope. Images were obtained at original magnification of 40. Bar = 50 µm. Hoechst 33342 was used as a nuclear counterstain. The pictures show fluorescence images taken at 490/509 nm (green fluorescence) for eGFP fluorescence, 510/650 nm (red fluorescence) for GFP-siRNA-Alexa555, and at 350/461 nm (blue fluorescence) for Hoechst 33342 nuclear staining, or overlays thereof. Fluorescence microscopy data prove the association of the magnetic transfection complexes with a majority of the cells already 1.5 h post-transfection and are indicative of internalization into cells. 48 h post-transfection fluorescently labeled siRNA triplexes comprising magnetic nanoparticles are localized predominantly around the nuclei.

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8. Centrifuge again at 300g for 5 min; discard supernatants carefully; resuspend the cells in 0.5 ml FACS buffer. 9. To stain the siRNA complexes that are associated with cells but are not internalized into cells, add 1 µl of the cellimpermeable intercalating nucleic acid stain YOYO-1 iodide (1 mM in DMSO); incubate for 10 min. 10. Analyze the cells on a flow cytometer: excite fluorescence with an argon laser >488 nm, and detect eGFP or YOYO-1 fluorescence using a 530/30-nm bandpass filter and Alexa555 fluorescence using a 575/26-nm bandpass filter. Analyze a minimum total of 10,000 events per sample. 11. The percentages of cells with internalized or associated Alexa555-labeled transfection siRNA complexes are determined as a percentage of gated fluorescent events detected with a 575/26-nm bandpass filter (FL2) using untreated cells as a reference (see Fig. 6.4a). The percentage of cells either expressing eGFP or associated with YOYO-1-labeled transfection complexes is determined as a percentage of gated fluorescent events detected with a 530/30-nm bandpass filter (FL1) using untreated cells as a reference. Examples of the results are shown in Fig. 6.4b. 3.10. Quantification of Transfection Complex Internalization into Cells Using Radioactively Labeled siRNA

1. To quantify transfection complex internalization into cells, prepare the transfection complexes with 125I-labeled siRNA solution from Sect. 2.10 according to the protocols described in Sect. 3.6 and perform transfection of the cells according to Sect. 3.8. Reserve 50 µl each of the transfection complexes as a reference. 2. After incubation at 37°C in a 5% CO2 atmosphere, wash the cells with 150 µl PBS per well at different time points posttransfection; aspirate supernatant with a Pasteur pipette. To remove extracellularly bound complexes, add 100 µl per well heparin solution containing 75 mM sodium azide to inhibit endocytosis (33). 3. After incubation at 37°C in a 5% CO2 atmosphere for 30 min, wash the cells with 150 µl PBS per well; aspirate supernatant; add 10 µl Trypsin–EDTA (0.25%) solution per well, and incubate the flask at 37°C for 2–3 min. 4. Observe the cells under a microscope. When the cells are completely detached, add 200 µl cell culture medium. 5. Carefully collect the cell suspension from each well using a pipette. Transfer each sample together with the pipette tip into a scintillation vial. Measure the radioactivity (CPM) in every vial using the gamma counter. 6. Calculate the siRNA associated with magnetic nanoparticles as follows: Internalized siRNA (%) = [CPMsample/CPMref] × 100, where CPMref is the measured radioactivity from the reference

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Fig. 6.4. Vector association/uptake in HeLa human cervical epithelial adenocarcinoma cells stably expressing eGFP (HeLa-GFP cells) and H441 human lung epithelial cells characterized by flow cytometry. HeLa-GFP cells and H441 cells were transfected in a 96-well plate as described in Sect. 3.9. 48 h post-transfection the cells were trypsinized, washed, and resuspended in 1% FCS in PBS. Vector cell/association/internalization was analyzed using a FACS Vantage microflow cytometer. (a) Density plots of untransfected cells (untx); cells transfected with siRNA-Alexa555 (siRNA*) alone without magnetic nanoparticles or enhancer (naked siRNA*); polyplexes of PEI-Br25 with siRNA-Alexa555 (PEI/siRNA*); magnetic triplexes comprising PEI-Mag2 magnetic nanoparticles, PEI-Br25, and siRNA-Alexa555 (PEI-Mag2/PEI/siRNA*) for vector uptake analysis. Transfected H441 cells were additionally incubated with YOYO-1 to stain the siRNA transfection complexes associated with the cells but not internalized into cells. The siRNA dose was 100 ng per well in the examples in Fig. 6.4a. The numbers in squares indicate the percentages of gated cells with untreated cells as a reference. (b) Percentage of Alexa555 positive HeLa and H441 cells associated with siRNA* in dependence of the siRNA concentration in a well (black lines) and percentage of H441 cells that have internalized complexes (red lines) for triplexes of PEI-Mag2 nanoparticles with Metafectene (Mf) or PEI vs. nonmagnetic duplexes of siRNA* and naked siRNA*. Plots assignment as shown in the figure. The results given here clearly show that more than 80% of the HeLa-GFP and H441 cells are associated with transfection complexes, at the same time association with cells and internalization of naked siRNA is significantly lower compared to both magnetic and nonmagnetic transfection complexes. There is no considerable difference between the percentages of cells associated with/or internalized magnetic and nonmagnetic transfection complexes.

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Fig. 6.5. Vector internalization in HeLa human cervical epithelial adenocarcinoma cells and H441 human lung epithelial cells. HeLa-GFP cells and H441 cells were transfected in a 96-well plate using 125I-labeled siRNA complexes. The siRNA dose was 100 ng per well. At time points 0.5 h, 1 h, 3 h, and 24 h post-transfection the cells were incubated with heparin solution in the presence of sodium azide to remove extracellularly bound complexes, washed, trypsinized, and collected. Cell-associated radioactivity was measured with a gamma counter. The applied dose of the radioactively labeled siRNA complexes was used as a reference. The results were recalculated in terms of the siRNA molecules internalized per seeded cell. siRNA alone without magnetic nanoparticles or enhancer (naked siRNA, squares); Metafectene lipoplexes (open circles); polyplexes with PEI (open triangles); triplexes comprising Metafectene, siRNA, and PEI-Mag2 (black circles); triplexes comprising PEI, siRNA, and PEI-Mag2 (black triangles). In contrast to the FACS data on the percentage of the cells that have internalized the siRNA complexes (shown in Fig. 6.4b for H441 cells), the results given here clearly show that cell internalization of naked siRNA is negligible; polyplexes and lipoplexes of the siRNA are internalized better compared to naked siRNA. Magnetofection results in significantly higher internalization levels of the siRNA compared to lipo- or polyfection with the same vector type.

sample. In the example shown in Fig. 6.5 the results are recalculated in terms of the siRNA molecules internalized per seeded cell. 3.11. Quantification of GFP Reporter Gene Downregulation in Living Cells and in Cell Lysate

1. To prepare cell lysates from adherent cells, wash transfected adherent cells (from Sect. 3.8) with 150 µl per well PBS using a multichannel pipette. Add 100 µl lysis buffer per well. Incubate for 10 min at RT, then place the culture plate on ice. 2. To quantify GFP expression in cell lysates, transfer 50 µl cell lysate from each well into a black 96-well plate with a transparent bottom (e.g., clear bottom black-walled plate, Greiner). Add 100 µl PBS per well and mix with the pipette. Measure the fluorescence intensity (485/535 nm, 1.0 s) using a microplate fluorescence reader, for example, a Wallac 1420 Multilabel counter. As blanks, measure wells with lysates of nontransfected cells. 3. To construct a calibration curve for the determination of the amount of GFP in transfected cell samples, in well A1

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of a 96-well clear bottom black-walled plate, add 3 µl GFP stock solution to 147 µl lysis buffer, and mix well. Add 50 µl lysis buffer each to wells A2–A12. Transfer 100 µl from A1 to A2, mix well, transfer 100 µl from A2 to A3, mix and so on down to A11. Discard the surplus 100 µl from well A11. A12 is left as a blank. Add 100 µl PBS to each well of row A and mix well. Using a microplate fluorescence reader (e.g., Wallac 1420 Multilabel counter), measure the fluorescence intensity of GFP (excitation 485 nm, emission 535 nm, measuring time 1 s per well). Plot the measured fluorescence intensity as a function of GFP content per well. Use linear regression to derive a calibration function from which the GFP content in the samples can be calculated. Examples of the results are shown in Fig. 6.6a. 4. Use a calibration curve, constructed as described, to calculate the amount of GFP in the transfected cell samples (see Note 18). 5. To allow the results of the reporter gene expression assays to be presented as weight GFP (or luciferase) per weight unit total protein, the total protein content of the samples can be determined as follows: first, add 150 µl water to each well in a flat-bottom 96-well plate. Using a multichannel pipette, transfer 10 µl each of the cell lysates (from Step 1 or from Step 1 in Sect.3.12) into the corresponding wells of the protein assay plate. Add 40 µl BioRad protein assay reagent to each well; mix carefully using a plate shaker or a multichannel pipette. Measure the absorbance at 590 nm using a microplate reader (e.g., a Wallac 1420 Multi-label counter; measuring time set to 0.1 s). 6. To construct a calibration curve for the determination of the amount of total protein in the transfected cell sample, add 25 µl lysis buffer per well in one row (e.g., row A) of a flat-bottom 96-well plate. Add 50 µl BSA stock solution to well 1 (e.g., A1). Mix well using a pipette. Transfer 50 µl from well 1 to well 2, mix, transfer 50 ml from well 2 to well 3, and so on down to well 11. Well 12 is left as a blank. Add 150 µl water per well in another row (e.g., row B). Transfer 10 µl from row A to row B. Add 40 µl BioRad reagent to each well and mix carefully using a plate shaker or a multichannel pipette. Measure the absorbance at 590 nm (or 570 nm) using a microplate reader (e.g., a Wallac 1420 Multilabel counter; measuring time set to 0.1 s). Plot the measured absorbance versus the protein content per well. Use linear regression to derive a calibration function from which the protein content in the samples can be calculated. 7. Calculate the total protein content per 10 µl cell lysates for every sample using the calibration curve (from Step 6).

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8. Calculate weight GFP per weight unit total protein (see Note 19); normalize results to the reference data determined for untransfected cells. The results can be plotted against time post-transfection to evaluate the time course of the silencing effect and to define the optimum exposition for screening experiments (see Fig. 6.6a and b) or against the siRNA concentration or dose per well in order to get a dose–response curve (examples of the results are given in Figs. 6.6c and 6.7a).

Fig. 6.6. Enhanced GFP (eGFP) reporter gene expression analysis in cell lysates and in living cells. (a) GFP stably transfected NIH-3T3 cells (NIH-3T3-GFP cells) were seeded in a 96-well plate and 24 h later transfected with 200 µl transfection volume of the magnetic complexes prepared with 0.5 µl of SilenceMag (OZ Biosciences) and 1 nM, 5 nM, or 10 nM siRNA (targeting GFP). GFP expression was monitored in cell lysates in function of post-transfection incubation time. Results show percentage of reporter gene expression. Untreated cells were used as a reference. (b) Shows a calibration curve for eGFP. (c) GFP stably transfected H441 human lung epithelial cells (H441-GFP cells) were seeded in a 96-well plate and 24 h later transfected with a 200 µl transfection volume of the magnetic triplexes PalD1-Mag1/PEI/siRNA (ironto-siRNA ratio of 0.5 to 1, PEI/siRNA ratio of N/P = 10). GFP expression was monitored in living cells (fluorescence intensity, IFL) at different time points post-transfection, and finally in cell lysate in function of the siRNA dose (concentration). Results show percentage of reporter gene expression. Untreated cells were used as a reference. (d) Shows the correlation between fluorescence intensity IFL registered in cell lysates against IFL registered for the same sample in wells with living cells for a set of samples. The results show minimum GFP expression (maximum down-regulation effect) 60–96 h post-transfection. Thus, in screening experiments for siRNA delivery, magnetic complex analysis of GFP expression can be performed after 60–90 (usually after 72) h post-transfection. The example given in (c) shows that GFP-expression monitoring in living cells provide realistic semiquantitative information on transfection efficiency.

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Fig. 6.7. Magnetofection versus lipofection and polyfection efficiency in HeLa-GFP cells. (a) GFP stably transfected HeLa cells (HeLa-GFP cells) were seeded in a 96-well plate and 24 h later transfected with a 200 µl transfection volume of the magnetic anti-GFP–siRNA complexes prepared with 0.5 µl of SilenceMag (OZ Biosciences) at different concentrations of siRNA or PEI/siRNA and Mf/siRNA poly- and lipoplexes, or magnetic duplexes PEI-Mag2/siRNA (Iron-to-siRNA ratio of 1) or magnetic triplexes PEI-Mag2/PEI/siRNA, PL-Mag1/Mf/siRNA, and PalD1/Mf/siRNA (iron-to-siRNA ratio of 0.5 to 1) (Mfto-siRNA vol/wt ratio of 4, PEI-to-siRNA ratio N/P = 10). GFP expression was monitored 72 h post-transfection. Results show percentages of reporter gene inhibition. Plots assignments as shown in the figure. (b) GFP expression was monitored 72 h post-transfection by fluorescence microscopy in HeLa-GFP cells transfected with SilenceMag as shown in (a) at 1, 5, or 10 nM siRNA. The results show that magnetofection results in significantly lower expression levels of the GFP (i.e., more efficient target gene downregulation) compared to lipo- or polyfection with the same vector type. Efficiency of the PEI.Mag2/PEI/siRNA complexes is comparable with that of a magnetofection-based formulation of OZ Biosciences called SilenceMag. Magnetic duplexes PEI-Mag2/siRNA (at iron-to-siRNA ratio of 1) deliver siRNA rather efficiently, but less efficient compared to the PEI-Mag2/PEI/siRNA magnetic triplexes formulated at an iron-to-siRNA ratio of 0.5:1.

9. To evaluate the time course of the silencing effect, measurements of the GFP expression in living cells can be performed as follows (see Note 20): aspirate cell culture medium, wash the cells twice with 150 µl per well PBS, and measure the fluorescence intensity (485/535 nm, 1.0 s) using a microplate fluorescence reader. As blanks, measure wells with nontransfected cells. Change PBS for the complete cell growth medium and continue incubation. At the end of

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observation after final measurements in living cells, perform cell lysis according to Step 1, and determine GFP concentration in cell lysates as described in Steps 2, 3, and 8. 10. To calculate absolute values of GFP concentration from measurements in living cells, plot fluorescence intensity IFL registered in cell lysates against IFL registered for the same samples in wells with living cells for a set of samples. Examples of the results are shown in Fig. 6.6d. Use linear regression to derive a function for calculating IFL in lysate. The function in combination with a calibration curve for lysates (shown in Fig. 6.6b) allows one to estimate GFP content in living cells. Examples of the results are shown in Fig. 6.6c. 3.12. Quantification of Luciferase Reporter Gene Downregulation in Cell Lysates

1. Prepare cell lysates from adherent cells as described in Sect. 3.11, Step 1. 2. To quantify luciferase reporter gene expression in cell lysates, transfer 50 µl cell lysate from each well into a 96-well black flat-bottom microplate. Add 100 µl luciferase buffer per well, optionally mix with a pipette. Measure the chemiluminescence intensity (count time 0.20 min with background correction) using a luminometer such as, for example, a microplate scintillation and luminescence counter (Canberra Packard) or a Wallac Victor 2 Multilabel Counter (Perkin Elmer). 3. To construct a calibration curve to determine the amount of luciferase in transfected cell samples, add 50 µl lysis buffer per well to columns 1 and 3 of a black 96-well plate and 40 µl lysis buffer per well in columns 2 and 4. To well A1, add 30 µl lysis buffer and 20 µl luciferase standard stock (0.1 mg luciferase per ml and 1 mg BSA per ml in 0.5 M Tris–acetate buffer, pH 7.5). Pipette 50 µl from A1 to B1, mix well, then from B1 to C1, etc., down to H1. From H1, continue the dilution series by transferring 50 µl to A3; continue in column 3 down to G3. H3 is a blank. Pipette 10 µl each from column 3 to 4, and from column 1 to 2. Add 100 µl luciferase buffer each to the wells of columns 2 and 4. Measure the chemiluminescence intensity as described above. Plot the logarithm of luciferase content in the dilution series as a function of the logarithm of measured luminescence intensity (light units). Use an approximation function (usually linear regression in this concentration range) for calculating the amount of luciferase in the transfected cell samples. An example of the results is given in Fig. 6.8. 4. Use a calibration curve, constructed as described in Step 3, to calculate the amount of luciferase in the transfected cell samples.

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5. To allow the results of the luciferase expression assays to be presented as weight luciferase per weight unit total protein, determine total protein content in the lysate as described in Sect. 3.11, Steps 3–5. 6. Calculate weight luciferase per weight unit total protein; normalize results to the reference data determined for untransfected cells. The results can be plotted against time post-transfection or against the siRNA concentration or dose per well in order to get a dose–response curve. Examples of the results are given in Fig. 6.8a and c. 3.13. MTT-Based Test for Toxicity of the Transfection Complexes

1. Wash transfected adherent cells with 150 µl PBS per well using a multichannel pipette and discard wash solutions. 2. Add 100 µl per well of MTT solution, and incubate in a cell culture incubator for 1.5–2 h. 3. Observe the accumulation of the insoluble violet formazan crystals. When necessary, continue the incubation to obtain an optical density of ~0.3–1.0 at 550–590 nm for untreated cells (as a reference) after product solubilization. 4. Add 100 µl MTT solubilization solution to dissolve formazan. 5. Seal the plate with parafilm or an adhesive film to avoid liquid evaporation, and incubate overnight at RT until complete dissolution of the formazan crystals. 6. Measure the optical density D of the MTT–formazan solution after solubilization in the range of the wide absorption spectrum maximum (550–590 nm), for example, at 590 nm, using a microplate reader (e.g., Wallac Multi-label Counter; measuring time 0.1 s). Use untransfected cells as a reference. Register the absorbance for one or several wells with a mixture of 100 µl MTT solution and 100 ml solubilization solution as a blank. 7. Cell viability in terms of cell respiration activity (34,35) normalized to the reference data (%) is expressed as: Cell viability (%) = (Dsample− Dblank)/(Dref − Dblank)·100%, here

Fig. 6.8. (continued) for luciferase. (c) Luciferase stably transfected H441 cells (H441-Luci-cells) were seeded in a 96-well plate and 24 h later transfected with naked siRNA, PEI/siRNA, and Mf/siRNA poly- and lipoplexes, or magnetic duplexes PEI-Mag2/siRNA (iron-to-siRNA ratio of 1) or magnetic triplexes PEI-Mag2/PEI/siRNA, PL-Mag1/Mf/siRNA, and PalD1/Mf/ siRNA. Luciferase expression in lysate and cell viability in terms of respiration activity was measured 48 h post-transfection as a percentage of a reference (untransfected cells). In (a) and (c) PEI/siRNA ratio of N/P = 10. Mf-to-siRNA vol/wt ratio of 4. Iron-to-siRNA ratio for magnetic triplexes 0.5:1. The results in (a) show minimum luciferase expression (maximum downregulation effect) in HeLa-Luci cells between 24 and 48 h post-transfection, i.e., in screening experiments for siRNA delivery by magnetic complex analysis of luciferase expression can be performed usually after 48 h posttransfection. The results in (c) imply that magnetic triplexes comprising PEI-Mag2 nanomaterial are considerably more efficient in downregulation of the target gene in H441-Luci cells compared to similar nonmagnetic vectors. The results of the MTT assay suggest relatively low toxicity of tested complexes within the tested siRNA concentration range.

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Fig. 6.8. Luciferase reporter gene expression analysis in cell lysates. Magnetofection versus lipofection and polyfection efficiency in H441-Luci cells and MTT-based toxicity test. (a) Luciferase stably transfected HeLa cells (HeLa-Luci cells) were seeded in a 96-well plate and 24 h later transfected with magnetic triplexes PL-Mag1/PEI/siRNA. Luciferase expression was monitored in cell lysates in function of post-transfection incubation time. (b) Shows a calibration curve.

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Fig. 6.9. Enhancement of the GFP downregulation by magnetic siRNA vectors modified with INF-7 fusogenic peptide in GFP stably transfected HeLa human cervical epithelial adenocarcinoma cells. The cells were seeded in a 96-well plate and 24 h later transfected with magnetic triplexes PalD1-Mag1/PEI/siRNA or modified with INF-7 peptide as described in Sect. 4.14 (INF-7-to-siRNA mol/mol ratio of 9.4) PalD1-Mag1/PEI/siRNA/ INF-7. Iron-to-siRNA ratio of 0.5 to 1. PEI-to-siRNA ratio N/P = 10. GFP expression was monitored 72 h post-transfection. Results show percentages of reporter gene inhibition. Plots assigned as shown in the figure. The results clearly demonstrate considerable GFP-gene silencing improvement resulting from modification of the magnetic transfection triplexes with INF-7 peptide.

Dsample, Dblank and Dref are optical densities at the maximum of the MTT–formazan absorption spectrum registered for a sample, blank and reference sample, respectively. Examples of the results are given in Fig. 6.8c. 3.14. Enhancement of the Reporter Gene Downregulation by Magnetic siRNA Vectors Modified with INF-7 Fusogenic Peptide

To test the efficiency of the magnetic transfection triplexes modified with INF-7 fusogenic peptide (see Note 21): 1. Add 10 µl INF-7 solution to the transfection complexes from Sect. 3.6, Step 4 (see Note 22). 2. Perform Steps 5 and 9, described in Sect. 3.6 in order to generate dilutions of the INF-7 modified complexes. 3. Perform Magnetofection according to Sect. 3.8 and evaluate the GFP expression in cell lysates as described in Sect. 3.11. An example of the data is shown in Fig. 6.9.

4. Notes 1. Unless stated otherwise, all solutions should be prepared in water that has a resistivity of 18.2 MΩ;·cm and total organic

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content of less than five parts per billion. This standard is referred to as “water” in this text. 2. Palmitoyl dextran PalD1 for PalD1-Mag1 nanomaterial stabilization can be synthesized using a modification of Suzuki’s procedure (36): Dry Dextran 10 (20 g; M w = 10500 ; Amersham Biosciences) is suspended in anhydrous formamide (300 ml); the mixture is stirred in a water bath at 70°C for 1.5 h until the polysaccharide dissolves completely. Anhydrous tributylamine (30 ml; Sigma) and palmitoyl chloride (6 ml; Sigma) are added, and warming and stirring is continued for a further 2 h. The reaction mixture is cooled to 20°C. The resultant solution is diluted with 1500 ml methanol; a white precipitate is formed, collected by centrifugation, washed with methanol, and dried in vacuum. The crude product is dissolved in 400 ml formamide by stirring and heating at 50°C. The solution is poured into 2000 ml methanol. The white precipitate is collected by centrifugation, washed with methanol, and dried in vacuum. The PalD1 product with an esterification degree of 10 palmitoyl groups per 100 dextran units (determined as described in ref. 37) is water soluble. 3. This iron-to-siRNA weight ratio has turned out useful for triplexes of a variety of magnetic nanoparticle types. To determine the optimal weight ratio for an unknown particle type, it is useful to carry out this protocol also with magnetic nanoparticle stock suspensions of 18, 72, or more µg iron per ml (resulting in wt/wt ratios of 0.25 and 1 or higher). 4. The particles may aggregate due to magnetization; therefore, do not magnetize magnetic nanoparticles before transfection. Do not freeze magnetic nanoparticle suspensions. Before use, always vortex magnetic nanoparticle suspensions very thoroughly. Optionally, sonicate magnetic nanoparticle suspension after longer periods of storage using a water bath sonicator. 5. Gamma sterilization is preferred as heat sterilization in an autoclave in the presence of air can result in at least partial desorption of the coating components as well as surface oxidation of the magnetite nanocrystals. It is important to avoid freezing and magnetization of the suspensions prior to magnetofection. 6. Appropriate dilutions for measurement have a concentration between 0.5 and 6 µg iron per ml. The suggested final dilution for measurement of the original 20 µl magnetic nanoparticle with iron concentration of 10–90 mg iron per ml is 1:15,000.

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7. The iron content of the magnetic nanoparticles varies from 0.41 to 0.56 g iron per g dry weight; aqueous suspensions after dialysis of the material contain usually ~10 mg iron per ml. 8. The method described in Sect. 3.1 yields materials with magnetite cores with a mean crystallite size of 8–11 nm. The mean magnetite crystallite size was calculated from the broadening of the x-ray diffraction peak using the Scherer formula. The mean hydrodynamic diameters of these particles vary from 63 to 101 nm, and the x-potentials of the materials vary from highly positive (+55 mV) to negative (−16 mV), depending on the coating material used. The hydrodynamic diameter and z-potential of the magnetic nanoparticles and transfection complexes (given in Table 6.2) were determined by photon correlation spectroscopy using, for example, a Malvern Zetasizer 3000 (UK). The average iron weight per particle is evaluated taking into account the magnetite crystallite size (core diameter). The effective magnetic moment of the insulated particle is evaluated taking into account the experimentally determined saturation magnetization of the core and average iron weight per particle. The parameters given here are usually not supplied by manufacturers of magnetic nanomaterials. The physical and chemical characteristics of the particles are listed in Table 6.1. Remarkably, the ξ-potentials of these particles range from highly negative to highly positive. Particles with a negative ξ-potential are not suitable to bind nucleic acids on their own. For this purpose, either a combination with enhancers or divalent cations are required. These magnetic particles are used in combination with either siRNA alone (in the latter case, only positively charged particles such as PEI-Mag2 are used) or in formulations with nucleic acids and enhancers (either a lipid transfection reagent or PEI). We have found that particles with a magnetite crystallite size of 9–11 nm are superior to smaller particles with 3–4 nm crystallite size as components of the magnetic transfection vectors for magnetofection. For more details concerning the synthesis and characterization of the magnetic nanoparticles as components of gene vectors, see also ref. (24). 9. This protocol has to be performed by authorized personnel and according to the rules and regulations for work with radioactive substances. Use pipette tips provided with an aerosol filter to avoid radioactive contamination of the pipette. This procedure can be accomplished during 2 h. 10. For complete dissolution of thallium chloride just before DNA labeling, heat the solution to 70°C using a water bath. Caution: Thallium chloride is highly toxic.

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11. The N/P ratio is a measure of the ionic balance of the complexes and refers to the number of nitrogen residues of PEI per DNA phosphate. 1 µg of DNA contains 3 nmol of anionic phosphate. For N/P = 10, 1 µg DNA (3 nmol phosphate) corresponds to 30 nmol PEI units, i.e., to 1.3 µg PEI. 12. Cell culture and plating should be performed under sterile conditions. Timing: 30 min cell plating plus 24 h cell culture in a plate before transfection. 13. Cell seeding in clear bottom black-walled plates enables GFP expression measurements in living cells. 14. The order of reagent mixing and the medium for reagent dilution can be critical for the sizes, charges, and compositions of the complexes, and thus for final transfection efficiencies. To optimize the conditions for a given cell line, magnetic nanoparticle type and enhancer reagent, also other mixing orders as described above should/could be tested. 15. Timing: 60 min. Prepare transfection complexes just before transfection—all stages should be undertaken under sterile condition. 16. To evaluate the magnetic moment M of the magnetic complex and estimate the number of magnetic nanoparticles N associated with the complex, an efficient velocity uzof the complexes under a gradient magnetic field evaluated from the magnetic responsiveness curves as uz = L / t 0.1 , where L = 1 mm is an average path of the particle movement in an optical cuvette and t0.1 is the time required for a 10-fold decrease in optical density. The magnetic moment M of the complexes is calculated from the efficient velocity uzof the ∂B complexes as M = 3phDuz / , where D is the average ∂z hydrodynamic diameter of the complexes determined using the dynamic light scattering method, and h = 8.9·10−4 Pa·s (kg·m−1·s−1) is the viscosity of water. The total magnetic moment of the complex M is the product of the effective magnetic moment m eff of the magnetic nanoparticle under the magnetic field B and the total number N of magnetic particles associated with the complexes as M = N ·meff . At a magnetic field of 213 mT, the magnetization of the magnetite nanoparticles according to the experimentally measured magnetization curve corresponds to 97% of its saturation value, Ms. Thus the effective magnetic moment meff of each Fe particle is meff = (0.97 Ms )Ppart , where Ms is the specific satFe uration magnetization per unit of iron weight and Ppart is the content of iron in one particle with the diameter of the magnetite core equal to the average crystallite size. Thus

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determination of the complex velocity uz leads to evaluation of the number of magnetic particles associated with the complex. For more details on the physical background for the estimations see also ref. (38). 17. The optimal incubation conditions may differ from one cell type to another and from one complex to another and must be determined experimentally. 18. Use the same type of 96-well clear bottom black-walled plate for both eGFP calibration curve measurements and experimental sample measurements. Make sure to measure equal volumes for the calibration curve and the experimental samples. 19. Bear in mind that the luciferase and GFP assays are carried out with 50 ml cell lysate, while the protein assay is carried out with 10 µl only. Correspondingly, the measured values for luciferase (or GFP) must be divided by 5 to obtain correct results, when normalizing per total protein determined in 10 µl cell lysate. 20. To allow repeated GFP expression measurements post-transfection in living cells, the cells have to be seeded in a 96-well clear bottom black-wall plate: Greiner Bio-One; catalogue no. 655090 (or similar clear bottom black-wall plate). 21. INF-7 endosomolytic peptide derived from the influenza virus INF7 containing 24 amino acids [GLFEAIEGFIENGWEGMIDGWYGG) (39)] can be synthesized using the published procedure (27) or purchased from NeoMPS (Strasbourg, France). 22. Addition of the 2.33 µg INF-7 (865 pmol) peptide to (1.44 µg) 92.1 pmol siRNA in a 360 µl transfection complex results in INF-7-to-siRNA mol/mol ratio of 9.4.

Acknowledgments The authors would like to thank Dr. Bob Scholte for transduction of the H441 cells with eGFP and luciferase using lentiviral vectors. This work was supported by the European Union through the FP6-LIFESCIHEALTH Project “Improved precision of nucleic acid based therapy of cystic fibrosis” under contract no. 005213 as well as by the German Ministry of Education and Research, Nanobiotechnology grants 13N8186 and 13N8538. Financial support of the German Excellence Initiative via the “Nanosystems Initiative Munich (NIM)” is gratefully acknowledged.

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20. Minami, R., Yamamoto, M., Takahama, S., Miyamura, T., Watanabe, H., and Suematsu, E. (2006) RCAS1 induced by HIV-Tat is involved in the apoptosis of HIV-1 infected and uninfected CD4+ T cells. Cell. Immunol., 243, 41–47. 21. Bonetta, L. (2005) The inside scoop—evaluating gene delivery methods. Nat. Methods, 2, 875–883. 22. Smith, C. (2006) Sharpening the tools of RNA interference. Nat. Methods, 3, 475–486. 23. Felgner, P.L., Barenholz, Y., Behr, J.P., Cheng, S.H., Cullis, P., Huang, L., Jessee, J.A., Seymour, L., Szoka, F., Thierry, A.R. et al. (1997) Nomenclature for synthetic gene delivery systems. Hum. Gene Ther., 8, 511–512. 24. Mykhaylyk, O., Vlaskou, D., Tresilwised, N., Pithayanukul, P., Moller, W., and Plank, C. (2007) Magnetic nanoparticle formulations for DNA and siRNA delivery. J. Magn. Magn. Mater., 311, 275–281. 25. Terebesi, J., Kwok, K.Y., and Rice, K.G. (1998) Iodinated plasmid DNA as a tool for studying gene delivery. Anal. Biochem., 263, 120–123. 26. Azzam, T. and Domb, A.J. (2004) Current developments in gene transfection agents. Curr. Drug Deliv., 1, 165–193. 27. Plank, C., Oberhauser, B., Mechtler, K., Koch, C., and Wagner, E. (1994) The influence of endosome-disruptive peptides on gene transfer using synthetic virus-like gene transfer systems. J. Biol. Chem., 269, 12918–12924. 28. Plank, C., Zauner, W., and Wagner, E. (1998) Application of membrane-active peptides for drug and gene delivery across cellular membranes. Adv. Drug Deliv. Rev., 34, 21–35. 29. Funhoff, A.M., van Nostrum, C.F., Lok, M.C., Fretz, M.M., Crommelin, D.J., and Hennink, W.E. (2004) Poly(3-guanidinopropyl methacrylate): a novel cationic polymer for gene delivery. Bioconjug. Chem., 15, 1212–1220. 30. Jiang, X., Lok, M.C., and Hennink, W.E. (2007) Degradable-brushed pHEMA-pD-

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Chapter 7 In Vitro and In Vivo Gene Silencing by TransKingdom RNAi (tkRNAi) Shuanglin Xiang, Andrew C. Keates, Johannes Fruehauf, Youxin Yang, Hongnian Guo, Thu Nguyen, and Chiang J. Li Abstract RNA interference (RNAi) is a potent and specific mechanism for eliminating the mRNA of specific genes. This gene silencing mechanism occurs naturally and is highly conserved from plants to human cells, holding promise for functional genomics and for revolutionizing medicine due to its unlimited potential to treat genetic, epigenetic, and infectious disease. However, efforts to unleash the enormous potential of RNAi have met with significant challenges. Delivery is problematic because short interfering RNAs (siRNA) are negatively charged polymers that inefficiently enter cells and undergo rapid enzymatic degradation in vivo. In addition, the synthesis of siRNAs is expensive for long-term research and therapeutic applications. Recently, we have shown that nonpathogenic bacteria can be engineered to activate RNAi in mammalian cells (TransKingdom RNA interference; tkRNAi). This new approach offers several advantages and has significant implications. First, this method allows the establishment of a long-term stable gene silencing system in the laboratory against genes of interests in vitro and in vivo, and enables high-throughput functional genomics screening in mammalian systems. RNAi libraries can be constructed, stored, reproduced, amplified, and used with the help of E. coli as currently done with gene cloning. Second, this technology provides a clinically compatible way to achieve RNAi for therapeutic applications due to the proven clinical safety of nonpathogenic bacteria as a gene carrier. tkRNAi also eliminates the siRNA manufacture issue, and may circumvent or mitigate host interferon-like responses since siRNA is produced intracellularly. Key words: RNAi, TransKingdomRNA interference, tkRNAi, gene silencing, functional genomics, RNAi therapy, shRNA, bacteria-mediated RNAi.

1. Introduction The recent discovery of RNA interference (RNAi), a potent gene silencing mechanism found in eukaryotic cells, promises to revolutionize medicine due to its unlimited potential to treat genetic, M. Sioud (ed.), Methods in Molecular Biology, siRNA and miRNA Gene Silencing, vol. 487 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-60327-547-7_7

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epigenetic, and infectious disease (1–3). RNAi is triggered when endogenous micro RNA, or exogenous double-stranded RNA (dsRNA) or short hairpin RNA (shRNA) are processed by the cytoplasmic enzyme Dicer into 21- to 23-nucleotide short interfering RNA (siRNA) duplexes (4–6). The processed siRNA duplexes are then loaded into a large multiprotein complex called RISC (RNA-induced silencing complex) where the siRNA duplex is unwound and the passenger (sense) siRNA strand is discarded (7, 8). The RISC complex then locates target mRNA using the incorporated guide (antisense) siRNA strand and cleaves them using the slicer activity of the Argonaute protein, thereby preventing protein production. RNAi has proven a powerful technology for laboratory research of gene functions. However, in vivo gene silencing and large-scale research use of siRNA are limited due to delivery challenge and cost of synthesis (9, 10). To overcome these obstacles, we have developed a bacteria-based RNAi technology called TransKingdom RNAi (tkRNAi) (Fig. 7.1) for in vitro and in vivo gene silencing in mammalian cells (11). This approach utilizes genetically engineered, nonpathogenic E. coli to simultaneously manufacture silencing shRNA and deliver them to target cells. Our method makes use of a bacterial vector, pT7RNAi-Hly-Inv, termed TRIP (TransKingdom RNAi plasmid) that, when introduced into E. coli containing endogenous T7 RNA polymerase activity (e.g., BL21 DE3), can produce high levels of silencing shRNA. To enable delivery to the gastrointestinal tract, the TRIP plasmid was engineered to express the invasin gene (Inv) from Yersinia pseudotuberculosis, which allows noninvasive E. coli to enter β1-integrin positive epithelial cells (12, 13). To facilitate efficient gene silencing following cell entry, the TRIP vector was also engineered to express the listeriolysin O gene (HlyA) from Listeria monocytogenes, which allows the bacterially produced shRNA to escape from entry vesicles (14, 15). Using this approach, we have shown that tkRNAi directed against the colon cancer oncogene β-catenin can induce significant gene silencing in vitro and in vivo (11). This technique offers a number of advantages over chemically modified siRNA and viral vector-mediated shRNA methods for biomedical research and development of RNAi-based medical therapies. First, tkRNAi has significant implications for high throughput functional genomics in mammalian systems. Bacteria, E. coli in particular, have served as a well-validated and versatile vector system for the revolution in molecular biology and biotechnology that has occurred over the last few decades. Using tkRNAi, a laboratory can easily establish E. coli-based RNAi against various genes of interest. Besides saving on the cost of siRNA manufacture, the bacterial gene silencing system can be reproduced and

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Fig. 7.1. Schematic representation of TransKingdom RNAi.

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stored for long term use, offering a stable and consistent gene silencing tool. Further, the bacterial vector can be used in a routine biological laboratory, rather than a BL2 laboratory, which is required for viral vector-based systems. The bacterial system can also conveniently be translated to in vivo systems in order to verify in vitro observations in live animals. Other advantages of using bacteria as a delivery vector for siRNA include the ability to control the vector using antibiotics and/ or auxotrophy, and the ease of engineering specific vectors for particular applications. Second, the tkRNAi system provides a practical and clinically compatible way to achieve RNAi for medical therapies. Although RNAi can theoretically be employed to target any disease gene with specificity and potency, the development of RNAi-based therapeutics has been impeded by challenges in delivery, manufacture, and the activation of host interferon-like responses (9, 10). Of these, delivery has proven to be the major stumbling block. In order to overcome these limitations, researchers have focused mainly on developing chemically modified siRNA to increase stability, and on complexing siRNA with liposomes or nanoparticles to promote cell uptake (9, 16, 17). The main disadvantage of these pharmaceutical approaches, however, is that they tend to have a limited ability to target specific cell types or tissues. Moreover, they typically require large quantities of siRNA that is expensive to manufacture. Viral vectors have also been explored as a means of delivering RNAi in vivo (9). While this approach has important research applications, problems associated with insertional mutagenesis, safety, lack of tropism, and the generation of host immune responses have significantly limited the utility of viral vectors for gene therapy. Unlike viral vectors, nonpathogenic bacteria (including E. coli) have been used safely for many years as probiotics or as vaccine vectors, and do not integrate into the human genome (18–21). Thus, the tkRNAi technique may help to achieve potent and therapeutic RNAi with versatility and less cost. This RNAi approach can be exploited clinically to silence genes of interest in the colonic mucosa, and possibly in other organs which can be colonized by commensal or nonpathogenic bacteria, including the oral cavity, urinary bladder, female genital tract, etc. Because shRNAs are released intracellularly by the engineered bacteria directly into the cytoplasm, this RNAi approach eliminates the siRNA manufacture issue and may have the advantage of mitigating the Toll-like receptor-mediated immunostimulatory effect of siRNA (see Chap. 8). Therefore, the TransKingdom system may provide a practical and clinically compatible way to achieve RNAi for medical indications.

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2. Materials 2.1. Bacterial Culture

1. Brain Heart Infusion (BHI) Broth (Remel, M & M Industries, Inc., Chattanooga TN): Dissolve 37 g BHI in 1 l of tissue culture water, dispense into appropriate containers, and sterilize by autoclaving at 121°C for 15 min. Store at room temperature. 2. Sterile disposable round-bottom plastic tubes with dualposition cap (14 ml: VWR International, West Chester, PA); KIMAX brand baffled culture flasks (500 ml; Fisher Scientific, Pittsburgh, PA). 3. Ampicillin 100 mg/ml (Sigma, St Louis, MO). Store at −20°C. 4. Isopropyl β-D-1-thiogalactopyranoside (IPTG; Sigma, St Louis, MO). Prepare 1 M working solution, filter through a 0.22 µm filter, and store at −20°C.

2.2. Cell Culture

1. SW480 colonic epithelial cells (American Type Culture Collection, Manassas, VA) are maintained in complete growth media in an atmosphere of 5% CO2 and 95% air. For longterm storage, cells are resuspended in complete growth medium supplemented with 5% (v/v) dimethyl sulfoxide and placed in liquid nitrogen. 2. Roswell Park Memorial Institute (RPMI) medium (Invitrogen, Carlsbad, CA) supplemented with 10% fetal bovine serum (Invitrogen, Carlsbad, CA). Store at 4°C. 3. Sterile plastic tissue culture flasks (75 cm2; BD Falcon, Franklin Lakes, NJ) and dishes (6 cm; Corning Life Sciences, Lowell, MA). 4. Penicillin–streptomycin solution: 10,000 U/ml penicillin, 10 mg/ml streptomycin in 0.9% sodium chloride (Sigma, St Louis, MO). Store at −20°C. 5. Amphotericin B solution: 250 mg/ml in tissue culture water (Sigma, St Louis, MO). Store at −20°C. 6. 0.25% Trypsin–EDTA solution (Sigma, St Louis, MO) is stored in aliquots at −20°C. 7. Gentamycin solution: 10 mg/ml in tissue culture water. Store at 4°C. 8. Ofloxacin 10 mg/ml in tissue culture water. Store at −20°C. All working solutions are prepared by diluting the corresponding stock solution 1000-fold.

2.3. Oral Administration of Bacteria to Mice

1. Female C57BL/6 mice (Charles River Laboratories, Wilmington, MA) are housed under conventional conditions in isolator cages (4 mice per cage). Mice are fed standard chow

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(Harlan Teklad, Indianapolis, IN) and provided with tap water ad libitum. 2. 10X phosphate buffered saline is stored at room temperature. A 1X working solution is prepared by diluting the stock solution 10-fold in sterile water. Store the solution at 4°C. 3. Feeding needle (PS 20; Poppers and Sons, Inc., New Hyde Park, NY). 4. 1 ml Norm-Ject tuberculin syringes (Henke Sass Wolf, Tuttlingen, Germany). 2.4. Intravenous Administration of Bacteria to Mice

1. Female nude Balb/c mice (Nu/Nu; Charles River Laboratories, Wilmington, MA) are housed under specific pathogenfree conditions in sterile isolator cages (4 mice per cage). Mice are fed with irradiated chow (Harlan Teklad, Indianapolis, IN) and provided with sterile water ad libitum 2. 1 ml Norm-Ject tuberculin syringes (Henke Sass Wolf, Tuttlingen, Germany). 3. 26G1/2 PrecisionGlide needles (Becton Dickinson, Franklin Lakes, NJ).

3. Methods In vitro and in vivo gene silencing via tkRNAi takes advantage of a “Trojan Horse” strategy. In this system, interfering shRNA are transcribed (from the T7 promoter) inside nonpathogenic bacteria (or commensal bacteria) that are engineered to actively invade target cells. This process is comprised of four steps: 1. Cell entry 2. Bacterial lysis and rupture of the entry vesicle 3. Release of interfering shRNA into the host cell cytoplasm 4. Gene-silencing through RISC As gene silencing relies on effective bacterial invasion and the intracellular release of interfering shRNA, the ability of the engineered bacteria to stimulate cellular uptake by endocytosis and subsequent intracellular are important. For systemic treatment, intravenous injection and oral administration of shRNA-expressing bacteria can be used. Intravenous injection of nude mice carrying human colon cancer xenografts with 1 × 108 c.f.u. of engineered E. coli resulted in bacterial delivery to the liver, spleen, and tumor tissue during the first 24 h. It should be noted that during the first week after treatment, bacterial numbers continue to increase in the tumor tissue but decrease

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in the liver and spleen. Interestingly, injection of live engineered bacteria did not generate a significant endotoxin response in nude mice; thus, repetitive treatment is feasible. The enrichment of the engineered E. coli in the xenograft tissue likely reflects the ability of this facultative anaerobe strain to colonize and replicate in the hypoxic environment present in tumor tissues. For oral administration of tkRNAi, attenuated versions of E. coli are developed, thus allowing the administration of large amounts of therapeutic bacteria (typically in the order of 1010 c.f.u.). Such attenuated tkRNAi bacteria do not colonize the gastrointestinal tract when administered orally. They are rapidly eliminated. 3.1. In Vitro tkRNAi

A typical in vitro tkRNAi experiment (see Fig. 7.2) consists of three experimental phases: 1. Preparation of shRNA-expressing bacteria 2. Target cell preparation and bacterial infection 3. Postinfection treatment and assessment of target gene silencing

3.1.1. Bacteria Preparation

1. Chemically competent E. coli BL21-Gold (DE3) (50~100 µl) cells are transformed with control or silencing TRIP plasmids (100 ng) according to the manufacturer’s instructions (Stratagene; see Note 1). Bacteria are then grown on BHI plates containing 100 µg/ml ampicillin overnight at 37°C until colonies appear. A single colony is then inoculated into BHI medium containing 100 µg/ml ampicillin, and grown overnight at 37°C (see Note 2). 2. The next day, 5 ml of each overnight culture is diluted 1:40 into fresh BHI medium containing 100 µg/ml ampicillin

Fig. 7.2. Flow chart outlining in vitro TransKingdom RNAi.

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and grown for a further 2–4 h (until the OD600 = 0.5). Each culture is then treated with IPTG (1 mM final concentration) for 2–4 h to induce transcription of interfering shRNA (see Note 3). To verify the identity of each bacterium, plasmid DNA is isolated from each overnight culture and sequenced using standard techniques. 3. After IPTG induction, the total number of bacteria in each culture is calculated by measuring the OD600 value (8 × 108 bacteria/ml culture has an OD600 = 1). The number of bacteria for cell treatment is then calculated according to cell confluency and the needed multiplicity of infection. We advise to use a range of 20:1 to 2000:1, bacteria to cells in appropriate reaction volumes. 4. The required volume of bacteria culture is then centrifuged at 2500g for 10 min at 4°C and the pellet is washed once with serum-free, RPMI 1640 medium containing 100 µg/ ml ampicillin and 1 mM of IPTG, and resuspended in the same medium at the required density for bacterial infection. 3.1.2. Cell Preparation and Bacterial Infection

1. SW480 human colon cancer cells are cultured in an atmosphere of 95% air, 5%CO2 at 37°C in RPMI 1640 medium containing 10% FBS, 10 U/ml penicillin G, 10 µg/ml streptomycin, and 250 µg/ml amphotericin. 24 h prior to bacterial infection, stock cell cultures are trypsinized, resuspended in complete RPMI 1640 medium, and plated on 6 cm tissue culture dishes at 20–30% confluency. 2. 30 min prior to bacterial infection, the cell culture medium is replaced with 2 ml of fresh serum-free RPMI 1640 medium containing 100 µg/ml of ampicillin and 1 mM IPTG. 3. Bacteria as prepared in Sect. 3.1.1 are then added to the cells at the desired MOI for 2 h at 37°C.

3.1.3. Postinfection Treatment and Assessment of Target Gene Silencing

1. After the infection period, the cells are washed 3 times using serum-free RPMI 1640 medium (see Note 4). The cells are then incubated with 2 ml of fresh complete RPMI 1640 medium containing 100 µg/ml of ampicillin and 150 µg/ ml of gentamycin for 2 h to kill any remaining extracellular bacteria. 2. After treatment with ampicillin and gentamycin, the cells are incubated with 3 ml RPMI 1640 medium containing 10 µg/ ml of ofloxacin in order to kill any intracellular bacteria. 3. The cells are then harvested at various time points (from 24 to 96 h) in order to assess the extent of target gene silencing by real-time PCR (for mRNAs) and Western blotting (for proteins). To illustrate the protocol, some experimental data are presented in Fig. 7.3.

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Fig. 7.3. TransKingdom RNAi in vitro. (a) The expression of beta-catenin protein was silenced in SW480 cells after treatment with E. coli expressing shRNAs against beta-catenin at various MOI (lanes 4–7). Lanes 2 and 3 show lack of gene silencing when either the hly-inv part of the plasmid (lane 2) or the shRNA encoding part (lane 3) of the plasmid are missing, even at very high MOI. (b) The gene-silencing effect depends on the co-culture time of human cells with the bacteria (lanes 3–6). E. coli containing a TRIP against wild-type k-Ras exerted no gene silencing effects against mutant k-Ras (V12G). (c) E. coli containing a TRIP against mutant k-Ras (V12G) silenced codon-matched mutant k-Ras expression in SW480 cells, but not in DLD1 cells containing a k-Ras mutation in a different codon.

Fig. 7.4. Flow chart outlining in vivo TransKingdom RNAi.

3.2. In Vivo tkRNAi

Similar to the in vitro tkRNAi protocol (Sect. 3.1), a typical in vivo tkRNAi experiment (see Fig. 7.4) also consists of three experimental phases (Fig. 7.4):

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1. Preparation of shRNA-expressing bacteria 2. Oral or intravenous delivery 3. Assessment of target gene knockdown 3.2.1. Bacteria Preparation

1. Transformed E. coli BL21 (DE3) bacteria containing control or silencing TRIP plasmids (verified by plasmid purification and DNA sequencing) are grown in BHI medium containing 100 µg/ml ampicillin at 37°C until early log phase (OD600 = 0.5). The bacteria are then harvested by centrifugation at 2500g for 10 min at 4°C, resuspended in 25 ml of BHI medium, aliquoted and stored in −80°C freezer as 15% glycerol stock (see Note 5). 2. One day prior to animal treatment, the bacteria stocks are thawed, inoculated into 50 ml of fresh BHI medium containing 100 µg/ml ampicillin, and incubated overnight with shaking at 37°C. 3. The next day, each overnight culture is re-inoculated into fresh BHI medium (at a 1:40 ratio) containing 100 µg/ml ampicillin and grown for a further 2–4 h (until OD600 = 0.5). IPTG is then added to a final concentration of 1 mM, and the bacteria are incubated at 37°C with shaking for another 2–4 h. 4. Subsequent to IPTG induction, the total number of bacteria in each culture is determined by measuring the OD600 value (8 × 108 bacteria/ml culture has an OD600 = 1). The volume of bacterial culture required for animal treatment is then centrifuged at 2500g for 10 min at 4°C and the pellet is washed once with 1X PBS, and then resuspended at the required density in 1X PBS. This preparation is ready for oral administration or intravenous injection. Keep the preparations at 4°C, but warm just prior to treatment.

3.2.2. Oral Treatment of Normal Mice

1. Age-matched female C57BL/6 mice are randomly divided into treatment and control experimental groups (typically consisting of 6–8 animals per group). 2. Animals in each treatment group are given 5 × 108 to 5 × 1010 c.f.u. of shRNA-expressing E. coli (in 200 µl PBS) via a PS-20 oral feeding needle fitted to a 1 ml syringe. Control animals are treated similarly except that the administered E. coli contains a TRIP vector without the shRNA insert, or an insert coding for an inactive (e.g., scrambled) shRNA sequence. 3. Oral administration of control or shRNA-expressing bacteria is then performed 5 days per week for a total of 4 weeks (see Note 6). Mice are sacrificed 2 days after the last treatment,

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and colonic tissues are collected for analysis of target gene silencing. 3.2.3. Intravenous Treatment of Nude Mice Bearing Colon Cancer Xenografts

1. Age-matched female nude Balb/c mice are randomly divided into treatment and control experimental groups (typically consisting of 6–8 animals per group). 2. Three weeks before bacterial treatment, animals in each experimental group are subcutaneously implanted in the right flank with 1 × 107 SW480 colon cancer cells (in 100 µl PBS) using a 1 ml syringe fitted with a 26G needle. 3. Bacterial treatments are initiated when the xenograft tumors reach approximately 10 mm in diameter. Animals are treated intravenously with 1 × 108 c.f.u. of shRNA-expressing or control E. coli (in 100 µl PBS) by tail vein injection using a 1 ml syringe fitted with a 26G needle. 4. Control or shRNA-expressing bacteria are administered to the animals every 5 days for a total of three treatments (see Note 7). Mice are sacrificed 5 days after the final treatment, and the tumor tissues are collected for analysis of target gene silencing.

3.2.4. Assessment of Target Gene Knockdown

1. For analysis of target gene mRNA levels by real-time PCR, colon and xenograft tissues are frozen and stored at −80°C. Total RNA isolation and real-time PCR analysis are then performed according to standard protocols. 2. For analysis of target gene protein levels by immunohistochemistry, colon and xenograft tissues are fixed in paraformaldehyde, paraffin-embedded, sectioned, and then stained according to standard procedures. For illustration of the protocol, some experimental data are presented in Fig. 7.5.

4. Notes 1. As TRIP plasmids are relatively large (~8.9 kb), the transformation efficiency using E. coli BL21-Gold (DE3) is quite low. To guarantee successful transformation, newly constructed TRIP plasmids (in the ligation mixture) should be immediately transformed into competent BL21-Gold bacteria. Plasmids purified from BL21-Gold bacteria can then be used to synthesize other engineered BL21 (DE3) strains. 2. Bacteria containing TRIP plasmids tend to grow very slowly in LB medium. Brain heart Infusion (BHI) medium, which is nutrient-rich, allows for faster growth of TRIP-containing bacteria. The expression of invasin (Inv) and listeriolysin

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Fig. 7.5. TransKingdom RNAi in vivo. (a) Oral administration of E. coli expressing shRNAs against beta-catenin in mice leads to significant reduction of beta-catenin expression in normal intestinal epithelium, especially in the regions of, or adjacent to, Peyer’s patches. (b) Representative view of intestinal epithelium from treated (right) and control (left) animals. (c–e) Intravenous administration of E. coli containing a TRIP against human beta-catenin in mice containing human colon cancer xenograft tumors resulted in a decrease in beta-catenin mRNA levels (c) and protein levels (d, e) in tumor tissues.

O (HlyA) is also better in BHI medium than LB medium. Maximal expression of both of these proteins is critical for stimulating bacterial endocytosis by target cells and endosomal escape of shRNA from entry vacuoles. 3. With most bacteria, IPTG induction is usually performed in the mid-log phase (OD600 > 0.7). However, with E. coli BL21-Gold (DE3), IPTG induction is best performed during early log phase (OD600 = 0.4 ~ 0.6). 4. Cells must be washed using serum-free RPMI 1640 medium. Washing the cells with PBS will cause the bacteria to firmly attach to the cell surface. This can negatively affect cell viability and enhance the probability of false positive data. 5. The use of aliquoted bacterial stocks gives more reproducible results for in vivo tkRNAi than growing single colonies from BHI-ampicillin plates.

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6. Oral administration is well tolerated with no gross or microscopic signs of epithelial damage or ulcerations. 7. Intravenous injection is generally well tolerated without adverse effects. However, mice should be closely monitored during treatment. If the nude mice get sick during treatment, 40 mg/kg of ofloxacin can be applied. Alternatively, one treatment with bacteria can be omitted.

Acknowledgements We thank Dr. J. T. LaMont for advice and discussions, and C. Griillot-Courvalin of the Pasteur Institute, Paris, France for providing the sequences for Inv and Hly (pGB2Ω) and also for helpful discussions.

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20. Daudel, D., Weidinger, G., and Spreng, S. (2007) Use of attenuated bacteria as delivery vectors for DNA vaccines. Expert Rev Vaccines 6, 97–110. 21. Kruis, W., Fric, P., Pokrotnieks, J., Lukas, M., Fixa, B., Kascak, M., Kamm, M. A., Weismueller, J., Beglinger, C., Stolte, M., Wolff, C., and Schulze, J. (2004) Maintaining remission of ulcerative colitis with the probiotic Escherichia coli Nissle 1917 is as effective as with standard mesalazine. Gut 53, 1617–1623.

Chapter 8 Bacterial Delivery of siRNAs: A New Approach to Solid Tumor Therapy De-Qi Xu, Ling Zhang, Dennis J Kopecko, Lifang Gao, Yueting Shao, Baofeng Guo, and Lijing Zhao Abstract RNAi is a powerful research tool for specific gene silencing and may also lead to promising novel therapeutic strategies. However, the development of RNAi-based therapies has been slow due to the lack of targeted delivery methods. The biggest challenge in the use of siRNA-based therapies is the delivery to target cells. There are many additional obstacles to in vivo delivery of siRNAs, such as degradation by endogenous enzymes and interaction with blood components leading to nonspecific uptake into cells, which govern biodistribution and availability of siRNA in the body. Naked unmodified synthetic siRNA including plasmid-carried-shRNA-expression constructs cannot penetrate cellular membranes, and therefore, systemic application is unlikely to be successful. The success of gene therapy by siRNAs relies on the development of safe, economical, and efficacious in vivo delivery systems into the target cells. Attenuated Salmonella have been employed recently as vectors to deliver silencing hairpin RNA (shRNA) expression plasmids into mammalian cells. This approach has achieved gene silencing in vitro and in vivo. The facultative anaerobic, invasive Salmonella have a natural tropism for solid tumors including metastatic tumors. Genetically modified, attenuated Salmonella have been used recently both as potential antitumor agents by themselves, and to deliver specific tumoricidal therapies. This chapter describes the use of attenuated bacteria as tumor-targeting delivery systems for cancer therapy. Key words: Bacterial delivery vector, Salmonella, siRNA, shRNA, RNAi, solid tumor.

1. Introduction RNAi is a eukaryotic intracellular process wherein a small interfering RNA (siRNA) directs a sequence-specific degradation of its target mRNA (1). Within the cytoplasm of a given cell, this RNAi duplex is recognized by a multiprotein complex called RISC M. Sioud (ed.), Methods in Molecular Biology, siRNA and miRNA Gene Silencing, vol. 487 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-60327-547-7_8

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(RNA-induced silencing complex). One of the RNA strands (the antisense strand) remains bound to the RISC complex and binds to complementary mRNA sequences within the cytoplasm. This mRNA–siRNA–RISC complex induces cleavage of the mRNA, thereby preventing its translation into protein and lowering genespecific expression. Many tumors are caused by mutations that abnormally upregulate specific gene expression. RNAi therapy has been used successfully to knockdown this abnormal gene expression and the associated tumor growth in animal studies, but the application of RNAi therapy in humans has faced practical difficulties. The biggest challenge to the systemic use of siRNA-based therapies is the difficulty of delivery, specifically across the plasma membrane to reach the cytoplasm of target cells. There are additional obstacles to in vivo delivery of siRNAs, such as enzymatic degradation in blood, interaction with blood components, and nonspecific uptake by cells, which affect siRNA availability and distribution in the body. Naked unmodified synthetic siRNA including plasmid-encoded-shRNA-expression constructs cannot, by themselves, penetrate cellular membranes and, therefore, direct systemic administration of purified siRNA is unlikely to be successful. Alternatively, shRNA can be expressed inside host cells using DNA templates that direct the synthesis of RNA duplexes (2, 3). In this latter case, the RNAi effect can be sustained for a longer term, depending upon the vector employed and optimal selective pressure for retention of the plasmid-borne expression cassettes. DNA-directed RNAi (ddRNAi) expression constructs can be directly administered intratumorally for surface nodes, but are not appropriate for systemic administration for the reasons indicated above. Systemic delivery systems that use viral vectors, such as retrovirus or adenovirus, have targeting advantages, but are limited to research use in animal models due to safety issues. Thus far, bacteria as a delivery system have great potential advantages over other delivery vectors and have already been employed in animals and humans (see Chap. 7). An ideal delivery system should be (a) nontoxic to normal cells, and (b) able to deliver the therapeutic efficiently and specifically to the tumor. Attenuated Salmonella have been employed recently as vectors to deliver silencing shRNA expression plasmids to mammalian cells and shown to achieve specific gene silencing in vitro in cultured cancer cell lines and in vivo in mice containing implanted tumors (4). The facultative anaerobic, invasive Salmonella have a natural tropism for solid tumors including metastatic tumors. Genetically modified, attenuated Salmonella have been used as potential antitumor agents, either to elicit direct tumoricidal effects and/or to deliver tumoricidal molecules (e.g., siRNA) (5–9). This chapter is focused specifically on the use of live bacterial vector systems for targeted delivery of cancer therapies.

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Bacterial delivery of siRNA is dependent upon the construction of shRNA-expression cassettes designed to induce sustained RNAi production. Plasmid-driven expression of siRNA in vivo has typically been based on the general design. A Pol III promoter is used to direct the synthesis of inverted RNA repeat sequences separated by a short spacer region. The expressed hairpin RNA contains two complementary nucleotide stems that encode a specific target sequence, a terminal loop sequence (4 to 9 nt) at one end, and a 3′-U overhang at the opposite end. These hairpin RNAs are processed in vivo by Dicer to leave only the duplex region (Fig. 8.1) (10). Since poly(T) is used as a transcriptional terminator, stretches of >4 Ts or As in the target sequence should be avoided. siRNA target regions with 30–50% GC content appear to be more active than those with a higher GC content. The selection of the target site can be aided by using the BLAST search tool (www.ncbi.nlm.nih.gov/BLAST). RNAimediated gene silencing can be assessed in cultured mammalian cells by plasmid delivery and endogenous expression of shRNA harboring a fold-back stem–loop structure. The advantage of this shRNA expression system is that the RNAi effects are prolonged when plasmid-based siRNAs are used. The selection of specific target sequence, the length of the inverted repeats that encode the stem of a putative hairpin, the order of the inverted repeats (i.e., sense vs. antisense), the length and the composition of the spacer sequence that forms the loop region, all can affect the functioning of the siRNA. It is necessary to check the specificity of the target sequence to confirm that there are no highly hom*ologous sequences in other genes. The selected target site should be devoid of strong secondary structures, which could impair the binding of the siRNAs; computerbased folding programs can be helpful in this selection. However, the actual site accessibility can only be evaluated empirically in biological function studies. The shRNA cassette can be designed with either the sense or the antisense strand, placed immediately after the promoter. However, since U6 initiates transcription with a G, it is preferable that this be the first base of the sense strand, since a G in the first position of the sense strand of siRNA is positively correlated with siRNA functionality (11). The length of the loop sequence of the hairpin-type siRNA is also important for silencing activity. Although a short 4-nt, 5′-UUCG-3′ could still support significant RNAi function (12) when a 19-bp stem sequence was used, the hairpin RNA with a standard 9-nt loop sequence (UUCAAGAGA) had greater silencing activity than the corresponding RNA with a shorter loop sequence (12–14). However, it has been shown that the natural loop sequence of microRNAs, which are endogenous shRNAs, are preferable sequences for shRNA production.

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Hairpin siRNA Contruction Example Target Sequence (from Stat3/TAD domain/19 nt) 5’- GCAGCAGCTGAACAACATG- 3’ Create 2 complementary oligonucleotides; each contains sense strand-loop sequence-antisense strand and 5 Ts (terminator) plus terminal restriction sites

Annealing shRNA Template Insert

Sense Strand Loop Antisense Strand Terminator 5’-GATCC GCAGCAGCTGAACAACATGTTCAAGAGACATGTTGTTCAGCTGCTGCTTTTTTGGAAA -3’ 3’-CGTCGTCGACTTGTTGTACAAGTTCTCTGTACAACAAGTCGACGACGAAAAAACCTTTTCGA-5’ BamH I Hind III

BamHI

HindIII

CMV

shRNA Structure-Expressed in Cells

Amp

GFP pGCsi-U6/Neo/GFP

(psi-Stat3-GFP)

Sense sequence UUCA 5’-GCAGCAGCTGAACAACATG A 3’-CGTCGTCGACTTGTTGTAC Antisense sequence UGUG

SV40 pUC ori

Processed in cell cytoplasm:

Dicer

Neo

Fig. 8.1. Strategies for hairpin siRNA plasmid construction for bacterial delivery of design target siRNA sequence using the web-based converter at www.ambion.com/techlib/misc/psilencer_converter.html. Here we chose a target sequence GCAGCAGCTGAACAACATG from the TAD domain of human oncogene Stat3 (corresponding to nucleotides 2,144 to 2,162; GenBank accession number NM_003150). The expression vector psi-Stat3-GFP was constructed with pGCsiU6Neo-GFP vector containing a U6 promoter, BamHI and HindIII cloning sites, a CMV promoter and GFP-encoding sequence, and a neomycin-resistant gene for selection. The details of construction are described in the text. The structure of psi-Stat3GFP plasmid containing the sequence of Stat3-specific hairpin RNA (shRNA-Stat3) or Scramble sequence insert from Ambion’s (negative control plasmid) are shown in the lower panel.

Vector-based RNAi synthesis also permits coexpression of reporter genes such as GFP or luciferase, which facilitate the tracking of transfected cells or evaluating the effect of a GFP-tagged shRNA expression vector. In any delivery system, coexpression of GFP can be very important to track bacterial distribution in vitro (in cell lines) or in vivo. For any RNAi experiment, it is necessary to include a negative control plasmid to rule out nonspecific gene

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knockdown. An optimal negative control plasmid should contain a scrambled sequence of your gene-specific siRNA. 1.2. Choice of Attenuated Bacterial Species or Strains

Any bacteria suitable for use as a tumor-specific delivery vector must possess several features: (i) It must be genetically or biologically traceable (e.g., through genetic markers or GFP production), (ii) it must be nonpathogenic with minimal toxicity, (iii) it must have reduced immunogenicity, (iv) it should have invasive and motile abilities (e.g., ability to invade epithelial cells and to spread from cell to cell), (v) it must have preference for tumor tissue over normal tissue, and (vi) it must be susceptible to antibiotics for eradication purposes. The discovery that genes vectored by bacteria can be functionally transferred to mammalian cells has suggested the possible use of bacterial vectors as vehicles for gene therapy. Several reports have demonstrated the utility of anaerobic bacteria (such as Clostridium spp.) as anticancer agents; their utility may be limited by the absolute requirement for anoxic conditions, restricting their use to large tumors (15). These restrictions do not apply to facultative anaerobes (e.g., Salmonella spp.) such as the gram-negative, bacterium Salmonella enterica serovar Typhimurium (S. typhimurium). S. typhimurium is known to colonize various types of human and murine tumors, allowing treated mice to survive for periods long after untreated mice have died. Salmonella can grow in the presence or absence of oxygen and can colonize both large and small tumors (16). Salmonellae have been shown to inhibit the number and size of melanoma micrometastases and to distribute throughout the tumor mass (17). Construction of attenuated bacterial strains is feasible with current DNA manipulation techniques and knowledge of molecular microbiology. A strain could be deleted for a single gene or mutated in multiple genes to avoid reversion to virulence. The following principles have played a role in the selection of bacterial genes to create attenuating mutations, which would be essential for use in cancer therapy. (A) Advances in microbiology and biotechnology have led to genetic modification of bacteria so that they can replicate and persist within tumors, but which limit their distribution to and survival in normal tissues. For example, bacterial mutations that result in nutritional auxotrophy generally restrict bacterial growth in normal tissues. Deletions in purI create a bacterial requirement for exogenous adenine and deletions in the aro pathway result in a requirement for aromatic amino acids. Auxotrophic Salmonella mutants disseminate to the nutrient-rich tumor environment where they replicate to levels exceeding 103–104 times the low concentration found in normal tissues (9). Combining auxotrophic mutants with mutations in the antigenic bacterial LPS genes can minimize immunogenic/ inflammatory potential. (B) Deletion of the msbB (renamed

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waaN) gene, which controls the terminal myristylation of lipid A, reduces strain toxicity. The msbB mutant is reduced in endotoxicity about 10,000-fold, which allows for the systemic administration of such attenuated strains. The mutation of this gene reduces tumor necrosis factor-α (TNF-α)-mediated septic shock and tremendously increases the LD50 of this pathogen in the mouse model (18). The msbB mutant is of clinical interest because it allows Salmonella to be administered safely to mammals, which is essential for the development of safe, live, attenuated bacterial delivery systems. However, S. typhimurium msbB-deletion strains have severe growth defects, in many laboratory media, that can be suppressed by extragenic compensatory mutations which arise at high frequency in related genes. Suppressor mutations for msbB allow Salmonella to grow well in culture media but still have reduced lipid A myristylation to avoid the septic shock response. By choosing a suppressor mutation that confers the desired characteristics on a given product strain, (e.g., the tumor-targeting Salmonella strain VNP20009 is nontoxic and retains tumor targeting and tumor inhibition abilities) (19). (C) However, another well-characterized transcriptional regulon required for Salmonella pathogenesis is the phoP/phoQ two-component regulatory system (20–21). This regulatory system controls the expression of more than 40 different genes required for virulence in mice and which promote resistance to innate immune defenses (e.g., resistance to defensins and low pH), induces spacious phagosomes and survival in macrophages, and is involved in nutrient scavenging and lipid A modifications (22). Due to the simultaneous altered expression of multiple virulence factors, phoP/phoQ mutants retain a high degree of attenuation in IFN-γ−/− mice after oral challenge with 5 × 109 cfu (23). Virulence attenuation of phoP/phoQ mutants can also be attributed to structural modification of the lipid A moiety. The structural modifications to lipid A, the host signaling portion of LPS, occur by the addition of aminoarabinose and 2-hydroxymyrisate to the mutant hexaacyl lipid A. This modification not only confers polymyxin resistance, but also affects LPS-induced host responses (i.e., decreased E-selectin and TNF-α expression (24). The bacterial load of S. typhimurium phoP/phoQ mutants in mice in infected organs is elevated during the first week postinfection, but total bacterial clearance is achieved by day 14. Thus, even lacking a specific mutation in msbB, the phoP/phoQ deletion mutant has great reduced endotoxicity. In our recent report, a phoP/phoQ deletion mutant of S. typhimurium (4) was employed as a carrier for shRNA or shRNA combined with an antitumor protein-expressing plasmid for use in prostate cancer therapy. In vivo mouse and in vitro tissue culture studies showed significant delivery efficacy for Salmonelladelivered therapy into mammalian tumor cells. Regardless of the route of administration (i.e., oral, intravenous, intratumoral, or

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intraperitoneal inoculation) Salmonella-delivered therapy resulted in significant antitumor efficacy with no toxic effects observed in the mouse model. By day 15, when no bacteria were detected in normal spleen or liver tissues, Salmonella remained within the tumor tissue. The S. typhimurium phoP/phoQ deletion mutant has been shown to accumulate preferentially >1,000-fold greater in tumors than in normal tissues and to disperse hom*ogeneously in tumor tissues. Salmonella-delivered siRNA was found to downregulate expression of Stat3 and to exert significant antitumor effects including reduction in metastases and reversal of tumorigenesis (4). 1.3. Administration of Vector-Based shRNA Expression Systems Delivered with Attenuated Salmonella

Tumor-targeting bacteria have been investigated intensively in recent years as anticancer agents by themselves or, more recently, as a delivery system for anticancer therapies. Systemic administration of live bacterial vectors by intravenous (i.v.) or intraperitoneal injection (i.p.) has been widely and effectively used in mouse tumor studies (24–26). Direct local administration, such as intratumoral injection, has been demonstrated to be practical but still is limited to use on surface tumors. Oral administration would be ideal if a reproducible amount of Salmonella could reach the target tumor cells. In mice, oral administration of S. typhimurium results in reproducible systemic inoculation. The intravenous administration of attenuated Salmonellae has already been employed in human use and allows for bloodstream delivery of a specific therapeutic dose to both primary and secondary tumors (18, 26). Bacterial motility should favorably disperse the vector throughout the tumor tissue, overcoming diffusion and distal pressure gradients to deliver the therapeutic agents to all regions of the tumor. Motility is a major advantage that bacteria have over viral vectors, as viruses exhibit a lack of tumor specificity and are poorly distributed throughout the tumor mass. In addition, metabolically active bacteria will replicate and continue to produce active antitumor agents, thus amplifying the dose to an effective level only within the target tumor.

2. Materials 2.1. Bacterial Strain, Growth Media, and Electrotransformation

1. Attenuated S. typhimurium phoP/phoQ null mutant LH 430 was derived from ATCC 14028 by deletion of the phoP/ phoQ locus (27, 28). 2. E. coli DH5α (Invitrogen). 3. Luria-Bertani (LB) broth, brain heart infusion (BHI) broth, (BD Diagnostic Systems).

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4. SOC medium (Quality Biological, Inc.). 5. 10% glycerol, sterilized and stored at 4°C. 6. Gene Pulser apparatus and cuvettes (0.2 cm electrode gap) (Bio-Rad, Hercules, CA). 2.2. Culture of Cell Lines, Cell Transfection, and Stable Cell Line Establishment

1. The human prostate cancer cell line PC-3M (29) (Xenogen Corporation) (see Note 1). 2. Clonal HeLa cell line that stably expresses the cycle 3 variant of GFP introduced via Invitrogen’s pTracer SV40 vector, which will be used to demonstrate the reduction of GFP expression after transfection with a plasmid containing a GFP-siRNA insert. 3. Iscove’s modified Dulbecco’s medium (IMDM) (Invitrogen). 4. Fetal bovine serum (FBS) (Invitrogen). 5. LipofectAMINE 2000 (Invitrogen). 6. Green fluorescent protein vector (pGCsiU6/Neo/GFP, Jikai Chemical, Inc). 7. Gentamicin, neomycin, tetracycline, penicillin, streptomycin (Sigma) and G418 sulfate (InvivoGen, San Diego, CA). 8. Trypsin/EDTA solution (Invitrogen). 9. Opti-MEM I medium (Invitrogen). 10. Tissue culture plate with 6, 24, or 96-well cluster flat bottom with lid (Corning Inc., NY). 11. 75 cm2 cell culture flask (Corning Inc., NY). 12. 5% CO2 humidified incubator.

2.3. Construction and Expression of shRNA and shRNA-GFP Expression Plasmids

1. Construct two complementary oligonucleotides containing target-siRNA-sequence referring to the principles described in Sect. 1.1. The target sequence, GCAGCAGCTGAACAACATG, corresponds to the nucleotides 2,144– 2,162 of the TAD domain of oncogene Stat3 (GenBank accession no NM_003150). The two complementary oligo DNAs are the sense strand 5′-GATCCGCAGCAGCTG AACAACATG TTCAAGAGA CATGTTGTTCAGCTGCTGCTTTTTGGAAA-3′, and the antisense strand 3′-CGTCGTCGACTTGT-TGTACAAGTTCTCTGTACAACA AGTCGACGACGAAAAACCTTTTCGA-5′. After annealing, the DNA sequence should contain the sense strand, a short spacer (loop shown in bold), the antisense strand, five Ts (Terminator), and BamHI and HindIII cutting sites (see Fig. 8.1). 2. The pGCsiU6/Neo/GFP vector contains a U6 promoter, polycloning sites, and the CMV promoter controlling the GFP gene (Jikai Chemical, Inc) (see Note 3)

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3. The pSilencer 2.1-U6 neo siRNA Expression Vector Kit contains the scramble negative control, the linearized pSilencer 2.1-U6 neo vector, and the GFP-siRNA control insert for GFP-expression knockdown. 4. Molecular Grade Water (DNase, RNase, and Protease-free) (Mediatech, Inc.). 5. 1 X DNA annealing solution: 0.1 M potassium acetate, 30 mM HEPES-KOH, pH 7.4. 6. T4 DNA ligase, 10 X T4 DNA ligase buffer (Roche). 7. Restriction enzymes: BamHI, HindIII (Roche). 8. 1 X TE: 10 mM Tris, 1 mM EDTA, pH 7.4. 9. Wizard Minipreps DNA Purification Systems (Promega). 10. ProbeQuant G-50 Micro Columns (Amersham Biosciences). 2.4. Titration and Distribution of Bacteria in Mouse Tumor Model

1. Luria-Bertani Broth (LB broth) and brain heart infusion (BHI broth) (BD Diagnostic Systems). 2. Waring Blender (Fisher Scientific). 3. Stirrer Motor hom*ogenizer with electronic speed controller (Cole Parmer). 4. Ultraviolet–visible spectrophotometer (Shimadzu, Japan). 5. Fluorescence microscope (Olympus, Japan). 6. Flow cytometer (Coulter, Hialeah, FL). 7. Frosted glass slides (Colorfrost/Plus—Fisher Scientific Co). 8. Tissue-Tek V.I.P (Electron Microscopy Science). 9. Sakura Tissue-Tek OCT Compound (International Equipment Inc). 10. 1 X PBS, pH 7.4, cold.

2.5. Northern Blot Assays for RNAi Effect Analysis 2.5.1. Isolation of Total RNA for Northern Blot

1. TRIzol Reagent (Invitrogen). 2. RNase-free deionized water. 3. Chloroform (Invitrogen). 4. 100% isopropanol. 5. 70% and 100% ethanol. 6. 3 M sodium acetate. 7. 1 X TE buffer: 10 mM Tris, 1 mM EDTA, pH 7.4. 8. MOPS (10 X) running buffer: 0.4 M morpholinopropanesulfonic acid, 0.1 mM. 9. Sodium acetate, 10 mM EDTA, adjust to pH 7.0 with 2 N NaOH. And store at room temperature protected from light.

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10. Transfer buffer (Ambion, NorthernMax Kit) (cat #1940). 11. ULTRAhyb buffer (Ambion). 12. Formaldehyde gel-loading buffer: 50% glycerol, 1 mM EDTA, 0.25% bromophenol blue, 0.25% xylene cyanol, 60 µg/ml ethidium bromide (EB). 13. 1 and 1.2% agarose gels (Sigma). 14. Hybond-N + membrane (Amersham Pharmacia Biotech, Piscataway, NJ). 15. α-32P dATP (6000 Ci/mmol) (Amersham). 16. X-ray film (Kodak). 2.6. Isolation for Small RNA and Detection of shRNAsby Northern Blot 2.6.1. Isolation of Small RNA from Total RNA Samples

1. mirVana miRNA isolation kit (Ambion) (contains miRNA wash solution I, wash solution 2/3, collection tubes, filter cartridges, lysis/binding buffer, miRNA hom*ogenate additive, acid-phenol: chloroform, gel loading buffer II, elution solution). 2. RNase-free 1.5 or 0.5 ml microfuge tubes. 3. 100% Ethanol (ACS grade) (Invitrogen). 4. Microcentrifuge capable of at least 10,000× g. 5. RNase-free water (Mediatec Inc. Herndon, VA).

2.6.2. Detection of shRNA by Northern Blot

1. 10 X TBE for 1 L: 109 g of Tris-base, 55 g of boric acid, 40 ml for 0.5 M EDTA (final concentration: 0.9 M, 0.9 M, and 20 mM, respectively). 2. BrightStar-plus nylon membrane (Ambion cat # 10100). 3. 50 X Denhardt’s solution: 10 g Ficoll 400, 10 g bovine serum albumin, 10 g polyvinylpyrrolidone, add nucleasefree water to 1 L. 4. 20 X SSC: 175.3 g NaCl, 88.2 g sodium citrate, 800 ml nuclease-free water, adjust to pH 7.0 with Tris, and then add nuclease-free water to 1 L. 5. Prehybridization solution: 6 X SSC, 10 X Denhardt’s solution, 0.2% SDS. 6. Hybridization solution: 6 X SSC, 5 × Denhardt’s solution, 1–5 × 106 cpm 5′end-labeled antisense probes, 0.2% SDS. 7. Wash solution: 6 x SSC, 0.2% SDS. 8. 15% denaturing polycrylamide gel (see Ambion’s reagents (www.ambion.com). 9. Novex mini-cell system (XCell II) (Invitrogen). 10. Oligonucleotide (refer to Sect. 2.3) containing 19-nt complementary to the antisense of target siRNA.

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11. T4 polynucleotide kinase (Takara Shuzo Co., Kyoto, Japan). 12. γ32P-ATP (6000 Ci/mmol) (Amersham). 13. PhosphorImager (Molecular Dynamics) or x-ray film (Kodark). 2.7. RT-PCR Assay for RNAi Effect Analysis at mRNA Level(see Note 2) 2.7.1. Total RNA Isolation (see Sect. 2.5.1)

We utilized the Superscript first-strand synthesis system for RTPCR (Invitrogen). 1. The Superscript first-strand synthesis system kit (Cat #11904018, Invitrogen)—this kit provides all components for first strand cDNA synthesis. 2. Ex-Taq DNA polymerase (Takara).

2.7.2. Reverse Transcription and PCR Amplification of cDNA

3. Two specific amplification primers selected from vector sites flanking the target mRNA sequence.

2.8. Western Blot Assay for the RNAi Effects at the Protein Level

1. Blocking solution (10% nonfat dry milk powder in TBST, pH 7.4).

4. Human β-actin primers and probes were obtained from Applied Biosystems.

2. TBST (pH 7.4): 0.2% Tween 20, 20 mM Tris-HCl, 150 mM NaCl. 3. Running buffer for Tris-glycine gels (10 X): 250 mM Tris base, 1.92 M glycine, 1%. 4. Sample buffer (2 X): 125 mM Tris-HCl, pH 6.8, 4% SDS, 20% glycerol, 0.1% bromophenol blue, 5% ß-ME* (2-mercaptoethanol; omit for native proteins). 5. Transfer buffer (for PVDF membrane): Add 18.2 g Tris base, 86.5 g glycine to 4 l of H2O. Add 1200 ml methanol and bring to 6 l with H2O (pH should be ~8.3–8.4). 6. Protein samples prepared from siRNA-treated cultured cell lines or tissue specimens from mouse model. 7. Primary antibody specific for protein of interest diluted in TBST buffer. 8. Secondary antibody: Horseradish peroxidase (HRP)- or alkaline phosphatase (AP)-anti-Ig conjugate (Santa Cruz, CA). 9. PBS (pH 7.1): 137 mM NaCl, 7 mM Na2HPO4, 1.5 mM KH2PO4, 2.7 mM KCl. 10. Trypsin-EDTA solution (Invitrogen). 11. 2 X Laemmli SDS sample buffer (Bio-Rad). 12. Lysis buffer: 1 M Tris-HCl, 0.5 M NaCl, 0.5 M EDTA, 1 mM phenylmethylsulfonyl fluoride, and 10 µg/ml each of aprotinin, pepstatin, and leupeptin (Sigma). 13. PVDF (Millipore Immunbilon P).

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14. ECL (enhanced chemiluminescent detection kit (Amersham). 15. Kodak X-OMAT (XAR-5, 18 × 24 cm). 16. Novex mini-cell system (XCell II) (Invitrogen) 17. 12–16% Tris-glycine gel (Pre-Cast, Noevx, Invitrogen) 2.9. Animal Model, Tumor Orthotopic Implantation, and Administration of Bacteria-Carrying siRNA-Expression Plasmid

1. 6-week-old male BALB/c nude mice weighing 18–24 g were raised in specific-pathogen free (SPF), and under controlled conditions of temperature (23 ± 3°C) and relative humidity (50 ± 20)%. 2. Dissecting microscope (Model MZ6, Leica, Nussloch, Germany). 3. Latex-free syringe (1–3 ml) and hypodermic needle (22– 30G) (Becton Dickinson). 4. Isopropyl alcohol. 5. Ketamine hydrochloride (100 mg/ml) and xylazine (20 mg/ml) at a mixture ratio of 7:3, respectively. 6. Surgical tools such as forceps and scissors. 7. Stainless steel gavage tube.

3. Methods 3.1. Construction of shRNA and shRNAGFP Expression Vectors 3.1.1. Cloning of the siRNA Target Insert into Vector

1. Two complementary oligonucleotides containing the sense and antisense siRNA sequences and other required elements to express shRNA (see Sects. 1.1 and 2.3) were synthesized and column-purified using ProbeQuant G-50 Micro Columns (Amersham). The oligonucleotides were designed with BamHI and HindIII sites at opposite ends (see Fig. 8.1). 2. Prepare a 1 µg/µl solution of each single-strand oligonucleotide in 1 X TE. 3. Anneal the two complementary template oligonucleotides. Assemble the annealing mixture as follows: 2 µl of sense siRNA template, 2 µl of antisense siRNA template oligonucleotides, and 46 µl 1 X DNA annealing buffer. Heat the mixture at 90°C for 3 min, then anneal at 37°C for 1 h. 4. Prepare the pGCsiU6/Neo/GFP backbone vector by digestion with BamHI and HindIII. (see Note 3). 5. Set up a ligation reaction containing insert double-stranded oligo-DNA-template and pGCsiU6/GFP vector-digested with BamHI and HindIII.

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6. Set up the positive control ligation reaction containing GFPsiRNA control insert, and linearized pSilencer 2.1-U6 neo vector using standard protocols, following manufacturer’s instructions (Ambion, pSilencer neo cat #5764). 3.1.2. Prepare DNA-Uptake Competent Cells of E. coli DH5α and S. typhimurium phoP/phoQ

1. The bacteria were routinely grown in 100 ml LB at 37°C to an OD600 of ~0.2–0.6. Bacterial cells should be collected in early to mid-log phase, and therefore the growth time will vary by strain and growth conditions. 2. Chill cells on ice, transfer the cells to prechilled centrifuge tubes, and centrifuge in cold conditions at 4,000× g for 4 min. 3. Resuspend the pellets in 50 ml of ice-cold sterile water and centrifuge as above. 4. Resuspend the cell pellet in a total of 2 ml of ice-cold sterile 10% glycerol in water, transfer to two 1.5 ml microcentrifuge tubes, and then centrifuge at 2,500× g for 5 min at 4°C. 5. Resuspend the cells in ~1 ml of ice-cold sterile 10% glycerol in water. The cell concentration should be ~2 × 1010 cfu bacteria/ml. 6. Aliquot the cell suspension in 0.1 ml quantities into prechilled sterile microcentrifuge tubes, freeze in a dry ice/ethanol bath, and store at −70°C until use.

3.1.3. Transform Competent Cells with Ligation Products by Electroporation

1. Prechill all tubes and electroporation cuvettes on ice, and mix 40 µl of competent cells with 1 µl of ligation products (do not exceed 1 µl of the ligation products if the ligation products were not desalted). Mix well, but gently, and incubate on ice for ~1 min. 2. Transfer the cell–DNA mixture to a cold 0.2 cm cuvette, shake to the bottom, and place in the Gene Pulser cuvette holder. 3. Apply one pulse with the Gene Pulser set at 2.5 kV and 25 uF and the pulse controller at 200 Ω. If using a 0.1 cm cuvette, set the apparatus at 1.5–1.8 kV. 4. Immediately suspend the electroporated cells in 1–2 ml of SOC, and incubate at 37°C with agitation for 1 h. 5. Plate the cells on selective agar growth medium (containing ampicillin at 50 µg/ml) in a volume of 100–200 µl to select for cells containing the vector plasmid.

3.1.4. Identify Positive Colonies and Prepare Plasmid DNA

1. Select at least 10 positive colonies (i.e., containing target insert sequence with vector DNA) grown on the selective agar plates. Grow culture in LB broth overnight. Prepare plasmid DNA

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using the standard method (Promega Mini-Prep), and check plasmid DNA for size on 1.0% agarose gel. 2. Digest the plasmid DNA with BamHI and HindIII, and check the digestion products on a 2% agarose gel. The target DNA fragments should be ~60–70 bp. 3. Confirm the insert DNA sequences: sequencing should be conducted using standard methods (ready reaction dideoxy terminator cycle sequencing kits (Applied Biosystems)). PCR primers for amplifying DNA fragments containing inserts depend upon the primer designed binding sites denoted by arrows on the plasmid map (Fig. 8.1). 3.1.5. Positive and Negative Controls

1. We chose the pSilencer neo with GFP-siRNA control insert as a positive targeting sequence, and a clonal HeLa cell line stably expressing the cycle 3 variant of GFP as controls to monitor GFP expression knockdown. 2. We utilize the pSilencer neo negative control plasmid containing scrambled sequences which are not found in the mouse, human, or rat genome databases. 3. To assess siRNA efficacy, cells transfected with the pSilencer neo plasmid expressing target-specific siRNA should be compared to cells transfected with the scrambled vector control. Also, the GFP-specific shRNA expressed vector should be used in experiments to demonstrate the RNAi efficacy in the HeLa cell line expressing GFP protein.

3.2. Cell Culture, Cell Transfection, and Stable Cell Line Establishment 3.2.1. Cell Culture Maintenance

1. Grow the PC-3M cell line in a 5% CO2 incubator at 37°C in IMDM (supplemented with 10% FBS, 100 µg/ml penicillin, and 100 µg/ml streptomycin). 2. Passage cells every 3–4 days by detaching with trypsinEDTA solution treatment. Do not exceed 30 passages after unfreezing the stock cells. Over-density of cell layer or overpassage may affect siRNA effects. 3. At least 24 h after seeding the cells, ensure that a confluency of 60–80% is reached. Begin transfection with your siRNA components or expression plasmids.

3.2.2. Cell Transfection with shRNA-Expression Plasmid and Stable Cell Line Establishment

1. Dilute 0.8 µg of shRNA-expression vector DNA into 50 µl of Opti-MEM I medium for transfection. 2. Trypsinize 90% confluent cells grown in a 175 ml cell culture flask with 10 ml of trypsin-EDTA for 10 min at 37°C. 3. Dilute the cell suspension 1:5 with fresh medium and seed into new culture flask or culture plate without serum in the well of tissue culture plate. Mix gently.

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4. After 5 min incubation, add the diluted Lipofectamine 2000 (2 µl of Lipofectamine in 50 µl of Opti-MEM I Medium) to the wells with the diluted shRNA expression vector DNA. Mix gently and incubate for 15 min at room temperature to allow complex formation to occur. 5. Add 100 µl complete growth medium without antibiotics with 20,000 cells to each well containing shRNA expression vector-Lipofectamine 2000 complexes. Mix gently by rocking the plate back and forth. 6. Incubate the cells at 37°C in a CO2 incubator until you are ready to harvest cells. 7. 24–48 h after transfection, start to assay for target gene knockdown. 8. You may select a stable cell line which is constantly expressing the selected shRNA. 9. Cells were passaged twice weekly with fresh selection medium containing 400 µg/ml G418 (see Note 4). Within two weeks, when individual transfectant foci developed, and each of these were individually dissociated, transfer the cells to a single well (6-well dish) and culture in selection medium containing 400 µg/ml G418. 10. Expand the desired clones using minimal concentration (200–400 µg/ml) of G418. 11. After 6–8 rounds of G418 selection, culture the cells in nonselective medium, collected two times per week and preserved in liquid nitrogen. 3.2.3. Infection of PC-3M Cells with S. typhimurium Carrying shRNA Expression Vector

1. In a 24-well plate, seed PC-3M cells at a density of 0.5–2 × 105 cells per well in 500 µl IMDM with10% FBS and grow at 37°C, 5% CO2 for 24 h. 2. Prior to coculture with bacteria, the cells should be first “conditioned” in 0.5% FBS without antibiotics for 24 h. (0.5% FBS was determined as an optimum by testing the effect of a range of FBS concentration on cell viability). 3. Next day, collect the mid-log phase bacteria-carrying siRNAexpression vector by centrifugation, resuspend the bacteria at 1 × 108 cfu/ml in IMDM without antibiotic, and coculture with PC-3M cells at 37°C for 30 min. 4. After exposure, wash the cell monolayers twice with serumfree IMDM containing 50 µg/ml gentamicin to kill all extracellular bacteria. 5. Add IMDM containing 10% FBS to the cells and then incubate at 37°C for 4 h.

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6. Next, add tetracycline (10 µg/ml) to the cells and incubate at 37°C for 72 h in order to prevent intracellular bacterial multiplication. Stable PC-3M cell clones containing the target plasmids are selected with G418 (800 µg/ml, see Note 4) or screened for GFP expression using a fluorescence microscope (when the recombinant vector carried the GFP gene). The cells were maintained by treating with 200 µg/ ml G418. 7. Use the transfected cells to assess for knockdown effects on the target gene or shRNA expression levels. 3.3. Animal Model: Nude Mice Tumor Orthotopic Implantation

1. Male BALB/c nude mice are inoculated with 2 × 106 PC-3M cells subcutaneously (s.c.) into the right flank of mice. 2. After the formation of palpable tumors (~5 mm by day 12), mice are killed by cervical dislocation. 3. Tumor tissues are excised, and cut into fragments (1 mm3), and then implanted by SOI (surgical orthotopic implantation) (30) onto the prostate in nude mice. After proper exposure of the prostate to be implanted, 8–0 surgical sutures are used to penetrate the tumor pieces and attach them to the appropriate orthotopic organ. The incision in the skin was closed with a 7–0 surgical suture in one layer. 4. The animals are kept under isoflurane anesthesia during surgery. All procedures of the operation described above are performed with a 7 X magnification microscope.

3.4. Administration with Bacteria Carrying shRNA-Expressing Vector(see Note 5) 3.4.1. Oral Administration

1. Mice are fasted for 3–4 h without food and water. 2. Mice are pretreated with 100 µl of 3% sodium bicarbonate in PBS orally. 3. Immediately following buffer administration, bacteria are administered orally using a stainless steel gavage tube,100 µl (2 × 109 cfu) of bacteria carrying the siRNA plasmid into the lower esophagus. 4. After treatment, mice are given food and drinking water ad lib.

3.4.2. Intravenous Inoculation

1. Prep tail with alcohol swab. 2. Needle placement should be no closer to the body than half the length of the tail. 3. With the tail under tension, insert needle approximately parallel to the vein. 4. Insure proper placement by inserting needle at least 3 mm into the lumen of vein. 5. Administer 100 µl bacterial suspension (i.e., 2 × 107 cfu) in PBS into the tail vein using a tuberculin syringe fitted with a 25-gauge needle in a slow, fluid motion to avoid rupture of vessel.

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1. Restrain mouse by grasping near base of tail; grasp nape of neck with the opposite hand. 2. Place tail between fingers to secure and control animal; prep the local area with alcohol swab. 3. Inoculate 0.5 ml bacterial suspension in PBS containing 5 × 105–106 cfu, using a 1 ml syringe.

3.5. Bacterial Titration and Bacterial Distribution in a Tumor Mouse Model 3.5.1. Direct CFU (Colony Forming Unit) Count

1. Inoculate a single colony of S. typhimurium carrying plasmid vectors into 5 ml LB with 50 µg/ml ampicillin and incubate at 37°C overnight. 2. Take 1 ml of the overnight culture and dilute 1:50 in fresh LB broth and restart incubation. Harvest bacteria when an A600 = 0.6 is reached. 3. Collect bacteria by centrifugation for 5 min at 4,000 rpm, and wash pellet twice in 2 ml cold PBS. 4. The washed bacteria were diluted to a final concentration of 108 cfu/100 µl cold PBS (an OD600 = 1 corresponds to 1 × 109 cfu/ml). Use diluted cells as soon as possible to avoid bacterial viability loss. 5. Inject 107 cfu/100 µl of bacteria via i.v. (tail vein) into tumor-bearing mice. 6. The mice were sacrificed at different times after the injection. 7. Tumors and relative normal tissues or organs such as liver, spleen, and kidney were removed, weighed aseptically, and hom*ogenized in five volumes of ice-cold, sterile PBS. 8. Larger organs are first chopped in a sterile warring blender on ice bath (Fisher Scientific) before hom*ogenization. 9. The hom*ogenates are serially diluted in BHI broth and plated onto BHI agar with ampicillin (50 µg/ml) in triplicate, and incubated for 24 h at 37°C. 10. Colony-forming units per gram of tissue were assessed and averaged.

3.5.2. Observation Under a Fluorescence Microscope to Determine the Extent of Bacterial Infection (Cryostat Sectioning)

1. Add Tissue-Tek OCT compound to infected host cells; allow to equilibrate for at least 30 min. 2. Prepare a block using Tissue-Tek. 3. Place the tissue block from the GFP control group of mice into the cryostat and allow it to equilibrate to the cutting temperature (i.e., −17°C). Adjust the positioning of the block to align the block with the knife blade. 4. Prepare 12–14 µm thick sections using a microtome and collect onto frosted glass slides. Observe immediately under a fluorescence microscope (see Fig. 8.2)

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tumor liver spleen

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Fig. 8.2. GFP-based detection for quantitation and distribution of 57 vector carrying different recombinant constructs based upon the pGCsi-U6/Neo/GFP vector in different samples. (a) Quantification of bacterial foci at specified times (hours or days) in the various tissues of tumor bearing mice (n = 3). (b) Localization of attenuated S. typhimurium in C57BL6 mice bearing RM-1 prostate tumor by using GFP expression as a marker. (c) Expression of GFP from the psiStat3-GFP and psi-Scrambled vector is shown in stable infected RM-1 cells versus mock uninfected cells (magnification 400 X).

3.5.3. Detection of Transfected Cells by Fluorescence-Activated Cell Sorting (FACS)

1. hom*ogenize the treated tumor tissues and dilute with cold PBS (five times volume/sample weight, (w/v)). 2. Adjust to 1 × 107 host cells/ml with PBS after lysis of the red blood cells with 0.85%NHCl4. 3. GFP-expressing cells are detected by flow cytometry.

3.6. Northern Blot Assays for Assessing RNAi effect at the mRNA level 3.6.1. Isolation of Total RNA

The RNA extraction protocol is designed to isolate total RNA from mammalian cells or tissue samples (e.g., frozen tissue specimens) from tumors. 1. If the samples are from cells grown in a monolayer, aspirate and discard the culture medium, wash the cells with PBS, place the culture dish on ice, and add directly 1 ml TRIzol reagent to a 3.5 cm diameter dish, and mix well. 2. If the samples are fresh tissues or frozen specimens from animals, perfuse the tissue with cold PBS and rinse one time to eliminate red blood cells. If necessary, quickly cut the tissue into small pieces, weigh the sample, add 1 ml of TRIzol reagent per 50–100 mg of tissue immediately, and hom*ogenize

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on ice. If you need to store the tissues, freeze the samples in liquid nitrogen first. When the liquid nitrogen stops churning, remove the samples from the liquid nitrogen and store them at −70°C. 3. Incubate the cell lysate or hom*ogenate for 5 min at room temperature. 4. Add 0.2 ml chloroform per ml of TRIzol. 5. Mix by shaking the tubes vigorously by hand for 15 s. 6. Incubate at room temperature for 2–3 min. 7. Centrifuge the sample at 12,000× g for 15 min at 4°C. 8. Transfer the colorless aqueous phase to a fresh tube. 9. Add 0.5 ml isopropyl alcohol per 1 ml TRIzol, and vortex. 10. Incubate at room temperature for 10 min. 11. Centrifuge the sample at 12,000× g for 10 min at 4°C. 12. Remove supernatant carefully without disturbing the RNA pellet. 13. Wash the RNA pellet once with 75% ethanol, adding at least 1 ml of ethanol per ml TRIzol, mix well by vortexing. 14. Air-dry the RNA pellet for 5–10 min. 15. Dissolve RNA in RNase-free water. 16. Read the OD260 of 1/500 dilution: 1 OD260 = 35 µg/ml. 17. Store immediately at −80°C. 3.6.2. Northern Blotting

1. Prepare a 1.2% agarose gel. Mix 1.2 g of agarose with 72 ml of water, and heat until boiling. Cool to 65°C and add 10 ml of 10 X running buffer and 18 ml of 37% formaldehyde. Mix and pour into gel box. 2. Prepare the RNA samples as follows: Mix the following to each RNA sample a. RNA (up to 30 µg) 11 µl b.

10 X MOPS running buffer 5 µl

Formamide 25 µl

d. Formaldehyde (12.3 M) 9 µl 3. Denature the RNA by heating at 65°C for 10 min. 4. After chilling on ice, add 10 µl formaldehyde loading dye. 5. Load the samples, and electrophorese in a fume hood at 100 V until the blue dye has migrated to two-thirds of the length of the gel. 6. Visualize the gel on a UV transilluminator and photograph the gel prior to transfer. 7. Measure the migration of the markers from the origin.

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8. Transfer the RNA onto Hybond-N + membrane (or other positively charged nylon membrane). To remove formaldehyde, the gel must be rinsed several times in RNase-free water. 9. The denatured RNA may be transferred immediately to the nylon membrane by capillary elution or electroblotting. Capillary elution is conducted following the description by Sambrook et al. (33) and electroblotting should be performed according to the instructions of the apparatus manufacturer. Usually, transfer for a typical 6 mm thick gel should be completed in 1.5–2 h, and does not exceed 4 h to avoid the hydrolysis of small RNAs. UV-crosslink the RNA to the membrane. 10. Prehybridization: treat the membrane for 1–2 h at 42°C in 50% formamide, 5 X SSPE, 2 X Denhardt’s reagent, and 0.1%SDS. 11. Probe design and labeling: the oligonucleotide probe should be designed to be hom*ologous to the antisense strand of the cDNA sequence. Radiolabel the oligonucleotide with 6000 Ci/mmol of α-32P dATP according to the manufacturer’s instructions. 12. Hybridization: add the denatured radiolabeled probe directly to the prehybridization fluid, and incubate for 16–24 h at 42°C. 13. Wash the filter for 30 min at room temperature in 1 X SSC, 0.1% SDS, followed by three washes of 30 min at 68°C in 0.2 X SSC and 0.1% SDS. 14. Expose the filter to x-ray film or a phosphorimager. Quantify signals with the phosphorimager software. 3.7. Detection of shRNAsby Northern Blot(see Note 6)

Determination of the siRNA expression levels is critical in the assessment of stability or half-life of siRNA after transfection in mammalian cells. A DNA (or RNA) oligonucleotide complementary to the antisense of the siRNA or target-mRNA sequence can be synthesized and used to probe the expression of siRNA. Usually, small RNAs are separated by 12.5–15% polyacrylamide gels. Northern blots can also be employed to detect the larger precursor of siRNAs (28, 29).

3.7.1. Isolation of Enriched Small RNAs

The procedure for isolation of small RNAs is adapted from the mirVana miRNA Isolation Kit manual. This procedure is designed for small scale RNA isolation from animal tissues or cultured cells. It can be also used with fresh or frozen cultured cells or animal tissues. The small RNA fraction can be enriched from the total RNA preparation.

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1 . Mix 50–100 µg of total RNA extracted using TRIzol reagent, with 5 volumes of lysis/binding buffer. 2. Add 1/10 volume of miRNA hom*ogenate additive to RNA mixture, mix well by vortexing, and leave the mixture on ice for 10 min. 3. Add 1/3 volume of 100% ethanol to the RNA mixture and mix by vortexing. 4. For each sample, place a filter cartridge into one of the collection tubes supplied by the Ambion’s kit. Pipet the RNA/ ethanol mixture onto the filter cartridge. Up to 700 µl can be applied to a filter cartridge at a time. 5. Centrifuge for 1 min at 5,000× g to pass the mixture through the filter. 6. Collect the filtrate, add 2/3 volume of room temperature 100% ethanol to the filtrate, and mix well. 7. Pipet the filtrate/ethanol mixture onto a second filter cartridge. 8. Centrifuge for 1 min (at 5,000× g), and discard the flowthrough. Reuse the collection tube for the washing steps. 9. Add 700 µl miRNA wash solution 1 to the filter cartridge and centrifuge for ~1 min at 5,000× g. 10. Discard the flow-through from the collection tube, and place the filter cartridge in the same collection tube. 11. Wash twice with 500 µl wash solution. 12. Discard the flow-through, replace the filter cartridge in the same collection tube, and spin for 1 min at 10,000× g to remove residual fluid from the filter. 13. Transfer the filter cartridge into a fresh collection tube, and apply 50 µl of 95°C elution solution, and close the tube. 14. Incubate at room temperature for 2 min and then spin for 1 min at 10,000× g to recover the RNA. Collect the eluate, which contains the small RNA-enriched fraction, and store it at −20°C. Repeat once the elution step in order to increase the RNA yield. 3.7.2. Separation on 15% Polyacrylamide Gel

1. Prepare denaturing 15% polyacrylamide gel as follows. Prepare 15 ml of 15% polyacrylamide gel mix with 8 M urea for use in the Bio-Rad Protean II minigel system. Use Ambion’s reagents for preparation of the gel as follows: Mix following reagents first: 7.2 g Urea, 1.5 ml of 10 X TBE, 5.6 ml 40% acrylamide (acryl:bis:acryl = 19.1), and add nuclease-free water to 15 ml. Stir to mix. Then add 75 µl 10% ammonium persulfate, and 15 µl TEMED. Mix briefly and pour the gel immediately.

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2. Mix 1–2 µg RNA with equal volume of gel loading buffer II. 3. Heat the sample for 2–5 min at 95–100°C, load the sample on the gel, and run at 30–45 mA. 4. Stop running when the bromophenol blue dye front has migrated to the bottom of the gel. 5. Soak the gel for 5 min in 0.5 µg/ml of EB solution in 1 X TBE. Wash the gel for 2–5 min in 1 X TBE buffer. 6. Visualize the RNA band using a UV transilluminator and collect the image on film or computer. 3.7.3. Detection of shRNA by Northern Blot

1. Run sample on gel (see Sect. 3.7.2) 2. After staining, transfer the RNA to a nylon membrane by electroblotting at 200 mA. Keep the membrane damp after blotting. 3. UV-crosslink the RNA to the membrane. 4. Specific DNA probe containing antisense shRNA target sequence should be designed and synthesized prior to the experiments. Use Ambion’s mirVana probe and marker kit for the 5′ end labeling (www.ambion.com). 5. Prehybridize for at least 1 h at 65°C. 6. Hybridize 8–24 h at room temperature. 7. Wash three times with wash solution at room temperature and once at 42°C. 8. Expose to x-ray film or a phosphorimager according to the manufacturer’s instructions.

3.8. RT-PCR for Quantification of Specific Target RNA(see Note 2)

3.8.1. Reverse Transcription

This procedure is adapted from the Invitrogen SuperScript FirstStrand Synthesis System for the RT-PCR manual. The procedures of RT-PCR involve (1) the isolation of total RNA or mRNA from the cultured cells or animal tissue sample, (2) first strand cDNA synthesis using poly dT, random hexamer, or gene-specific primers, (3) removal of complementary template RNA, and (4) amplification of target gene cDNA by PCR using gene-specific primers. For efficient measurement of gene silencing, the positions of the designed primers should flank the sites of cleavage of target mRNA. 1. Prepare RNA/primer mixture for first-strand cDNA synthesis: 3 µg/(1–4 µl) of total RNA (isolated from the samplestreated with your siRNA including control), 1 µl of random hexamers (50 ng/µl), 1 µl of 10 mM dNTP mix, and DEPCtreated water to 10 µl. Mix and spin the tube briefly. Incubate at 65°C for 5 min.

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2. Add the following: 2 µl of 10× RT buffer, 4 µl of 25 mM MgCl2, 2 µl of 0.1 M DTT, and 1 µl of RNaseout recombinant ribonuclease inhibitor. 3. Mix and spin briefly. 4. Incubate at 25°C for 2 min. 5. Add 1 µl (50 units) of SuperScript II RT. 6. Incubate at 25°C for 10 min. 7. Transfer the tube to 42°C and incubate for 50 min. 8. Stop reaction at 70°C for 15 min. Chill on ice. 9. Spin briefly and add 1 µl of RNase H and incubate at 37°C for 20 min. 3.8.2. PCR Amplification of the Target cDNA

1. Prepare PCR master mixture on ice: 5 µl of 10 X PCR buffer, 1.5 µl of 50 mM MgCl2, 1 µl of 10mM dNTP mix, 1 µl of 10 µM sense primer, 1 µl of 10 µM of antisense primer, 0.5 µl of Taq DNA polymerase (5 units/µl), 2 µl of cDNA from the first-strand reaction, 38.1 µl of autoclaved, and distilled water to 50 µl of final volume. 2. Run the following program: 94°C for 2 min, then perform 20–35 cycles of PCR with optimized conditions for the sample. 3. Analyze 10 µl of the amplified sample by agarose gel. 4. The bands can be quantified with ImageQuant 5.0 software (Molecular Dynamics).

3.9. Western Blot 3.9.1. Lysis of Sample Containing Target Protein from Cultured Cells

This procedure is adapted for minigels using the Bio-Rad Mini Trans-Blot cell for electrophoresis and electroblotting steps. 1. Remove the tissue culture medium from the siRNA-treated cells cultured in a 24-well plate. 2. Rinse the cells once with 200 µl PBS, add 200 µl trypsin– EDTA, and incubate for 1 min at 37°C. Subsequently, add 800 µl DMEM medium to inactivate the trypsin. 3. Transfer the suspended cells to a chilled 1.5 ml microfuge tube and collect the cells by centrifugation at 3,000 rpm (700× g) for 4 min at 4°C. 4. Resuspend the cell pellet in ice-cold PBS and centrifuge again. 5. Remove the supernatant and add 25 µl of hot (90°C) 2 X concentrated Laemmli sample buffer to each cell pellet obtained from one well of a 24-well plate. 6. Incubate the samples for 3 min in a boiling water bath and vortex.

3.9.2. Extracting Proteins from Animal Tissues

1. Remove tissues from target organs; weigh and hom*ogenize at 4°C in lysis buffer containing protein inhibitors.

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2. Centrifuge lysate at 15,000× g for 30 min at 4°C. 3. Determine the protein content using the Bradford reagent (Bio-Rad). 3.9.3. Protein Analysis by SDS-PAGE Gels

1. Aliquot 15 µl of each protein lysate into a microcentrifuge tube, add 15 µl of 2 X SDS sample buffer, heat at 100°C for 3 min, and chill and load 10–15 µl into 12.5–15% SDSPAGE. 2. Electrophorese at 15 to 20 mA until the tracking dye reaches the end of the gel (or 3 to 5 mA overnight). 3. Cut the membrane to same size as the gel plus extra 1–2 mm on each edge. 4. For PVDF membranes, first incubate the membrane for 1–2 s in 100% methanol, then equilibrate 10–15 min with the transfer buffer. 5. Electrophoretically transfer the proteins to the membrane for 30–60 min at 100 V at 4°C (or overnight at 14 V). 6. Place the membrane in heat-sealable plastic bag with 5 ml blocking buffer and seal bag. Incubate 30–60 min at room temperature with agitation. 7. Incubate the membrane with diluted primary antibody in blocking buffer (usually 1: 50–1000) for 30 min at room temperature with constant agitation. 8. Wash the membrane four times by agitating with 200 ml TBST, 10–15 min each wash. 9. Incubate the membrane with HRP- or AP-conjugated secondary antibody diluted in blocking buffer (1:10,000), and incubate for 30–60 min at room temperature with constant agitation. 10. Remove the membrane from bag and wash as in Step 8. 11. Develop the membrane with ECL according to the manufacturer’s instructions.

4. Notes 1. PC-3M cell line was isolated from liver metastases produced in nude mice subsequent to intrasplenic injection of the androgen-insensitive PC-3 human prostate carcinoma cell line. It has enhanced tumorigenicity and produces a high incidence of well disseminated metastases (29).

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2. RT-PCR (reverse transcription-polymerase chain reaction) is the most sensitive technique for mRNA detection and quantitation currently available. RT-PCR can be used with as little as 1 ng or as much as 5 µg of total RNA. RT-PCR is more sensitive compared to Northern technique when studying the efficacy siRNAs. 3. pGCsiU6/Neo/GFP vector contains U6 promoter, polycloning sites (BamHI and HindIII, CMV promoter-driven GFP protein expression, which can be used to assess transformed bacteria. 4. G418 is used for the selection and maintenance of eukaryotic cells expressing the neoR gene. G418 is an aminoglycoside antibiotic similar in structure to gentamicin B1, produced by Micromonospora rhodorangea. Unlike gentamicin, G418 blocks polypeptide synthesis in eukaryotic cells by binding irreversibly to 80S ribosomes. Resistance to G418 is conferred by the neoR gene from transposon Tn5 encoding an aminoglycoside 3″-phosphotransferase, APH 3′ II. This protein inactivates G418 by covalently modifying its amino or hydroxyl function therefore inhibiting the antibioticribosome interaction (31, 32). The concentration of G418 used for the selection varies with the cell type. Thus, working concentration should be determined for each cell type. G418 is a hazardous compound. Avoid contact with eyes, skin and clothes, harmful if swallowed. 5. Bacterial dose administered to animals varies and depends on the mouse type, age and weight, bacterial strain, and method of delivery. 6. Currently, two methods are available for shRNA detection: Northern blot on solid support using denaturing 15% polyacrylamide gel or ribonuclease protection assay (RPA). Ambion recommends the RPA using mirVana miRNA Detection kit, and indicates that RPA is 100–500 times more sensitive than Northern analysis. Please refer to Ambion Catalog 2005–2006, page 105–109, and the mirVana miRNA Isolation Kit, and the mirVana Probe and Marker kit. The mirVana miRNA Detection Kit provides a fast and sensitive method for detecting small RNAs. References 1. Hannon, G.J. (2002). RNA interference. Nature 418, 244–251. 2. Antoszczyk, S., Taira, K., and Kato, Y. (2006). Correlation of structure and activity of short hairpin RNA. Nucleic Acids Symp Ser (Oxf) 50, 295–296.

3. Hiroaki, K.H. and Taira, K. (2003). Short hairpin type of dsRNAs that are controlled by tRNAVal promoter significantly induce RNAi-mediated gene silencing in the cytoplasm of human cells. Nucleic Acids Res. 31, 700–707.

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4. Zhang, L., Gao, L., Guo, B., et al. (2007). Intratumoral delivery and suppression of prostate tumor growth by attenuated Salmonella enterica serovar Typhimurium carrying plasmid- based siRNAs. Cancer Res. 67, 5859–5864. 5. Bermudes, D., Zheng, L.M., and King, L.C. (2002). Live bacteria as anticancer agents and tumor-selective protein deliver vectors. Curr. Opin. Drug. Discov. Devel. 5, 194–199. 6. Tjuvajev, J., Blasberg, R., Luo, X., et al. (2001). Salmonella-based tumor-targeted cancer therapy: tumor amplified protein expression therapy (TAPET) for diagnostic imaging. J. Control Release 74, 313–315. 7. Zheng, L., Luo, X., Feng, M., et al. (2000). Tumor amplified protein expression therapy: Salmonella as a tumor-selective protein delivery vector. Oncol. Res. 12, 127–135. 8. Forbes, N.S., Munn, L.L., f*ckumura, D., et al. (2003). Sparse initial entrapment of systemically injected Salmonella typhimurium leads to heterogeneous accumulation within tumors. Cancer Res. 63, 5188–5193. 9. Pawelek, J.M., Low, K.B., and Bermudes, D. (2003). Bacteria as tumour-targeting vectors. Lancet Oncol. 4, 548–556. 10. Grosshans, H. and Slack, F.J. (2002). Micro-RNAs: small is plentiful. J. Cell Biol. 156, 17–21. 11. Amarzguioui, M. and Prydz, H. (2004). An algorithm for selection of functional siRNA sequences. Biochem. Biophys. Res. Commun. 316, 1050–1058. 12. Paul, C.P., Good, P.D., Winer, I., et al. (2002). Effective expression of small interfering RNA in human cells. Nat. Biotechnol. 20, 505–508. 13. Brummelkamp, T.R., Bernards, R., and Agami, R. (2002). A system for stable expression of short interfering RNAs in mammalian cells. Science 296, 550–553. 14. Brummelkamp, T.R., Bernards, R., and Agami, R. (2002). Stable suppression of tumorigenicity by virus-mediated RNA interference. Cancer Cell 2, 243–247. 15. Lambin, P., Theys, J., Landuyt, W., et al. (1998). Colonization of Clostridium in the body is restricted to hypoxic and necrotic areas of tumours. Anaerobe 4, 183–188. 16. Luo, X., Li, Z., Lin, S., et al. (2000). Antitumor effect of VNP20009, an attenuated Salmonella in murine tumor models. Ocol. Res. 12, 501–508. 17. Jazowiecka-Rakus, J. and Szala, S. (2004). Antitumour activity of Salmonella typh-

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imurium VNP20047 in B16(F10) murine melanoma model. Acta Biochim. Pol. 51, 851–856. Khan, S.A., Everest, P., and Servos, S. (1998). A lethal role for lipid A in Salmonella infections. Mol. Microbiol. 29, 571–579. Low, K.B., Ittensohn, M., Le, T., et al. (1999). Lipid A mutant salmonella with suppressed virulence and TNF-α induction retain tumortargeting in vivo. Nat. Biotech. 17, 3–41. Miller, S.I., Kukral, A.M., and Mekalanos, J.J. (1989). A two-component regulatory system (phoP phoQ) controls Salmonellatyphimurium virulence. Proc. Natl. Acad. Sci. USA 86, 5054–5058. Ronson, C.W., Nixon, B.T., and Ausubel, F.M. (1987). Conserved domains in Bacterial regulatory proteins that respond to environmental stimuli. Cell 49, 579–581. Ernst, R.K., Guina, T., and Miller, S.I. (1999). How intracellular bacteria, survive: surface modifications that promote resistance to host innate immune responses. J. Infect. Dis. 179, S326–S330. VanCott, J.L., Chatfield, S.N., Roberts, M., et al. (1998). Regulation of host immune responses by modification of Salmonella virulence genes. Nat. Med. 4, 1247–1252. Guo, L., Lim, K.B., Gunn, J.S., et al. (1997). Regulation of lipid A modifications by Salmonella typhimurium virulence genes phoP-phoQ. Science 276, 250–253. Dang, L.H., Bettegowda, C., Huso, D.L., et al. (2001). Combination bacteriolytic therapy for the treatment of experimental tumors. Proc. Natl. Acad. Sci. U S A 98, 15155–15160. Low, K.B., Ittensohn, M., Luo, X., et al. (2004). Construction of VNP20009: A novel, genetically stable antibiotic-sensitive strain of tumor-targeting Salmonella for parenteral administation in humans. Methods Mol. Med. 90, 47–60. Hohmann, E.L., Oletta, C.A., Killeen, K.P., et al. (1996). phoP/phoQ-deleted Salmonella typhi (Ty800) is a safe and immunogenic single-dose typhoid fevr vaccine in volunteers. J. Infect. Dis. 173, 1948–1014. Hohmann, E.L., Oletta, C.A., and Miller, S.I. (1996). Evaluation of a phoP/phoQdeleted, aroA-deleted live oral Salmonella typhi vaccine strain in human volunteers. Vaccine 14, 19–24. Koslowski, J.M., Fidler, I.J., Campbell, D., et al. (1984). Metastatic behavior of human tumor cell lines grown in the nude mice. Cancer Res. 44, 3522–3529.

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Chapter 9 The Therapeutic Potential of LNA-Modified siRNAs: Reduction of Off-Target Effects by Chemical Modification of the siRNA Sequence Kees Fluiter, Olaf R. F. Mook, and Frank Baas Abstract Post-transcriptional gene silencing mediated by double-stranded RNA represents an evolutionarily conserved cellular mechanism. Small dsRNAs, such as microRNAs (miRNAs), are part of the main regulatory mechanisms of gene expression in cells. The possibilities of harnessing this intrinsic natural mechanism of gene silencing for therapeutic applications was opened up by the discovery by Tom Tuschl’s team a few years ago that chemically synthesized small 21-mers of double-stranded RNA (small interfering RNA, siRNA) could inhibit gene expression without induction of cellular antiviral-like responses. siRNAs are especially of interest for cancer therapeutics because they allow specific inhibition of mutated oncogenes and other genes that aid and abet the growth of cancer cells. However, recent insights make it clear that siRNA faces some major hurdles before it can be used as a drug. Some of these problems are similar to those associated with classic antisense approaches, such as lack of delivery to specific tissues (other than the liver) or tumors, while other problems are more specific for siRNA, such as stability of the RNA molecules in circulation, off-target effects, interference with the endogenous miRNA machinery, and immune responses toward dsRNA. Chemical modifications of siRNA may help prevent these unwanted side effects. Initial studies show that minimal modifications with locked nucleic acids (LNA) help to reduce most of the unwanted side effects. In this chapter we will explore the limitations and possibilities of LNA-modified siRNA that may be used in future therapeutic applications. Key words: RNA interference, siRNA, locked nucleic acid, off-target effects.

1. Introduction The discovery of gene silencing by double-stranded RNA, known as RNA interference (RNAi), is one of the biggest breakthroughs of the last decade in the life sciences. This is reflected by the M. Sioud (ed.), Methods in Molecular Biology, siRNA and miRNA Gene Silencing, vol. 487 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-60327-547-7_9

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recent Nobel Prize awarded to Andrew Fire and Craig Mello, the two scientists who discovered the basic mechanism of RNAi in C. elegans (1). As can now be found in any biology textbook, doublestranded RNA binds to a protein complex, Dicer, which cleaves it into fragments. These fragmented double RNA strands are then bound to another protein complex, RISC. The two RNA strands are then separated and one of the RNA strands is eliminated while the other remains bound to the RISC complex and serves as a probe to detect mRNA molecules. When an mRNA molecule can pair with the RNA fragment on RISC, it is bound to the RISC complex, and then RISC directs either RNA cleavage, mediate translational repression, or induce chromatin modification. RNA interference is important in the defense against viruses and limits the effects of transposons. In both cases, doublestranded RNA from a virus or a transposon is immediately cleaved by Dicer, RISC is activated, and the viral or transposon RNA is degraded (2). However, small noncoding dsRNA is now also recognized as one of the key regulators of gene expression in general. Even before the advent of whole genome sequencing we have known that the majority of the transcribed genome is actually not coding for proteins. Most of us assumed that all the noncoding RNA was just evolutionary junk piled up in the genomes. Now we start to realize that many of these junk RNAs are actually involved in the RNAi pathway to regulate gene expression (3). A conservative estimate is that thousands (and maybe many more) of noncoding microRNAs (miRNA) use the RNAi machinery to regulate gene expression, allowing very specific local regulation of mRNA levels, and thus the evolution of highly organized tissues within organisms such as the brain (4). After the discovery by Tuschl in 2001 (5) that synthetically synthesized double-stranded RNA 21-mers (siRNAs) are very effective in mediating RNAi and can be used as a research tool to study gene function in mammalian cells, siRNA technology is nowadays utilized as a “standard tool” in most labs for genefunction analysis, drug-target discovery and validation. But one of the more attractive features of siRNA is that this research tool in theory can also be used directly as a therapeutic approach. Any disease-causing gene can potentially be targeted. However, harnessing RNAi for therapy is not so straight forward as many initially have thought. There remains important obstacles for effective therapeutic use of siRNA: stability, efficacy, and specificity (see Chaps. 1 and 2). However, there is also progress and pilot siRNA clinical studies were started just 3 years after the initial discovery that RNAi can be used in mammalian cells for specific gene knockdown.

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History tends to repeat itself over and over again. Also, in this new field of RNA interference we are facing exactly the same issues as antisense oligonucleotides were facing 30 years ago. Luckily, we have learned from the past and can use many of the solutions scientists working in the antisense field have found over the last three decades. Antisense oligonucleotides are normally defined as short, single-stranded DNA (-like) molecules that bind to mRNA through Watson–Crick base paring and then inhibit translation either through a steric block of the translation machinery or more effectively by recruiting an intracellular enzyme called RNase H, which recognizes DNA–RNA hybrids and will cleave the RNA in such a DNA–RNA hybrid (reviewed in Ref. 6). This is very hom*ologous with RNAi since both processes use short nucleotide sequences, use Watson–Crick base pairing to capture their target mRNAs, and both use intrinsic RNase activity in the cell to degrade the target mRNAs. Also hom*ologous is the first hurdle both approaches face when used as a therapeutic molecule: stability. When the first antisense experiments were performed over three decades ago it quickly became clear that nature was equipped with formidable barriers that prevent foreign strands of DNA (or RNA) to muck about with gene expression within a cell. Not only it proved quite difficult to get antisense DNA inside cells, for which all kinds of transfection technologies were invented (7), but even more problematic for in vivo usage were the wide range of nucleases that just destroyed the antisense strands in circulation before they could get to the intended tissues and could find a target mRNA. Progress in DNA amadite synthesis technology provided a real breakthrough in this field allowing the synthesis of chemically modified DNA that could resist nuclease activity. However, for effective use in siRNA the chemical modifications must not hinder incorporation into RISC and the activity of RISC. So the chemistry used must mimic the function of RNA but should add increased enzymatic and chemical stability. The most promising generation of modifications which are of interest for siRNA are those involving substitutions of the 2′ position of the ribosyl ring in RNA analogues. These modifications of the sugar moiety are in general extremely resistant against nucleases and can be incorporated into RNA strands without altering the backbone conformation of the RNA, making them prime candidates for use in siRNAs. In vitro studies showed that several of these modifications are allowed in functional siRNAs (8, 9, 10, 11). Phosphorothioates (PS) (9, 11) 2′-O-methyl (2′-O-Me) (8, 10), 2′-O-allyl (8), and 2′-deoxy-fluorouridine (9, 11) modifications have been examined for potential in vivo use. Some of the modified siRNAs were found to exhibit enhanced serum stability (10) and longer duration

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of action (8). Modification of the 5′ end of the antisense strand with 2′-O-allyl (8) or chemical bloc-king of the 5′-hydroxyl group (10) resulted in a dramatic loss in activity, consistent with the proposed in vivo requirement for 5′ end phosphorylation. Also, more substantial modifications, such as total modification by 2′-OMe (9) or PS modifications of every second or all internucleoside linkages (9, 11) increased cytotoxic effects and resulted in a significant decrease or complete loss of activity.

3. LNA-Modified siRNA Modifications with conformational restricted ribosyl groups such as bridged- and locked nucleic acids may be even more promising for use in siRNAs. In locked nucleic acid (LNA) analogs, the ribose is locked with an extra methylene bridge that fixes the ribose moiety either in the C3′-endo (beta-d-LNA) or C2′-endo (alpha-l-LNA) conformation (Fig. 9.1). The beta-d-LNA modification results in significant increases in melting temperature Tm of up to several degrees per LNA residue and a very high resistance against nuclease activity. Introduction of LNA into classical antisense oligos has been shown to increase its serum stability (12). In analogy, Braasch et al showed that LNA can be used to stabilize siRNAs without losing their function (9). Recently, a systematic study on LNA-containing siRNAs has identified the number and positions of LNA molecules within the siRNA which still allow a functional siRNA (13). Incorporation of LNA molecules into siRNA significantly increased its serum stability which

Fig. 9.1. Chemical structures of the four main LNA derivatives.

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potentially favors successful in vivo applications (13). Some LNA-stabilized siRNAs were still compatible with the intracellular siRNA machinery. We have recently demonstrated that LNA-modified siRNA can mediate specific target knockdown with comparable efficacy as unmodified siRNA in vivo (14). We designed LNA-modified siRNA (Fig. 9.2) and compared their in vitro and in vivo properties with unmodified siRNA in a mouse model system using GFP as target. Our studies on LNA-modified siRNA have shown that LNA offers the means to improve the serum half-life of siRNAs significantly. The introduction of only a few LNA moieties into siRNA is enough to protect siRNA against degradation (Fig. 9.2). However, introduction of LNA molecules in siRNA can lower the efficacy in target knockdown. In general, like most other chemical modifications of siRNA LNA modifications of the antisense strand should be minimized to the 3′ overhangs. The sense strand can tolerate more modifications but there are also limitations. We found that heavily modified siRNA was not effective in GFP knockdown. Importantly, it is needed to leave the RISC cleavage site of the antisense strand unmodified (15). Although the additional modifications in the heavily modified siRNA are only present in the sense strand we still see loss of efficacy (14). Minimal modified siRNA retained its efficacy (albeit slightly lower than unmodified). Recent findings have demonstrated that RISC incorporates dsRNA and becomes active after passenger (sense) strand degradation in the RISC complex (16). This may explain why heavily modified siRNA was inactive as possibly passenger strand degradation and, therefore, RISC activation could not occur. Nevertheless, using only a few LNA modifications as in our end-modified siRNA dramatically increased its half-life in serum while its efficacy in vivo was about equal as compared with nonmodified siRNA (Fig. 9.3). This suggests that lowered efficacy as observed with in vitro assays is compensated for

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Fig. 9.2. (a) The three siRNA designs and DNA nucleotides are depicted in italic. LNA molecules are depicted in capitals. SS = sense sequence; AS = antisense sequence. (b) stability of the siRNA designs in mouse serum: unmodified siGFP, end-modified siGFP, and heavily modified siGFP were incubated in 100% fresh mouse serum and the degradation was monitored on a PAGE gel.

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Fig. 9.3. (a) In vivo efficacy of the siRNA: unmodified siGFP, end-modified siGFP, and heavily modified siGFP were administered at a dosage of 0.25 mg/kg/day via osmotic minipumps. Seven days after implantation of the pumps, eGFP fluorescence in the tumors was analyzed using whole body imaging. (b) Using Affymetrix human whole genome arrays (U133 plus 2), we examined the number of differentially regulated genes in these tumor xenografts (P < 0.05 and over 3-fold difference). The complete list of genes can be found in Mook et al. (14).

by increased in vivo stability. These characteristics potentially improve this molecule for therapeutic application but maybe more importantly we noticed that off-target effects in vivo were reduced with LNA modified siRNA in analogy with studies presented by others using 2′O-Me-modified siRNA (17).

4. Nonspecific Effects of siRNA and How Chemical Modifications may Help Prevent These?

Now that siRNA has become a common technique in the life sciences we are becoming aware that siRNA can cause several types of nonspecific effects which will hinder its application in the clinic. The best known nonspecific effect associated with RNA interference is the interferon response. When a mammalian cell encounters a double-stranded RNA it is recognized as a viral byproduct and an immune response is mounted, causing widespread regulation of gene transcription. The short length of the siRNA designs as reported first by the Tuschl group prevented the activation of the interferon response in mammalian cells (5). Many thought that with the siRNA design the nonspecific responses towards RNAi were solved. When first reported, siRNAs seemed highly specific, as a single mutation in the target site could be demonstrated to completely abolish silencing. However, it was soon demonstrated that mutations in the target region of several siRNAs did not always inhibited knockdown (18). Similar findings

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were reported in studies that indicated tolerance to wobble base pairing (19, 20). Jackson et al. (21) reported severe nonspecific effects for genes with only a stretch of 11 nucleotides of sequence similarity between the target and the guide strand of the siRNA and it became clear that there were more problems with siRNA specificity. These effects were not related to the interferon effect, as was observed for both siRNAs (22) and short hairpin RNAs (23), but in many cases were dependent on the sequence of the siRNA. In the last 2 years there is a growing understanding of the mechanisms causing the sometimes widespread differential regulation of seemingly unrelated genes. Nonspecific effects of siRNA in an animal model or patient can be caused by: 1. Sequence dependent off-target effects by either accidental incorporation of the sense (passenger) strand into RISC or partial seed region hom*ology of the siRNA towards a miRNA target. 2. Sequence independent side-effects such as interference with the miRNA machinery itself and inadvertent activation of the immune system which in vivo is quite sensitive in detecting dsRNA through RNA-sensing immunoreceptors such as Toll-like receptors. Judicious use of chemical modifications of the siRNA might help to minimize these nonspecific effects as is discussed below.

5. SequenceDependent Off-Target Effects

Microarray studies by Jackson et al. (21, 24) showed substantial off-target effects. They tested 24 siRNAs against two genes in their initial study, followed by a second study involving 6 other target genes. Importantly, they found that many of the genes that were downregulated showed sequence complementarity to the siRNAs in the 3′ UTRs of the transcripts. Moreover, the most enriched hexamers in the 3′ UTRs were complementary to nucleotides 2–7, the seed region shown to be important for microRNA (miRNA) targeting (25, 26). They also noted that similarity to hexamers in positions 1–6 and 3–8 was also highly enriched (21); this means that complementarity to the first 8 nucleotides of the 5′ end of siRNAs is most important for off-targeting. These results confirmed other reports in the literature (27–30). Therefore, it is now clear that only limited complementarity between the target and the seed region of a miRNA (nucleotides 2–7/8) is sufficient to effect downregulation of the target sequence in itself. And additional binding of sequences in the miRNA 3′ end

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can increase the probability of silencing, determining target specificity within miRNA families (31). Chemical modifications may provide a solution to circumvent off-targeting. 2′O-Me modifications of the second base of the guide strand of the siRNA were reported to reduce off-targeting (17). Importantly, this common RNA oligonucleotide modification does not affect the degree of silencing of the intended target, as could also be expected from previous reports (32). Interestingly, the 2′-O-methyl modification was much more effective at reducing the levels of off-targeting when it was added to position 2 than when it was added to position 1 of the siRNA. We have found that LNA modifications also limit the off-target effects of siRNA. But in contrast, in our design we placed the LNA only in the overhangs and in the passenger strand. Initially we were afraid that the altered Tm of LNA-modified siRNA might affect the specificity for the target sequence, i.e., that the increase in Tm conferred by the LNA moieties would increase off-target effects. To test this assumption we performed whole genome expression profiling of treated tumor xenografts in vivo. The mice that had GFP-expressing tumor xenografts were systemically treated with unmodified siRNA and minimal LNA-modified siRNA against GFP and following GFP knockdown in the tumors the expression of the whole genome was analyzed using Affymetrix microarrays. When nonmodified siRNA was administered we found that in the tumors 93 genes were differentially regulated (Fig. 9.3). In contrast when LNA end-modified siRNA was administered (which resulted in similar GFP knockdown in the tumors) only 7 genes were found to be differentially regulated and there was no overlay between the differentially regulated genes in both experiments (14). Therefore, cautious LNA modifications of the siRNA ends are sufficient to increase the biostability of the siRNA and LNA modifications limit the extent of off-target effects without affecting the on-target efficacy in vivo. Elmen et al. (13) also showed that LNA placement at the 5′ end of the passenger strand limited the off-target preference in an in vitro assay. The reduction in off-target effects using chemically modified siRNA may be explained by several mechanisms. In the study by Jackson et al. (17) where they incorporated the 2′-O-methyl modification at the 2 position of the guide strand, it was suggested that there is some sort of steric hindering effect of the methyl group because position 2 has limited space in the RISC complex in which to accommodate a methyl group, forcing a conformational adjustment of the RISC complex, which limits the tolerance for mismatches in the seed region. The other mechanism that may explain less off-target effects in our LNA modified siRNA is the limitation of the chance of effective incorporation of the sense strand into RISC. The placement of a chemical modification at the 5′ end of the sense strand

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forces preferential loading of the antisense strand in to RISC. And, if despite this guided RISC loading, the sense strand is incorporated into RISC, then 5′ phosphorylation, which is needed for RISC activation of the sense strand, may be prevented by the chemical modifications rendering RISC inactive.

6. sisiRNA LNA enables a novel siRNA design invented by Jesper Wengel and Jorgen Kjems (33) that not only limits the chance of sense strand incorporation into RISC but may also allow more flexibility for the incorporation of chemical modifications into the siRNA. Small internally segmented interfering RNA (sisiRNA) entails an siRNA design where the passenger strand is nicked into two short separate strands and these are hybridized to the antisense strand using the high Tm inferred to by LNA modifications (Fig. 9.4). This “nicked” design seems equally effective in target knockdown as do conventional siRNA designs (33). Interestingly, sisiRNA is reported to allow more chemically modified antisense strands, which are known to be nonfunctional with standard siRNA designs. Even quite bulky chemical modifications are tolerated (33), which will open up new avenues for pharmacological applications. The nicked passenger strand has two advantages. First, the nicked strand is forced to become the passenger strand; it can never function as a functional guiding strand, thus the risk of passenger strand mediated off-target effects is reduced. Second, the passenger strand cleavage is not a rate-limiting step anymore limiting the risk of swamping the RISC machinery with a chemically modified passenger strand which is very stable but which may clog up the RISC complex. For siRNA to function the passenger

Fig. 9.4. Schematic representation of an siRNA, siLNA, and sisiRNA design. The positions of the DNA or LNA moieties are indicated.

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strand must be cleaved (34). Grimm et al. (35) demonstrated in vivo with short hairpin RNA (shRNA) constructs that high levels of shRNA resulted in dose-dependent liver injury and high morbidity. This was associated with the downregulation of liverderived microRNAs (miRNAs), indicating possible competition of the latter with shRNAs for limiting cellular factors required for the processing of small RNAs. Since siRNA and miRNA essentially use the same molecular pathways there is a distinct possibility (but not yet proven) that a very stable chemically modified siRNA can cause problems similar to “throwing a spanner in the works.” However, this fear is so far mainly theoretical. Indeed we ourselves have observed that a high level of modification of an siRNA is only detrimental for its efficacy, but in general we do not observe toxicity issues in cell lines with highly modified siRNAs. But the sisiRNA design might allow for more chemical modifications to be used and this may help to further open up therapeutic possibilities.

7. Preventing Inadvertent Stimulation of the Immune System

Negating off-target effects might be done using a clever strategy of chemical modifications and siRNA design. However, this is not the last concern that threatens the therapeutic use of siRNA. Nonspecific and toxic effects can also be caused by inadvertent stimulation of immune responses. The now classic 21-mer siRNA design as proposed by Tuschl (5) prevented the most prominent dsRNAdependent protein kinase (PKR) interferon response in mammalian cells. However, it is becoming clear that in vivo the innate mammalian immune system is very adapt in recognizing nucleic acid species as signatures of potential pathogens, including the short siRNAs. A number of recent studies have pointed to immunological effects of siRNAs, including the induction of pro-inflammatory cytokines and type I interferon (as reviewed in Ref. 36). These immunological responses are mainly observed when delivery vehicles are used (37). Immuno-recognition of RNA depends on certain molecular features such as length, double-strand configuration, sequence motifs, and nucleotide modifications. Judge et al. (37) described immunostimulatory motifs in siRNA. Avoiding these sequences in the design of siRNAs might yield molecules that induce less immune activation. RNA-sensing immunoreceptors include three members of the Toll-like receptor (TLR) family (TLR3, TLR7, TLR8) and cytosolic RNA-binding proteins like PKR and the helicases RIG-I and Mda5 (38–40). Detection of RNA molecules normally occurs during viral infection and triggers antiviral innate defense mechanisms including

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the induction of type I interferons and downregulation of gene expression. Type I interferon induction by synthetic siRNAs requires TLR7 and is sequence dependent, similar to the detection of CpG motifs in DNA by TLR9 (36, 41). Identification of the exact molecular mechanisms of immune-recognition of RNA will allow the development of methods to avoid immune stimulation by siRNA. Importantly, Hornung et al. (41) demonstrated in a mouse model that TLR7 stimulation by siRNA could be inhibited by modification with LNA. Therefore LNA modifications in siRNA can be used to limit the inflammatory response towards the siRNA. Similar finds were reported for 2′-O-Me (42) and a recent report by the same group demonstrated that 2′-O-Memodified RNA may have a general utility as a potent inhibitor of TLR7 and can function as an antagonist of immunostimulatory RNA, even when only a very limited percentage of positions are modified (80%) with negligible (60 kDa glycoprotein, which consists of three cysteine-rich domains (D1, D2 and D3) connected by short linker regions (8). The amino-terminus of the uPAR domain 1 is the primary site for the binding of uPA. However, domains 2 and 3 may be important for high affinity binding of uPA as purified domain 1 has a 1,500-fold lower ligand affinity than the complete tri-domain uPAR (7). uPAR is a multifunctional protein and is believed to play a role in the regulation of several physiological and pathological conditions that exploit cell adhesion and migration including wound healing, neutrophil recruitment during inflammation as well as tumor invasion and metastasis (8, 9). Several recent studies have shown that the various functions of uPAR are invoked not only by proteolysis but also by intracellular signaling (3, 10, 11, 12). uPAR levels have been strongly correlated with metastatic potential and advanced disease as has been demonstrated in tumor samples obtained from patients with colon and breast cancers (13–15). For example, uPAR is overexpressed in invasive breast cancer tissues, but not in normal and benign breast tumors (16). The importance of the uPAR system makes it a potential target for cancer therapy.

5. Therapeutic RNAi-Mediated Strategies for Targeting the uPAR-uPA System

Several approaches have been employed to target uPAR as a means of cancer therapy. These approaches include small molecule and peptide antagonists of the uPA-uPAR interactions as well as the uPAR interactions that are downstream of uPA binding; these include antibiotics, monoclonal antibodies and antisense technology. To date, antisense technologies used against uPAR include either the classic antisense oligodeoxynucleotides technology, which consists of the injection of antisense DNA strands complementary to uPAR mRNA, or the antisense RNA technology based on cell transfection with a vector capable of expressing the antisense transcript complementary to uPAR mRNA. Research groups investigating antisense RNA technology for downregulation of uPAR in vivo have employed both plasmid and adenovirus constructs for this purpose (17–26). RNAi has provided new avenues for the treatment of cancer. Small interfering RNAs (siRNAs) are believed to be more potent inhibitors of gene expression with less toxicity (27). Our laboratory had already employed shRNA-based RNAi plasmid system for the downregulation of uPAR in prostate cancer (28), glioma (11–33) and meningioma (34, 35). We have utilized a plasmid

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construct expressing the same small hairpin RNA (shRNA) to target uPAR. For the above-referenced studies, human glioblastoma cells were intracranially injected into athymic nude mice. Eight to ten days after tumor growth, mice were implanted with mini-osmotic pumps with a sustained release of 0.25 µl/h of 150 µg of the shRNA-expressing plasmid construct in a subcutaneous sac with a catheter to the intracranial tumor site. The mice were sacrificed and analyzed at the end of the 5-week follow up period or when the control mice started showing symptoms. We reported a 65% regression of pre-established intracranial tumor growth (23). These findings were further confirmed in our laboratory where we reported a 70% inhibition of pre-established intracranial tumor growth (26). Pulukuri et al. (28) reported that intratumoral injection with a plasmid construct expressing shRNA for uPAR resulted in partial reduction of pre-established orthotopic prostate tumor in athymic male nude mice with no observable secondary tumor. Downregulation of more than one component involved in tumor invasion and metastasis may possibly have a synergistic or additive effect in impeding tumor dissemination. We have reported that intracranial injection of human glioma cells infected with a bicistronic adenoviral construct capable of simultaneously expressing antisense uPAR and antisense matrix metalloproteinase-9 (MMP-9) resulted in decreased invasiveness and tumorigenicity in mice (26). Further, subcutaneous injections of the bicistronic construct into established tumors caused tumor regression. MMP-9 is involved with metastasis of various types of cancers, though its inhibition has not led to significant improvements in clinical trials as yet. We therefore hypothesized that a dual targeted approach combining MMP-9 downregulation with that of uPAR has the potential for efficient tumor targeting. Indeed, we found that a bicistronic plasmid construct expressing shRNA simultaneously targeting both uPAR and MMP-9 resulted in total regression of pre-established intracerebral tumor growth in mice. We have also shown that RNAi-mediated downregulation of uPAR and cathepsin B reduced glioma cell invasion and angiogenesis in in vivo models (29). In addition, intratumoral injections of these plasmid vectors expressing shRNA for uPAR and cathepsin B resulted in the regression of pre-established intracranial tumor. Similarly, we have also demonstrated that intraperitoneal injection of a bicistronic plasmid construct expressing shRNA for uPA and uPAR caused the regression of pre-established, intracranial tumors in mice (33). Despite our success thus far, it should be noted that the delivery of uPAR downregulation constructs, whether plasmid vectors, adeno-viral vectors or synthetic strands, still needs to be assessed appropriately in human systems.

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hom*ozygous uPAR-deficient mice display normal growth and fertility, do not show histological abnormalities in tissues and do not differ from wild-type mice for spontaneous lysis of experimental pulmonary plasma clot. This is very similar to what is also noted in uPA-deficient mice. Thus, the apparent lack of toxicity from inhibiting this proteolytic system makes it an ideal candidate for targeting as a cancer therapeutic agent. The uPA-uPAR system plays a very important role in cancer metastasis and may function via a number of signaling pathways. Binding of uPA with its receptor uPAR can activate downstream signaling molecules, including the mitogen-activated protein kinase, signal transducer and activator of transcription (Stat) and the Ras/extracellular signal-regulated kinase pathway, which in turn, lead to cell proliferation, migration and invasion (3). uPA–uPAR-mediated signaling can upregulate the production of MMPs, which induce ECM degradation and, in turn, tumor invasion and metastasis (3). Since uPA-uPAR and their downstream signaling pathways are implicated in many cancers, including essential cellular functions that contribute to the malignancy of tumor cells, targeting uPA–uPAR-mediated signaling pathways may be promising for the treatment of metastatic disease. In addition, targeting uPA/ uPAR may provide additive or synergistic treatment benefits if used in combination with conventional therapeutics such as chemotherapy or radiation.

6. Potential for Targeting MMPs Matrix metalloproteinases (MMPs) are a family of structurally related and highly conserved zinc-dependent endopeptidases collectively capable of degrading most components of the basem*nt membrane and ECM. MMP substrates also include a wide variety of proteins, such as chemotactic molecules, adhesion molecules, proteinase inhibitors, cell-surface receptors, blood clotting factors, latent growth factors and growth factor-binding proteins. Most human MMPs can be divided according to their sequence hom*ology, substrate specificity and cellular location into the following subclasses: collagenases, gelatinases, stromelysins, matrilysins, membrane-type MMPs and others (36). The basic multi-domain structure of MMPs comprises the following: (1) an amino-terminal domain, (2) a catalytic domain and (3) a carboxy-terminal domain. To date, we know of a minimum of 25 secreted or membranebound human MMPs. The expression, secretion and activity of MMPs in normal tissues are subject to tight control. Data generated from intensive studies on MMP activities in different cells and tissues, as well as studies from knock-out

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animals, illustrate the importance of these enzymes in many normal physiologic processes (e.g., embryonic development, bone resorption, angiogenesis and wound healing) and pathologic processes (rheumatoid arthritis, multiple sclerosis, periodontal disease and tumor growth and metastasis) (37). MMPs are secreted as pro-MMPs and then activated by sequential cleavage steps (38, 39), which involve the removal of signal peptide and pro-peptide domains or a change in configuration, which activates the enzymes. MMP expression and proteolytic activity are tightly regulated at three stages: gene transcription, pro-enzyme activation and activity of natural inhibitors (tissue inhibitors of metalloproteinase: TIMPs). The balance between production, activation and inhibition prevents excessive proteolysis or inhibition. Several factors like cytokines, growth factors, phorbol esters, cell-cell and cell-matrix interactions are thought to control MMP expression (40). Most MMPs are secreted as inactive zymogens, which may be proteolytically activated by different proteinases such as other MMPs, plasmin, trypsin, chymotrypsin and cathepsins. Several cell types produce MMPs including monocytes, macrophages, neutrophils (41, 42), T-lymphocytes (43), endothelial cells (44), fibroblasts (45) as well as microglia, astrocytes, oligodendrocytes and neurons in the CNS (46–48). In particular, MMP-2 and MMP-9 are secreted by microglia and astrocytes as active forms (49). MMPs have been shown to regulate tumor cell invasion through their interactions with extracellular matrix components including cell matrix embedded growth factors and cell adhesion molecules (15, 50). The most important of these metalloproteases are MMP-9 and MMP-2, which have shown to be involved in glioma invasion and angiogenesis (51, 52). MMPs are controlled by enzyme activation to produce a functional form and at the level of gene expression (53). There are also other underlying mechanisms that affect mRNA stability, protein secretion and specific degradation and clearance (53). Growth factors, such as endothelial growth factor (EGF), basic fibroblast growth factor (b-FGF), transforming growth factor (TGF-b1 and b2) and vascular endothelial growth factor (VEGF) have been shown to upregulate MMP-2 and MMP-9 (54). MMP-9 and stromelysin (MMP-3) have been shown to be chiefly transcribed under the influence of various transcription factors commonly found to be involved with cellular stress responses and tissue morphogenesis, including NF-kb, ETS family members and AP-1 (55). Epidermal growth factor variant subtype III promoted activation of MMP-9, possibly through the activation of MAPK/ERK in glioblastoma (56). We have previously shown that MMP-9 production is induced by cytoskeletal changes involving protein kinase C activation mediated by NF-kb (57). The mitogen-activated

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kinase/extracellular signal-regulated kinase (MEK/ERK) signaling pathway is essential for MMP-9 upregulation in astrocytes after PKC induction and TNF-a (cytokine) stimulation. It has been reported that SNB19 glioma cells transfected with dominant negative JNK, MEKK and ERK1 expression vectors decreased MMP-9 expression as well as promoter activity (58). The mtERK stable SNB19 cells showed decreased levels of MMP-9 and less invasiveness as compared to parental and vector-transfected stable clones (25). All these studies point to the fact that specifically targeting MMP-9 or MMP-2 singly or in combination with other proteases could have specific therapeutic implications in the treatment of cancer.

7. RNAi-Mediated Strategies for Targeting MMPs

We have previously demonstrated that inhibition of cathepsin B and MMP-9 gene expression via RNA interference reduced tumor cell invasion, tumor growth and angiogenesis in a glioblastoma cell line (59). We have also demonstrated that specific interference of uPAR and MMP-9 gene expression induced by double-stranded RNA resulted in decreased invasion, tumor growth and angiogenesis in gliomas (52). Other researchers have silenced MMP-1 to elucidate the mechanism involved in signaling (60). Wyatt et al. concluded that MMP-1 expression is essential for the ability of MDA-231 cells to invade and destroy a collagen matrix. In vivo experiments suggest an important role for MMP-1 in breast tumor growth and have demonstrated the potential of RNAi-mediated targeting of MMP-1 (61). Researchers have also demonstrated that siRNA-mediated blocking of either membrane type-1 MMP (MT1-MMP) or MMP-2 were effective in reducing the hypoxiainduced invasion in MDA-MB-231 and MDA-MB-435 breast carcinoma cell lines (62). Knocking down of MMP-7 by small interfering RNA was shown to suppress lysophosphatidic acid (LPA)-induced invasion in two EOC cell lines (DOV13 and R182). These results show that MMP-7 expression is correlated with EOC invasiveness and LPA-induced MMP-7 secretion/activation and may represent new mechanisms that facilitate ovarian cancer invasion besides the well-known induction of MT1-MMPmediated pro-MMP-2 activation by LPA (63). Our studies have reported that the simultaneous targeting of two or more components involved in invasion or migration is significantly more relevant therapeutically than concentrating on one component alone. For example, RNAi-mediated targeting of uPAR and MMP-9 gene expression in the IOMMLEE malignant meningioma cell line inhibited tumor growth,

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tumor cell invasion and angiogenesis both in vitro and in vivo. Our results show that downregulation of uPAR and MMP-9 leads to a decrease in the activation of some of the important enzymes participating in the MAPK and PI3 kinase pathways, which in turn, might decrease cell survival and proliferation. In addition, we have demonstrated the efficiency of RNAi-mediated targeting of uPAR and MMP-9 in pre-established tumor growth in vivo. We observed a significant regression of pre-established orthotopic tumors upon RNAi-mediated targeting of uPAR and MMP-9 and have also demonstrated that targeting both the proteins simultaneously augmented the therapeutic treatment of human meningiomas (35). In another study, we introduced small interfering RNA to downregulate the expression of uPAR and MMP-9 in breast cancer cell lines (MDA MB 231 and ZR 75 1). In vitro angiogenesis studies indicated a decrease in the angiogenic and invasive potential of the treated cells. These results suggest a synergistic effect from the simultaneous downregulation of uPAR and MMP-9. We also assessed the levels of phosphorylated forms of MAPK, ERK and AKT signaling pathway molecules and found reduced levels of these molecules in cells treated with the bicistronic construct as compared to the control cells. Furthermore, targeting both uPAR and MMP-9 using RNAi totally regressed orthotopic breast tumors in nude mice, thereby providing evidence that the simultaneous downregulation of uPAR and MMP-9 using RNAi technology may provide an effective tool for breast cancer therapy (64). In another study, we have demonstrated that the simultaneous targeting of more than two components is significantly superior to targeting two alone. We have showed that direct intratumoral injections of plasmid DNA expressing hpRNA for uPA, uPAR and MMP-9 significantly regressed pre-established intracranial tumors in nude mice as compared to the controls. In addition, cells treated with RNAi for uPAR, uPA and MMP-9 showed reduced pERK levels when compared to parental and EV/SV-treated SNB19 cells. Our results support the therapeutic potential of RNAi as a method for gene therapy in treating gliomas (30). A brief schematic representation of the possible mechanisms involved in MMP-9 and uPAR targeted RNAi therapy is given in Fig. 13.4. The main objective of cancer therapy is to arrest tumor invasion and convert it to a controlled, localized disease. Accumulated lines of evidence indicate that MMPs and the uPAR system play an essential role in tumor invasion and metastasis. Therapeutic strategies that can inhibit a broad spectrum of MMPs and the uPAR system may be beneficial for retarding and preventing tumor progression. To achieve this, further investigation and understanding of proteases at the molecular

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Fig. 13.4. The uPAR system and its association with MMPs. The uPAR system and MMPs are involved in multiple intracellular survival and proliferative signaling events.

level should play an important role in the future development of new, target-selective treatments. It is sufficiently clear that the simultaneous targeting of multiple systems is more synergistic than additive.

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Chapter 14 Silencing of HIF-1a by RNA Interference in Human Glioma Cells In Vitro and In Vivo David L. Gillespie, Jeannette R. Flynn, Brian T. Ragel, Maria Arce-Larreta, David A. Kelly, Sheryl R. Tripp, and Randy L. Jensen Abstract Higher-grade gliomas are distinguished by increased vascular endothelial cell proliferation and peritumoral edema. These are thought to be instigated by vascular endothelial growth factor, which in turn is regulated by cellular oxygen tension. Hypoxia inducible factor-1α (HIF-1α) is a main responder to intracellular hypoxia and is overexpressed in many human cancers, including gliomas. Here we present methods for investigating the role of HIF-1α in glioma growth in vivo and in vitro using RNA interference in U251, U87, and U373 glioma cells. Key words: HIF-1, glioma, VEGF, RNA interference, siRNA, hypoxia, gene therapy.

1. Introduction Overexpression of hypoxia-inducible factor-1α (HIF-1α) has been described in many common human cancers and their metastases (1, 2, 3). The role of HIF-1 in solid tumor growth is still not entirely clear, but previous work suggests that this transcription factor is necessary for growth and angiogenesis of these tumors (4, 5). A direct correlation between tumor grade and HIF-1 expression in glioblastoma multiforme (GBM) has been demonstrated (6). We and others have published data supporting the proposition that HIF-1α expression represents an “angiogenic switch” that facilitates the progression of a low-grade astrocytoma to a GBM and promotes cell survival in hypoxic conditions by elevating glycolysis and angiogenesis (2, 7). It has also been M. Sioud (ed.), Methods in Molecular Biology, siRNA and miRNA Gene Silencing, vol. 487 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-60327-547-7_14

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demonstrated that reducing HIF-1α expression using RNAi and other methods leads to a reduction of glioma growth in vivo and in vitro (5, 8–10). The transcription factor HIF-1 is composed of two heterodimeric subunits, HIF-1α and HIF-1β (known as aryl hydrocarbon receptor nuclear translocator (ARNT)). At the mRNA levels, HIF-1α and HIF-1β are both constitutively expressed and do not seem to be significantly modified by hypoxia (11). Whereas HIF-1β protein is found in normoxic cells, HIF-1α is rapidly degraded by proteasomal degradation. During low oxygen tension conditions (1–2% O2), this degradation is inhibited, leading to increased HIF-1 (12). HIF-1 binds to DNA hypoxia response elements (HREs) and induces the transcription of a number of well-characterized genes that help cells survive low oxygen conditions (13). These genes include vascular endothelial growth factor (VEGF), erythropoietin, transferrin, GLUT-1, and almost every gene in the glycolytic pathway (14). Stabilization of HIF1α promotes cell survival via adaptive modifications of cellular metabolism that increase these glycolytic enzymes and hence the glycolysis rate. This adaptation of cancer cells through increased glycolysis was first proposed by Warburg (15) as a necessary step toward an aggressive phenotype. The results of recent studies of HIF-1 have indicated a possible link between it and the Warburg effect in various cell types (16, 17) and prompted the proposal that aerobic glycolysis could be controlled by dysregulation of HIF-1α (18). Here we describe methods for promoting HIF-1α stabilization using hypoxia to assist in studying the effects of treatment and subsequent detection and analysis of HIF-1α and other downstream targets in response to RNAi targeting HIF-1α.

2. Materials 2.1. Cell Culture and Hypoxia Chamber

1. Dulbecco’s Modified Eagle’s Medium (DMEM) supplemented with 10% fetal bovine serum. 2. BD BBL GasPak Jar system. 3. GasPak Plus Hydrogen-CO2 generator envelopes. 4. Oxygen indicator strips.

2.2. Cell Lysis and Protein Isolation

1. Phosphatase inhibitor buffer (PIB), 40x: 250 mM NaF, 500 mM β-glycerophosphate, 50 mM Na3VO4. Filter and store in aliquots at −20°C indefinitely or at 4°C for up to 1 month. 2. Hypotonic buffer (HB): 0.5 M HEPES, pH 7.5, 0.5 M NaF, 0.5 M EDTA, 0.1 M Na2MoO4. Filter and store at 4°C for up to 6 months.

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3. 10% NP-40. 4. Lysis buffer: 400 mM NaCl, 20 mM HEPES (pH 7.5), 10 mM NaF, 10 mM PNPP, 1 mM Na2VO4, 0.1 mM EDTA, 10 µM Na2MoO4, 10 mM β-glycerophosphate, 20% glycerol. Store at 4°C for up to 6 months. Directly before use, add 0.1 M dithiothreitol (DTT), 100x protease inhibitor co*cktail (Sigma-Aldrich, St Louis, MO) for final concentrations of 1 mM DTT and 1x protease inhibitor. Lysis buffer that has had DTT and protease inhibitor added is referred to as “complete lysis buffer” in this protocol. Complete lysis buffer should be discarded after 24 h (see Note 1). 5. Polyethylene cell lifters (Corning, Corning, NY). 2.3. Protein Isolation from Tumor Tissues

1. Mihir’s hom*ogenation buffer: 10 mM HEPES (pH 7.6), 1 mM Na2VO4, 100 mM NaF, 0.4 mM PMSF, 0.1 mM EGTA, 10 mM Na4P2O7. Store at 4°C for 6 months. Just prior to using add 100x protease inhibitor (Sigma-Aldrich) for final concentration of 1x and 0.1 M DTT for a final concentration of 2 mM DTT. Buffer should be discarded after 24 h once the protease inhibitors are added. 2. Lysis buffer: see Sect. 2.2. 3. Scalpel blades, size #10. 4. Polytron PT-1200 hom*ogenizer (or similar rotor-stator hom*ogenizer).

2.4. SDSPolyacrylamide Gel Electrophoresis (SDS-PAGE)

1. Running buffer: NuPAGE MOPS 20x SDS running buffer diluted to 1x. 2. Loading buffer: 4xNuPAGE LDS buffer, 1% β-mercaptoethanol (BME). 3. Precast NuPAGE Bis-Tris 4–12% gradient gel, 12 well. 4. Prestained molecular weight markers: Kaleidoscope markers (Bio-Rad, Hercules, CA).

2.5. Western Blot

1. Transfer buffer: NuPAGE 20x Transfer buffer. 2. Hybond-P PVDF membrane from Amersham Biosciences, Piscataway, NJ, and Gel Blot paper from ISC Bioexpress (Kaysville, UT). 3. TBS-T (Tris-buffered saline with Tween-20: Prepare 10x stock with 1.37 M NaCl, 27 mM KCl, 250 mM Tris-HCl (pH 7.4), 1% Tween-20. Dilute 100 ml with 900 ml of water for use. 4. Blocking buffer: Make up 5% non-fat powdered milk in 1x TBS-T. Need 100 ml/membrane (small gel). Keep at 4°C for no more than 48 h.

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5. Primary antibody: Anti-human HIF-1 mouse monoclonal IgG from Novus Biologicals (Littleton, CO). 6. Secondary antibody: Anti-mouse IgG conjugated to horse radish peroxidase (Amersham Biosciences). 7. Enhanced chemiluminescent plus (ECL+) reagents from Amersham Biosciences (Piscataway, NJ) and Bio-Max light film (Kodak, Rochester, NY). 8. Straight-edge platform paper cutter. 2.6. Enzyme-Linked Immunosorbent Assay for VEGF

1. R&D systems Quantikine VEGF Enzyme-linked immunosorbent assay (ELISA) kit (Minneapolis, MN). 2. Benchmark microplate reader (Bio-Rad Laboratories, Hercules, CA). 3. Bio-Rad 1575 ImmunoWash plate washer.

2.7. RNAi Design

1. Silencer siRNA construction kit (Ambion, Austin, TX). 2. http://www.rnaiweb.com/ 3. http://www.ncbi.nlm.nih.gov/projects/genome/RNAi/ 4. http://rnai.cs.unm.edu/projects/ 5. http://www.ambion.com/techlib/resources/RNAi/index.html

2.8. Immunohistochemistry for MIB-1 and GLUT-1

1. Citrate buffer: 0.01 M citric acid (Sigma, St. Louis, MO), 0.025 M NaOH (Sigma), dH2O (pH 6.0). 2. 4% potassium iodine (Fisher), 2% iodine, dH2O. 3. 5% sodium thiosulfate, dH2O. 4. Antibodies for Ki-67 (MIB-1) and GLUT-1 (Dako Cytomation, Carpinteria, CA). 5. IView DAB detection kit (Ventana Medical Systems, Tucson, AZ). 6. Electric pressure cooker DC2000 (BioCare Medical, Concord, CA). 7. Dawn dish-washing detergent, 0.2%. 8. Automated BenchMark XT immunostainer from Ventana Medical Systems (Tucson, AZ).

2.9. Cell Transfection and Imaging

1. Lipofectamine 2000 (Invitrogen, Carlsbad, CA). 2. Dead Cell Reagent (Invitrogen). 3. Silencer siRNA Labeling kit (Ambion). 4. Optimem serum-free media (Invitrogen).

2.10. Real Time PCR

1. RNeasy Mini spin column kit (Qiagen, Valencia, CA). 2. Superscript III first-strand synthesis for RT-PCR kit (Invitrogen).

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3. IQ SYBR Green supermix kit (Bio-Rad, Hercules, CA). 4. Bio-Rad MyIQ iCycler. 2.11. Cell Inoculation

1. Matrigel (BD Biosciences, Franklin Lakes, NJ). 2. Lo-dose 0.5 ml insulin syringe (BD Biosciences). 3. Trypsin-EDTA, 0.25% (Invitrogen).

2.12. siRNA Treatment

1. Annealed siRNAs at 150 µM in sterile annealing buffer (50 mM Tris, 150 mM NaCl). 2. 4x JetPEI (Q-Biogene, Irvine CA). 3. Sterile 150 mM NaCl. 4. Lo-dose 0.5 ml insulin syringe (BD Biosciences).

3. Methods Because HIF-1α is a nuclear protein that is almost completely regulated post-translationally (11), it is important to assay nuclear protein levels. Assays involving transcription activity or mRNA levels (such as Real Time PCR) are only informative when they are used to assess RNAi knockdown effects or treatments that affect general transcription activity. Because HIF-1α is degraded quickly in the presence of oxygen, it is extremely important to do all steps quickly with protease inhibitors and on wet ice to slow down the degradation process. To increase HIF-1α levels and ease detection, it is important to expose cells to hypoxia during experimentation. This also creates a broader range of HIF-1α protein levels to observe response to treatments. The GasPak jar and pouch systems have been used for many years to produce an inexpensive, reliable hypoxic atmosphere for short-term cell culture and HIF-1α stabilization (8, 19, 20). Many more intricate and expensive oxygen-controlling environments are also commercially available. Because inappropriate sample handling can have significant effects on the results, all experiments should be repeated several times. It is also helpful to verify the results by assessing downstream HIF-1 transcription targets such as VEGF, Ki-67 (MIB-1), and GLUT-1, which are much more stable and easily measured (see Note 2). There are several methods for creating a positive HIF-1α control. We have used 18 h hypoxic U251 glioma cells or treatment with cobalt chloride. 3.1. Hypoxia Chamber Cell Growth Studies

1. Cells are plated on 60 or 100 mm tissue culture dishes until they reach 70–80% confluence. It is important that the cells be actively growing to get maximum HIF-1α production. We use

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glioma cell lines U251 and U87 for most hypoxia experiments because they produce high levels of HIF-1α after a short 4 h exposure. Maximum levels for these cells are observed between 12 and 18 h exposures, which are best done overnight. 2. The culture medium is changed immediately before the dishes are placed in a GasPak Plus anaerobic culture chamber. Two GasPak catalyst envelopes are taped to the inside of the chamber to reduce the oxygen concentration below 1%. One oxygen indicator strip is taped to the inside to verify the oxygen concentration. Cells can be treated for 4–48 h. 3.2. Protein Isolation from Cell Lines

This procedure can be used for an 80% confluent cell layer of 21 cm2 (plate 5–8 × 106 cells/60 mm dish). Volumes can be scaled up for use with 100 mm plates. It is absolutely essential to proceed quickly and keep everything on ice. The following steps should be done before starting: (a) Locate/reserve 4°C swing-bucket centrifuge and 4°C fixedrotor centrifuge, and make up phosphate-buffered saline (PBS)/PIB using cold PBS. Put in ice bucket. Add 250 µl of PIB to 10 ml of PBS, needs 8 ml/plate. (b) Make complete lysis buffer, 30 µl/sample. Add 1 µl of 0.1 M DTT and 1 µl of protease inhibitor to 100 µl of lysis buffer). Put HB buffer on ice. (c) Make up ice/water bath in flat ice tray (see Note 3). Label one 1.7 ml tube/sample, and one 15 ml tube/sample, and put on ice (see Note 4). (d) Assemble one 25 ml pipette (for adding PBS/PIB), four 1 ml pipettes (for aspirating media/wash), one 5 ml pipette, and one cell lifter/plate, 10% NP-40. 1. Put cell plates in ice tray and aspirate media. 2. Wash cells with 4 ml of ice-cold PBS/PIB. Swirl and aspirate (see Note 5). 3. Add 4 ml of ice-cold PBS/PIB. 4. Scrape the cells off the dish with a cell lifter. Tip plate and use a 5 ml pipette to transfer cells into a pre-chilled 15 ml tube (see Note 6). 5. Centrifuge at 386 g for 5 min at 4°C, then aspirate the supernatant. 6. Resuspend the pellet in 0.6 ml of ice-cold HB buffer by gentle pipetting. 7. Allow the cells to swell on ice for 15 min. 8. Add 30 µl of 10% NP-40 (0.5% final), and vortex for 10 s. 9. Centrifuge the hom*ogenate at 500 g for 5 min at 4°C (see Note 7). For a cytoplasmic VEGF assay, transfer 600 µl of

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supernatant to 1.5 ml tube, keep on ice for protein assay (Step 12). Place tubes upside down in ice to drain all residual liquid, aspirate supernatant from side of tube and lid. 10. Resuspend the nuclear pellet in 30 µl of complete lysis buffer and shake vigorously on ice (or in cold room) for 30 min (use a vortexer with tube holder, medium setting). During the incubation time, label one 1.7 ml tube/sample for protein, and keep in ice. 11. Centrifuge at 30,000 g for 20 min at 4°C and save the supernatant (nuclear cell extract). Transfer into prechilled tube. 12. Assay protein concentration using the standard Bradford method. 13. Aliquot if desired and store at −80°C. Avoid freeze/thaw cycles. 3.3. Protein Isolation from Tumors

Tumors should be placed in liquid nitrogen immediately after resection. If the tumors are harvested from mice or other small animals, the animals should not be sacrificed using CO2 because this method can lead to elevated levels of HIF-1 in tumor samples. All steps are to be done on ice. 1. Place frozen tissue (300–500 mg) in 3 ml of ice cold Mihir’s hom*ogenation buffer in a 15 ml centrifuge tube on ice. Slice into small pieces with a #10 scalpel (see Note 8). 2. hom*ogenize for 2 min. If using a variable speed hom*ogenizer, start at the lowest speed and gradually increase to the maximum. 3. Centrifuge at 850 g for 10 min (see Note 9). 4. Remove supernatant to clean 15 ml tube, save pellet. 5. Repeat spin with the supernatant. During this step, make up complete nuclear lysis buffer, add 300 µl to pellet and resuspend. 6. Remove supernatant to clean 15 ml tube containing 600 µl 100% glycerol (final glycerol concentration is 20%). Vortex for 30 s, then hold on ice for 5 min. 7. Combine pellet with resuspended pellet from first spin, and hold on ice. 8. Centrifuge supernatant and glycerol at 15,344 g for 15 min. Save the pellet (this is the nuclei) and supernatant if needed for VEGF assay (see Note 10). 9. Resuspend pelleted nuclei in three volumes complete nuclear lysis buffer (~200 µl). 10. Mix vigorously at 4°C for 30 min.

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11. Spin at 30,000 g for 20 min. Aliquot supernatant into 1.7 ml tubes, determine protein concentration of nuclear and whole cell extract (from Step 7), and store at −80°C. 3.4. SDS-PAGE

These instructions assume the use of a Novex X-cell SureLock electrophoresis system (Invitrogen, Carlsbad, CA) using the NuPAGE pre-cast 12-well gel and buffers. 1. Calculate protein volume for 25–50 µg of total protein. Use lysis buffer to adjust for a final volume of 16 µl per well. 2. Make up loading dye—50 µ l 4x LDS dye + 5 µ l BME per gel. 3. Label one 0.7 µl tube/sample. Add 4 µl loading dye to each tube. 4. Add complete lysis buffer, then add protein to tubes. Heat at 75°C for 10 min. 5. While heating, make up running buffer—40 ml 20x MOPS NuPage buffer + 760 ml dH2O. Remove gel from wrapper, rinse in dH2O, remove well comb and bottom adhesive strip. Assemble gel, fill inner chamber with buffer, and rinse out wells with a transfer pipette. 6. Spin down samples to collect condensation and load gel. 7. Fill outer gel box chamber with remaining buffer. 8. Run at 200 V until blue (20 kD) standard runs off gel— about 2 h.

3.5. Western Blot

All rinses and washes are done at room temperature using TBS-T, unless otherwise noted. HIF-1α appears as a doublet at ~120 kD (Fig. 14.1). Do not mistake the two bands at ~100 kD for HIF-1α. 1. Cut one corner of the membrane for ease in orientation. 2. Activate membrane in 100% MEOH for 20 s, then rinse in dH2O for 2 min. Make up 250 ml of NuPage transfer buffer: 25 ml MeOH and 12.5 ml 20x buffer. 3. Equilibrate membrane for 2 min in 10 ml of transfer buffer. While equilibrating, assemble pads and filter paper in the cathode side of the transfer module and soak in transfer buffer. 4. Place gel large plate down, break open, and remove the small plate, being careful to leave gel on large plate. Cut off and remove wells. 5. Place wet filter paper on gel, remove bubbles, and flip over. Push the “foot” of the gel out of slot and carefully remove gel from plate. Cut off the gel “foot.” 6. Assemble module in the following order, from the bottom (cathode) side—2 pads, paper, gel, membrane, paper, 2 pads, top cover (anode).

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Fig. 14.1. (a) Western blot of protein isolated from transfected cells. The cells were allowed to recover for 30 h posttransfection and then exposed to hypoxia (75 kD) is probed for HIF-1, the bottom ( 250 nM in human T cells) was able to cause strong cytotoxicity similar to that of long double-stranded dsRNAs (46,47). This toxicity is due to the double-stranded structure of siRNAs and dsRNAs, which activates the interferon-mediated non-specific RNA degradation and programmed cell death through the signaling of PKR and 2-5A systems. It is known that interferon-induced protein kinase PKR can trigger cell apoptosis, while activation of the interferon-induced 2′,5′-oligoadenylate synthetase (2-5A) system leads to extensive cleavage of single-stranded RNAs (i.e., mRNAs) (48). High siRNA/shRNA concentrations generated by the Pol-III-directed RNAi systems can also over-saturate the cellular microRNA pathway and thus cause global miRNA inhibition and cell death (49). In contrast, a Pol-II-directed intronic miRNA expression system does not show these problems due to their precise regulation under cellular RNA splicing and nonsensemediated decay (NMD) mechanisms (50, 51, 52, 53, 54), which degrade excessive RNA accumulation to prevent potential cytotoxicity. For therapeutic purpose in vivo, the Pol-II-directed intronic miRNA expression system is likely a better solution than Pol-III-based siRNA/shRNA expression systems. The intron-derived miRNA system is activated in a specific cell type under the control of a type-II RNA polymerases (Pol-II)-directed transcriptional machinery. To overcome the Pol-III-mediated siRNA side-effects, we have successfully developed a novel Pol-II-based miRNA biogenesis strategy, employing intronic miRNA molecules

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(2) to knock down more than 85% of selected oncogene function or viral genome replication (22,23,47). Because of its flexibility in binding with partially complementary mRNA targets, miRNA can serve as an anti-cancer drug or vaccine to achieve a major breakthrough in the treatments of cancer polymorphisms and viral mutations. We are the first research group who discovered the biogenesis of miRNA-like precursors from the 5′-proximal intron regions of gene transcripts (pre-mRNAs) produced by the mammalian Pol-II. Depending on the promoter of the miRNA-encoded gene transcript, intronic miRNA is co-expressed with its encoding gene in the specific cell population, which activates the promoter and expresses the gene. It has been noted that a spliced intron was not completely digested into monoribonucleotides for transcriptional recycling since approximately 10–30% of the intron was found in the cytoplasm with a moderate half-life (6,55). This type of miRNA generation has been found to rely on the coupled interaction of nascent Pol-II-mediated premRNA transcription and intron excision, occurring within certain nuclear regions proximal to genomic perichromatin fibrils (22,56, 57, 58). After Pol-II RNA processing and splicing excision, some of the intron-derived miRNA fragments can form mature miRNAs and effectively silence the target genes through the RNAi mechanism, while the exons of pre-mRNA are ligated together to form a mature mRNA for protein synthesis (Fig. 19.2a). Because miRNAs are single-stranded molecules insensitive to PKR- and 2-5A-induced interferon systems, the utilization of this Pol-II-mediated miRNA generation can be safe in vitro and in vivo without the cytotoxic effects of dsRNAs and siRNAs. These findings indicate new functions for mammalian introns in intracellular miRNA generation and gene silencing, which can be used as a tool for analysis of gene functions and development of gene-specific therapeutics against cancers and viral infections. Using artificial introns carrying hairpin-like miRNA precursors (pre-miRNA), we have successfully generated mature miRNA molecules with full capacity in triggering RNAi-like gene silencing in human prostate cancer LNCaP, human cervical cancer HeLa and rat neuronal stem HCN-A94-2 cells (2,59,60) as well as in zebrafish, chicken and mouse in vivo (30,61). As shown in Fig. 19.2b, the artificial intron (SpRNAi) was co-transcribed within a precursor messenger RNA (pre-mRNA) by Pol-II and cleaved out of the pre-mRNA by RNA splicing. Then, the spliced intron containing a pre-miRNA structure was further processed into mature miRNAs capable of triggering RNAi-related gene silencing effects. Based on this artificial miRNA model, we have tested various pre-miRNA constructs, and observed that the production of intron-derived miRNA fragments was originated from the 5′-proximity of the intron sequence between the 5′-splice site and the branching point. These miRNAs were able to trigger strong

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Fig. 19.2. Biogenesis and function of intronic miRNAs. (a) The native intronic miRNA is co-transcribed with a precursor messenger RNA (pre-mRNA) by Pol-II and cleaved out of the pre-mRNA by an RNA splicing machinery, spliceosome. The spliced intron with hairpin-like secondary structures is further processed into mature miRNAs capable of triggering RNAi effects, while the ligated exons become a mature messenger RNA (mRNA) for protein synthesis. (b) We designed an artificial intron containing pre-miRNA, namely SpRNAi, mimicking the biogenesis processes of the native intronic miRNAs. (c) When a designed miR-EGFP(280–302)–stemloop RNA construct was tested in EGFP-expressing Tg(UAS:gfp) zebrafishes, we detected a strong RNAi effect only on the target EGFP (lane 4). No detectable gene silencing effect was observed in other lanes from left to right: 1, blank vector control (Ctl); 2, miRNA–stemloop targeting HIV-p24 (mock); 3, miRNA without stemloop (anti); and 5, stemloop–miRNA* complementary to the miR-EGFP(280–302) sequence (miR*). The off-target genes such as vector RGFP and fish actin were not affected, indicating the high target specificity of miRNA-mediated gene silencing. (d) Three different miR-EGFP(280–302) expression systems were tested for miRNA biogenesis from left to right: 1, vector expressing intron-free RGFP, no pre-miRNA insert; 2, vector expressing RGFP with an intronic 5′-miRNA-stemloop-miRNA*-3′ insert; and 3, vector similar to the 2 construct but with a defected 5′-splice site in the intron. In Northern bolt analysis probing the miR-EGFP(280–302) sequence, the mature miRNA was released only from the spliced intron that resulted from the vector 2 construct in the cell cytoplasm.

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suppression of genes possessing over 70% of complementarity to the miRNA sequences, whereas non-hom*ologous miRNAs, i.e., empty intron without the pre-miRNA insert, intron with an offtarget miRNA insert (negative control), and splicing-defective intron showed no silencing effects on the targeted gene. The same results can also be reproduced in the zebrafish directed against target EGFP expression (Fig. 19.2c), indicating the consistent preservation of the intronic miRNA biogenesis system in vertebrates. Further, no effect was detected on off-target genes, such as RGFP and β-actin, suggesting the high specificity of miRNA-directed RNA interference (RNAi). We have confirmed the identity of the intron-derived miRNAs, which were sized about 18–27 base nt, approximately similar to the newly identified Piwi-interacting RNAs. Moreover, the intronic small RNAs isolated by guanidinium chloride ultracentrifugation can elicit strong, but short-term gene silencing effects on the hom*ologous genes in the transfected cells, indicating their temporary RNAi effects. However, the long-term (>1 month) gene silencing effect that we observed in vivo occurs only in nuclear transfection of the Pol-II-mediated intronic miRNA system by retrovectors. The components of the Pol-II-mediated SpRNAi system include several consensus nucleotide elements consisting of a 5′-splice site, a branch-point domain, a poly-pyrimidine tract and a 3′-splice site (Fig. 19.3). Additionally, a pre-miRNA insert sequence is placed within the artificial intron between the 5′-splice site and the branchpoint domain. This portion of the intron would normally form a lariat structure during RNA splicing and processing. We currently know that spliceosomal U2 and U6 snRNPs, both helicases, may be involved in the unwinding and excision of the lariat RNA fragment into pre-miRNA; however, the detailed processing remains to be elucidated. Further, the SpRNAi contains a translation stop codon domain (T codon) in its 3′-proximal region to facilitate the accuracy of RNA splicing, which if present in a cytoplasmic mRNA,

Pre-mRNA construct with SpRNAi: 5’-promoter –

exon 1 –artificial intron (SpRNAi)– exon 2

5’ splice site – pre-miRNA insert

After intronic insert is spliced:

– 3’ T codons

– BrP – PPT – 3’-splice site –3’ T codon

5’-UTR – exon 1– exon 2 (mRNA) –3’-UTR

+ Intronic microRNAs Fig. 19.3. Schematic construct of the artificial SpRNAi intron in a recombinant gene SpRNAi-RGFP for intracellular expression and processing. The components of the Pol-II-mediated SpRNAi system include several consensus nucleotide elements consisting of a 5′-splice site, a branch-point domain (BrP), a poly-pyrimidine tract (PPT), a 3′-splice site and a pre-miRNA insert located between the 5′-splice site and the BrP domain. The expression of the recombinant gene is under the regulation of either a mammalian Pol-II RNA promoter or a compatible viral promoter for cell-type-specific effectiveness. Mature miRNAs are released from the intron by RNA splicing and further Dicer processing.

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would signal the diversion of a splicing-defective pre-mRNA to the nonsense-mediated decay (NMD) pathway and thus cause the elimination of any unspliced pre-mRNA in the cell. For intracellular expression of the SpRNAi, we need to insert the SpRNAi construct into the DraII cleavage site of a red fluorescent membrane protein (RGFP) gene from mutated chromoproteins of coral reef Heteractis crispa. The cleavage of RGFP at its 208th nt site by the restriction enzyme DraII generates an AG–GN nucleotide break with three recessing nucleotides in each end, which will form the 5′ and 3′ splice site, respectively after SpRNAi insertion. Because this intronic insertion disrupts the expression of functional RGFP, it becomes possible to determine the occurrence of intron splicing and RGFP-mRNA maturation through the appearance of red fluorescent emission around the membrane surface of the transfected cells. The RGFP also provides multiple exonic splicing enhancers (ESEs) to increase RNA splicing efficiency.

5. Strand-Specific Gene Silencing in Zebrafish

The foregoing establishes the fact that intronic miRNAs can be used as an effective strategy to silence specific target gene in vivo (27). We firstly tried to resolve the structural design of pre-miRNA inserts for the best gene silencing effect and found out that a strong structural bias exists in the selection of a mature miRNA strand during assembly of the RNAi effector, RNA-induced gene silencing complex (RISC). RISC is a protein-RNA complex that directs either target gene transcript degradation or translational repression through the RNAi mechanism. Formation of siRNA duplexes has been reported to play a key role in assembly of the siRNA-associated RISC. The two strands of the siRNA duplex are functionally asymmetric, but assembly into the RISC complex is preferential for only one strand. Such preference is determined by the thermodynamic stability of each 5′-end base-pairing in the strand. Based on this siRNA model, the formation of miRNA and its complementary miRNA (miRNA*) duplexes was thought to be an essential step for the assembly of miRNA-associated RISC. If this were true, no functional bias would be observed in the stemloop of a pre-miRNA. Nevertheless, we observed that the stemloop of the intronic pre-miRNA was involved in the strand selection of a mature miRNA for RISC assembly in zebrafish. In these experiments, we constructed miRNA-expressing SpRNAirGFP vectors as previously described (2) and two symmetric pre-miRNAs, miRNA-stemloop-miRNA* [1] and miRNA*stemloop-miRNA [2], were synthesized and inserted into the vectors, respectively. Both pre-miRNAs contained the same

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double-stranded stem arm region, which was directed against the EGFP nts 280–302 sequence. Because the intronic insert region of the SpRNAi-RGFP recombined gene is flanked with a PvuI and an MluI restriction site at the 5′- and 3′-ends, respectively, the primary insert can be easily removed and replaced by various gene-specific inserts (e.g., anti-EGFP) possessing cohesive ends. By changing the pre-miRNA inserts directed against different gene transcripts, this intronic miRNA generation system provides a valuable tool for genetic and miRNA-associated research in vivo. To determine the structural preference of the designed pre-miRNAs, we have isolated the zebrafish small RNAs by mirVana miRNA isolation columns (Ambion, Austin, TX) and then precipitated all potential miRNAs complementary to the target EGFP region by latex beads containing the target RNA sequence. One full-length miRNA identity, miR-EGFP(280–302), was verified to be active in the transfections of the 5′-miRNA-stemloop-miRNA*-3′ construct, as shown in the Fig. 19.4a (gray-shading sequences). Since the mature miRNA was detected only in the zebrafish transfected by the 5′-miRNA-stemloop-miRNA*-3′ construct, the miRNA-associated RISC tends to preferably interact with the construct [2] rather than the [1] pre-miRNA. The green fluorescent protein EGFP expression was constitutively driven by the β-actin promoter located in almost all cell types of the zebrafish, while Fig. 19.4b shows that transfection of the SpRNAi-RGFP vector into the Tg(UAS:gfp) zebrafish co-expressed the red fluorescent protein RGFP, serving as a positive indicator for the miRNA generation in the transfected cells. This approach has been successfully used in several mouse and human cell lines to show RNAi effects (59,60). We applied the liposomecapsulated vector (total 60 µg) to fishes and found that the vector easily penetrated almost all tissues of two-week-old zebrafish larvae within 24 h, reaching fully systemic delivery of the miRNA effect. The indicator RGFP was detected in both of the fishes transfected

Fig. 19.4 (continued) pre-miRNA insert resulted in no gene silencing significance. (d)–(g) Silencing of endogenous β-catenin and noggin genes in chicken embryos. (d) The pre-miRNA construct and fast green dye mixtures were injected into the ventral side of chicken embryos near the liver primordia below the heart. (e) Northern blot analysis of extracted RNAs from chicken embryonic livers with anti-β-catenin miRNA transfections (lanes 4–6) in comparison with wild types (lanes 1–3) showed a more than 98% silencing effect on β-catenin mRNA expression, while the house-keeping gene, GAPDH, was not affected. (f) Liver formation of the β-catenin knockouts was significantly hindered (upper right 2 panels). Microscopic examination revealed a loose structure of hepatocytes, indicating the loss of cell-cell adhesion due to breaks in adherin junctions formed between β-catenin and cell membrane E-cadherin in early liver development. In severely affected regions, feather growth in the skin close to the injection area was also inhibited (lower right 2 panels). Immunohistochemistry staining of β-catenin protein expression (brown) showed a significant decrease in the feather follicle sheaths. (g) The lower beak development was increased by the mandibular injection of the anti-noggin pre-miRNA construct (down panel) in comparison to the wild type (up panel). Right panels showed bone (alizarin red) and cartilage (alcian blue) staining to demonstrate the out growth of bone tissues in the lower beak of the noggin knockout. Northern blot analysis (small windows) confirmed a ~60% decrease of noggin mRNA expression in the lower beak area.

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Fig. 19.4. Intronic miRNA-mediated gene silencing effects in vivo. (a)–(c) Different preferences of RISC assembly were observed by transfection of 5′-miRNA*-stemloop-miRNA-3′ [1] and 5′-miRNA-stemloop-miRNA*-3′ [2] pre-miRNA structures in zebrafish, respectively. (a) One mature miRNA, namely miR-EGFP(280/302), was detected in the [2]-transfected zebrafishes, whereas the [1]-transfection produced another kind of miRNA, miR*-EGFP(301–281), which was partially complementary to the miR-EGFP(280/302). (b) The RNAi effect was only observed in the transfection of the [2] pre-miRNA, showing less EGFP (green) expression in the transfectant [2] than [1], while the miRNA indicator RGFP (red) was evenly present in all vector transfections. (c) Western blot analysis of the EGFP protein levels confirmed the specific silencing result of (b). No detectable gene silencing was observed in fishes without (Ctl) and with liposome only (Lipo) treatments. The transfection of either a U6-driven siRNA vector (siR) or an empty vector (Vctr) without the designed

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by either 5′-miRNA*-stemloop-miRNA-3′ or 5′-miRNA-stemloopmiRNA*-3′ pre-miRNA, whereas the silencing of target EGFP expression (green) was observed only in the fish transfected by the 5′-miRNA-stemloop-miRNA*-3′ pre-miRNA (Fig. 19.4b and c). The suppression level in the gastrointestinal (GI) tract was found to be less effective, probably due to the high RNase activity in this region. Because thermostability in the 5′ end of the siRNA duplexes resulting from both of the designed pre-miRNAs is the same, we suggest that the stemloop of pre-miRNA is involved in strand selection of mature miRNA during RISC assembly. Given that the cleavage site of Dicer in the stemarm determines the strand selection of mature miRNA (62), the stemloop may function as a determinant for the recognition of a special cleavage site. Therefore, the different stemloop structures among various species may also provide a clue for the prevalence of native siRNAs in invertebrates but rarely in mammals.

6. Intronic piRNAMediated RNA Interference in Chicken

The in vivo model of chicken embryos has been widely utilized in research in developmental biology, signal transduction and flu vaccine development. We have successfully tested the feasibility of localized gene silencing in vivo using the intronic miRNA approach and also discovered that the interaction between premRNA and perichromatin DNA may be essential for the intronic miRNA biogenesis. As an example, the β-catenin gene was selected because its products play a critical role in the biological development and ontogenesis (63). The β-catenin is known to be involved in the growth control of skin and liver tissues in chicken embryos. The loss-of-function of β-catenin is lethal in transgenic animals. As shown in Fig. 19.4d, e, f g, experimental results demonstrated that the miRNAs derived from a long RNA–DNA hybrid construct (≥150 bp) were capable of inhibiting β-catenin gene expression in the liver and skin of developing chicken embryos. This mimics the mechanism by which interaction between the intronic miRNA precursor and genomic DNA may account for a part of its specific gene silencing effect (22,23,51). We have demonstrated that the [P32]-labeled DNA component of a long RNA–DNA duplex construct in cell nuclear lysates was intact during the effective period of miRNA-induced RNA interference (RNAi) phenomena, while the labeled RNA part was replaced by cold hom*ologues and excised into small 18~27-nt RNA fragments in a 3-day incubation period (22). Since the intronic miRNA generation relies on a coupled interaction of nascent Pol-II-directed pre-mRNA transcription and intron excision occurring proximal to genomic perichromatin fibrils, the

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above observation indicates that the pre-mRNA–perichromatin DNA interaction may facilitate new intronic miRNA generation by the Pol-II RNA transcription and excision for long-term gene silencing. Thus, Pol-II may actually function as an RNAdependent RNA polymerase (RdRp) for producing more small miRNAs. Recent studies have shown that the pre-mRNA–perichromatin DNA interaction results in the generation of Piwi-interacting RNAs (piRNA), which are similar to intronic miRNAs but distinct from other small double-stranded siRNAs and shRNAs by their relatively larger size (approximately 26–31 nucleotides), single-strandedness and strand-specificity as well as by the clustered arrangement of their origins (58). The piRNA class of small RNAs is likely transcribed by an intracellular RNA polymerase, similar to RdRp, from the pre-mRNA–perichromatin DNA duplex region of a replicating cell genome during mitosis or meiosis. Mammalian type-II RNA polymerases (Pol-II) have been observed to possess the RdRp-like activities (4,64,65). Nuclear transfection of long DNA-RNA duplex templates has also been shown to trigger piRNA-like gene silencing effects against viral infection and retrotransposon activity (22). In Drosophila and zebrafish, Piwi proteins are recently found to be directly implicated in piRNA biogenesis to maintain transposon silencing in the germline genome (24,66). This function may be conserved in mice as loss of Miwi2, a mouse Piwi hom*olog, leads to germline stem cell and meiotic defects correlated with increased transposon activity (67). Because the RNAi effector of the piRNA-mediated gene silencing requires Piwi proteins rather than siRNA/shRNAassociated Dicer RNases, this suggests that the piRNA-mediated RNAi mechanism is slightly different from the siRNA/shRNAmediated RNAi pathway. In an effort to examine the pre-mRNA–perichromatin DNA interaction theory, we tested intracellular transfection of a long RNA–DNA hybrid construct containing a hairpin anti-β-catenin intronic pre-miRNA, which was directed against the central region of the β-catenin coding sequence (aa 306–644) with perfect complementarity. A perfectly complementary miRNA theoretically directs target mRNA degradation more efficient than translational repression. Using embryonic day 3 chicken embryos, a dose of 25 nM of the pre-miRNA construct was injected into the ventral body cavity, which is close to where the liver primordia would form (Fig. 19.4d). For efficient delivery into target tissues, the pre-miRNA construct was mixed with a liposomal transfection reagent (Roche Biomedicals, Indianapolis, IN). A 10% (v/v) fast green solution was concurrently added during the injection as a dye indicator. The mixtures were injected into the ventral side near the liver primordia below the heart using heat pulled capillary needles. After injection, the embryonic eggs were sealed with

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sterilized scotch tapes and incubated in a humidified incubator at 39–40°C until day 12 when the embryos were examined and photographed under a dissection microscope. Several malformations were observed, while the embryos still survived and there was no visible overt toxicity or overall perturbation of embryo development. The liver was the closest organ to the injection site and thus was most dramatically affected in its phenotypes. Other regions, particularly the skin close to the injection site, were also affected by the diffused miRNA effects. As shown in Fig. 19.4e, Northern blot analysis detecting the target β-catenin mRNA expression in the dissected livers showed that β-catenin expression in the wild-type livers remained normal (lanes 1–3), whereas those of the miRNA-treated samples was decreased dramatically (lanes 4–6). The miRNA silencing effect degraded more than 98% of β-catenin mRNA expression in embryonic chicken, but has no influence in the house-keeping gene GAPDH expression, indicating its high target specificity and very limited interferonrelated cytotoxicity in vivo. After 10 days of primordial injection with the anti-β-catenin pre-miRNA template, the embryonic chicken livers showed an enlarged and engorged first lobe, but the size of the second and third lobes of the livers were dramatically decreased (Fig. 19.4f). Histological sections of normal livers showed hepatic cords and sinusoidal space with few blood cells. In the anti-β-catenin miRNA-treated embryos, the general architecture of the hepatic cells in lobes 2 and 3 remained unchanged; however, there were islands of abnormal regions in lobe 1. The endothelium development appeared to be defective and blood leaked outside of the blood vessels. Abnormal types of hematopoietic cells were also observed between the space of hepatocytes, particularly dominated by a population of small cells with round nuclei and scanty cytoplasm. In severely affected regions, hepatocytes were disrupted (Fig. 19.4f, small windows) and the diffused miRNA effect further inhibited the feather growth in the skin area close to the injection site. The results discussed above showed that the anti-β-catenin miRNA was very effective in knocking out the targeted gene expression at a very low dose of 25 nM and was effect over a long period of time (≥10 days). Further, the miRNA gene silencing effect appeared to be very specific as off-targeted organs appear to be normal, indicating that the small single-strand composition of miRNA herein possessed no overt toxicity. In another attempt to silencing noggin expression in the mandible beak area using the same approach (Fig. 19.4g), it was observed that an enlarged lower beak morphology is reminiscent of those of BMP4-overexpressing chicken embryos reported previously (68,69). Skeleton staining showed the outgrowth of bone and cartilage tissues in the injected mandible area (Fig. 19.4g, right panels) and Northern blot analysis further confirmed that about

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60% of noggin mRNA expression was knocked out in this region (small windows). Since bone morphogenetic protein 4 (BMP4), a member of transforming growth factor-β (TGF-β) superfamily, is known to promote bone development and that noggin is an antagonist of BMP2/4/7 genes, it is not surprising to find out that our miRNA-mediated noggin knockouts created a morphological change, resembling the BMP4-overexpression results as previous reported in chicken and other avian models. Thus, the gene silencing in chicken by the pre-miRNA transfection presents a great potential of localized transgene-like approach in creating animal models for developmental biology research.

7. Localized RNA Interference Effects on Mouse Skin

To evaluate the efficacy and safety of intronic miRNA in animals, we have tested the vector-based intronic miRNA transfection in mice as previously described (61). As shown in Fig. 19.5, patched albino (white) skins of melanin-knockout mice (W-9 black) were created by a succession of intracutaneous (i.c.) transduction of anti-tyrosinase (Tyr) pre-miRNA construct (50 µg) for 4 days (total 200 µg). Tyr, a type-I membrane protein and coppercontaining enzyme, catalyzes the critical and rate-limiting step of tyrosine hydroxylation in the biosynthesis of melanin (black pigment) in skin and hair. After 13-day incubation, the expression of melanin has been blocked in the miRNA transfections due to a significant loss of its intermediates resulting from the antiTyr miRNA-triggered gene silencing effect. Contrarily, the blank control and the Pol-III (U6)-directed siRNA transfections presented normal black skin color under the same dosage. Northern blot analysis using RNA–PCR-amplified mRNAs from hair follicles showed a 76.1 ± 5.3% reduction of Tyr expression 2-day after the miRNA transfection, consistent with the immunohistochemical staining results from the same skin area, whereas mild, nonspecific degradation of common gene transcripts was detected in the siRNA-transfected skins (seen from smearing patterns of both house-keeping control GAPDH and targeted Tyr mRNAs). Given that Grimm et al. (49) have recently reported that high siRNA/ shRNA concentrations generated by the Pol-III-directed RNAi systems could over-saturate the cellular microRNA pathway and caused global miRNA dysregulation, the siRNA pathway may be incompatible with the native miRNA pathway in some tissues of mammals. Therefore, these findings have shown that the utilization of intronic miRNA expression systems provides a powerful new approach for transgenic animal generation and in vivo gene therapy. It was noted that non-targeted skin hair appears to be

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Fig. 19.5. In vivo effects of anti-tyrosinase miRNA on the mouse pigment production of local skins. Transfection of the miRNA induced strong gene silencing of tyrosinase (Tyr) mRNA expression but not house-keeping GAPDH, whereas that of U6-directed siRNA triggered mild non-specific RNA degradation of both Tyr and GAPDH gene transcripts. Since Tyr is an essential enzyme for black pigment melanin production, the success of gene silencing can be observed by a significant loss of the black color in mouse hairs. The red circles indicate the location of i.c. injections. Northern blot analysis of Tyr mRNA expression in local hair follicles confirmed the effectiveness and specificity of the miRNA-mediated gene silencing effect (small windows).

normal after miRNA transfection. This underscores the fact that the intronic miRNA is safe and effective in vivo. The results also indicated that the miRNA-mediated gene silencing effect is stable and efficient in knocking down the targeted gene expression over a relatively long period of time since the hair re-growth takes at least 10 days. Taken together, the intronic miRNA-mediated transgene approach may offer relatively safe, effective and longterm gene manipulation in animals, preventing the non-specific lethal effects of the conventional transgenic methods. More recent advances of the utilization of intronic miRNA expression systems have been reported in mice. Chung et al. (53) have succeeded in expression of a cluster of polycistronic miRNAs using the Pol-II-mediated intronic miRNA expression system. A polycistronic miRNA cluster can be processed into multiple miRNAs via the cellular miRNA pathway. This new RNAi approach has a few advantages over the conventional Pol-III-mediated shRNA expression systems. First, Pol-II expression is tissue-specific,

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whereas Pol-III expression cannot. Second, Pol-II expression is compatible with the native microRNA pathway, while Grimm et al. (49) have reported some incompatibility between the Pol-III-mediated shRNA and Pol-II-mediated native miRNA pathways. Third, excessive RNA accumulation and toxicity can be prevented by the NMD mechanism of a cellular Pol-II-mediated intron expression system, but not a Pol-III exon-like expression system (70). Lastly, one Pol-II is able to express a large size cluster (>10 kb) of polycistronic shRNAs, which can be further excised into multiple shRNAs via the native miRNA pathway, preventing the promoter conflict that often occurs in a multiple promoter vector system. For example, in many commercial U6-mediated shRNA expression systems, a self-inactivated vector promoter is often used to increase the U6 promoter activity.

8. Development of MicroRNA/ piRNA-Based Gene Therapy

The following experimentations have shown the preliminary success of silencing exogenous retrovirus replication in an ex vivo cell model of patient-extracted CD4+ T lymphocytes. The specific anti-HIV SpRNAi-rGFP vectors were designed to target the gag-pol region of about the nts +2113 to +2450 of HIV-1 genome. This region is relatively conserved and can serve as a good target for anti-HIV treatment (71). The viral genes located in this target region include 3′-proximal Pr55gag polyprotein (i.e., matrix p17 + capsid p24 + nucleocapsid p7) and 5′-proximal p66/p51pol polyprotein (i.e., protease p10 + reverse transcriptase); all these components have critical roles in viral replication and infectivity. During the early infection phase, the viral reverse transcriptase transcribes the HIV RNA genome into a double-stranded cDNA sequence, which forms a pre-integration complex with the matrix, integrase and viral protein R (Vpr). This complex is then transferred to the cell nucleus and integrated into the host chromosome, consequently establishing the HIV provirus. We hypothesized that although HIV carries few reverse transcriptase and matrix proteins during its first entry into host cells, the co-suppression of Pr55gag and p66/p51pol gene expressions by miRNAs is expected to eliminate the production of infectious viral particles in the late infection phase. Silencing Pr55gag may prevent the assembly of intact viral particles due to the lack of matrix and capsid proteins, while suppression of protease in p66/p51pol can inhibit the maturation of several viral proteins. HIV expresses about nine viral gene transcripts, which encode at least 15 various proteins; thus, the separation of a polyprotein into individual functional proteins requires the viral protease activity. As shown in Fig. 19.6, this therapeutic approach has been reported to be feasible (22,47).

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Fig. 19.6. Silencing of HIV-1 genome replication using anti-gag/pro/pol miRNA transfections into CD4+ T lymphocytes isolated from the acute and chronic phases of AIDS infections. (a) Northern blot analysis showed about 98 and 70% decreases of HIV genome in the acute and chronic infections after miRNA treatments (lanes 4), respectively. No effect was detected in the T cells transfected by miRNA* targeting the same gag/pro/pol region of the viral genome (lane 5). The size of pure HIV-1 provirus was measured about 9,700 nucleotide bases (lanes 1). RNA extracts from normal noninfected CD4+ Th lymphocytes were used as a negative control (lanes 2), whereas those from HIV-infected T cells were used as a positive control (lanes 3). (b) Immunostaining of HIV p24 marker confirmed the results of (a). Since the ex vivo HIV-silenced T lymphocytes were resistant to any further infection by the same strains of HIV, they may be transfused back to the donor patient for eliminating HIV-infected cells.

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The anti-HIV SpRNAi-rGFP vectors were tested in the CD4+ T lymphocyte cells from HAART-treated, HIV-seropositive patients. Because only partial complementarity between miRNA and its target RNA is needed to trigger the gene silencing effect, it would be very advantageous to overcome the daunting challenge of high HIV mutations, which frequently generate new drug resistance to current small molecule drugs. Northern blot analysis in Fig. 19.6a demonstrated the ex vivo gene silencing effect of anti-HIV miRNA transfections (n = 3 for each set) on HIV-1 replication in CD4+ T lymphocytes from both acute and chronic phase AIDS patients. In the acute phase (≤1 month), the 50 nM miRNA vector transfection degraded an average of 99.8% viral RNA genome (lane 4), whereas the same treatment knocked down only an average of 71.4 ± 12.8% viral genome replication in the chronic phase (about 2-year infection). Immunocytochemical staining of HIV p24 marker protein confirmed the results of Northern blot analysis (Fig. 19.6b). Sequencing analysis has revealed that at least two HIV-1b mutants in the acute phase and seven HIV-1b mutants in the chronic phase were found within the targeted HIV genome domain. It is likely that the higher genome complexity of HIV mutations in chronic infections is able to counteract the miRNA-mediated silencing efficacy. Transfection of 50 nM miRNA* vector hom*ologous to the HIV-1 genome failed to induce any RNAi effect on viral genome, indicating the specificity of the miRNA effect (Fig. 19.6b, lane 5). Expression of cellular house-keeping gene, β-actin, was at a normal level and showed no interferon-induced non-specific RNA degradation. These results suggest that the designed anti-HIV SpRNAi-rGFP vector is highly specific and efficient in suppressing HIV-1 replication in the early infections. In conjunction with an intermittent interleukin-2 therapy (47), we may stimulate the growth of non-infected CD4+ T lymphocytes to eliminate the HIV-infected cells.

9. Conclusions The consistent evidence of miRNA-induced gene silencing effects in zebrafish, chicken embryos, mouse stem cells and human diseases demonstrates the preservation of an ancient intronmediated gene regulation system in eukaryotes. In these animal models, the intron-derived miRNAs determine the activation of RNAi-like gene silencing pathways. We herein provide the first time evidence for the biogenesis and function of intronic miRNAs in vivo. Given that natural evolution gives rise to more complexity and more variety of introns in higher animal and plant species for coordinating their vast gene expression volumes and interac-

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tions, dysregulation of these miRNAs due to intronic expansion or deletion will likely cause genetic diseases, such as myotonic dystrophy and fragile X mental retardation. Thus, gene expression produces not only gene transcripts for its own protein synthesis but also intronic miRNAs, capable of interfering with other gene expressions. Based on this concept, the expression of a gene results in gain-of-function of the gene and also loss-of-function of some other genes, which contain complementarity to the mature intronic miRNAs. An array of genes can swiftly and accurately coordinate their expression patterns with each other through the mediation of their intronic miRNAs, bypassing the time-consuming translation processes under quickly changing environments. Conceivably, intron-mediated gene regulation may be as important as the mechanisms by which transcription factors regulate the gene expression. It is likely that intronic miRNA is able to trigger cell transitions quickly in response to external stimuli without the tedious protein synthesis. Undesired gene products are reduced by both transcriptional inhibition and/or translational suppression via miRNA regulation. This could enable a rapid switch to a new gene expression pattern without the need to produce various transcription factors. This regulatory property of miRNAs may serve as one of the most ancient gene modulation systems before the emergence of proteins. According to the variety of microRNAs and the complexity of genomic introns, a thorough investigation of miRNA variants in the human genome will markedly improve the understanding of genetic diseases and also the design of miRNA-based drugs. Learning how to exploit such a novel gene regulation system in future therapy will be a forthcoming challenge. References 1. Ambros, V., Lee, R.C., Lavanway, A., Williams, P.T., and Jewell, D. (2003). MicroRNAs and other tiny endogenous RNAs in C. elegans.Curr Biol. 13, 807–818. 2. Lin, S.L., Chang, D., Wu, D.Y., and Ying, S.Y. (2003). A novel RNA splicing-mediated gene silencing mechanism potential for genome evolution.Biochem Biophys Res Commun. 310, 754–760. 3. Rodriguez, A., Griffiths-Jones, S., Ashurst, J.L., and Bradley, A. (2004). Identification of mammalian microRNA host genes and transcription units.Genome Res. 14, 1902–1910. 4. Lin, S.L., Chuong, C.M., and Ying, S.Y. (2001). A Novel mRNA-cDNA interference phenomenon for silencing bcl-2 expression in human LNCaP cells.Biochem. Biophys. Res. Commun. 281, 639–644.

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Chapter 20 Emergence of a Complex Relationship Between HIV-1 and the microRNA Pathway Dominique L. Ouellet, Isabelle Plante, Corinne Barat, Michel J. Tremblay, and Patrick Provost Abstract Recent experimental evidences support the existence of an increasingly complex and multifaceted interaction between viruses and the microRNA-guided RNA silencing machinery of human cells. The discovery of small interfering RNAs (siRNAs), which are designed to mediate cleavage of specific messenger RNAs (mRNAs), prompted virologists to establish therapeutic strategies based on siRNAs with the aim to suppress replication of several viruses, including human immunodeficiency virus type 1 (HIV-1). It has been appreciated only recently that viral RNAs can also be processed endogenously by the microRNAgenerating enzyme Dicer or recognized by cellular miRNAs, in processes that could be viewed as an adapted antiviral defense mechanism. Known to repress mRNA translation through recognition of specific binding sites usually located in their 3¢ untranslated region, miRNAs of host or viral origin may exert regulatory effects towards host and/or viral genes and influence viral replication and/or the host response to viral infection. This article summarizes our current state of knowledge on the relationship between HIV-1 and miRNA-guided RNA silencing, and discusses the different aspects of their interaction. Key words: HIV-1, RNA silencing, microRNA, small interfering RNA, gene expression.

1. Biology of microRNAs 1.1. microRNAs as Key Regulators of Gene Expression

microRNAs (miRNAs) are short ~21 to 24-nucleotide (nt) RNA species expressed in most eukaryotes and are known as key regulators of gene expression that act through imperfect base pairing with their target messenger RNA (mRNA) (1,2). According to the latest update of miRBase (release 11.0, April 2008), the repository of miRNA data on the Web, more than 678 human

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miRNA sequences have been identified so far, among a total of 6396 entries (3,4). A recent study suggested that miRNAs may regulate up to 92% of the genes in humans (5)! The panoply of small gene regulatory RNAs has recently gained in complexity with the discovery in eukaryotic organisms of additional classes distinct from miRNAs, such as repeatassociated small interfering RNAs (rasiRNAs) (6), tiny noncoding RNAs (tncRNAs) (7) and Piwi-interacting RNAs (piRNAs) (6). 1.2. The Endogenous microRNA-Based RNA Silencing Machinery

Encoded by the genome of most eukaryotes examined so far, miRNA genes are transcribed by RNA polymerase (pol) II into stem-loop structured primary miRNAs (pri-miRNAs). Harboring a 5´m7G cap and a 3´ poly(A) tail (8,9), these pri-miRNAs are then trimmed into ~60–70-nt miRNA precursors (pre-miRNAs) (see Fig. 20.1a) by the nuclear ribonuclease (RNase) III Drosha (10), acting in concert with the DiGeorge syndrome critical region 8 (DGCR8) protein within the microprocessor complex (11–14). The pre-miRNAs are subsequently exported to the cytoplasm via Exportin-5 (15–18) and the base of their stem recognized by the PAZ domain of Dicer (19). Acting as an intramolecular dimer, the RNase IIIa and IIIb domains cleave the stem at the base of the loop to generate miRNA:miRNA* duplexes (19, 20, 21, 22). Dicer was recently shown to operate with the transactivating response RNA-binding protein (TRBP) (23) within a pre-miRNA processing complex (24,25). Following a strand selection and separation step, which is based on the thermodynamic stability of the RNA duplex (26), the miRNA strand (~21 to 24-nt) with the least stable 5´ end pairing (called the guide strand) is incorporated into effector miRNAcontaining ribonucleoprotein (miRNP) complexes, containing Argonaute 2 (Ago2), TRBP and Dicer (25), and guiding them towards specific messenger RNAs (mRNAs). The opposite miRNA* strand (also called passenger strand) is encountered much less frequently and is presumably degraded (27). miRNA assembly on specific mRNA sequences may be facilitated by the fragile X mental retardation protein, which can accept and use miRNAs derived from Dicer (28). The targeted mRNA will be primarily subjected to translational repression, although mRNAs containing partial miRNA complementary sites may also be targeted for degradation in vivo (29). These regulatory events may occur at specific cytoplasmic foci referred to as processing bodies (P-bodies) (30,31), or GW182-containing bodies (GW-bodies) (32), which are formed as a consequence of the presence of miRNAs (33). P-bodies are enriched in proteins involved in RNA-mediated gene silencing, such as Ago2 (30) , mRNA degradation (34) and nonsense-mediated mRNA decay (35,36)

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Fig. 20.1. Schematic representation of the intricate relationship between the miRNA silencing pathway and HIV-1. (a) In HIV-1-infected cells, full-length and short viral RNA transcripts are produced from the integrated HIV-1 genome. In these cells, like in normal non-infected cells, miRNA genes are transcribed into primary miRNAs (pri-miRNAs), which are trimmed into miRNA precursors (pre-miRNAs) in the nucleus. Pre-miRNAs are then exported to the cytoplasm where they are cleaved by Dicer to generate miRNA:miRNA* duplexes. Following a strand selection and separation step, the mature miRNA is incorporated into effector complexes to mediate recognition and translational repression of specific cellular mRNAs, as reviewed recently (107). (b) Secondary structures in HIV-1 mRNAs may themselves be recognized and processed by Dicer into viral miRNAs or siRNAs, which could act on gene regulation in the cell, i.e., by restricting LTR-driven transcription, recruit HDAC-1 to the LTR or repress cellular mRNAs. (c) TAR recognition by the Dicer•TRBP complex could be hampered by the viral transactivating protein Tat, which has been suggested to inhibit Dicer activity and could lead to suppression of silencing. This inhibitory effect of Tat remains unclear. (d) In infected cells, HIV-1 could alter host gene expression through the modulation of miRNA expression. For instance, downregulation of the miR-17/92 cluster, via miR-17-5p and miR-20, has been reported to increase expression of a Tat cofactor, the P/CAF protein. This regulation could lead to an enhanced transactivation of the TAR element and could contribute to activating latent reservoirs. (e) A 1.2-kb fragment present in the 3´UTR of almost all HIV-1 mRNAs can be recognized by cellular miRNAs with a negative impact on viral protein production in CD4+ T cells, following a process that could contribute to keep the virus in its latency phase.

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1.3. Biological Roles of microRNAs

miRNAs have been shown to control various processes, such as cell proliferation and apoptosis in flies, and hematopoietic cell differentiation in mice (37). Their biological role is linked to their ability to initially repress translation of specific mRNAs, although a case of enhancement of translation mediated by miRNAs/miRNPs during the cell cycle was recently reported (38). The action of miRNAs is accomplished through recognition of specific miRNA binding sites usually located in the mRNA 3´ untranslated region (UTR), thereby inhibiting translation initiation (39). Because recognition by miRNAs is mainly based on imperfect sequence complementarity, the identification of their physiological mRNA targets is difficult to predict and is rather arduous. Characterization of a few experimentally validated miRNA:mRNA interactions (e.g., let-7 and lin-41) (40) allowed to establish a context in which this interaction is favored. For example, the critical miRNA:mRNA pairing region, referred to as the “miRNA seed,” involves nucleotides 2–8 of the miRNA in the 5´ to 3´ orientation. Although it appears to be less important, pairing of the miRNA 3´ region may compensate a weaker binding of the 5´ region (40). A better understanding of mRNA recognition by miRNAs helped develop bioinformatic approaches that have proven to be instrumental for identifying potential miRNA targets and initiating characterization of miRNA function.

1.4. A Role for Small RNAs in Antiviral Host Defenses

In addition to fulfilling important gene regulatory functions in their eukaryotic hosts, small RNAs may also help defend against invasion of the host genome by RNAs of foreign origin, such as viruses. Initial evidences for such a role came from observations made by plant biologists. Indeed, while investigating the natural antiviral defense mechanism known as posttranscriptional gene silencing (PTGS), Hamilton and Baulcombe detected the presence of antisense viral RNA of ~25-nt in virus-infected plants by Northern blot (41). The authors noted that these small RNAs, which were later found to originate from viral double-stranded RNA (dsRNA) processing by Dicer or DICER-LIKE 1 (DCL1 in Arabidopsis) (42), were long enough to convey sequence specificity and suggested their probable role in limiting virus infection in plants. The antiviral function of small RNAs and their biosynthetic machinery in plants has recently been extended to insects, nematodes (43,44) and mammals.

2. Biology of HIV-1 2.1. HIV-1 Life Cycle

HIV-1 is an enveloped virus that binds cell receptor CD4 and most common coreceptors C-X-C motif receptor 4 (CXCR4) in lymphocytes or C-C motif receptor 5 (CCR5) in macrophages. For more details about HIV-1 life cycle, please refer to recent reviews

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(45, 46, 47). Viral gp120 is responsible for binding, whereas gp41 is essential for fusion of the viral particle to the cellular membrane. The genome of HIV-1 is composed of two identical single-stranded 9.6 kb RNA molecules. After cell entry and uncoating, the viral genetic material is reverse transcribed into cDNA by the HIV-1 viral reverse transcriptase (RT) enzyme and integrated as double-strand DNA into the host genome before directing viral gene expression. HIV-1 relies heavily on the cellular transcription and translation machineries for the synthesis of viral genomic RNA and proteins. As partly illustrated in Fig. 20.1a, the full-length RNA is transcribed by RNA pol II and serves both as genomic RNA and as a template for expression of the structural proteins Gag and Gag/Pol. The singly spliced 4 kb mRNA encodes Vif, Vpr, Vpu and Env and the fully spliced 2 kb mRNAs encode Tat, Rev and Nef. Some HIV-1 RNA transcripts are produced from the DNA bipartite element, known as the inducer of shorts transcripts (IST), located downstream of the start site of transcription into the long terminal repeat (48,49). The subpopulation of non-polyadenylated short transcripts have heterogenous 3´ nucleotide ends situated around position +60 and contain the Trans-Activation Responsive (TAR) element (49). All these RNA species adopt complex and dynamic secondary structures that are reminiscent of pre-miRNAs and thus could potentially be targeted by the host miRNA-guided RNA silencing machinery. Although the high variability of the HIV-1 genome gives rise to a multitude of RNA folding possibilities, a number of structures are very well conserved because of their essential function in the virus life cycle. Among these are the dimerization sites TAR region and the Rev-Responsive Element (RRE). These elements all have in common dsRNA structures (dimerization or stem-loop), which may potentially be processed into miRNAs. Whether these structures are recognized and processed by the Drosha•DGCR8 or Dicer•TRBP complexes remains to be determined. 2.2. HIV-1 Dimerization Initiation Site

In virus particles, the genome consists of two identical molecules of RNA that are non-covalently linked near their 5´ ends. The dimerization process involves a series of conformational changes of the untranslated leader region in which a first structure referred as the kissing-loop complex is rearranged into a more extended molecular duplex (50) that can be targeted by synthetic molecules for potential inhibition of the dimerization initiation site (DIS). However, these conformations possess a number of dimer RNA molecules and stem-loop structures that could potentially be recognized by Drosha and/or Dicer, whereby the latter preferentially cleaves dsRNAs at their termini (22), to generate miRNAs.

2.3. HIV-1 TAR

The TAR region is a 59-nt stem-bulge-loop structure located at the 5´ end of all spliced and unspliced HIV-1 transcripts found in the nucleus and cytoplasm, which is essential for efficient viral

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transcription. TAR is a short leader RNA structure targeted by the viral transactivating protein Tat that is known to act at the RNA level to enhance virus gene expression by more than 100fold (51). Upon binding to the trinucleotide pyrimidine-rich bulge of TAR, Tat recruits the positive transcription-elongation factor b (P-TEFb), a complex made of cyclin T1 and the cyclin dependent kinase 9 (CDK9), to the initiation complex. The CDK9 then phosphorylates the C-terminal domain (CTD) of the RNA polymerase II, which promotes the formation of an efficient elongation transcription complex (52). Apart from this nuclearbased function, TAR is also important after RNA export to the cytoplasm since it inhibits translation by two mechanisms, i.e., through a direct block of translation initiation by its secondary structure and by activation of the dsRNA binding protein kinase R (PKR) which, in turn, phosphorylates the eukaryotic initiation factor 2 alpha (eIF2a), leading to an arrest of translation initiation. Both of these negative effects are alleviated by TRBP, which inhibits PKR (53) and releases the translational block due to the TAR structure (54). (For more details about HIV-1 TAR RNA and Tat protein, please refer to these reviews. (52,55,56)) 2.4. HIV-1 RRE

The RRE domain is a large RNA structure present in all 9 kb and 4 kb RNAs located within the Env intron. Through its interaction with RRE, Rev protein is responsible for the nuclear export of these unspliced or singly spliced RNAs. In the absence of Rev, these RNAs are sequestered in the nucleus and only the multiply spliced 2 kb RNA encoding the regulatory proteins Tat, Rev and Nef are exported in the cytoplasm and translated. The interaction between Rev and RRE promotes the transition between this early phase of the viral life cycle to the late phase where structural proteins are produced (57). The RRE is a 351 nt complex structure that comprises five stem-loop structures on which Rev assembles as a multimeric complex (58,59). This structure may resemble pri-miRNAs, which are often composed of multiple stem-loop structures, and represent a very good candidate for the source of viral miRNA. Although it is interesting to note that, like TAR, RRE interacts with TRBP (53), the latter does not appear to influence the effect of Rev on RRE-containing sequence.

3. Relationship Between HIV-1 and RNA Silencing 3.1. Small RNAs Directed Against HIV-1

Small interfering RNAs (siRNAs) are synthetic 21-nt RNA duplexes that have been designed to mimic the endogenous miRNAs or Dicer-generated siRNAs. Their efficiency in downreg-

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Fig. 20.2. siRNAs and shRNAs expressed from viral or mammalian expression vectors can mediate cleavage of HIV-1 RNAs and inhibit viral protein synthesis.

ulating expression of specific genes in cultured mammalian cells (60) established the basis for the use of siRNAs or RNA interference (RNAi) technology in therapeutics. This strategy was exploited successfully to inhibit the replication of several viruses, including HIV-1 (see Fig. 20.2, and Chap. 16). Approaches based on siRNA targeting of host genes have been also used to restrict HIV-1 production (61). This anti-HIV-1 therapy is one of the RNA-based strategies that include antisenses, ribozymes and aptamers. Some of them are currently being tested in clinical trials (62). Short hairpin RNA (shRNA) precursors are also used to trigger RNAi against HIV-1. shRNAs are produced either from mammalian expression vectors or viral vectors bearing H1, U6 or 7SK promoters for expression (63) (see Fig. 20.2). The gene inhibitory potency of shRNAs, which requires prior processing into effector siRNAs by Dicer, is superior to siRNAs themselves, presumably because they enter the miRNA pathway upstream to siRNAs. Such approaches may be combined, for example, with protein-based anti-HIV-1 agents, for increased therapeutic efficiency (64). Although RNAi-based antiviral therapies are promising, HIV-1 has been shown to escape RNAi induced by a specific siRNA. In these cases, the emergence of mutants was observed, either showing nucleotide substitutions or deletions within the targeted sequence (65), or evolving an alternative structure in its RNA genome occluding the siRNA binding site (66). A single substitution in the targeted sequence is sometimes sufficient to abolish the antiviral activity of siRNAs (67). Such problems may be circumvented by targeting the most conserved sequences at multiple locations in the HIV-1 genome (68).

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3.2. Viral microRNAs

The observed sequence-specific HIV-1 RNA degradation induced by siRNAs implies that certain HIV-1 RNA sequences are accessible to the RNA silencing machinery in vivo. Likely to be applicable to other viruses, this concept is supported by the demonstrated improvement of RNA-induced silencing complex (RISC)-mediated target RNA cleavage when the target site access is increased (69). Accessibility to the viral RNA is mandatory for an antiviral function of the miRNA-guided RNA silencing pathway, a research theme that has attracted the interest of several laboratories. A research group led by Thomas Tuschl was the first to investigate the role of RNA silencing in human cells infected with viruses. Regarding the small RNA profile of a Burkitt’s lymphoma cell line latently infected with Epstein-Barr virus (EBV), they found that this large DNA virus expresses several miRNAs (70). Bioinformatic analysis of the genomic sequences flanking the cloned RNAs, which were detectable by Northern blot, unveiled fold-back structures characteristic of miRNA precursors. As for plant virus-derived siRNAs (71), EBV miRNAs may originate predominantly from Dicer processing of highly structured single-stranded RNA. EBV is now known to express in latently infected cells at least 17 distinct miRNAs that are originating from two clusters located in the introns of the viral BART and adjacent BHRF1 genes. Differential regulation of EBV miRNA expression is observed and implies distinct roles during infection of different human tissues (72). Recently, EBV LMP1 was discovered as a target for miRNAs derived from BART cluster 1. This protein is implicated in the activation of cell signaling and gene expression in infected cells (73). Using similar approaches, investigation of several other viruses identified miRNAs encoded in the Kaposi’s sarcoma-associated herpesvirus (KSHV or HHV8), mouse gammaherpesvirus 68 and human cytomegalovirus (also called HHV5) (74). However, viral miRNAs derived from HIV-1 were neither predicted (using an algorithm identifying genomic regions that may assume a secondary structure similar to that of pri- or pre-miRNAs) nor found among 260 cloned miRNA sequences derived from HeLa cells stably expressing CD4 and CXCR4, and infected by HIV-1, isolate Bru (LAV-1) (74). These findings suggested that HIV-1 may effectively hide its highly structured RNA from RNase III cleavage.

3.3. HIV-1-Derived microRNAs

However, this assertion is being challenged, as concurrent studies about HIV-1 miRNAs have been reported. Using a computational method designed to uncover well-ordered folding patterns in nucleotide sequences, five candidate pre-miRNAs encoded by different regions of the HIV-1 genome were flagged (75). Omoto and colleagues (76) reported a miRNA (miR-N367) derived from the nef region, an accessory gene partially overlapping with the 3´ long terminal repeat (LTR) (see Fig. 20.1b). This HIV-1 miRNA could be detected by Northern blot analysis in MT-4 T cells per-

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sistently infected with HIV-1 IIIB and cloned from a ~25-nt RNA sub-population. Overexpression of miR-N367, which shows perfect complementarity with nef, seemed to suppress HIV-1 LTRdriven transcription in reporter gene assays (77), suggesting that this nef-derived miRNA could act as a negative regulator of HIV-1 transcription. The biogenesis and action of this particular miRNA require further investigation. Another study reported that the HIV-1 RNA genome also encodes an siRNA derived from the env gene (78). The authors observed that two RNA strands forming a perfect 19-bp duplex, and joined by an extended 198-nt loop, could be converted into siRNAs upon incubation with recombinant Dicer in vitro. A probe specific for the viral siRNA detected a ~24-nt signal not seen in mockinfected cells by Northern blot analysis (78). Overexpression of this viral siRNA effectively reduced Env mRNA levels and viral replication, whereas its neutralization with complementary 2´-O-methyl oligonucleotides led to a dose-dependent increase in HIV-1 replication in human cells (78). These results suggest that an HIV-1derived siRNA can reduce virus production (see Fig. 20.1b). Another natural HIV-1 RNA structure, the TAR element, was recently reported to be cleaved by Dicer to generate a miRNA that has been suggested to recruit the histone deacetylase HDAC-1 to the HIV-1 LTR promoter to silence transcription by chromatin remodeling (79) (see Fig. 20.1b), a concept that has been proposed previously (80). The authors hypothesize that this sequence of events may suppress transcription of viral as well as cellular genes, thereby influencing particular steps of HIV-1 pathogenesis, such as latency. Whether or not HIV-1 miRNAs are effectively produced in infected cells and fulfill important biological roles warrants further experimental validation and confirmation. A recent study by Lin and Cullen (81) is challenging the existence of miRNAs derived from primate retroviruses, such as HIV-1 and human T cell leukemia/lymphoma virus type 1 (HTLV-1), and is questioning the suppressive properties of HIV-1 Tat on RNA silencing that has been reported by Bennasser and colleagues (78) (see Fig. 20.1c). It may be that the identification of some miRNAs are restricted to specific viral strains or that miRNAs may escape detection by standard small RNA cloning strategies, since methylation of the 2¢ hydroxyl of the terminal ribose significantly reduces the cloning efficiency of silencing-associated small RNAs (82). This would explain some of the discrepancies observed between laboratories using different techniques to identify viral miRNAs. 3.4. Biosynthetic Mechanism of HIV-1 microRNAs

The controversy surrounding the existence of miRNAs derived from HIV-1 may be related to their levels of expression that may be barely detectable using the techniques currently available. If their existence is proven unequivocally, their biosynthesis would

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merit due mechanistic considerations. For instance, are these viral miRNAs processed by the successive action of Drosha and Dicer like cellular miRNAs, by Dicer only or, as in plants, by an RNAi pathway adapted to viruses? How does the cleavage of an RNA substrate flanked by genomic single-stranded RNA sequences occur, knowing the preference of Dicer for RNA duplexes bearing terminal 2-nt 3¢overhangs (22)? Is this processing linked in any way to the infectious state, considering that the latency period of HIV-1 is associated with the expression of aborted mRNA transcripts? How does the presence of an exceedingly expanded loop in env siRNA precursor influence its processing? Is the expression level of viral miRNAs related to the relatively inefficient processing of HIV-1 dsRNA substrates by RNases III and/or to the limited access to a structure embedded within the HIV-1 RNA? Although regions of the HIV-1 genome show structures relatively close to that of pri- or pre-miRNAs, the fact that they are decorated with many cellular and viral proteins may also affect their recognition and processing by Drosha and/or Dicer. 3.5. Biological Significance of HIV-1 microRNAs

The possibility that HIV-1 miRNAs exert significant biological roles and directly influence viral pathogenesis and persistence in human cells is appealing. The results of recent studies have suggested a role for HIV-1 miRNAs in transcriptional repression induced either by miR-N367 (76) or TAR miRNA binding to the LTR-driven promoter (79). Cellular mRNAs that could potentially be regulated by these viral miRNAs have been tentatively identified (75). Investigation of the human and viral genes, as well as the processes possibly regulated by HIV-1 miRNAs, which have been the subject of speculations, awaits their prior experimental confirmation and validation.

3.6. Cellular microRNAs and HIV-1 Infection

A few years ago, candidate HIV-1 genes that could be controlled by host miRNAs have been predicted in view of thermodynamically favorable miRNA:target pairing (83). In addition, changes in miRNA expression profiles, i.e., downregulation of a large pool of miRNAs, have been observed in human HeLa cells transfected with the infectious molecular clone pNL4-3 (84). A more recent study explored the importance of the miRNA pathway in the control of HIV-1 replication (85). Using siRNAs against Drosha and Dicer in peripheral blood mononuclear cells (PBMCs) from HIV-1-infected patients, Triboulet et al. (85) noticed a faster virus replication kinetic in Drosha- or Dicer-depleted cells, as compared to cells treated with a control siRNA. The authors also confirmed in latently infected U1 cells that both Drosha and Dicer contribute to the suppression of HIV-1 replication. HIV-1 infection was also associated with either up- or down-regulation of specific miRNA clusters. For instance, the miR-17/92 cluster, which encode for seven miRNAs, among

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which miR-17-5p and miR-20 may target the histone acetyltransferase and HIV-1 Tat cofactor p300/CBP-associated factor (PCAF), was substantially decreased (see Fig. 20.1d). The authors proposed that this gene regulatory axis may help understand how latent virus reservoirs could be activated. A more recent study explored the possible contribution of cellular miRNAs to HIV-1 infection. Huang et al. (86) showed that the 3´ UTR of almost all HIV-1 mRNA produced during latency in resting primary CD4+ T lymphocytes contain a 1.2-kb fragment that can be recognized by cellular miRNAs, with a negative impact on viral protein production (see Fig. 20.1e). Combined with the relatively inefficient synthesis of Tat and Rev, miRNAs harbored by resting CD4+ T cells may participate in post-transcriptional regulation of HIV-1 mRNA and contribute to keep the virus in its latency phase, as observed in patients with suppressive highly active antiretroviral therapy (HAART) (86). These new elements contribute to our understanding of the molecular basis of viral latency and help us design therapeutic strategies aimed at purging HIV-1-infected patients of the quiescent virus. 3.7. HIV-1, RNA Silencing, and RNA Editing

The susceptibility of viral RNAs to RNases III may also be modified by structural changes produced by adenosine deaminases that act on RNA (ADARs) (87,88). The predominant form of RNA editing in human consists in the specific conversion of adenosine (A) into inosine (I) within largely double-stranded cellular and viral RNAs [reviewed in Ref. (89)]. A-to-I RNA editing may thus alter base pairing of a dsRNA substrate and reduce its susceptibility to Dicer cleavage, preventing it from initiating RNAi (87). Several viral genomes or transcripts show sequence changes consistent with such modification, including HIV-1. Indeed, TAR was previously reported to be a substrate for ADAR in Xenopus oocytes and edited in a process dependent on Tat (90).Whether editing of viral RNAs may lead to viral persistence, as speculated previously (89), remains to be confirmed. Another group of deaminases from the APOBEC3 family has been reported to counteract HIV-1 replication. For example, APOBEC3G is incorporated into HIV-1 particles and acts to restrict HIV-1 replication in infected cells by deaminating dC to dU in the first (minus)-strand cDNA replication intermediate during the viral reverse transcription process, (91) which is correlated with a G-to-A modification of the second (positive)strand. Despite its action on cDNA, APOBEC3 family members may induce mutations in the HIV-1 genomic DNA prior to its integration and thereby contribute to a pool of mutant RNA transcripts (92). The viral accessory protein Vif is known to be an inhibitor of APOBEC3 (93). The effect of these editing events on the generation of mutations in dsRNA structures remains to be elucidated.

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3.8. Interaction between HIV-1 Tat and the microRNABased RNA Silencing Machinery

In addition to protein-RNA and RNA-RNA interactions, recent studies have revealed an intriguing link between protein components involved in HIV-1 pathogenesis and RNA silencing, such as the virally encoded Tat protein and the cellular TRBP. Overexpression of Tat in mammalian cells was shown to attenuate silencing of reporter genes induced by short hairpin RNAs (shRNAs), but not siRNA (78). Knowing that the former elicits RNAi upon Dicer processing, the authors investigated and determined that Tat could inhibit Dicer activity in vitro (see Fig. 20.1b). However, prior to qualifying HIV-1 Tat as a proven inhibitor of Dicer function, it would be prudent (i) to determine if the Dicer inhibitory effect of Tat can be extended in vivo and occurs at physiological expression levels, (ii) to confirm that the observed inhibitory effects of Tat are specific and not due to random binding to dsRNAs, (iii) to verify if RNAi proceeds normally in the context of HIV-1 infection, and (iv) to assess whether Dicer function is indeed inhibited by HIV-1 in infected cells. Whether HIV-1 Tat suppresses RNA silencing remains controversial. (81).

3.9. TRBP and PACT Function in RNA Silencing

HIV-1 TAR RNA-binding protein (TRBP) was originally discovered as a cellular protein that cooperates synergistically with viral Tat function and enhances transactivation of the HIV-1 5¢ LTR (see Fig. 20.1a) (23). TRBP is also known to inhibit the interferon (IFN)-induced dsRNA-regulated protein kinase R (PKR) (94), and to be involved in miRNA-guided RNA silencing, more specifically, in assisting Dicer function within a pre-miRNA processing complex (see chapter 18, section 3.3) (24,25). Immunoprecipitation approaches identified TRBP as a Dicer-interacting protein (24,25). The Dicer-binding region on TRBP could be delineated to its third C-terminal dsRNA-binding domain (dsRBD) and depletion of TRBP was found to negatively affect pre-miRNA processing using cell extracts in vitro (24). Similar to TRBP, PKR-activating protein (PACT) (95) has been found to interact with the N-terminal domain of Dicer via its third dsRBD (96). In fact, PACT can bind directly to TRBP and form a ternary complex with Dicer and TRBP to facilitate the production of siRNAs by Dicer. Knockdown of both TRBP and PACT in cultured mammalian cells led to a significant inhibition of gene silencing mediated by shRNAs, but not by siRNAs, suggesting that TRBP and PACT function primarily at the step of siRNA production (97). Despite exerting opposite effects on PKR, PACT and TRBP may thus play a similar, possibly redundant, role in miRNA biogenesis and function. The exact role of PACT in HIV-1 pathogenesis and RNA silencing remains to be clearly defined.

3.10. A Dual Role for TRBP – Implications for HIV-1

TRBP may thus exert a dual role in HIV-1 pathogenesis and RNA silencing, as recently discussed (98). The requirement of TRBP to achieve a higher virus production may have forced the virus to

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evolve under selective pressure from the RNA silencing machinery. There is a possibility that TAR and RRE RNA structures could also compete with Dicer for TRBP binding, and thus inhibit RNA silencing (98). The delicate balance thereby created may have conferred to HIV-1 the ability to replicate in TRBP-expressing cells and be responsible, to some extent, for the low virus load and persistence in HIV-1-infected individuals. Pharmacological interventions aimed at dissociating TRBP functions may represent a relevant therapeutic area to combat the HIV-1 pandemic. 3.11. Perspectives for HIV-1

A number of studies published recently have provided key insights into the increasingly complex interaction between HIV-1, miRNAs and host RNA silencing machineries. It has been known that HIV-1 induces drastic changes in gene expression programming of infected cells. With the recent idea that HIV-1 may encode miRNAs, the identification and validation of the complete HIV-1 miRNA array as well as their cellular and viral mRNA targets, which pose a considerable challenge, may significantly improve our understanding of HIV-1 pathogenesis. In particular, it may help determine to what extent the perturbed gene expression profiles in HIV-1-infected cells (99, 100, 101) can be related to virus-derived miRNAs and how it ultimately influences viral replication, latency as well as the efficiency of host defenses. This raises the attractive hypothesis that HIV-1 replication may result, at least in part, from a delicate balance between the structural requirements to support HIV-1 replication versus the potential beneficial role of HIV-1 miRNAs in conferring an advantage to the virus and/or thwarting host defenses.

3.12. Applicability to Other Viruses

The various aspects of the interaction between HIV-1 and the RNA silencing machinery may also be applicable to other viruses of global importance for human health. In turn, mechanisms described for other viruses may ultimately be transposed to HIV-1. A few examples that may be relevant include (i) herpesvirus Kaposi’s sarcoma-associated herpesvirus (KSHV), which encodes as much as 11 distinct miRNAs that may play critical roles in the establishment and/or maintenance of KHSV latent infection (102), (ii) EBV, which expresses miRNAs from its BART and BHRF transcripts that either target the viral protein LMP1 (73) or characterize type III latency in infected B lymphocytes (103), (iii) simian virus 40 (SV40), which encodes miRNAs that regulate viral gene expression and reduce susceptibility to cytotoxic T cells (104), (iv) a cellular miRNA, miR-32, that was recently shown to restrict the accumulation of the retrovirus primate foamy virus type 1 (PFV-1) (105), and, in contrast, (v) an abundant miRNA specifically expressed in the human liver, miR-122, that has been shown to assist hepatitis C virus (HCV) replication through a genetic interaction with the 5¢ noncoding region of the viral genome (106).

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4. Conclusion Further investigation on the relationship between HIV-1 and RNA silencing pathways may unveil key aspects of host-pathogen interactions, provide new insights into the persistence of the virus in infected patients and offer novel basis for anti-HIV-1 therapies.

5. Notes During the preparation of this manuscript, Ouellet et al. (108) reported the identification of two miRNAs originating from the HIV-1 TAR element, namely miR-TAR-5p and miR-TAR-3p. The functional implication of these miRNAs in HIV-1 pathogenesis remains to be elucidated.

Acknowledgments We express our gratitude to Gilles Chabot for the graphic illustrations. P. P. is a New Investigator of the Canadian Institutes of Health Research (CIHR) and Junior 2 Scholar from the Fonds de la Recherche en Santé du Québec. M.J.T. is the recipient of the Canada Research Chair in Human Immuno-Retrovirology (senior level). This work was financially supported by grant HOP83069 from Health Canada/CIHR (P.P. and M.J.T.).

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Chapter 21 Synthetic microRNA Targeting Glioma-Associated Antigen-1 Protein Naotake Tsuda, Takahi Mine, Constantin G. Ioannides, and David Z. Chang Abstract The transcription factor glioma-associated antigen-1 (Gli-1) mediates activation of the sonic hedgehog (Shh) pathway, a process that precedes the transformation of tissue stem cells into cancerous stem cells and that is involved in early and late epithelial tumorigenesis. Hypothesizing that targeting the 3′-untranslated region (3′-UTR) of Gli-1 mRNA would effectively inhibit epithelial tumor cell proliferation, we evaluated several complementary miRNA molecules for their ability to do so. The synthetic miRNAs and corresponding duplex/small temporal RNAs were introduced as 3-nucleotide (nt) loops into GU-rich portions of the 3′UTR Gli-1 sequence. One particular miRNA (miRNA Gli-1-3548) and its corresponding duplex (Duplex 3548) significantly inhibited proliferation of Gli-1 + ovarian (SK-OV-3) and pancreatic (MiaPaCa-2) tumor cells by delaying cell division and activating late apoptosis in MiaPaCa-2 cells. Here, we describe the design of effective miRNA sequences and their applications as anti-gene agents. Key words: Glioma-associated antigen-1, microRNA, ovarian cancer, pancreatic cancer, apoptosis

1. Introduction Small RNAs are a novel class of active molecules, which effectively control mRNA translation, mRNA-stability and are abundantly expressed in cells (1–3). To date this class of RNA contains, the short double stranded-RNAs, (siRNA), the single strandednegative strand microRNAs (miRNA), the Piwi-interacting RNA (piRNA) and the small RNAse L activators (4–5). Most siRNAs originate from the in-frame mRNA-sequence, while miRNAs contain 3′-UTR mRNA sequences and piRNAs are found

M. Sioud (ed.), Methods in Molecular Biology, siRNA and miRNA Gene Silencing, vol. 487 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-60327-547-7_21

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only in spermatocytes (see Chaps. 18 and 19). RNase L activators are collectively referred to as 2–5A (px5′A (2′p5′A)n; x = 1–3; n > 2 (5). Animal miRNAs, siRNAs and piRNAs contain 5′-phosphate and 2′3′-hydroxy-termini. piRNA and rasiRNAs are 2′-O-methylated, which protects small RNAs from degradation by 3′-5′ exonucleases. Small molecule activators of RNAse L are cyclical RNA forms (5). In addition to classical small RNAs, a large number of intermediates are present in cells. Delivery of small RNAs to cells in vitro uses transfection. Novel approaches for RNA delivery are being developed (see Chap. 3). Linking of small RNAs to peptides, proteins, and antibodies to extracellular receptors or particles can ensure targeted delivery. For example, endocytosed proteins via heat-shock protein (HSP) receptors or specific cancer cell receptors can be delivered to the endoplasmic reticulum (ER), and in interfere in situ with protein synthesis and translation (6, 7). Phagocytosed particles by macrophages and dendritic cells are delivered to cytoplasm. In contrast to siRNAs, miRNAs are genome encoded as long RNA precursors that are processed into 21–22 short RNAs (see Chaps. 18 and 19). These non-coding RNAs have been identified in the genomes of a wide range of multicellular life forms, includ ing plants and animals. The function of miRNAs in vertebrates and mammals is largely unknown, but studies in Caenorhabditis elegans and Drosophila melanogaster have revealed that miRNAs can bind to target sites in mRNAs with imperfect base pairing and, by unknown mechanisms, significantly reduce translational efficiency (8, 9). miRNAs are found both in normal tissues and cancer cells. More than half of the human miRNA genes are located at sites known to be involved in cancers, such as fragile sites, minimal regions of loss of heterozygosity, minimal regions of amplification or common breakpoint regions. Such locations suggest that some miRNAs are involved in tumorigenesis (10). miRNA expression profiles of a large number of human tumor samples show that miRNAs are generally, though not always, downregulated in tumors and that such downregulation is often associated with poor prognosis (11). For example, miR-143 and miR-145 expression is downregulated in colorectal neoplasia and miRNA let-7 expression in lung cancer, whereas expression of the precursor of miR-155 is upregulated in pediatric Burkitt lymphoma. These observations, coupled with the observation that naturally occurring miRNAs also regulate stem cell division (12, 13), imply that miRNAs may act as both tumor suppressors and oncogenes. Although several studies reported on gene silencing by miRNAs, RNA sequences that generate miRNAs inhibiting the expression of regulatory proteins have not yet been described for the glioma-associated antigen 1 (Gli-1) gene. Approaches for synthetic

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miRNA design targeting cancer genes have not been yet established. Here we describe our approaches of designing novel miRNA targeting Gli-1. We engineered Gli-1 miRNA to downregulate protein expression. Autonomous activation of the Gli-1/2 pathway provides metastatic tumor cells with a means of efficiently proliferating at a distance from continually Shh-expressing epithelium (14). Because Gli function is a last and essential step of the Shh pathway, its inhibition may inhibit Shh signaling at any level (15, 16). Thus, Gli-1/2 pathway-blocking agents might be used to combat epithelial cancers. For example, cyclopamine and other small molecule inhibitors have been used in animal models to inhibit Smo activity in prostate and pancreatic cancer xenografts with good results (17). We found that our engineered Gli-1 miRNA and Gli-1 small temporal/duplex miRNA (Duplex 3548) (to which we gave the name “butterfly RNA”) significantly inhibited the proliferation and division of the tumor cells. We also unexpectedly observed that Duplex 3548 inhibition led to a decrease in the number of tumor cells overexpressing the HER-2 receptor (HER-2hi). Together, our findings suggested that in vitro miRNA inhibition of Gli-1 might be a potentially novel approach to the treatment of pancreatic cancer. Unlike naturally occurring RNA, our mutant synthetic miRNAs may be able to activate cytokine responses in tumors via RNA receptors (6). A number of cancer cells are sensitive to cytokines, such as IFN-α and TNF-α/β and IFN-γ, which may induce apoptosis in cancer cells. However, systemic administration of cytokines resulted in high toxicity and weak effects because of their very short life. Therefore, if our synthetic miRNAs approach turns out to be clinically feasible, it might help overcome the known shortcomings of systemic cytokine therapy for cancers. Potential target RNA receptors include the intracellular Tolllike receptors (TLRs) (see chapt. 2). The TLRs are intracellular receptors; some are located in the endoplasmic reticulum. TLR-3 binds dsRNA while TLR-7 and TLR-8 bind single-stranded RNA. Activation of signaling by TLRs results in cytokine production. TLRs, which bind TLR-3, TLR-7 and TLR-8, activate signaling through the adaptor MyD88. The activation of MyD88 enhances TLR expression and production of cytokines which include IFN-α and TNF-α, and in some instances IL-12. These cytokines may promote autoimmunity (18). Small-nuclear RNAs complexed with proteins (RNP) perform similar functions. The reverse phenomenon is the regulation of TLR-expression levels by miRNAs (19). Foreign RNAs from negative strand viruses such as influenza are well-known activators of cytokine responses in cancer cells and activate TLR7 and -8. The cytoplasmic RNA receptors

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include retinoic-acid-inducible-gene-I (RIG-1/DDX58) and melanoma differentiation-associated gene (MDA-5/IF1H1) (20). These receptors contain N-terminal caspase activation and recruitment motifs. RIG-1 and MDA-5 relay their signals to interferon regulatory factor-3 (IRF-3) and NF-κB. Recent studies showed that the 5′-triphosphate is essential for RIG-1 recognition of viral RNAs (21). Notably, RNase L is a peculiar RNA-response receptor. It becomes activated by increase in the concentration of RNA and functions to degrade RNA. The small self-RNAs produced by action of RNase L on cellular RNA induce IFN-β expression (22, 23). The ability of miRNAs to activate IFNs but not IL-6 or IL-1 is important for their anticancer effects. This difference may explain the double-faced “Janus” effects of miRNA described above. Activation of IL-6 became a topic of recent concern in liver and colorectal cancers, because it promotes cancer growth (24, 25). As such it will be important to design small RNAs which can activate more IFN production than IL-6 and IL-1 production. How this can be accomplished is still unknown. Based on the differential cytokine responses to viruses (26), we predict that changes in RNA sequence and structure will be beneficial to learn how to activate anti-cancer responses with small-synthetic RNAs. The methodology for functional small RNA design is presented below.

2. Materials 2.1. Reagents

1. RPMI 1640 medium. 2. Fetal calf serum. 3. Propidium iodide. 4. Ribonuclease A. 5. Lipofectamine-2000. 6. Control siRNA-FITC (Dharmacon, Chicago, IL). 7. MTT cell proliferation assay kit (Molecular Probes, Eugene, OR). 8. TACS Annexin V-FITC apoptosis detection kit (R&D Systems, Minneapolis, MN).

2.2. Buffers

1. Permeabilization buffer: 4X eBioscience permeabilization solution. 2. Phosphate-buffered saline (PBS): 137 mM NaCl, 10 mM Phosphate, 2.7 mM KCl, pH 7.4.

2.3. Cell Culture

MiaPaCa-2 (pancreatic cancer) and SK-OV-3 (ovarian cancer) were obtained from the American Type Culture Collection (ATCC)

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(Manassas, VA) and maintained in culture in complete RPMI 1640 medium containing 10% FCS and antibiotics. Cell lines were screened for mycoplasma contamination and were found mycoplasma free. Cells were treated periodically with BM-cyclin (Boehringer) to maintain them mycoplasma free. 2.4. Immunofluorescence

1. Antibodies against Gli-1 and Gli-2 (Santa Cruz Biotechnology, CA). The anti-Gli-1 antibody was raised against a peptide mapping the C-terminus of Gli-1 of human origin, and the antibody against Gli-2 was raised against a peptide mapping near the N-terminus of Gli-2 of human origin. 2. Human IgG (Sigma Chemical Co. St. Louis, MO). 3. Mouse anti-HER-2-PE-conjugated mAb (Sigma chemical Co. St. Louis, MO). 4. Mouse IgG1-PE conjugated.

2.5. microRNAs

1. The used microRNAs are synthesized as a custom order by Dharmacon with 5′-phosphate groups added, and they are purified by high-pressure liquid chromatography (HPLC). 2. microRNAs are dissolved in RNAse-free water at 20 nM, stored in aliquots at −20°C. 3. Lipofectamine2000 transfection reagent (Invitrogen). 4. Negative control siRNA-FITC (fluorescein-labeled luciferase GL2 Duplex) was also obtained from Dharmacon.

3. Methods 3.1. Immunofluorescence

1. Expression of HER-2 protein is determined using the PEconjugated HER-2-specific mAb clone Neu 24.7 (IgG1κ). Expression of Gli-1 is detected with mAb listed in Sect. 2. 2. To increase the sensitivity of staining and reduce background, tumor cells are first incubated with 5 µl of a solution of 1 mg/ml of purified human IgG per 106 tumor cells, then washed and stained with mouse anti-HER-2-PE-conjugated, mAb or with isotype control mouse IgG1-PE conjugated. 3. These antibodies react with Gli-1 and Gli-2 by immunofluorescence. 4. To detect and quantitate the intracellular Gli-1 and Gli-2 proteins, tumor cells are permeabilized using the eBioscience permeabilizationn buffer. Subsequently, to permeabilization, tumor cells are then incubated for 1 h with human IgG1 as blocking antibody followed by Gli-1- and Gli-2- specific Abs. The incubation was done for 1 h at room temperature.

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5. The fluorescence of cells should be determined over a range of Ab concentrations. However, 1.0 µg anti-Gli-1/50,000 cells, and 0.25 µg secondary Ab-FITC conjugated gave the lowest background staining. 6. The sensitivity of the staining can be verified by fluorescent or confocal microscopy. 7. Cells are then analyzed in a Becton Coulter, Excalibur, flow cytometer, and the data are analyzed with a Cell Quest software program. 8. The effects of miRNAs on cell sizes are determined from the forward scatter (FS) geometrical means values (x2). 9. Cells of FS > 500 are designated as large-size (LS), while cells of FS < 500 are designated as intermediate size (IS). 10. The relative density of the HER-2 and Gli-1 protein in tumor cells is determined from the geometrical mean (y2) of fluorescence intensity (MFI) of Ab staining. 3.2. Design of miRNAs

1. The sequences of Human HER-2/neu and human Gli-1 cDNA containing 5′- and 3′-UTR are present in NCBI database (accession numbers HER2/neu: M11730//Gli-1: NM 005269). 2. Both UTR sequences are analyzed for the presence of candidate siRNAs using the algorithms listed by Dharmacon on its Web site (www.dharmacon.com). 3. According to the criteria of Dharmacon siDESIGN Center, the following design criteria are used. For more detail, see Chap. 1. (a) 30–52% GC Content—add 1 point for satisfying this criterion. (b)

At least 3 A/Us at positions 15–19 (sense)—add 1 point for each A/U for a total up to 5 points. At least 3 points are required to be scored as positive in the final output.

(c)

Absence of internal repeats—add 1 point for satisfying this criterion.

(d)

A at position 19 (sense)—add 1 point for satisfying this criterion.

(e)

A at position 3 (sense)—add 1 point for satisfying this criterion.

(f)

U at position 10 (sense)—add 1 point for satisfying this criterion.

(g)

No G/C at position 19 (sense)—subtract 1 point for satisfying this criterion.

(h)

No G at position 13 (sense)—subtract 1 point for satisfying this criterion.

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All siRNA candidates are evaluated and scored by the SMART selection algorithm and then filtered after a modified BLASTn analysis. The maximum score is 10. 4. Although siRNA binds in the translated region of the gene, microRNA binds in the 3′untranslated regions (3′UTR). 5. Candidate siRNA sequences in 3′ UTR selected by algorithms 1 and 2 are then manipulated to design miRNA. 6. Modification of anti-sense RNA is performed by targeting its central region. We also modified positions 10–11 of the anti-sense to mismatch (if sense is A, it was changed to C or G, if sense is G, it was changed to G or A to avoid G-U wobble) to the identical position of sense, and one base is inserted between position 10 and 11 of anti-sense, which would make a loop toward identical sense sequence (see Note 1). 7. This inserted base was designed such that it should not bind to either position 10 or 11 of sense strand. For example, in Gli-1 3548miRNA, AU in position 10–11(counting from 5′end) of the anti-sense strand replaced to UCA, CCC, CCA and UCC to create a loop between the anti-sense and sense strands RNA. 8. After defining these candidates, each GC% and ∆G are calculated. 9. Among these candidates, the ones which have lower ∆G than the unmutated sense and antisense strands but still are closer to the ∆G of the unmutated sense and antisense, and 36–50% GC content were selected for experimental testing (see Note 2). 10. Verification of the specificity for the target sequences of the designed miRNA is performed with the algorithms provided by Drs. Peter Scacheri and Frances Collins, Human Genome Program, National Institute of Health (31). 11. In brief, algorithm 1 identifies the miRNAs which are hom*ologous in the first 5′ nt and the last 3′ nt. 12. Then, if miRNA is 19 nt long, 7 + 7 = 14, 19 − 14 = 5, it follows that only two miRNA differ in the central 5 nt left from Gli-1 3418. 13. Consequently only one miRNA is hom*ologous with Gli-1 3418 (8 + 8 = 16, 19 − 16 = 3). 14. Algorithm 3 searches for matches between the 3′UTRs and the sequence of the 5′ anchor of each miRNAs strand. 15. The miRNA candidates have at least exact match in 7 bases anchor length in the first 5′ nt and the last 3′ nt through algorithm 1, or 12 bases anchor length in the first 5′ nt through

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Table 21.1 Selected 3′ UTR mRNA sequences are Gli-1 specific Start(sense)

Sequence (5′→3′))

Number of mRNA of hom*ologous sequence in the first and last (5–9) nt 5

Gli-1 3418

GGAAATGCGATCTGTGATG

333

Gli-1 3414

ATGCGATCTGTGATGGATG

Gli-1 3548

CATTATCAAATTTCTCCTC

209

HER-2 4350

AGCCUGGAUGGAUGACACCA

350

Number of mRNA of hom*ologous sequence in the first (8–14) nt 8

Gli-1 3418

GGAAATGCGATCTGTGATG

508

Gli-1 3414

ATGCGATCTGTGATGGATG

Gli-1 3548

CATTATCAAATTTCTCCTC

410

126

HER-2 4350

AGCCUGGAUGGAUGACACCA

723

124

algorithm 3 were selected as a best candidate miRNAs for in vitro testing (see Note 2). The results of algorithm were shown in Table 21.1. 16. All of these candidates are checked whether they have any conserved sequence in the 5′ends of miRNA (see Note 3). 17. All miRNA sequences are BLAST searched in the National Center for Biotechnology Information’s (NCBI) “search for short nearly exact matches” mode against all human sequences deposited in the GenBank and RefSeq databases and are not found to have significant hom*ology (>17 contiguous nucleotides of identity) to genes other than the targets (see Note 4). The design of synthetic miRNA targeting 3′-UTR Gli-1 mRNA are shown in Table 21.2. 3.3. dsRNA and miRNA Transfection

1. dsRNA and miRNAs are transfected in SKOV3 and MiaPaCa-2 cells using Lipofectamine-2000. At the time of transfection, the cells are 50~70% confluent. 2. miRNAs are added to each well in 100 micro of serum-free RPMI medium/well at final concentrations of 83, 167, 250

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Table 21.2 Design of synthetic microRNAs targeting 3′UTR Gli-1 mRNA Free energy(kcal/mol) Anti-sense : sense Design

Gli-1 mRNA

Sequence (5′→3′)

3414 sense

CATCCATCACAGATCGCAT-3′

3414 anti-sense

AUGCGAUCU GU GAUGGAUG

−30.9

3414 microRNA

AUGCGAUCUAAAGAUGGAUG

−21.5 (−9.4)

3418 sense

CATCACAGATCGCATTTCC

3418 anti-sense

GGAAAUGCG AU CUGUGAUG

−26.4

3418 microRNA

GGAAAUGCGUCACUGUGAUG

−18.9 (−7.5)

3548 anti-sense

CAUUAUCAAA UUUCUCCUC UUAUCA AAUCUCCAGGGGUAC G

1: Selected the 3’UTR region with best hom*ology with miR-361

GAGGAGAAAUUUGAUAAUG ACUCCAUUUGUUUUGAUGAUGGA UUUGAUAA

2: The sense strand contains a conserved miR 8-mer.

miR-361*

3548 sense miR-136** Conserved 8-mer 3548 anti-sense 3548 microRNA

3548 Duplex microRNA

CAUUAUCAA A UUUCUCCUC 5’-CAUUAUCAAUCCUUCUCCUC-3’

−21.9

3-nt Loop in Position 10

3: Loop introduction in the anti-sense strand.

−16.0 (−5.9)

−27.9 (+6.0) 3’-GUAAUAGUUAGGAAGAGGAG-5’ 5’-CAU UAUCAAUCCUUCUCCUC-3’

4: Complementary sequence with microRNA 3548 for the loop in the sense strand.

and 500 nM. The volume of the Lipofectamine 2000 is maintained at a constant 0.0025% of the total volume. The miRNA-cationic liposome complexes are incubated together for 30 min at room temperature prior to adding the tumor

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cells. Subsequently, the cells are incubated for 4 h in 200 µl serum-free RPMI medium/well. 3. Control transfections are performed in parallel with negative control siRNA-FITC. 4. After 4 h of incubation, 200 µl of RPMI medium containing 20% FCS is added to each well. The final volume is now 400 µl/well. 5. At concentrations below 83 nM, the siRNA-FITC failed to transfect more than 65% of SKOV3 cells. 6. To transfect SKOV3 cells with the same amount of RNA, we used 83 nM of dsRNA and 167 nM of single-stranded miRNAs (see Note 5). The decreased expression of Gli-1 protein after transfection of Gli-1 miRNA-3418, Gli-1 duplex 3548 or HER-2 miRNA-10GGA is shown in Fig. 21.1.

% Live gated cells of total cells

IS-cells

LS-cells

D 80

80 % Decrease in Gli-1 positive cells

* 60

Duplex 3548

250

miRNA-3418 or 10GGA (nM)

167

500

167

500

Fig. 21.1. Gli-1 miRNA decreases expression of Gli-1 protein in tumor cells. (a) and (b) Gli-1 miRNAs increased the number of intermediate-size (IS) tumor cells and decreased the number of large-size (LS) cells. (c) and (d) Gli-1 miRNAs decreased the number of IS and LS Gli-1+ cells. () Gli-1 Duplex 3548, (●) Gli-1 miRNA 3418,. *, p < 0.05, at least 50% decrease in cell numbers compared with nontransfected miRNAs.

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1. Prior to transfection with miRNAs, plate the cells for 24 h. 2. After transfection, add 10% FCS and incubate the cells for 48–72 h. Subsequently, detach and determine the cell number. 3. Non-adherent cells are separated from adherent cells by gentle pipetting. 4. Adherent cells are detached by incubation with 10 mM EDTA in phosphate-buffered saline (PBS). 5. Inhibition of tumor cell proliferation is determined using the classic MTT cell proliferation assay kit (Molecular Probes, Eugene, OR).

3.5. Cell Division Assay

1. Tumor cells are resuspended in PBS at a final concentration of 1 × 106 cells/ml. 2. Then, 2 µl of a 5 mM stock CFSE solution is added to each milliliter of cells in solution to a final working concentration of 10 µM. The mixture is then incubated at 37°C for 7 min. 3. Staining is quenched by adding five volumes of ice-cold RPMI medium containing 10% FCS to the cells and incubating the cells on ice for 5 min. 4. The cells are then pelleted by centrifugation. 5. The resulting pellet is washed by being suspended in fresh media. This is done three times. 6. The washed cells are plated onto 24-well plates in 1 ml of RPMI medium containing 10% FCS per well and then cultured for 24 h. 7. The cells are transfected with miRNAs for 48 h, after which they are subjected to flow cytometric analysis. 8. The number of generations and the number of cells in each generation are calculated using Flow-Jo software for Windows (Tree Star, Inc., Ashland, OR).

3.6. Apoptosis Assay

1. To address whether miRNAs induced death by apoptosis, we determined the number of early and late apoptotic MiaPaCa-2 and SKOV-3 cells in response to miRNAs and to duplex-3548. 2. The percentage of early and late apoptotic cells are determined with the TACS Annexin V-FITC apoptosis detection kit. 3. The percentage of apoptotic cells are quantitated by flow cytometry after fluorescent staining with Annexin-FITC and PI. The results of the apoptosis assay are shown in Fig. 21.2.

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Fig. 21.2. (a–f), Gli-1 miRNA 3548 (3548) and Duplex 3548 induced late apoptosis. Cells were stained 48 h after transfection. “Cells,” control untransfected MiaPaCa-2 cells. Double-positive cells in the upper right quadrant [PI (Red)+Annexin-V-FITC(Green)+] represent late apoptotic cells. Cells in the bottom right quadrant [PI(Red)+-AnnexinV-(FITC) +] represent early apoptotic cells.

3.7. Cell Cycle Assay

Cell cycle analysis is performed by flow cytometry after staining with propidium iodide. 1. Cells are harvested, washed twice with 1X PBS, resuspended in 200 µl of 1X PBS, and then fixed with 10 ml of cold 75% ethanol at 4°C for a minimum of 4 h and then washed twice with 1X PBS. 2. Subsequently, cells are resuspended in 500 µl of 1X PBS and stained with 200 µl of propidium iodide (50 µg/ml) and 20 µl of RNase (1 mg/ml) in a 37°C water bath for 15–20 min. 3. Cell cycle analysis is done with a FACStation equipped with CellQuest software.

4. Notes 1. Since in Drosophila embryo extracts the antisense strand of the siRNA sets the ruler for cleavage of target mRNA, at the ninth nucleotide from its paired 5′ end (33), the position of the bulge may be a critical determinant of translational repression activity.

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2. The core algorithm predicts a minimum free energy, G, as well as minimum free energies for foldings that must contain any particular base pair (34). The ability of a miRNA to translationally repress a target mRNA is largely dictated by the free energy of binding of the first eight nucleotides in the 5′ region of the miRNA. The mutations creating mismatches with the 5′ region of the miRNA inactivated the repression, whereas the order mutations had no effect. The interactions in the 3′ regions were not important, the stability of the miRNAmRNA interaction in the 5′ region was high. Also G:U wobble in the 5′ region of the miRNA hinders repression despite its favorable contribution to RNA:RNA duplexes (29,30). 3. The 5′ ends of related miRNAs tend to be better conserved than the 3′ end, further supporting the hypothesis that these segments are most critical for mRNA recognition. The critical importance of pairing to segment 2.8 for target identification in silico reflects its importance for target recognition in vivo and speculates that this segment nucleates pairing between miRNAs and miRNAs (35). The 3′UTR analysis yields 106 motifs likely to be involved in post-transcriptional regulation. Nearly onehalf are associated with miRNAs, leading to the discovery of many new miRNA genes and their likely target genes (36) 4. Studying RNA expression alone may seriously underestimate off-target effects. Once the rules for siRNA and miRNA sequence context are better defined experimentally, improved computational resources will be needed to aid in design of miRNAs, to minimize the potential for off-target interactions. 5. RNAi is often very effective at minimal concentrations, and using the lowest possible concentration of miRNA or siRNA has been suggested to prevent saturation of the RNAi machinery and unwanted side effects.

Acknowledgements This work has been supported in part by the Topfer Pancreatic Cancer Research Fund (DZC, NT), Grant DOD-01-1-299 (CGI, NT), and grant from the 21st Century COE program from the Kurume University (NT, TM). References 1. Chu C.Y. and Rana T.M. (2007) Small RNAs: Regulators and guardians of the genome. J. Cell. Physiol. 213:412–419.

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Chapter 22 Therapeutic Targeting of Gene Expression by siRNAs Directed Against BCR-ABL Transcripts in a Patient with Imatinib-Resistant Chronic Myeloid Leukemia Michael Koldehoff and Ahmet H. Elmaagacli Abstract Within the recent years, RNA interference (RNAi) has become an almost-standard method for in vitro knockdown of any target gene of interest. Now, one major focus is to further explore its potential in vivo, including the development of novel therapeutic strategies. From the mechanism, it becomes clear that small interfering RNAs (siRNAs) play a pivotal role in triggering RNAi. Thus, the efficient delivery of target gene-specific siRNAs is one major challenge in the establishment of therapeutic RNAi. Here we show that in vivo application of targeted nonvirally delivered synthetic bcr-abl siRNA in a female patient with recurrent Philadelphia chromosome positive chronic myeloid leukemia (CML) resistant to imatinib (Y253F mutation) and chemotherapy after allogeneic hematopoietic stem cell transplantation can silence the expression of bcr-abl gene. We found a remarkable inhibition of the overexpressed bcr-abl oncogene resulting in increased apoptosis of CML cells. In vivo siRNA application was well tolerated without any clinically adverse events. Our findings imply that the clinical application of synthetic siRNA is feasible, safe and has real potential for genetic-based therapies using synthetic nonviral carriers. Key words: bcr-abl, small interfering RNA, RNAi, CML, oligonucleotherapy, oncogenes.

1. Introduction Chronic myeloid leukemia (CML) is a relatively well-differentiated myeloproliferative disorder originating from transformed hematopoietic stem cells. The disease arises as a consequence of a rare mutational event resulting in a reciprocal translocation between the long arms of chromosomes 9 and 22. The shortened chromosome 22 formed by this translocation is the Philadelphia (Ph1) chromosome, named after the city in which it was discovered. CML was the first neoplastic process to be linked to M. Sioud (ed.), Methods in Molecular Biology, siRNA and miRNA Gene Silencing, vol. 487 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-60327-547-7_22

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a consistent acquired genetic abnormality and currently it is the best-studied molecular model of leukemia. The translocation creates the chimeric oncogene bcr-abl with the protein product BCR-ABL, a tyrosine kinase with constitutive activity. The expression of these consistent molecular changes has been shown to be necessary and sufficient for the transformed phenotype of CML cells. BCR-ABL is responsible for the pathogenesis of CML, demonstrated by the transforming ability of BCR-ABL expression in cell lines and within mice. The natural clinical course in human CML can be divided into three phases. It follows a fairly benign course for several years (chronic phase) before transforming into the more aggressive accelerated phase and life-threatening blast crisis. At the time of diagnosis the majority of patients are in the chronic phase and are asymptomatic. The blast crisis occurs when there is a failure of maturation of the malignant precursors, often accompanied by additional cytogenetic abnormalities and resulting in a disease resembling acute myeloid or lymphoblastic leukemia (1 – 4). Treatment with imatinib mesylate, a selective inhibitor of ABL, the chimeric BCR-ABL fusion protein, platelet-derived growth factor receptor (PDGF-R) α and β, and c-kit has shown remarkable clinical activity with minimal side effects in bcr-abl(+), c-kit(+) or PDGF-R(+) leukemias and has resulted in complete cytogenetic remission in a majority of chronic phase CML patients, and in fewer patients with accelerated and blast phase disease. Disease progression remains low but detectable, though this risk may decline over time (5– 7). Although allografting is still considered to be the only potentially curative approach in CML patients, transplantation numbers have dropped significantly in the imatinib era due to transplantation-associated mortality and morbidity. Furthermore, as patients may relapse after allogeneic transplantation or develop imatinib-resistant disease, additional CML therapies are required (8,9). In 1998 Andrew Fire and Craig Mello discovered, in a series of experiments in Caenorhabditis elegans, that injection of sense or antisense RNAs led to negligible decreases of target RNA, whereas introduction of double-stranded RNA (dsRNA) resulted in effective and specific degradation of cytoplasmic mRNA. Furthermore, the aforementioned silencing effects of dsRNA in C. elegans were systemic and heritable. An evolutionary conserved cellular mechanism to protect against viral infections, RNA interference (RNAi) inhibits gene expression by degrading mRNA in a sequence-specific post-transcriptional manner, upon introduction of dsRNA. This long dsRNA is cut into 21–23-mer active intermediates, termed small interfering RNAs (siRNAs). siRNAs are the mediators of mRNA degradation in the process of RNAi. Different ways of producing these small interfering molecules, including chemical synthesis, in vitro transcription, or vector-based delivery into mammalian cell lines have been successfully developed over

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the last few years (10– 13). Regardless of the transfection method, the efficiency of the siRNAs is mostly dependent on the successful rational design of the 21-mer sequences (14, see Chap. 1). Scherr et al. recently published a study showing that siRNA directed against bcr-abl can specifically inhibit bcr-abl expression in the Ph1(+) cell lines and the cells of CML patients (15, see Chap. 11). Furthermore, Wohlbold et al. showed that siRNA treatment might sensitize cells to imatinib mesylate contributing to its therapeutic potential (16). We demonstrated that combined transfection with the Wilm’s tumor gene (WT1) siRNA and bcr-abl siRNA in the Ph1(+) cell lines and the cells of CML patients increased the inhibition of the rate of proliferation and the rate of induced apoptosis compared to transfection with bcr-abl siRNA or WT1 siRNA alone (17). Also we showed that bcr-abl siRNA had antiproliferative and pro-apoptoic effects on Ph1(+) AML cells in vitro (18). Limitations in the use of oligonucleotides as gene expression inhibitors include their poor stability in biological medium, their weak intracellular penetration, and the poor cytoplasmic delivery when they have to reach their complementary target. It is, however, possible to bypass this problem by using synthetic nonviral carriers, such as cationic liposomes and polymers (19,20). RNAi therapeutics represents a fundamentally new way to treat human disease by addressing targets that are otherwise undruggable with existing medicines. We recently demonstrated the application of targeted, nonviral delivery of bcr-abl siRNA as a therapeutic approach in a female CML patient with imatinib-resistant medullar and extramedullar relapse after allogeneic hematopoietic stem cell transplantation (HSCT) and report here in vivo evidence of the efficacy of RNAi-based therapeutics efficacy in this CML disease (21).

2. Methods of Inducing RNAi One remarkable property of RNAi and cosuppression is that in both processes a signal appears to be generated, which travels through the organism to induce sequence-specific gene silencing at a considerable distance. The discovery that siRNA is the effector mechanism of endogenous RNAi prompted investigation into the utilization of exogenously administered siRNA, or vectors inducing the expression of siRNA, for gene-specific silencing. Genetic manipulation using such a strategy raises several issues that need to be addressed: (1) stability of siRNA; (2) ability to constitutively express the siRNA; (3) possibility of tissue-specific delivery; and (4) finding the best method for identifying effective silencing sites on the mRNA transcript. In order to answer these questions various versions of siRNA have been developed (22,23).

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3. RNAi Delivery in Mammalian Cells The three most important attributes to take into account when designing and selecting siRNA are potency, specificity ,and nuclease stability. As our understanding of the molecular and structural mechanism of RNAi has grown, it is now possible to identify potent, specific, and stable siRNA candidates targeting any gene of interest. RNAi in mammalian cells can be triggered by direct introduction through injection, electroporation, lipid-mediated transfection, nanoparticles, or antibody bound enzymatically generated or chemically synthesized siRNAs, among others. Alternatively, siRNAs or small hairpin RNAs (shRNAs) can be delivered by vector-based intracellular expression. SiRNAs can be synthesized chemically, generated enzymatically through in vitro transcription by T7 phage polymerase, or through endonuclease digestion by recombinant Dicer of in vitro transcribed long dsRNAs. In mammalian cells, direct delivery of siRNAs can only induce transient silencing due to their limited half-life and to their dilution during cell division. Chemical modifications are required to potentiate siRNA nuclease and thermodynamic stability in vivo without compromising their efficacy. Recently, several groups reported different approaches for systemic in vivo delivery of siRNAs. Soutschek et al. described the intravenous injection in mice of chemically modified naked siRNAs coupled to a cholesterol group chemically linked to the terminal hydroxyl group of the sense string to promote entry into the cells (24). In vivo delivery of chemically modified siRNAs encapsulated into liposome particles has been recently reported by Morrisey et al., and Song et al. described an antibody-based delivery system which offered the possibility of systemic, cell-type-specific siRNA delivery (25,26).

4. In Vivo siRNA Delivery Through Dispersion Lipid Solution (DLS) Complexation

Naked siRNAs are degraded in human plasma with a half-life of minutes. To convert siRNAs into optimized drugs, the modification of chemically synthesized siRNAs need to be protected from nuclease digestion and last longer than naked siRNAs, especially when it is exposed to nuclease-rich environments, such as blood to target human leukemias (23,27). Our recently described method for the efficient protection and delivery of siRNAs in vitro and in vivo relies on anionic liposome complexation in leukemic malignancies (21). In brief, the sequence of our siRNA targeted against bcr-abl was AAGCAGAGTTCAAAAGCCCTT (from Qiagen-Xeragon, Germany), as published previously by Scherr

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and coworkers (14). Chemically synthesized naked siRNA was dissolved in a suspension buffer (100 nM potassium acetate, 30 mM HepesKOH, 2 nm Mg-acetate, pH 7.4), heated at 90°C for 1 min, and then incubated for 1 h at 37°C. Then the solution was diluted 1:10 in a 10% lipid solution (Lipovenös) purchased from Fresenius Kabi (Bad Homburg, Germany) containing, per liter: 100 g soya bean oil, 25 g glycerol, and 6 g phospholipids from egg. The solution was incubated for 15 min at room temperature and mixed multiple times by passing it through a small lumen syringe generating negative pressure using a three-way valve to form the dispersion lipid solution (DLS) and reconstitute siRNA chylomicrons in the lipid solution.

5. Efficiency of siRNA Transfection with DLS Ex Vivo

In vitro transfections with 0.8 µg siRNA were performed in 24-well plates using TransMessenger transfection reagent (Qiagen, Hilden, Germany) or DOTAP Liposomal transfection reagent (Roche Diagnostics, Mannheim, Germany) following the manufacturer’s protocol, or using DLS (1 × 105/well) following the above described delivery protocol. Using fluorescence-marked, nonsilencing siRNA (Qiagen) we evaluated the transfection rate of DLS, TransMessenger or DOTAP Liposomal in K562 cells. Twenty-four hours after transfection the number of fluorescently marked cells was evaluated using fluorescence microscopy. 5 × 100 cells were counted per sample. With DLS we found a mean transfection rate of 70% (range 61–78%), with TransMessenger or DOTAP Liposomal we found a mean transfection rate of 50% (range 44–62%) and 59% (range 54–66%), respectively. Cell proliferation was determined by 5-bromo-2-deoxyuridine (BrdU) incorporation assay and apoptotic cells were determined using the in situ cell Death detection kit, both following the manufacture’s instructions from Roche Diagnostics (Mannheim, Germany). Twenty-four hours after transfection with bcrabl siRNA we observed a moderate inhibition of proliferation in the CML cells of the patient prior to siRNA therapy from either spontaneously 36.8 ± 4.6% or nonsilencing siRNA 34.8 ± 3.2% to 14.6 ± 2.1% (reduction of 76%, p < 0.002) and a strong induction of apoptosis from spontaneously 8.1 ± 1.7 or nonsilencing siRNA 8.6 ± 2.2% to 16.1 ± 1.2 (induction of 99%, p < 0.05) as shown in Table 22.1. Transfection with siRNA in K562 cells induced an increased rate of apoptosis from spontaneously 13.0 ± 2.5% or nonsilencing siRNA 13.5 ± 2.7% to 18.5 ± 4.4% (induction of 42%, p < 0.01) and an inhibition of proliferation from 48.6 ± 4.2 or nonsilencing siRNA 50.6 ± 3.2% to 28.3 ± 4.9% (reduction of 42%, p < 0.05). The effects on bcr-abl siRNA amounts were

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Table 22.1 Effects of bcr-abl siRNAs on apoptosis and proliferation in patient leukemic cells and K562 cell line Effects of bcr-abl siRNA on apoptosis bcr-abl siRNA

Nonsilencing siRNA

Control (spontaneous)

P-value

CML cells of the patient

16.1 ± 1.2

8.6 ± 2.2

8.1 ± 1.7

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