Gene Therapy of Cancer: Methods and Protocols, 2nd Edition (Methods in Molecular Biology, Vol. 542)

i METHODS IN MOLECULAR BIOLOGY™ Series Editor John M. Walker School of Life Sciences University of Hertfordshire Ha...

Author: Wolfgang Walther | Ulrike S. Stein

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i

METHODS

IN

MOLECULAR BIOLOGY™

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

For other titles published in this series, go to www.springer.com/series/7651

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Gene Therapy of Cancer Methods and Protocols Second Edition

Edited by

Wolfgang Walther and Ulrike S. Stein Max-Delbrück-Center for Molecular Medicine, Berlin, Germany

Editors Wolfgang Walther Max-Delbrück-Center for Molecular Medicine Berlin, Germany

Ulrike S. Stein Max-Delbrück-Center for Molecular Medicine Berlin, Germany

ISBN: 978-1-934115-85-5 e-ISBN: 978-1-59745-561-9 ISSN: 1064-3745 e-ISSN: 1940-6029 DOI: 10.1007/978-1-59745-561-9 Library of Congress Control Number: 2008942247 © 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 springer.com

Preface When the first edition of Gene Therapy of Cancer as part of the series Methods in Molecular Medicine series was released in 1999/2000, this research field was still filled with euphoric moments and great expectations for almost unlimited success. Due to decisive drawbacks in clinical use of gene therapy, the voices of those demanding gene therapy go back to the benches became more intense. In fact, during the last decade many important issues for gene therapy, including cancer gene therapy, have been investigated with great efforts to ameliorate vector safety, transfer efficiency, improve vector targeting, find more effective and specific therapeutic genes, etc. In parallel, numerous gene regulatory issues have evolved and been enhanced for the benefit of patients treated with gene therapy. Because gene therapy of cancer is still the field of greatest efforts, representing more than 60% of all gene therapy trials, this field has accumulated a tremendous amount of preclinical, and, more importantly, of clinical data. Such increment in gene therapeutic experience did promote and will further accelerate the development of cancer gene therapy into a safe and clinically applicable treatment option. This second edition was facing the difficulty of potentially being just one more of those countless books aiming at some coverage of cancer gene therapy. However, the editors felt the responsibility to create something slightly different. Therefore, contributions were selected that cover both experimental and clinical approaches to cancer gene therapy. These were carefully chosen with special emphasis on presentation of established and, more importantly, novel protocols to at least in part reflect all of the efforts made for the improvement of cancer gene therapy. Furthermore, this edition provides state-of-the-art overviews of new concepts and strategies in cancer gene therapy. Because regulatory and ethical issues are of pivotal importance for clinical gene therapy, these topics are acknowledged for their impact in the field as separate chapters in this new edition. Furthermore, the inclusion of chapters that cover the developments, problems, and possible limitations of design and production of gene therapeutics for the clinic broaden the insights into the very complex field of cancer gene therapy, also comprising such translational issues. Taken together, this second edition has been developed with the intent of providing more than merely a remake of the first edition of the book. This edition has been made for those who are working in the field and are strongly interested in receiving an interesting, spotlighted overview of nonviral, viral, experimental, and clinical cancer gene therapy. In parallel, this edition certainly also addresses those who are interested in the field and are willing to dig into this exciting research area. The editors thank all contributors for their valuable chapters, which will further stimulate the interest of the readers in the field of cancer gene therapy. Wolfgang Walther and Ulrike S. Stein

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

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SECTION I: EXPERIMENTAL APPROACHES IN CANCER THERAPY SUBSECTION A: INTRODUCTION 1

The Development of Gene Therapy: From Monogenic Recessive Disorders to Complex Diseases Such as Cancer . . . . . . . . . . . . . . . . . . . . . . . . . . . Jean-Pierre Gillet, Benjamin Macadangdang, Robert L. Fathke, Michael M. Gottesman, and Chava Kimchi-Sarfaty

5

SUBSECTION B: VECTOROLOGY FOR CANCER GENE THERAPY 2 3 4 5

6 7

Designing Adenoviral Vectors for Tumor-Specific Targeting . . . . . . . . . . . . . . . . . 57 Ramon Alemany Analysis of HSV Oncolytic Virotherapy in Organotypic Cultures. . . . . . . . . . . . . . 75 Giulia Fulci and Brent Passer Use of Minicircle Plasmids for Gene Therapy . . . . . . . . . . . . . . . . . . . . . . . . . . . . 87 Peter Mayrhofer, Martin Schleef, and Wolfgang Jechlinger Transposable Elements as Plasmid-Based Vectors for Long-Term Gene Transfer into Tumors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 105 John R. Ohlfest, Zoltán Ivics, and Zsuzsanna Izsvák Designing Plasmid Vectors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 117 Oleg Tolmachov Development of Bacterial Vectors for Tumor-Targeted Gene Therapy . . . . . . . . . . 131 Li-Jun Jia and Zi-Chun Hua

SUBSECTION C: NONVIRAL TRANSFER TECHNOLOGIES IN CANCER GENE THERAPY 8

Electroporative Gene Transfer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Marco Schmeer 9 Gene Gun Delivery Systems for Cancer Vaccine Approaches . . . . . . . . . . . . . . . . . Kandan Aravindaram and Ning Sun Yang 10 Ultrasound-Mediated Gene Transfection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Loreto B. Feril Jr. 11 Nonviral Jet-Injection Technology for Intratumoral In Vivo Gene Transfer of Naked DNA. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Wolfgang Walther, Iduna Fichtner, Peter M. Schlag, and Ulrike S. Stein

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157 167 179

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Contents

SUBSECTION D: EXPERIMENTAL STUDIES IN CANCER GENE THERAPY 12 13 14 15

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17 18

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20

21

Methods for Constructing and Evaluating Antitumor DNA Vaccines . . . . . . . . . . Brian M. Olson and Douglas G. McNeel Immunity of Lentiviral Vector-Modified Dendritic Cells . . . . . . . . . . . . . . . . . . . . Shuhong Han and Lung-Ji Chang Saporin Suicide Gene Therapy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Natasa Zarovni, Riccardo Vago, and Maria Serena Fabbrini Using In Vivo Biopanning for the Development of Radiation-Guided Drug Delivery Systems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jerry J. Jaboin, Zhaozhong Han, and Dennis E. Hallahan Chemosensitization of Tumor Cells: Inactivation of Nuclear Factor-Kappa B Associated with Chemosensitivity in Melanoma Cells After Combination Treatment with E2F-1 and Doxorubicin . . . . . . . . . . . . . . . . . . . . . Hongying Hao, H. Sam Zhou, and Kelly M. McMasters Induction of Tumor Cell Apoptosis by TRAIL Gene Therapy. . . . . . . . . . . . . . . . Thomas S. Griffith Silencing Epidermal Growth Factor Receptor by RNA Interference in Glioma. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Chunsheng Kang, Peiyu Pu, and Hao Jiang Delivery of Phosphorodiamidate Morpholino Antisense Oligomers in Cancer Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Gayathri R. Devi Use of RNA Aptamers for the Modulation of Cancer Cell Signaling . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sunjoo Jeong, Hee Kyu Lee, and Mee Young Kim G-Rich Oligonucleotides for Cancer Treatment . . . . . . . . . . . . . . . . . . . . . . . . . . Paula J. Bates, Enid W. Choi, and Lalitha V. Nayak

211 245 261

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301 315

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363 379

SECTION II: CLINICAL APPROACHES IN CANCER GENE THERAPY SUBSECTION A: REQUIREMENTS FOR CLINICAL GENE THERAPY TRIALS 22

Regulatory Aspects for Translating Gene Therapy Research into the Clinic . . . . . . Carolyn M. Laurencot and Sheryl Ruppel 23 Ethics of Cancer Gene Transfer Clinical Research . . . . . . . . . . . . . . . . . . . . . . . . . Jonathan Kimmelman 24 Virus Production for Clinical Gene Therapy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Tiago Vicente, Cristina Peixoto, Manuel J.T. Carrondo, and Paula M. Alves 25 Production of Plasmid DNA as Pharmaceutical. . . . . . . . . . . . . . . . . . . . . . . . . . . Martin Schleef and Markus Blaesen

397 423 447

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SUBSECTION B: CLINICAL PROTOCOLS IN CANCER GENE THERAPY 26

Gene Immunotherapy for Non-Small Cell Lung Cancer . . . . . . . . . . . . . . . . . . . . 499 John J. Nemunaitis

Contents

27

Gene Therapy for Antitumor Vaccination . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Seunghee Kim-Schulze and Howard L. Kaufman 28 HSV-tk/IL-2 Gene Therapy for Glioblastoma Multiforme . . . . . . . . . . . . . . . . . . Luisa Barzon, Monia Pacenti, Elisa Franchin, Federico Colombo, and Giorgio Palù 29 Construction and Characterization of an Oncolytic HSV Vector Containing a Fusogenic Glycoprotein and Prodrug Activation for Enhanced Local Tumor Control . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Guy R. Simpson and Robert S. Coffin 30 Newcastle Disease Virus: A Promising Vector for Viral Therapy, Immune Therapy, and Gene Therapy of Cancer . . . . . . . . . . . . . . . . . . . . . . . . . . Volker Schirrmacher and Philippe Fournier 31 Oncolytic Viral Therapy Using Reovirus. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Chandini Thirukkumaran and Don G. Morris 32 Design and Testing of Novel Oncolytic Vaccinia Strains . . . . . . . . . . . . . . . . . . . . Steve H. Thorne 33 Tumor-Targeted Salmonella typhimurium Overexpressing Cytosine Deaminase: A Novel, Tumor-Selective Therapy . . . . . . . . . . . . . . . . . . . . . . . . . . Ivan King, Martina Itterson, and David Bermudes 34 Chemoprotection by Transfer of Resistance Genes . . . . . . . . . . . . . . . . . . . . . . . . Tulin Budak-Alpdogan and Joseph R. Bertino 35 Phase I Clinical Trial of Locoregional Administration of the Oncolytic Adenovirus ONYX-015 in Combination with Mitomycin-C, Doxorubicin, and Cisplatin Chemotherapy in Patients with Advanced Sarcomas . . . . . . . . . . . . . Mateusz Opyrchal, Ileana Aderca, and Evanthia Galanis Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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515 529

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565 607 635

649 661

705 719

Contributors ILEANA ADERCA • Division of Medical Oncology, Mayo Clinic, Rochester, MN, USA RAMON ALEMANY Translational Research Laboratory, Catalan Institute of Oncology, L´Hospitalet de Llobregat, Barcelona, Spain PAULA M. ALVES • Instituto de Biologia Experimental e Tecnológica (IBET) and Instituto de Tecnologia Química e Biológica – UNL (ITQB-UNL), Oeiras, Portugal KANDAN ARAVINDARAM • Agricultural Biotechnology Research Center, Academia Sinica, Nankang, Taipei, Taiwan, Republic of China LUISA BARZON • Department of Histology, Microbiology and Medical Biotechnologies, University of Padova, Padova, Italy PAULA J. BATES • University of Louisville, Louisville, KY, USA DAVID BERMUDES • Celator Pharmaceuticals Corp., Vancouver, BC, Canada JOSEPH R. BERTINO • Departments of Medicine and Pharmacology, The Cancer Institute of New Jersey, Robert Wood Johnson Medical School, University of Medicine & Dentistry of New Jersey, New Brunswick, NJ, USA MARKUS BLAESEN • PlasmidFactory GmbH & Co KG, Bielefeld, Germany TULIN BUDAK-ALPDOGAN • Departments of Medicine and Pharmacology, The Cancer Institute of New Jersey, Robert Wood Johnson Medical School, University of Medicine & Dentistry of New Jersey, New Brunswick, NJ, USA MANUEL J. T. CARRONDO • Instituto de Biologia Experimental e Tecnológica (IBET) and Instituto de Tecnologia Química e Biológica-UNL (ITQB-UNL), Oeiras, Portugal LUNG-JI CHANG • Department of Molecular Genetics and Microbiology, Powell Gene Therapy Center and McKnight Brain Institute, University of Florida, College of Medicine, Gainesville, FL, USA ENID W. CHOI • Department of Biochemistry & Molecular Biology, Brown Cancer Center, University of Louisville, Louisville, KY, USA ROBERT S. COFFIN • Biovex, Inc., Woburn, MA, USA FREDERICO COLOMBO • Division of Neurosurgery, Robotic Radiosurgery Unit, San Bortolo Hospital, Vicenza, Italy GAYATHRI R. DEVI • Comprehensive Cancer Center, Duke University Medical Center, Durham, NC, USA MARIA S. FABBRINI • Istituto Biologia Biotecnologia Agraria, IBBA, National Research Council, CNR, Milano, Italy 208 ROBERT L. FATHKE • Center for Biologics Evaluation and Research, Food and Drug Administration, Bethesda, MD, USA LORETO B. FERIL, JR. • Department of Anatomy, Fukuoka University School of Medicine, Fukuoka City, Fukuoka, Japan IDUNA FICHTNER • Max-Delbrück-Center for Molecular Medicine, Berlin, Germany ELISA FRANCHIN • Department of Histology, Microbiology and Medical Biotechnologies, University of Padova, Padova, Italy xi

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Contributors

GIULIA FULCI • Brain Tumor Research Center, Simches Research Building, Neurosurgery Service, Massachusetts General Hospital, Boston, MA, USA PHILIPPE FOURNIER • German Cancer Research Center (DKFZ), Heidelberg, Germany EVANTIA GALANIS • Division of Medical Oncology and Department of Molecular Medicine, Mayo Clinic, Rochester, MN, USA JEAN-PIERRE GILLET • Laboratory of Cell Biology, National Cancer Institute, National Institutes of Health, Bethesda, MD, USA MICHAEL M. GOTTESMAN • Laboratory of Cell Biology, National Cancer Institute, National Institutes of Health, Bethesda, MD, USA THOMAS S. GRIFFITH • Department of Urology, University of Iowa, Iowa City, IA, USA DENNIS E. HALLAHAN • Department of Radiation Oncology, Department of Developmental and Cell Biology, Department of Cancer Biology, School of Medicine, Vanderbilt University, Nashiville, TN, USA; Vanderbilt-Ingram Cancer Center, Nashville, TN, USA SHUHONG HAN • Department of Molecular Genetics and Microbiology, Powell Gene Therapy Center and McKnight Brain Institute, University of Florida, College of Medicine, Gainesville, FL, USA ZHAOZHONG HAN • Department of Radiation Oncology, Vanderbilt University, Nashville, TN, USA; Vanderbilt-Ingram Cancer Center, Nashville, TN, USA HONGYIN HAO • Surgical Oncology, University of Louisville, Louisville, KY, USA ZI-CHUN HUA • The State Key Laboratory of Pharmaceutical Biotechnology and Department of Biochemistry, College of Life Sciences, Nanjing University, Nanjing, P. R. China MARTINA ITTERSON • Vion Pharmaceuticals, Inc., New Haven, CT, USA ZOLTÁN IVICS • Max-Delbrück-Center for Molecular Medicine, Berlin, Germany ZZUZSA IZSVÁK • Max-Delbrück-Center for Molecular Medicine, Berlin, Germany JERRY J. JABOIN • Department of Radiation Oncology, Vanderbilt University, Nashville, TN, USA; Vanderbilt-Ingram Cancer Center, Nashville, TN, USA WOLFGANG JECHLINGER • Mayrhofer & Jechlinger OEG, Vienna, Austria (died Dec. 2007) SUNJOO JEONG • National Research Laboratory for RNA Cell Bioloigy and Department of Molecular Biology, Dankook University, Gyeonggi–do, Republic of Korea LIJUN JIA • Comprehensive Cancer Center, University of Michigan, Ann Arbor, MI, USA HAO JIANG • Department of Neurology, Henry Ford Hospital, Detroit, MI, USA CHUNSHENG KANG • Department of Neurosurgery, Tianjin Medical University General Hospital, Laboratory of Neuro-Oncology, Tianjin Neurological Institute, Tianjin, People’s Republic of China HOWARD L. KAUFMAN • Division of Surgical Oncology, Columbia University, New York, NY, USA MEE YOUNG KIM • National Research Laboratory for RNA Cell Bioloigy and Department of Molecular Biology, Dankook University, Gyeonggi–do, Republic of Korea

Contributors

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CHAVA KIMCHI-SARFATY • Center for Biologies Evaluation and Research, Food and Drug Administration, Bethesda, MD, USA JONATHAN KIMMELMAN • Department of Social Studies of Medicine, Clinical Trials Research Group, Biomedical Ethics Unit, McGill University Faculty of Medicine, Montreal, QB, Canada SEUNGHEE KIM-SCHULZE • The Tumor Immunology Laboratory, Division of Surgical Oncology, Columbia University, New York, USA IVAN KING VION • Pharmaceuticals, Inc., New Haven, CT, USA CAROLYN M. LAURENCOT • Surgery Branch, Center for Cancer Research, National Cancer Institute National Institutes of Health, Bethesda, MD, USA HEE KYO LEE • National Research Laboratory for RNA Cell Bioloigy and Department of Molecular Biology, Dankook University, Gyeonggi–do, Republic of Korea BENJAMIN MACADANGDANG • Laboratory of Cell Biology, National Cancer Institute, National Institutes of Health, Bethesda, MD, USA PETER MAYRHOFER • Mayrhofer & Jechlinger OEG, Vienna, Austria KELLY M. MCMASTERS • Surgical Oncology, University of Louisville, Louisville, KY, USA DOUGLAS G. MCNEEL • Department of Medicine, Section of Medical Oncology, University of Wisconsin–Madison, Madison, WI, USA DON G. MORRIS • Department of Oncology, Tom Baker Cancer Centre, Department of Medicine and Oncology, University of Calgary, Calgary, Alberta, Canada LALITHA V. NAYAK • Department of Medicine, Brown Cancer Center, University of Louisville, Louisville, Kentucky, USA JOHN NEMUNAITIS • Mary Crowley Medical Research Center, Dallas, Texas, USA JOHN OHLFEST • Department of Pediatrics, University of Minnesota, Minneapolis, MN, USA BRIAN OLSON • Department of Medicine, Section of Medical Oncology, University of Wisconsin–Madison, Madison, WI, USA MATEUSZ OPYRCHAL • Division of Medical Oncology, Mayo Clinic, Rochester, MN, USA MONIA PACENTI • Department of Histology, Microbiology and Medical Biotechnologies, University of Padova, Padova, Italy BRENT PASSER • Brain Tumor Research Center, Simches Research Building, Neurosurgery Service, Massachusetts General Hospital, Boston, MA, USA GIORGIO PALÙ • Department of Histology, Microbiology and Medical Biotechnologies, University of Padova, Padova, Italy CRISTINA PEIXOTO • Instituto de Biologia Experimental e Tecnológica (IBET) and Instituto de Tecnologia Química e Biológica – UNL (ITQB-UNL), Oeiras, Portugal PEIYU PU • Department of Neurosurgery, Tianjin Medical University General Hospital, Tianjin, People’s Republic of China SHERYL RUPPEL • Regulatory Affairs Consulting, Damascus, MD, USA VOLKER SCHIRRMACHER • Division of Cellular Immunology (D010), German Cancer Research Center (DKFZ), Heidelberg, Germany PETER M. SCHLAG • Clinic for Surgery and Surgical Oncology, Charité University Medicine Berlin, Berlin, Germany

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Contributors

MARTIN SCHLEEF • PlasmidFactory GmbH & Co KG, Bielefeld, Germany MARCO SCHMEER • PlasmidFactory GmbH & Co. KG, Bielefeld, Germany GUY SIMPSON • Department of Oncology, Postgraduate Medical School, University of Surrey, Manor Park, Surrey, UK ULRIKE S. STEIN • Max-Delbrück-Center for Molecular Medicine, Berlin, Germany STEVE H. THORNE Division of Surgical Oncology, University of Pittsburgh, Pittsburgh, PA, USA CHANDINI THIRUKKUMARAN • Tom Baker Cancer Centre, Department of Medicine and Oncology, University of Calgary, Calgary, Alberta, Canada OLEG TOLMACHOV • National Heart and Lung Institute, Faculty of Medicine, Imperial College London, South Kensington, London, UK RICARDO VAGO • Department of Biological and Technological Research, Dibit, San Raffaele H Scientific Institute, Milano, Italy TIAGO VICENTE • Instituto de Biologia Experimental e Tecnológica (IBET) and Instituto de Tecnologia Química e Biológica – UNL (ITQB-UNL), Oeiras, Portugal WOLFGANG WALTHER • Max-Delbrück-Center for Molecular Medicine, Berlin, Germany NING SUN YANG • Agricultural Biotechnology Research Center, Academia Sinica, Nankang, Taipei, Taiwan, Republic of China NATASA ZAROVNI • Department of Biological and Technological Research, Dibit, San Raffaele H Scientific Institute, Milano, Italy SAM H. ZHOU • Surgical Oncology, University of Louisville, Louisville, KY, USA

Section I Experimental Approaches in Cancer Therapy

Subsection A Introduction

Chapter 1 The Development of Gene Therapy: From Monogenic Recessive Disorders to Complex Diseases Such as Cancer* Jean-Pierre Gillet, Benjamin Macadangdang, Robert L. Fathke, Michael M. Gottesman, and Chava Kimchi-Sarfaty Summary During the last 4 decades, gene therapy has moved from preclinical to clinical studies for many diseases ranging from monogenic recessive disorders such as hemophilia to more complex diseases such as cancer, cardiovascular disorders, and human immunodeficiency virus (HIV). To date, more than 1,340 gene therapy clinical trials have been completed, are ongoing, or have been approved in 28 countries, using more than 100 genes. Most of those clinical trials (66.5%) were aimed at the treatment of cancer. Early hype, failures, and tragic events have now largely been replaced by the necessary stepwise progress needed to realize clinical benefits. We now understand better the strengths and weaknesses of various gene transfer vectors; this facilitates the choice of appropriate vectors for individual diseases. Continuous advances in our understanding of tumor biology have allowed the development of elegant, more efficient, and less toxic treatment strategies. In this introductory chapter, we review the history of gene therapy since the early 1960s and present in detail two major recurring themes in gene therapy: (1) the development of vector and delivery systems and (2) the design of strategies to fight or cure particular diseases. The field of cancer gene therapy experienced an “awkward adolescence.” Although this field has certainly not yet reached maturity, it still holds the potential of alleviating the suffering of many individuals with cancer. Key words: Antiangiogenic therapy, chemoinducible gene therapy, gene delivery systems, gene silencing, history of gene therapy, immunotherapy, oncolytic viruses, suicide gene therapy, tumor suppressors.

* The review material, findings, and conclusions in this book chapter have not been formally disseminated by the US Food and Drug Administration (FDA) and should not be construed to represent any Agency determination or policy.

Wolfgang Walther and Ulrike S. Stein (eds.), Methods in Molecular Biology, Gene Therapy of Cancer, vol. 542 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-561-9_1

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1. The Checkered History of Gene Therapy 1.1. The Birth of an Elegant but Provocative New Theory

1.2. From Theory to Bedside

The early 1960s were characterized by the emergence of a revolutionary treatment concept termed gene therapy, defined as the delivery and expression of genetic information in cells of a patient to restore health or to alleviate the consequences of a disease. The first speculation discussing this new scientific approach appeared in 1963, when Joshua Lederberg commented on the future of medicine: “The ultimate application of molecular biology would be the direct control of nucleotide sequences in human chromosomes, coupled with recognition, selection, and integration of the desired genes” (1). Edward Tatum extended the idea during a symposium entitled “Reflections on Research and the Future of Medicine” in 1966: “Finally we can anticipate that viruses will be used effectively for man’s benefit, in theoretical studies concerning somatic cell genetics and possibly in genetic therapy.… We even can be somewhat optimistic about the long-range possibility of therapy based on the isolation or design, synthesis, and introduction of new genes into defective cells of particular organs” (2, 3). In the same year, Lederberg discussed the theory further in a manuscript entitled “Experimental Genetics and Human Evolution” (4). Arthur Kornberg (5) and French Anderson (6) also participated actively in the delineation of the theoretical basis of engineering human cells, in highly speculative and provocative papers appearing in the 1960s. Although the first ethical debates on the subject arose by the end of that decade (7, 8), gene therapy was still entirely based on theory, and was considered by most of the scientific community as entirely fanciful. A strange combination of events significantly influenced this nascent field. Lederberg read a manuscript published in The Lancet (9) describing two sisters who suffered from a genetic defect in which toxic levels of arginine increased in their bloodstream. He was intrigued by this, because he was familiar with the work of his collaborator, Stanfield Rogers, who had studied the Shope rabbit papilloma virus for several years. Rogers reported that animals infected with this virus had, among other characteristics, a decreased level of blood arginine. In an article published in Nature (10), Rogers pointed out that numerous researchers working with the Shope virus had reduced blood arginine levels but experienced no other side effects from exposure to the virus. They hypothesized that the Shope virus could be an effective treatment for the two sisters. In 1970, Rogers carried out the first human genetic engineering experiment by administering the virus to the sisters. However, for various reasons, the dose level administered was not sufficient to be therapeutic.

The Development of Gene Therapy: From Monogenic Recessive Disorders

7

The science leading to gene therapy was advanced significantly by the characterization of restriction endonucleases (11). Subsequent findings of cleavage site-specific endonucleases became the cornerstone of the emerging revolution of molecular biology (12, 13). Research on recombinant DNA, and the analysis of viral genomes such as that of SV40, which became an important actor in gene therapy, became conceivable (14). 1.3. The First Promising Steps Toward Gene Therapy

Gene transfer into mammalian cells to correct a genetic defect had its first success in 1977 when Wigler and colleagues transferred the purified herpes virus thymidine kinase (TK) gene into TK− mouse cells (15). Later, in 1979, Anderson performed the first transfer of two functional genes into TK− mouse cells using microinjection. One plasmid contained the thymidine kinase gene of herpes simplex virus type I (HSV-I) and the other contained the human beta globin gene. The genetic defect in the cells was corrected by the microinjected thymidine kinase gene, and the coinjected human beta globin gene was replicated and weakly expressed in the TK+ cells, demonstrating the principle of co-selection of selectable and unselected genes (16). Ten years after the first human gene therapy engineering, Martin Cline carried out an experiment on patients with beta-thalassemia that was followed by a debate concerning the ethics of gene therapy in humans (17).

1.4. The 1980s: Development of Vectors and First In Vivo Studies

The first viral system for gene transfer was developed in the early 1980s. The human gene for hypoxanthine phosphoribosyltransferase (HPRT) was successfully transduced into HPRT− rodent and human cells via a retrovirus (18–20). Subsequent studies reported the development of a helper-free retroviral packaging cell line with avian (21) and murine (22, 23) retroviruses. These were a new step forward for gene therapy, because helper viruses can cause substantial cytotoxicity to infected healthy cells. In the meantime, additional viral systems for gene transfer were developed, such as the adenovirus (24) and adeno-associated virus (25, 26). In 1985, the first in vivo expression of a transduced gene in an animal model (mouse) was reported by Eglitis and colleagues, using the retroviral vector N2 carrying the neomycin resistance (NeoR) marker (27). This was followed by the first human gene transfer clinical trial demonstrating the feasibility of using retroviral gene transduction for human gene therapy. The study was carried out at the National Institutes of Health (NIH) in 1989 by Rosenberg and coworkers (28). They used retroviral-mediated gene transduction to introduce the gene coding for resistance to neomycin into human tumor-infiltrating lymphocytes (TILs) before infusion of the TILs into patients with melanoma. The neomycin gene was then used as a marker for the infused cells.

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1.5. The 1990s: More In Vivo Studies and First Clinical Trials

The Anderson group also started the first approved gene therapy clinical trial in the early 1990s. Two children with adenosine deaminase-deficiency (ADA)–severe combined immunodeficiency (SCID) were treated with their own T cells engineered with a retroviral vector carrying a normal adenosine deaminase gene. Gene treatment ended after 2 years, but integrated vector and ADA gene expression in T cells persisted. Although the process was not optimally effective, this study demonstrated that gene therapy can be a safe addition to treatment for some patients with this severe immunodeficiency disease (29). At the same time, Wilson and colleagues used an animal model, the Watanabe Heritable Hyperlipidemic rabbit (WHHL), to develop gene therapies for the homozygous form of familial hypercholesterolemia (FH). Liver tissue was removed from WHHL rabbits and used to isolate hepatocytes and establish primary cultures. A functional rabbit low-density lipoprotein (LDL) receptor gene was transduced into a high proportion of hepatocytes using recombinant retroviruses, and the genetically corrected cells were transplanted into the animal from which they were derived. Transplantation of the genetically corrected, autologous hepatocytes was associated with a 30–40% decrease in serum cholesterol that persisted for the duration of the experiment. Recombinant-derived LDL receptor RNA was detected in the animals’ livers for at least 6 months without apparent immunological response to the recombinant-derived LDL receptor (30). Later, an ex vivo approach to gene therapy for FH was developed in which autologous hepatocytes that were genetically corrected with recombinant retroviruses carrying the LDL receptor were transplanted into the patient (31). The Wilson team also reported the outcome of the first pilot study of liver-directed gene therapy. Five patients with familial hypercholesterolaemia were enrolled; each patient tolerated the procedure well without significant complications. Transgene expression was detected in a limited number of hepatocytes of liver tissue harvested 4 months after gene transfer from all five patients. Significant and prolonged reductions in LDL cholesterol were reported in three of the five patients. This study demonstrated the feasibility of engrafting limited numbers of retrovirus-transduced hepatocytes without morbidity and achieving persistent gene expression lasting at least 4 months after gene therapy (32).

1.6. The New Century: From Severe Doubt to Real Success

Since its inception, gene therapy has been filled with promise. However, its history has been marred by failures, false hopes, and even death. Perhaps the most dramatic example of this concerns the X-linked form of severe combined immunodeficiency (SCID-X1), caused by mutations in the gene encoding the common γ chain (γc) of several interleukin receptors. Affected


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infants typically lack both T and natural killer (NK) cells, and have normal or elevated levels of functionally deficient B cells that are unable to undergo immunoglobulin class switching and antibody production. In 2000, Fischer and colleagues reported the successful use of gene therapy to treat infants with SCID-X1 (33). However, the enthusiasm generated by the apparent cure of nine of the ten infants turned to alarm when, nearly 3 years after treatment, T cell leukemia developed in two of the boys (34). Subsequently, a third child in the initial study developed leukemia (35, 36). In these patients, the retrovirus carrying the γc gene had inserted itself near LMO2, an oncogene that is activated as a result of translocations in acute lymphoblastic leukemia. The insertion caused overexpression of the LMO2 protein, implicating the LMO2 gene as the cause of the leukemia (37). Two years later, Aiuti and coworkers reported that gene therapy of hematopoietic stem cells (HSCs) improved the status of children with adenosine deaminase deficiency (ADA)–severe combined immunodeficiency (SCID) (38). Their study indicated the efficacy of CD34+ hematopoietic stem cell retroviral-mediated gene therapy combined with nonmyeloablative conditioning for the treatment of SCID (38). These results were recently confirmed by Gaspar and colleagues (39). Two years after a similar procedure was performed in a child with ADA-SCID, the immunological and biochemical correction had been maintained, with a progressive increase in lymphocyte numbers, reinitiation of thymopoiesis, and systemic detoxification of ADA metabolites (39). These results suggest that gene therapy is an effective treatment option for individuals with ADA-SCID. Initially designed for monogenetic recessive disorders, the concept of gene therapy was later extended to tackle complex pathologies such as degenerative diseases (e.g., Parkinson’s, Alzheimer’s, and heart disease), infectious diseases (e.g., human immunodeficiency virus [HIV]), and cancer. As researchers gained a fuller understanding of the genetic basis of cancer, an entirely new approach to the treatment of cancer using genetransfer techniques evolved. Among many examples of the use of gene therapy to treat cancer, in the late 1990s, Rainov reported a phase III clinical evaluation of herpes simplex virus type 1 thymidine kinase (TK) and ganciclovir gene therapy as an adjuvant to surgical resection and radiation in adults with glioblastoma multiforme (40). The authors reported that this adjuvant treatment improved neither time to tumor progression nor overall survival time, although the feasibility and good biosafety profile of this gene therapy strategy were further supported. The authors suggested that the failure of this specific protocol may have been mainly caused by the poor rate of delivery of the HSV-TK gene to tumor cells. They addressed several concerns to refine the procedure, such as a better transgene delivery system and an improved

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delivery of the prodrug across the blood–brain and blood–tumor barrier to the transduced tumor cells (40). In 2000, Khuri and colleagues reported a phase II clinical trial based on a combination of intratumoral ONYX-015 injection with cisplatin and 5-fluorouracil in patients with recurrent squamous cell cancer of the head and neck (41). ONYX-015, an adenovirus with the E1B 55-kDa gene deleted, was engineered to replicate selectively in and lyse p53-deficient cancer cells while sparing normal cells (41). 1.7. Hopeful Progress

During the last 4 decades, gene therapy has moved from preclinical to clinical studies for many diseases ranging from monogenic recessive disorders such as hemophilia, cystic fibrosis, and Duchenne muscular dystrophy, to more complex diseases such as cancer, cardiovascular disorders, and HIV. Recently, 20 leaders in the field summarized and discussed the gene therapy clinical trials performed in their respective areas of expertise (42, 43). To date, more than 1,340 gene therapy clinical trials have been completed, are ongoing, or have been approved in 28 countries, using more than 100 genes (44) (Fig. 1). Most of the clinical trials (66.5%) were aimed at the treatment of cancer. Early hype, failures, and tragic events have largely been replaced by the necessary stepwise progress needed to realize clinical benefits. The understanding of the strengths and weaknesses of gene transfer vectors and the choice of the appropriate vector for the individual disease has advanced (42). Gene therapy for severe combined immunodeficiency (SCID) is the most significant success story to date, but progress in many other areas has kept the promise

USA 64.2% (n=864) UK 11.1% (n=150) Germany 5.5% (n=74) Canada 4% (n=54) Switzerland 3.1% (n=42) France 1.5% (n=20) Belgium 1.4% (n=19) Australia 1.3% (n=17) Japan 1.2% (n=16) Italy 1.1% (n=15) Other countries 5.6% (n=75)

Fig. 1. Geographical distribution of completed or ongoing clinical trials in gene therapy (Reproduced from Edelstein et al. (44) J. Gene Med., 2007. Copyright John Wiley & Sons, Ltd., with permission).


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of gene therapy alive. The ability to prevent adverse events will depend on the availability of improved vectors, the findings of additional studies involving appropriate models, and a thorough assessment of the putative oncogenic capacity of genes, including marker genes, that are incorporated in various gene therapy vectors (45).

2. Vectors and Delivery Systems for Gene Therapy

2.1. Viral Delivery Systems 2.1.1. Adenoviral Vectors and Structure

One of the most difficult challenges in gene therapy is the development of vectors and delivery systems that are safe, efficient, and targeted. Figure 2 shows the proportionate application of various vectors and delivery techniques in gene therapy trials to date (44). The structure and biological properties of adenoviruses have made them the most commonly used vectors for gene therapy clinical trials worldwide (46). Adenoviral particles consist of a 90-nm icosahedral protein capsid that surrounds an inner DNA–protein complex (47). Their protein capsids consist of a penton base at the 12 vertices with each penton base surrounded by five hexon proteins (48). Projecting away from each of the penton bases is a fiber protein, whose C-terminus is called the knob and is responsible for binding to different receptors (49). The primary receptor recognized by most adenovirus serotypes, including the two most common serotypes used in gene therapy, serotypes 2 and 5, is the coxsackie–adenovirus receptor (CAR), which is present in many tissues of the human body (50, 51). After binding, to

Adenovirus 24.7% (n=331) Retrovirus 22.8% (n=305) Naked/plasmid DNA 18% (n=241) Lipofection 7.6% (n=102) Vaccinia virus 6.8% (n=91) Poxvirus 6.4% (n=86) Adeno-associated virus 3.5% (n=47) Herpes simplex virus 3.2% (n=43) RNA transfer 1.3% (n=17) Other categories 2.7% (n=36) Unknown 3% (n=40)

Fig. 2. Gene therapy vectors used in clinical trials (Reproduced from Edelstein et al. (44) J. Gene Med., 2007. Copyright John Wiley & Sons, Ltd., with permission).

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Table 1 The early transcriptional units of the adenoviral vector Unit

Function of protein encoded by unit

Other notes

E1A

Activates other promoters Induces cell into S phase (332)

First unit expressed

E1B

Interacts with p53 to prevent p53-induced apoptosis Interacts with the Bak and Bax proteins to prevent tumor necrosis factor-related apoptosisrelated ligand (333–337)

Protein called E1B 55K Helps cell survive longer so that virus can use cell’s machinery

E2

Regulates viral replication

Proteins include single-stranded DNA binding protein, the precursor terminal protein, and DNA polymerase (338)

E3

Suppresses the infected cell’s immune response, specifically by inhibiting the presentation of the class I MHC peptides and inhibiting cell death by TNFα, Fas, or TRAIL, giving the virus more time to replicate (339, 340)

E4

Affects the infected cell’s metabolism and regulates the cell cycle and DNA repair (341)

enter the cell, the arginine, lysine, aspartic acid (RGD) motif of the penton base associates with integrins αvβ3 and αvβ5, which are located on the cell surface, leading to endocytosis of the viral particle (52). Fifty-one different adenoviral serotypes have been identified, ranging from a genomic size of 24 to 45 kb, with the genome organized into early, intermediate, and late transcriptional units (48, 53). Of the three types of transcriptional units, the early units (E1A, E1B, E2, E3, and E4) are typically the ones most modified during gene therapy (Table 1) (54–57). The majority of the capsid proteins are encoded by the region regulated by the major late promoter located in the late region genes (58). Advantages of Using Adenoviruses for Gene Delivery

Adenoviral vectors have many attractive qualities, making them a popular choice in both in vitro and in vivo studies. The particles are able to accommodate up to 37 kb of DNA inserts. They also are able to efficiently infect and deliver genes to both proliferating cells and resting cells. The genes delivered do not integrate into the host’s genome, but rather provide only a transient cell response (59). This transient expression of genes is normally satisfactory, especially if a toxic chemotherapeutic is used in conjunction with the therapy. Adenoviral vectors can be made in


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high titers and can be modified to infect tumor cells while leaving normal, quiescent cells unaffected (54, 60, 61). Improvements in the Adenovirus Structure Help to Overcome Some of Their Main Disadvantages

One of the biggest drawbacks to adenoviral-based gene therapy is that adenoviruses are only able to enter cells through certain receptors, such as CAR receptors. These receptors are absent on tumor cells (59, 62), so it is difficult for adenoviral vectors to transduce those cells, and this limits their efficiency in gene therapy. To overcome this obstacle, a high concentration of the adenoviral vector is needed, which creates two other problems. First, the high concentration needed produces an innate immune response against the virus, leading to toxic side effects. A 1999 case resulting in the death of an 18-year-old patient from a toxic dose of adenoviral vectors highlighted the importance of this aspect of adenoviral-based gene therapy (63). Second, the adenoviral vector naturally accumulates in the liver in vivo, and also accumulates in the heart, spleen, lung, and kidneys of mice (48). However, the immunogenic response of patients to adenoviral treatment has improved over time as researchers learned from their mistakes. In first-generation adenoviral vectors, E1 genes were deleted to keep the virus from replicating, and often the nonessential E3 genes were also deleted to provide more room for foreign genes (64). Although it was thought that E1-deleted viruses were incapable of replication, instances were found in which the virus would replicate, inducing an immune response and limiting the effectiveness of those vectors (65, 66). In a cystic fibrosis study, the first-generation adenoviral vectors produced pulmonary inflammation in two of three patients (67). In the next two generations of adenoviruses, the E1, E2, and E4 genes were deleted (65, 68, 69). These deletions effectively reduced the immune response to the vectors in vivo while providing more room to insert foreign genes (70). The fourth generation of adenoviral vectors, termed “gutless” or “high-capacity” vectors, can accommodate up to 37 kb (71, 72). Not only do these vectors provide the highest transgene capacity, but they also produce a much lower immune response (73). Several modifications have been made to overcome the adenoviral tropism problem experienced during many cancer gene therapy trials. One approach was the modification of the fiber proteins of the adenoviral vector using other serotypes to allow binding with other non-CAR receptors (74–78). Suominen et al. created an Ad5/35 hybrid vector that was able to infect head and neck tumors more efficiently. Another approach was to modify the HI loop or the C terminus of the fiber knob with different peptides (79, 80). For example, Denby et al. modified the HI loop of Ad19p to accommodate insertions and clones, which resulted in renal targeting of the virus. Yet another approach was to modify the amount of CAR expressed in cells through

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chemotherapeutics such as cisplatin (81, 82). Administration of cisplatin leads to upregulation of CAR in certain cells, which allows Ad vectors to transfect them more efficiently. Researchers have also retargeted adenoviral vectors to other cell receptors through adapter molecules. The adapter molecules contain two parts, an anti-fiber antibody and a cell-binding component. Oh et al. demonstrated that using a folate–polyethyl glycol conjugate could retarget adenoviral vectors to cells expressing the folate receptor (FR) rather than CAR (83). The FR, a tumor-associated glycosylphosphatidylinositol-anchored protein, can actively internalize bound folates and folate-conjugated compounds via receptor-mediated endocytosis (84). 2.1.2. Retroviral Vectors Characteristics of Retroviruses

Advantages of Using Retroviruses

The retroviral family is a group of viruses that use only RNA for genomic material. Like other viruses, retroviruses infect cells to use the cell’s machinery to synthesize its viral proteins. Because retroviruses use RNA for this genomic material, the RNA must first be processed by reverse transcriptase (RT) proteins also found within the virus. These RT proteins convert RNA to fulllength DNA that can be integrated into the cell’s own genome by a protein called integrase to express the viral genes (85–87). There are seven different genera of retroviridae, the α, β, γ, δ, and ε retroviruses, as well as the spumaviruses and lentiviruses (88). Each type of retrovirus contains two copies of a 5′ capped and 3′ polyadenylated RNA molecule. The RNA molecules are positive sense and are joined together at their 5′ ends through a dimerization process (89). Then nucleoplasmid proteins coat the RNA strands (89). There are three open-reading frames within the retrovirus genome, named gag, pol, and env (90). The gag proteins, or group antigens, are internal structural proteins, the pol proteins are polymerase proteins, and the env proteins are envelope proteins. To gain entry into a cell, retroviruses use a variety of cell surface receptors (91). The next step involves the fusion of the cellular membrane with the membrane of the virus, which allows the genetic material and proteins of the virus to enter the cell (92). Retroviruses have been one of the most popular vectors used in gene therapy because their mode of action and biology is well understood (93). They are able to accommodate up to 8 kb of therapeutic genetic material. Simple retroviruses can only attack and infect dividing cells (93). Therefore, retroviruses are suited for many cancer gene therapy trials because many tumor cells are actively dividing during the cancer. This selectivity is a desired quality among therapeutic vectors in cancer gene therapies. Another advantage of retroviruses is that their genome gets integrated into the cell’s genome, thereby providing stable expression of the therapeutic genes (94). This long-term expression may be needed to provide effective therapy. The long-term expression


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also reduces the immune response of the host compared with other types of vectors used for gene therapy (95). Disadvantages of Using Retroviruses

Insertion of the retroviral genomic material into a nonspecific portion of the target cell genome is a concern in retroviral gene therapy (96). Nonspecific insertion can disrupt a current gene or its promoter, thereby affecting the cell negatively. The viral insert also may enhance other genes in its vicinity through the promoters located within the insert, causing those genes to be expressed more or less than normal. Another disadvantage of retroviral vectors is that they cannot be produced at very high titers when compared with other viral vectors, such as adenoviruses. A recent paper has shown that retroviral vectors can be prepared at up to 3.2 × 108 infectious particles/mL, but adenoviruses can be produced at up to 1013 infectious particles/mL (97).

Types of Retroviral Therapy

There are two main strategies involving retroviral gene therapy: treatment either with replication-competent retroviruses (RCRs) or with replication-defective retroviruses. Replication-defective retroviruses were the first to be developed because there was less chance that genes would be inserted in nontargeted cells (98, 99). There was initial hype surrounding replication-defective retroviruses because of a human trial in which X-linked severe combined immunodeficiency (X-SCID) was cured in nine infants by ex vivo gene transfer using retroviruses (33). These infants would have had an extremely low survival rate had it not been for the gene therapy trial. However, leukemia developed in three of the nine infants later in life, and the leukemia was thought to be triggered by the retroviral insertion of a proto-oncogene (100).

Replication-Defective Retroviruses

Several approaches have been used in the use of replication-defective retroviruses in gene therapy, some of which are similar to those used with other vectors. One of these approaches is to modify the retroviral envelope proteins to make them more selective toward specific cancer cells. Because the envelope proteins bind to different receptors within cells, by changing those proteins, different cancer cells with certain overexpressed receptors can be targeted. Table 2 lists commonly modified envelope proteins (94). Fear of random integration of the retroviral genes into the genome of the targeted cells, as in the case of the X-SCID patients, has prompted researchers to develop better methods for targeted integration. Certain sites within the cell genome, called hot spots, are more common retroviral insertion points than others, indicating that the process of insertion is not a random event and might be affected by transcription (101). To target the site of integration, researchers have combined a DNA-recognition sequence with the viral integrase. For example, the polydactyl zinc finger protein E2C was fused with HIV I integrase to produce

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Table 2 Retroviral vectors with modifications* Modification

Target cell

Reference

EGF-MMP cleavable linker chimeric env

Cancer invasion, angiogenesis, inflammation

(342–344)

IL-2 chimeric env

IL-2 R

(345)

EGF chimeric env

EGFR

(346)

SCF-Factor Xa chimeric env

Stem cell (Kit)

(347)

vWF (collagen binding) chimeric env

Cancer (collagen expressing), vascular lesion

(348, 349)

Single-chain variable fragmented antibody Cancer (brain, breast, lung, ovary) (scFv) for EGFRvIII

(350)

scFv for HMWMAA

Cancer

(351)

scFv from phage display

T cell

(352)

scFv for carcino-embryonic antigen (CEA)

Cancer

(353)

Receptor pseudotyping (CD4 and CXCR4)

HIV-1 infected cell

(354, 355)

*Reproduced from Yi et al. (94) Curr. Gene Ther., 2005, by permission of Bentham Science Publishers, Ltd.

retroviral transgene vectors that integrated very near the E2C binding site in vitro; this produced promising results (102). Another major problem associated with retroviral insertion is the silencing of retroviral transgenes (103). To prevent this silencing from occurring, special segments of DNA, called chromosomal insulators, can be used. These insulators protect the retroviral transgene from being affected by other promoters and genes in its vicinity (104). Replication-Competent Retroviruses

In contrast to replication-defective retroviruses, RCRs are not made with packaging cells and can replicate in infected cells. The main advantage of using RCRs is that they can increase the efficiency of transgene integration into tumor cells (105). This is because each time a cell is infected, the infection spreads to other tumor cells. For example, Wang et al. demonstrated that replication-defective retroviruses used in U-87 tumor cells infected only 0.2% of the tumor after 6 weeks. On the other hand, RCRs injected into the same type of cells infected more than 97% of the tumor, and most importantly, no RCRs were detected in nontumor tissues (106). Another advantage of retroviral gene therapy, and specifically RCRs, is that the host immune response is often low when compared with other methods of gene therapy, such


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as adenoviruses (95, 107). Finally, fewer RCR viral particles are needed for infection. For instance, Solly et al. report that RCRs were 1,000 times more efficient at delivering suicide genes to brain tumors than adenoviral vectors (108). The major disadvantage when using RCRs is the unchecked spread of the virus particles throughout the body. However, like replication-defective retroviruses, RCRs also cannot infect cells unless they are actively dividing because they contain no nuclear localization signals (105). It has been shown that RCRs are detectable in tumor cells not only when injected directly into the tumor (106), but also when injected intravenously in a nearby area (109). Also, because RCRs replicate, and thus insert their genome into more cells than replication-defective retroviruses, there exists a greater chance for a nonspecific insertion to have negative effects. Leukemia has developed not only in humans, but also in rats and primates that had retroviral gene therapy (33, 110, 111). 2.1.3. Lentiviral Vectors

Lentiviruses are members of the Retroviridae family. There are various types of lentiviruses, which can be classified according to the animal species from which they are derived. Although nonhuman lentiviral vectors, such as simian (112) and feline (113) vectors have been used in research, HIV-derived vectors were the first to be developed and have been most intensively studied because of their greater potential for use in human therapy (114) and their high-titer vector stocks (115). The unique ability of lentiviruses to infect both dividing and quiescent cells (115, 116) has facilitated their usage in targeting cells of the nervous system, muscles, lungs, and liver. The use of lentiviral vectors in animal models has met with the most success in treating neurological disorders such as Alzheimer’s disease, amyotrophic lateral sclerosis (ALS), Huntington disease, and Parkinson’s disease. Early efforts at transducing rodent neurons (117) and retinal cells (118) in vivo with lentiviral vectors revealed not only the feasibility of transfecting quiescent cells, but also the prolonged expression of lentivirally delivered genes. Sustained expression is the result of stable integration of lentiviral genes within the host genome. This relatively long period of lentiviral genome expression (“lenti” is derived from the Latin word for “slow”) reduces the need for reintroduction of the genes of interest, which makes lentiviruses an attractive clinical prospect. Recent studies continue to demonstrate the ability of lentiviral vectors to transduce an array of animal and human cell types. Recent advances in pseudotyping, which involves substituting envelope glycoproteins that act as cellular ligands, have made possible the targeting of specific cell types (119). Pseudotyping with certain envelope proteins has also allowed modification of mechanisms of transport inside transduced cells. For example, several groups have pseudotyped equine infectious anemia virus (EIAV) vectors with a rabies G-envelope protein, which has made

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it possible to target distant neuronal locations via retrograde transport (120, 121). The comparatively low immunogenicity of lentiviral vectors with respect to other vectors is yet another distinct advantage of their use. Many studies have reported an absence of rodent immune response (122–124) or toxicity (125) after in vitro introduction of HIV vectors. Because lentiviral vectors do not encode viral proteins after transduction, as most viral genes can be removed (126), only the viral particle or the transgene product may be targeted by an immune response. In clinical trials using HIV as the vector of choice, it is suspected that only patients with HIV and the circulating antibodies against protein in the vector would elicit an immune response. Further, the HIV capsid breaks down rather quickly, thus reducing the timeframe during which the immune system can recognize the transduced cells (127). The ability of lentiviral vectors to efficiently deliver, into both dividing and nondividing cells, a set of genes capable of prolonged expression and little immunogenicity has made this vector an attractive option in clinical trials. However, as with all forms of gene therapy, lentiviral vectors come with functional shortcomings and biosafety concerns. An overarching limitation of the lentiviral system is the relatively short lentivirus genome (around 8–10 kb), which restricts the amount of transferable genetic material. Depending on the nature of the targeted disease, the poor diffusion of lentiviruses after injection may also pose a problem, because a multitude of concurrent injections may be required to sufficiently target a large cell population. However, the limited diffusion of lentiviral vectors has proven useful in targeting the small, restricted cell populations implicated in Parkinson’s disease. Insertional mutagenesis, which may disrupt the expression of essential genes, and the potential for replication competency represent the main biosafety concerns associated with lentiviral vectors. Various degrees of “minimal” packaging systems for HIV (128–130) as well as EIAV (112, 131) and FIV (113) have been developed to increase safety by reducing the likelihood that the intact virus will be reconstituted via recombination. One such “third-generation” system consists of only three of the nine original genes comprising HIV-1 (128). The deleted virulence genes can be replaced with transgenes of interest, and the remaining HIV-1 genes regulate the transfer of genetic material. Thus, the threat of reconstituting a full HIV genome is largely circumvented. Recent advances in developing “self-inactivating” vectors have also increased the safety of this gene delivery system (132, 133). These vectors permit regulation of or even switching off transgene expression, which may be necessary in the event that negative side effects occur. The ability to downregulate or terminate expression may also prove useful in treating a temporary illness that may not require prolonged expression of the therapeutic transgenes (134).


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Overall, the ability of lentiviruses to stably integrate their viral genome into a wide range of host cells provides an important advantage over other types of therapy. This innate flexibility of lentiviral vectors, along with the added flexibility obtained through pseudotyping and self-inactivation, makes lentiviruses promising vehicles for gene therapy. 2.1.4. Herpes Simplex Vectors

Recent scientific advances have revealed the means by which to convert the dreaded destructive power of herpes simplex virus strains into life-saving medical applications. Among the three known forms of human herpes simplex virus, type I (HSV-I), which is known to cause cold sores, has proven the most promising for use as a vector in gene therapy. There is currently great potential for the use of HSV-1 to selectively destroy tumor cells and treat various neurological diseases. Several properties of HSV render it an important vector for gene therapy. HSV-1 has a large, 152-kb genome, 30–50 kb of which can be replaced by desired transgenes (135, 136). The herpes simplex virus (HSV) replicates lytically in an array of cell types (137), with minimal toxicity to normal tissue, which enables it to kill various types of cancer cells. The innate tendency of HSV to form latent infections in neurons without destroying them makes this vector particularly useful in cases in which a persistent expression of the transferred genetic material is preferred. The ability of HSV-1 to spread retrogradely and anterogradely within the nervous system via neuronal transport may be useful in treating diseases with an expansive spatial distribution, because the therapeutic genetic material can be delivered over a broad spatial range (138). Together, these factors also reduce the need for repeated injections, which may damage surrounding tissue and dissuade patients from continuing treatment. Unlike lentiviral vectors, herpes vectors pose no threat of insertional mutagenesis because they remain episomal within the infected cell (136, 139). However, this comparatively heightened degree of safety may come at the cost of a reduced period of transgene expression, which may limit the usefulness of HSV treatment for chronic conditions (134). The possibility of inciting an immune response in patients previously exposed to the herpes virus stands as an apparent shortcoming to the use of herpes viruses in gene therapy. Attenuated forms of the virus containing fewer genes can reduce the threat of immune response (139), but the risk of recombination of a herpes virus genome in patients is not yet fully understood. Because HSV-1 crosses the blood–brain barrier and naturally persists in neurons, in animal models, gene therapy vectors based on this virus have proven very effective at treating neurological diseases such as brain tumors (140), various brain diseases (141–143), and disease within the peripheral nervous system (PNS), including

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Parkinson’s disease (144). There is concern that introducing HSV-1 into the brain may initiate an immune response and lead to encephalitis. To circumvent this threat, the viral (immediate-early) genes that encode lytic functions of HSV can be deleted, leaving a recombinant noncytotoxic vector with the ability to express transgenes and the capacity for latency (145). The inherent cytotoxicity and replicative power of the herpes virus can be easily modified to target a focal set of cells, such as a localized brain tumor, while leaving the surrounding tissue unscathed. This can be achieved by targeting the surface of the tumor cell or by benefiting from aberrant intracellular signaling, which results in tumor cells with a defective antivirus response (146). Several studies have used nonpathogenic, replicationcompetent viruses in animal studies to selectively target and kill cancer cells in tissues ranging from brain tumors (147) to diffuse liver metastases from colon carcinoma (148). The application of HSV to the treatment of cancer in clinical trials has provided encouraging results. Numerous oncolytic HSV vectors with varying degrees of genomic attenuation have been created, and many forms, including G207, 1716, and NV1020, have proven safe in phase I clinical trials (149). A study that tested the conditionally replicative G207 form of HSV-1 in early clinical trials against malignant glioblastoma reported antitumor activity and an absence of induced toxicity or encephalitis (150). Early clinical trials with 1,716 subjects have also proven these vectors to be safe in treatments of glioma (151, 152) and melanoma (153). Current efforts at improving the efficacy of oncolytic HSV cancer treatments include testing alternative routes of virus delivery, improving virus specificity for the target tumor or tissue, and considering effective methods of combining traditional cancer therapy, such as chemotherapy, with HSV gene therapy (149). Ongoing work also involves improving the sophistication and intricacy of attenuated HSV genomes, because overly attenuated genomes may reduce desired levels of viral spread and replication (146). The combined characteristics of the herpes virus render it a promising option for treating cancer and various neurological diseases. As with other gene therapy vectors, much remains to be accomplished before the potential of this powerful virus can be fully harnessed. Future clinical applications of HSV will undoubtedly involve variants of the virus modified for more specific cases, involving tailoring the virus toward distinct tissue types and optimization of treatment duration and potency. 2.1.5. SV40 In Vitro Packaging Vectors: An Efficient Method to Deliver DNA and RNA into Cells

Simian virus 40 (SV40) is a non-enveloped polyomavirus with a double-stranded circular DNA of 5.2 kb. SV40 infection differs from most other viral infection pathways. It begins with the virus binding to its primary receptor, the major histocompatibility complex (MHC) class I. Then the virus binds to GM1 gangliosides,


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and travels with them to the endoplasmic reticulum (ER). This transport to the ER is mediated by caveolae and facilitated by ER chaperones. Following disassembly, nuclear pore complexes assist in delivery of the viral DNA to the nucleus for replication. Wildtype SV40 encodes two early genes, the large T antigen (Tag), which is essential for viral genome replication and for expression of the late gene, and the small t antigen (tag) (154, 155). Tag has been reported to have the ability to bind and inactivate p53 and pRb tumor suppressors; hence, removal of Tag produces a safer recombinant virus that is nonreplicating and has not been reported to be immunogenic. When prepared in vitro, the delivery system is composed of only the SV40 envelope major protein (VP1), which is produced in Sf9 insect cells (156). These particles do not harbor any of the wild-type SV40 genes, and no encapsidation sequences are required, which reduces the potential adverse effects that accompany any viral-based delivery system. The lack of viral genetic material allows room for larger DNA plasmids (up to ~17 kb) to be packaged very efficiently (157). In this delivery system, it is possible to package large genes together with a selective marker (such as the ABCB1 gene). When prepared in vivo, removal of the large T antigen makes space for a transgene of up to 5.3 kb. The rSV40 particles are propagated using either a wild-type or a temperature-sensitive mutant of SV40 as a helper via a viral producer cell line, COS7 (158). The packaging cell line COS7 stably expresses an origin-defective SV40 mutant and is capable of supporting the lytic cycle of SV40. With its unique pathway, gene delivery via SV40 has proved to be very suitable for numerous gene therapy approaches (159). Table 3 summarizes SV40 characteristics and advantages as a gene delivery system. We and others have demonstrated several possible applications of SV40 gene delivery, such as: 1. Delivery of drug-selectable markers such as ABCB1 (160) and ABCC1 into various murine and human cell types, including primary human bone marrow cells. The short-term expression of the SV40/ABCB1 in vitro vectors may be an advantage for use in chemoprotection. 2. Delivery of in vitro-packaged GFP vectors or lethal genes through the SV40 delivery system into live animal models. SV40 pseudovirion delivery of Pseudomonas exotoxin A (PE38) was found to be effective in the treatment of human adenocarcinomas growing in mice either by direct injection or systemically. Using a combined treatment of SV40-PE38 with doxorubicin reduces the side effects of chemotherapy (161). 3. Delivery of clotting factors—the Von Willebrand Factor gene, or its cleavage protease, the ADAMTS13 gene, into erythroleukemia

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Table 3 Characteristics and advantages of the SV40 gene delivery vehicles* Subject

In vitro packaging and recombinant vectors

Tropism

Virus particles are able to infect a wide variety of dividing as well as nondividing mammalian cells, including human cells (356, 357), and to express genes in these cells with high efficiency. Successful infection of live animal models

Delivery of various nucleic acids

In addition to circular DNA, the SV40 vector system is able to deliver untranslated RNA products (358); in vitro particles carrying siRNA were shown to silence green fluorescent protein (GFP) expression with better efficiency than with lipid transfection protocols

High capacity

In vitro-packaged SV40 vectors that do not contain SV40 sequences permit packaging of plasmid DNA up to 17 kb in size (359)

Persistence of expression

In vitro preparation results in short-term expression only, which is beneficial in many therapeutic environments; long-term expression can be achieved either by using a selective marker that conjugates to the gene; or by delivery of the LTR promoter, which can maintain the transgene expression (156). In vivo preparation results in long-term expression (360)

High titers

In vitro preparation produces approximately 106 infectious units/mL. Multiple infections result in higher titers of the virus—up to 1010 infectious units/mL (361)

Expression levels

Introduction of the in vitro-packaged genes at 24-h intervals results in greater expression than a single transduction. Using the SV40 in vivo system results in numerous copies of recombinant plasmids per cell (362)

Straightforward preparation of vectors

The in vitro packaging system preparation is a very straightforward process; the particles can be prepared in large batches; virions can be stored for years at −20°C. No packaging cell line is required (156)

*Reprinted from Devi (283) by permission of Macmillan Publishers, Ltd.: Cancer Gene Ther., copyright 2006.

K562 cells. Resting cells as well as dividing cells can express these genes. 4. Delivery and expression of small interfering RNA (siRNA) into cells in suspension as well as adherent cells. In conclusion, SV40-based vectors have promising potential in gene therapy; they are highly efficient vehicles that can infect a wide variety of cells with relatively large amounts of genetic material. 2.2. Nonviral Delivery Systems

As discussed above, although viruses are highly efficient in the delivery of a transgene, their use suffers from several severe drawbacks, such as toxicity, immunogenicity, and limited packaging capacity. Therefore, nonviral delivery systems represent an interesting alternative to viral vectors. They are easier to prepare and scale-up at the manufacturing stage, more flexible regard-


23

ing the transgene size being transferred, and are less immunogenic than viral vectors. There are three main nonviral vector systems: naked DNA, cationic liposomes, and cationic polymers. For excellent reviews, see Kodama et al. (162), Park et al. (163), and Gardlik et al. (164). 2.2.1. Naked DNA

The simplest nonviral treatment is the transfer of naked plasmid DNA by injection to local tissues. Physical delivery of the DNA directly into the cytoplasm, bypassing the endosomes and lysosomes, avoids enzymatic degradation. Naked DNA has been successfully used for DNA vaccination. An antigen-encoding DNA is intramuscularly administered to produce a protective immune response to the transgene antigenic product (165, 166). Success has also been reported after direct injection of naked DNA into the epidermis and hair follicles (167), into the liver (168), and into solid tumors (169). However, low transfection efficiency has often been reported, because the lack of DNA protection results in rapid degradation of the DNA. Several methods, such as electroporation (170, 171), gene-gun (172–174), and ultrasound (175, 176), were developed to increase the transfection efficiency and expression level. Although these methods may be highly toxic to the cells, recent reports indicate interesting prospects for the success of this type of treatment (172–176).

2.2.2. Cationic Lipids

Uptake-enhancing chemicals, such as lipids and synthetic polymers, were developed in the late 1980s. They not only facilitate the transfer of the DNA into the cell, but also protect the DNA from degradation. A lipid-mediated DNA-transfection procedure was developed in 1987 (177). Since then, numerous synthetic cationic lipids have been used as nonviral carriers for gene delivery. These lipids, termed liposomes, form large complexes (lipoplexes) (178) in which their positive charge chains interact with DNA and their hydrophobic portions interact with the negatively charged cell membrane (179) (Fig. 3). Most of the liposomes used are mixed with a neutral helper lipid (co-lipids) such as dioleoyl phosphatidylethanolamine (DOPE) (180) and cholesterol (181, 182). The co-lipid facilitates the formation of a stable lipid bilayer and improves in vivo transfection by facilitating endosomal escape (183–185). Although cationic liposomes prevent DNA degradation in the plasma (186), they are rapidly cleared by the lungs (187). This serious drawback can be overcome by incorporating polyethylene glycol (PEG) lipids into the lipoplexes; this diverts lipoplexes from the lungs (188, 189). Cationic lipids forming micellar structures called liposomes are complexed with DNA to create lipoplexes. The liposomes sometimes fuse with the cell membrane after interactions with surface proteoglycans. The complexes are internalized by endocytosis, resulting in the formation of a double-layer inverted micellar

24

Gillet et al. Cell membrane

Nucleus

– –

Proteoglycan

–

– – –

+

– – –

–

Lipoplex

+

DNA

+

+

–

Lipoplex

+ +

+

+

Endosome with lipid mixing

Endocytosis

+

Liposome

– –

+ +

–

+

DNA

Escape from endosome

–

Endosome

Endosomal maturation

–

DNA fragmentation

DNA

+

Lysosome

Fig. 3. Lipoplex-mediated transfection and endocytosis. Cationic lipids forming micellar structures called liposomes are complexed with DNA to create lipoplexes. The structures sometimes fuse with the cell membrane after interactions with surface proteoglycans. The complexes are internalized by endocytosis, resulting in the formation of a double-layer inverted micellar vesicle. During the maturation of the endosome into a lysosome, the endosomal wall might rupture, releasing the contained DNA into the cytoplasm and potentially toward the nucleus. DNA imported into the nucleus might result in gene expression. Alternatively, DNA might be degraded within the lysosome (Reproduced from Parker et al. (331) Expert Rev. Mol. Med., 2003, by permission of Cambridge University Press).

vesicle. During the maturation of the endosome into a lysosome, the endosomal wall might rupture, releasing the contained DNA into the cytoplasm and potentially toward the nucleus. DNA imported into the nucleus might result in gene expression. Alternatively, DNA might be degraded within the lysosome. 2.2.3. Cationic and Nanoparticles Polymers

Cationic polymers are other chemicals that can condense DNA in complexes termed polyplexes. The most studied polyplexes are polylysine-, polyethylenimine (PEI)-, and polyamidoamine-based polymers. Polylysine polymers can be used to mediate gene transfer in a target-specific manner using ligands such as asialoorosomucoid (190), transferrin (191), and epidermal growth factor (192) to facilitate cellular uptake. In contrast to polylysine, polyethylenimine polymers do not require endosomolytic (inactivated adenovirus) or lysosomotropic (e.g., chloroquine) agents to facilitate the intracellular release of DNA (193, 194). Like polylysine polymers, PEI efficiency can be enhanced by incorporation of a cell-binding ligand such as transferrin (195). As is the case with cationic lipids, both polylysine and PEI polymers are rapidly cleared from the plasma after intravenous injections (196). Their opsonization by plasma proteins can be circumvented by coating polyplexes with a hydrophilic polymer such as hydroxypropyl methacrylic acid


25

or PEG (197). Polyamidoamine dendrimers, highly branched cationic polymers, have also been shown to be efficient gene delivery vectors (198, 199). Polymer nanoparticles such as gelatin (200), methacrylate/ methacrylamide polymers (201), and pyridinium surfactants (202) are an additional set of nonviral vectors used for gene delivery. One nanoparticle, chitosan (203), was successfully used as an oral delivery vector in several applications (204, 205). 2.2.4. Prospects

Besides some toxicity and relatively low transfection efficiency, the main limitation of nonviral delivery systems is the absence of stable and long-lasting expression. The addition of an SV40 or Epstein-Barr virus (EBV) replicon to the nonviral vector can circumvent this issue. Indeed, these sequences ensure the presence of the vector as a stable episome and thus allow extrachromosomal replication. For this purpose, artificial chromosomes are of great interest. Several studies have recently demonstrated the potential of such approaches to gene therapy using human cell lines and mice models (206–210).

2.3. Bacterial Delivery Systems

Bacteria can also mediate gene transfer to mammalian cells (211, 212) and can be designated as a third type of gene delivery system termed bactofection (for reviews, see Gardlick et al. (164), Vassaux (213), and Palffy et al. (214)). Various attenuated bacteria strains are used in gene therapy. They include Escherichia coli, Salmonella, Yersinia, Shigella, and Listeria. These bacteria can enter cells by phagocytosis, pinocytosis (Salmonella, Shigella), or by bacterial protein binding to a host cell ligand (Yersinia, Listeria). Once they reach the cell, they localize primarily in the cytoplasm (Listeria, Shigella), in vacuoles (Salmonella, Yersinia), or in extracellular space (agrobacterium). Bacterial delivery systems have been successfully used in oral vaccination. For this purpose, the bacteria, which carry a plasmidencoded antigen under the control of a eukaryotic promoter, target the macrophages or the dendritic cells and enter these cells by phagocytosis. Shigella, Salmonella, and Yersinia have been used to induce immune responses against bacterial and viral antigens such as brucella (215), hepatitis B (216), and herpes simplex virus type 2 (217). Many Salmonella strains have also proved to be very useful for vaccination against a tumor antigen. Among several examples, Niethammer and colleagues reported an exciting and promising study (218) using an oral DNA vaccine mediated by Salmonella enterica serovar typhimurium to target proliferating endothelial cells in tumor vasculature through upregulated vascular endothelial growth factor receptor 2 (FLK-1). This vaccine effectively protected mice from lethal challenges with melanoma, colon,

26

Gillet et al.

and lung carcinoma cells and reduced the growth of established metastases (218). In addition, the antiangiogenic and antitumor effects were enhanced by co-delivery of a plasmid encoding the cytokine interleukin (IL)-12 (219). Finally, some anaerobic bacteria strains such as Clostridium (220) and Bifidobacterium (221) have the ability to specifically colonize the tumor site, leaving normal tissues unaffected. Although an oncolytic effect was observed for large tumors with hypoxic centers, the anaerobic bacteria had little effect on small metastatic lesions. In contrast to these bacteria, auxotrophic Salmonella strains have the potential to colonize both large tumors and small metastatic lesions (222). Nemunaitis and colleagues reported a promising pilot clinical trial in three refractory cancer patients using attenuated Salmonella expressing the cytosine deaminase gene, which can sensitize tumors to 5-fluorocytosine by converting 5-fluorocytosine to 5-fluorouracil. Two patients showed evidence of bacterial colonization of the tumor with conversion of 5-FC to 5-FU (223). Although bacteria are characterized by high selectivity in tissue colonization, this selectivity and, thus, the therapeutic efficiency, can be improved by the generation of a transgene-expressing vector under the control of a radiation-inducible promoter (224). This strategy allows highly specific expression of the transgene in bacteria colonizing irradiated tissues and avoids expression in nontumor and hypoxic tissues (e.g., infarction).

3. Strategies for Gene Therapy Developing successful treatments for particular cancers depends, to a large extent, on our knowledge concerning those diseases. Continuous advances in the understanding of tumor biology refine our understanding of cellular pathways and thus also highlight numerous new chemotherapeutic targets. This knowledge has allowed the development of elegant, more efficient, and less toxic strategies for the patient. Figure 4 summarizes the main gene types used as of July 2007 (44). Immunotherapy was used in approximately 50% of the trials, whereas 8.2% of the trials were based on suicide gene therapy. In 2.1% of trials, oncolytic viruses were transferred, whereas 1.7% of trials involved RNA interference (RNAi) therapy (44). 3.1. Introduction of Tumor Suppressors

Cells are continuously assaulted by stress and this leads to a dilemma: repair the DNA damage and live, or die. A defect in this decision process can contribute to the development of cancer. This important choice is made by tumor suppressors such as p53,


27

Antigen 20.3% (n=266) Cytokine 18.9% (n=247) Tumor supressor 12% (n=157) Growth factor 8.2% (n=107) Suicide 8.2% (n=107) Deficiency 7.9% (n=103) Receptor 5.1% (n=67) Marker 4.1% (n=54) Replication inhibitor 3.7% (n=48) Other categories 8.6% (n=115) Unknown 2.9% (n=38)

Fig. 4. Gene types transferred in gene therapy clinical trials (Reproduced from Edelstein et al. (44) J. Gene Med., 2007. Copyright John Wiley & Sons, Ltd., with permission).

which is mutated in most cancers. The restoration of p53 function in tumor cells has been considered an obvious way to treat cancer (for review, see Kumar et al. (225)). Both retroviral and adenoviral vectors have been used to restore p53 function in vitro in p53-defective tumor cell lines (226, 227) or in vivo in mice (228, 229), resulting in growth arrest or apoptosis. As discussed earlier, the random integration of retroviral genetic material into the genome and the mutations generated through their replication process severely hamper the use of retroviruses (230). However, two commercial adenoviruses, termed Advexin/INGN 201 (Introgen Therapeutics, Houston, TX, USA) and Gendicine (SiBiono Genetech Co., Shenzhen, China), have entered clinical trials using this approach. Gendicine is the world’s first commercially available gene therapy product. Although it was approved by China’s State Food and Drug Administration (SFDA) in 2003 for clinical use and licensed for commercial production in 2004, questions remain concerning its efficacy (231). Advexin is now in phase III clinical trials in the USA as a monotherapy or in combination with other treatments (232–234), and the evaluation of the efficacy of Gendicine is still underway. Another example of a tumor suppressor approach that has had success is an adenoviral vector with the retinoblastoma (Rb)binding CR2 region of E1A deleted. Viruses can only replicate in cells with a defective Rb pathway, because the Rb protein normally blocks the genes needed for the S phase of cell division. When Rb is removed, the genes can be transcribed (55). Success using the E1A deletion mutant dl922-947 has been shown for ovarian cancer, where the median survival was increased from 20 to 96 days in nude mice (235). Recent studies have highlighted additional tumor suppressors that could be used in gene therapy, especially in lung cancer.

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Indeed, it has been shown that a genomic aberration in the human chromosome 3p is the most frequent and earliest genetic event in lung tumorigenesis (236). This genomic aberration affects several tumor suppressor genes, such as FUS1, 101F6, NPRL2 (237), and FHIT (238). Finally, this type of gene therapy may be combined with treatments such as radiotherapy and chemotherapy or other strategies such as the induction of apoptosis to yield more effective results. For example, Guo et al. used both tumor necrosis factor-related apoptosis-inducing ligand (TRAIL)-induced gene therapy and Ad-Δ24, another adenoviral vector with the Rb-binding region of E1A deleted, to produce greater antitumor activity than either therapy alone in murine breast cancer cells (239). 3.2. Suicide Gene Therapy

Suicide gene therapy, also termed prodrug activation or gene-directed enzyme–prodrug therapy, involves the use of an enzyme-encoding transgene that transforms a prodrug into a toxic chemical inside the cell, with the added advantage of the bystander effect, in which cells nearby are also affected. Enzyme/prodrugs used include herpes simplex virus thymidine kinase (HSV-tk)/ganciclovir (240), E. coli cytosine deaminase (CD)/5-fluorocytosine (241), and cytochrome P450 2B (CYP2B)/cyclophosphamide (242), among others. When herpes simplex virus thymidine kinase (HSV-tk) is delivered to tumor cells by adenoviral vectors, followed by injection of the prodrug ganciclovir (243), the HSV-tk protein converts ganciclovir to a ganciclovir triphosphate nucleotide, which inhibits DNA replication (59). Although initial studies did not show that HSV-tk and ganciclovir could cure malignant tumors, recent studies have shown more promise. Mathis et al. have described a new system that regulates the translation of the enzyme in cells with high levels of eIF4E, as is the case in most tumor cells. They report that this system is effective for cells in vitro and in vivo (244). HSV-tk therapy can also be combined with other methods of gene therapy for increased efficiency. Kagaya et al. have shown that colon cancer treatment using HSV-tk and ganciclovir can be enhanced by monocyte chemoattractant protein (MCP)-1 in nude mice (245). Another enzyme–prodrug combination is E. coli cytosine deaminase (CD), which converts 5-fluorocytosine into 5-fluorouracil, a toxic metabolite. This approach may have more potential than other suicide gene therapies because CD transgene expression led to antitumor efficacy in mouse tumor xenografts for both prostate cancer (246) and melanoma (247). Retroviral gene therapy transducing herpes simplex virus thymidine kinase (HSV-tk) was also investigated (248). However, when replication-defective retroviruses were used, the results were disappointing because of low efficiency of the vector in a mouse model for pancreatic cancer (249) and in a phase I/II clinical trial (250) and a phase III clinical trial (251), both to treat glioblastoma


29

multiforme. To solve this problem, replication-competent retroviruses (RCRs) were used with cytosine deaminase and the prodrug 5-fluorocytosine in U-87 intracranial gliomas with a 100% survival rate during a 60-day period compared with a 0% survival rate in the control group within the same period (106). Replication-competent retroviruses were also used to treat bladder tumor xenografts and were able to infect up to 93% of the tumor (252). Some studies have shown that suicide gene therapy can be improved by more accurate targeting of tumors through “transcriptional targeting.” This approach involves replacing the viral promoters and enhancers with promoters and enhancers specific for or overexpressed in certain tissues, thereby inducing the transgene to be expressed only in those tissues and minimizing the chance of its expression in nontargeted tissues. Examples include the Co11a1 promoter, which has been used to target bone cells (253); the tyrosinase promoter, used to target melanoma cells (254); the human T cell-specific CD2 enhancer, used to target T lymphocytes (255); and MCK enhancers, which have been used to target skeletal muscle cells (256). Another interesting approach is the improvement of the tumor distribution of the prodrug by using suicide gene mutants that activate the compound with higher catalytic efficiency (for review, see Griesenbach (43)). 3.3. Oncolytic Viruses

Oncolytic viruses have the ability to selectively replicate and cause tumor cell death. Once the tumor cells undergo lysis, the release of mature viral particles can infect neighboring cells. Two types of oncolytic viruses have been tested: (1) viruses showing inherent tumor selectivity such as Newcastle disease virus and reoviruses and (2) engineered viruses such as ONYX-015 and HSV1-G207 (for reviews, see Woo et al. (257) and Crompton and Kiru (258)). ONYX-015 (dl1520) is the first oncolytic virus that has been tested in a clinical trial. This engineered adenovirus, in which the E1B-55K gene is deleted, can replicate in and selectively kill p53-negative cells (54). E1A viral proteins induce the infected cell to enter the S phase, simultaneously stimulating apoptosis through p53, while E1B proteins bind to p53 to prevent cell death. Therefore, a virus without E1B will not be able to replicate in cells with a normal p53 mechanism, but will be able to replicate in cells with a defective p53 mechanism, as is the case in many tumor cells. A phase II clinical trial involving a treatment using ONYX-015, cisplatin, and 5-fluorouracil for head and neck carcinomas produced some promising results (41). HSV1 engineered viruses are also being studied in ongoing clinical trials. The double-mutant G207 herpes simplex virus and HSV1716 (similar to G207) were administered in phase I dose escalation studies in patients with recurrent gliomas (150, 151). No toxicities have been reported (see Barzon et al. for review (259)).

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Newcastle disease virus is an avian paramyxovirus that exploits defects in the interferon response pathway to achieve tumor selectivity. The oncolytic NDV-PV701 strain has been tested in several phase I trials (260–263). Pecora and colleagues reported a phase I trial including 79 patients with various advanced solid cancers (263). The patients exhibited adverse events such as flulike symptoms, but these adverse events decreased in number and severity with each subsequent dose (263). The treatment was later improved by a two-step desensitization (261) and by slowing the intravenous infusion rate (260), both improving the tolerability of repeated high-dose administration. Early clinical trial results have been very promising and have demonstrated the relative safety of these viruses. As discussed previously, additional research is needed to define the most efficient treatment modalities. Other viruses are also currently under scrutiny, such as reoviruses, another type of intrinsically tumorselective oncolytic virus that targets Ras-activated tumors (264). 3.4. From Radioinducible to Chemoinducible Gene Therapy

Radiation-targeted gene therapy is based on the identification of radiation-inducible genes such as TNFα and Egr-1 (265–267). It has also been shown that the CArG box, a promoter element of the Egr-1 gene, was the only necessary sequence for radioinduction (267). Technically, this led to a strategy based on the cloning of a radiation responsive sequence such as the CArG box upstream from the transgene. Ionizing radiation can then regulate the spatial distribution and maintain temporal control of transgene expression through the radiation responsive sequence (for review, see Mezhir et al. (268)). Because early studies revealed the expression of TNFα after radiation treatment, it made this gene a good candidate for radiation-targeted therapy (265, 266). In radioresistant animal tumors, the direct injection of an Egr-1/TNFα construct first (269, 270), followed by the use of a replication-incompetent adenovirus that carries the Egr1/TNFα construct (Ad.Egr-TNF), showed tumor regression with neither local nor systemic toxicity (271, 272). Senzer et al. conducted the first phase I study in which TNFerade, a secondgeneration replication-deficient Ad.Egr-TNF, was administered (273). They reported that 21 out of 30 patients with solid tumors responded well to the treatment, with 5 of the patients having complete regression of the tumor. Another phase I trial assessed the tolerance of this treatment using an intravenous administration of TNFerade in contrast to the intratumoral administration reported by Senzer and colleagues (274). The treatment was well tolerated and the patients’ response was encouraging. Once the virus carrying the transgene is administered, ionizing radiation generates radical oxygen intermediates (ROI),


31

which activate the Egr-1 promoter and thus results in localized transgene expression. Chemotherapeutic drugs such as cisplatin or temozolomide are known to induce ROI. Therefore, studies have been undertaken to investigate their application as chemoinducible gene therapies (for review, see Mezhir et al. (275)). It has been clearly demonstrated that anticancer drugs can induce transgene expression (276, 277). Moreover, using a murine model of glioblastoma, it was shown that the combination of both modalities, Ad.Egr-TNF and the alkylating agent temozolomide, results in a higher cytotoxicity than either treatment alone and significantly increased the survival time (278, 279). The promoter of MDR1 harbors drug-responsive elements mediating the drug inducibility of MDR1 gene expression. Walther and colleagues exploited this characteristic to develop a construct expressing TNFα under the control of MDR1 promoter elements (280). The in vivo study highlighted a high expression of the transgene after doxorubicin and vincristine induction followed by a significant tumor decrease (280). The same group also exploited the heat-responsive elements of the MDR1 promoter (281). A more recent study reported that in vivo heat-induced TNFα expression combined with Adriamycin treatment led to the inhibition of tumor growth in animals (282). Although these results support the idea that MDR1 promoterdriven expression of therapeutic genes is efficient and feasible for this particular gene therapy approach, this idea has not yet been evaluated in clinical trials. 3.5. Posttranscriptional Gene Silencing or RNA Interference

RNA interference (RNAi) is a posttranscriptional gene-silencing pathway that has opened new avenues in numerous fields, especially in gene therapy. It is now conceivable to downregulate a gene specifically involved in the development or the progression of cancer without any toxicity in normal cells. Other approaches targeted to RNA downregulation include hammerhead ribozymes, antisense oligonucleotides, and peptide nucleic acids (for reviews, see Devi (283) and Karagiannis and Assam (284)). Preclinical cancer studies have shown inhibition of tumor growth by RNAimediated downregulation of various genes such as a translocated oncogene (BCR-abl), angiogenic factors and their receptors, etc. Table 4 highlights some of the studies showing RNAi efficacy in targeting cancer-specific genes (283). This field is expanding dramatically. RNAi has been used to unravel the function of genes in both in vitro and in vivo studies. Its specificity and potency are major assets that encourage its use for gene therapy and the development of efficient delivery methods. We expect that RNAi for gene therapy will enter clinical trials in the near future.

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Table 4 Cancer-associated genes targeted by RNAi Pathway

Target gene

References

Apoptosis

Bax

(363)

Bcl-2

(364, 365)

Angiogenesis

Focal adhesion kinase (FAK)

(366)

Adhesion

Matrix metalloproteinase

(367)

Cell–cell communication

VEGF

(365, 368)

Lipid metabolism

Fatty acid synthase

(369)

Transport

MDR

(370)

Signaling

H-Ras

(365)

K-Ras

(371)

PLK-1

(372)

TGF-B

(365)

STAT3

(373)

EGFR

(374, 375)

PKC-α

(365)

Viral

Epstein-Barr virus, HPV E6

(376, 377)

Oncogenes

BCR-Abl

(378, 379)

Nuclear

Telomerase

(380)

4. Strategies Altering the Host Response 4.1. Antiangiogenic Cancer Therapy

Antiangiogenic therapy by use of antibodies and gene therapy is one of the most promising strategies to inhibit the growth and the dissemination of tumors. This approach is based on negative regulators of neovascularization that suppress pro-angiogenic signals such as vascular endothelial growth factor (VEGF) or increase the expression of angiogenic inhibitors such as angiostatins (285), endostatins (286), and pigment epithelial-derived factor (PEDF) (287). Inhibition of the VEGF pathway can be achieved by blocking the receptor VEGFR2 (VEGFR2 in humans; Flk1 in mice), which is a tyrosine kinase receptor. Several approaches have been


33

investigated to block the VEGF signaling pathway, including expression of dominant-negative receptor mutants (288–290), a monoclonal antibody (bevacizumab) (91) and a tyrosine kinase inhibitor (vatalanib) (292). Phase III clinical trials using bevacizumab demonstrated that the efficacy of angiogenesis inhibition in metastatic colorectal cancer was associated with an increase in survival time when combined with a chemotherapy regimen (293, 294). The FDA has recently approved bevacizumab administered in combination with FOLFOX4, a regimen of 5-FU, leucovorin, and oxaliplatin for second-line treatment of metastatic carcinoma of the colon or rectum (295). The bevacizumab antibody was also approved in combination with carboplatin and paclitaxel as a first-line treatment of advanced/metastatic recurrent nonsquamous non-small cell lung cancer (296). The efficacy of angiogenic inhibitors was also successfully tested on animal models (286, 297). A recent study evaluated the use of adenoviral-mediated delivery of a mutated P125A endostatin in a mouse model of ovarian cancer (298). Inhibition of angiogenesis was clearly demonstrated. A subsequent study reported the impressive efficacy of P125A endostatin delivery combined with carboplatin (299). More recently, the combination of recombinant adeno-associated virus (rAAV)-mediated endostatin and 3TSR, the antiangiogenic domain of thrombospondin-1, showed a significant effect on the inhibition of angiogenesis and then on tumor growth in a mouse model of pancreatic cancer (300). Preclinical studies using inhibitors of angiogenesis are promising and clinical trials are now conceivable. Finally, the antiangiogenic properties of the immunomodulatory cytokines such as IL4, IL12, and interferon (IFN)-α and -β have also been investigated (301–303). This particular strategy will be discussed in Subheading 4.2. 4.2. Immunotherapy

Although tumoricidal approaches attempt to destroy cancer cells directly, immunogenic approaches work indirectly to illicit an immune response to the tumor cells through vaccination or modification of effector T cells (for reviews, see Xue and Stauss (304), Sobol (305), and Thomas et al. (306)). The development of preventive rather than curative vaccines seems to be the most promising immunotherapeutic approach.

4.2.1. Active Immunotherapy

Cancer vaccination consists of the administration of tumor antigen(s) either by inactivated tumor cells or tumor-specific antigens (TSAs), which are presented to CD8+ (cytotoxic T lymphocytes [CTLs]) or CD4+ T helper cells by antigen-presenting cells (APCs). If available, TSAs are used for vaccination. A second, less desirable option is to use tumor-associated antigens (TAAs), because these T cellrecognized tumor antigens can also be present at low levels in normal cells.

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Success in animal models has been achieved using adenoviruses delivering TAAs. For example, mice with breast cancer were shown to respond well to treatment of adenoviral vectors with TAAs (307). However, initial clinical trials did not have much success due, in part, to natural adenoviral-neutralizing antibodies in humans (308). In addition, certain tolerance mechanisms inactivate CTLs that are able to recognize the “self” antigen with high avidity. Consequently, the remaining CTLs with low avidity are also less sensitive to tumors expressing the same TAA and thus present a weak protection against tumors. In contrast to TAA vaccination, tumor-specific antigens generate a strong immune response that is harmless to normal tissue. The most striking illustration of this approach is the FDA-approved human papilloma virus (HPV) vaccine (309, 310), which is directed to the primary cause of cervical cancer (311). Besides viral protein, tumor cells also express specific proteins that result from mutations. Although these products are in theory the most appropriate to induce a strong CTL response, they are often invisible to CTLs because of antigen-presenting MHC class I mechanisms. 4.2.2. Adoptive Immunotherapy

In the adoptive immunotherapy approach, effector T cells are engineered ex vivo and then adoptively transferred to the patient. The efficacy of autologous tumor-infiltrating lymphocyte (TIL)-based treatment was demonstrated in patients with refractory metastatic melanomas (312). Under this procedure, T cells are expanded ex vivo under IL-2 stimulation and then are transferred to the patient after lymphodepleting chemotherapy (312). Another example of the clinical success of this strategy is the transfer of allogeneic T cells in immunosuppressed patients with leukemia (for an extensive review, see Xue and Stauss (304)). However, the allogeneic T cells are highly reactive and can be extremely toxic to normal tissue because of graft versus host complications. The transduction of a T cell receptor, which solely determines the specificity of the T cell tumor, into the host T cell is a very elegant method developed to treat the severe problems related to allogeneic T cell toxicity (for review, see Thomas et al. (306)). Recently, Morgan and colleagues reported the ability to specifically confer tumor recognition by autologous lymphocytes using a retrovirus encoding an anti-MART1 T cell receptor (313). Of the 15 patients enrolled in this study, two showed complete tumor regression. Although the response rate was low, this approach could be used with patients for whom other strategies would not be available.

4.2.3. Protection of Normal Cells from Chemotherapy

The expression of drug resistance genes is a common survival response of malignant cells to cancer chemotherapy. Certain types of cells such as hematopoietic stem cells (HSCs) are highly


35

chemosensitive and express these genes at a low level or not at all. The use of drug resistance genes to protect HSCs and to give them a selectable advantage is of particular interest to individuals undergoing chemotherapy. Numerous drug resistance genes have been investigated for gene therapy of malignant hematopoietic systems (for an extensive review, see Gillet et al. (314)). Examples of genes mediating single-agent resistance include: methotrexate resistance caused by mutant dihydrofolate reductase (DHFR) (315), alkylating-agent resistance mediated by methyl-guanine methyl transferase (MGMT) (316), and also glutathione transferase (GST Yc) (317), and aldehyde dehydrogenase (ALDH) (318), which confers resistance to cyclophosphamide, etc. (for review, see Zaboikin et al. (319)). Multidrug resistance is mediated by ATP-binding cassette transporters such as ABCB1/MDR1 (320), ABCC1/MRP1 (321), and ABCG2/BCRP (322) that recognize and remove from the cell a broad range of structurally unrelated compounds via an ATP-dependent transport process (for reviews, see Szakacs et al. (323) and Gillet et al. (324)). Early clinical trials transferring the ABCB1 gene to hematopoietic progenitor cells have been conducted (325, 326). Bone marrow or peripheral blood progenitor cells from patients suffering from advanced neoplastic diseases were retrovirally transduced and reinfused after high-dose chemotherapy (327–329). Although these studies confirmed that the human multidrug resistance gene can serve as a drug-selectable marker gene in vivo in the hematopoietic system, they also revealed that transduction efficiencies using ABCB1 vectors as detected in the bone marrow or peripheral blood of patients tended to be low, and varied from one patient to another. These data suggested that gene transfer procedures and selection strategies need to be improved to efficiently and safely protect human hematopoietic cells from the cytotoxicity of drug treatment.

5. Conclusions In this chapter, we introduced two major recurring themes in gene therapy: (1) the development of vector and delivery systems, and (2) the design of strategies to fight or cure a particular disease. We have previously noted that the field of cancer gene therapy experienced an “awkward adolescence” (330). Although the field has certainly not yet reached maturity, it still holds the potential of alleviating the suffering of many individuals with cancer. In many ways, scientists reflect society. The highly optimistic individual strongly believes that gene therapy will be able to cure

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targeted diseases. For the pragmatic individual, it is more realistic to envision gene therapy as one weapon in a multimodality treatment strategy that could help to control a disease, rather than be a magic bullet that will cure cancer or other diseases. Nevertheless, gene therapy has been and will continue to be a gold mine for science. Research in this field has not only contributed to a better understanding of molecular biological pathways but has also provided an abundance of elegant and challenging new strategies to struggle against disease.

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Subsection B Vectorology for Cancer Gene Therapy

Chapter 2 Designing Adenoviral Vectors for Tumor-Specific Targeting Ramon Alemany Summary Adenovirus provides an attractive candidate tool to destroy tumor cells. However, to fulfill the expectations, selective targeting of tumor cells is mandatory. This chapter reviews critical aspects in the design of tumor-targeted adenovirus vectors and oncolytic adenoviruses. The review focuses on genetic modifications of capsid and regulatory genes that can enhance the therapeutic index of these agents after systemic administration. Selectivity will be considered at different levels: biodistribution selectivity of the injected virus particles, transductional selectivity defined as cell receptor interactions and trafficking that lead to virus gene expression, transcriptional selectivity by means of tumor-selective promoters, and mutationrescue selectivity to achieve selective replication. Proper assays to analyze selectivity at these different levels are discussed. Finally, mutations and transgenes that can enhance the potency and efficacy of tumor-targeted adenoviruses from virocentric or immunocentric points of view will be presented. Key words: Adenovirus, oncolytic, targeting, tumor, vector.

1. Introduction Adenoviruses are non-enveloped viruses with a linear doublestranded DNA genome of 36 kb. The capsid is an icosahedron formed mainly by trimers of the hexon protein. At the vertices, five units of the penton base polypeptide hold a protruding trimer of fiber polypeptides. The distal knob of the fiber binds to the cellular receptors for virus attachment. When present, an arginine– glycine–aspartic acid (RGD) motif on the penton base binds to cellular integrins to promote entry via clathrin-mediated endocytosis (for a review on entry steps see (1)). Endosomal acidification triggers a membrane-lytic activity of capsid protein VI, and the particle at the cytoplasm binds dynein to reach the nucleus pore. Only the virus DNA is imported into the nucleus, where Wolfgang Walther and Ulrike S. Stein (eds.), Methods in Molecular Biology, Gene Therapy of Cancer, vol. 542 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-561-9_2

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transcription of early genes starts. Certain early gene products derived from E1 and E4 genomic regions control the cell cycle and prepare the cell for virus replication. Early 2 genes encode the proteins for virus DNA replication, and Early 3 genes mostly produce proteins that counteract the immune response against the virus. With the onset of replication, the major late promoter switches on and produces a long transcript that is spliced in the different RNAs of the capsid proteins. The capsid is assembled in the nucleus and a packaging signal sequence at the left end of the newly formed DNA is recognized for packaging. The release of the progeny of viruses from the cell through autophagy is quite inefficient (2, 3). Adenovirus vectorology applied to cancer gene therapy has advanced for more than 2 decades. During the first decade, up to the late 1990s, major efforts in adenovectorology were not addressed toward cancer gene therapy but instead toward the application of adenovectors to gene therapy of diseases such as cystic fibrosis or hemophilia. From first generation to second generation and gutless vectors, a stepwise removal of virus genes from the vector was achieved (4). However, the requirements for cancer gene therapy are very different from the requirements for other genetic diseases. Cancer is a systemic disease and, to reach metastases, the vector should be administered systemically. With the exception of immunotherapy approaches, a therapeutic intention requires reaching most tumor cells. Finally, the transduction event may be transient because the outcome is the destruction of the transduced cell. Tumor targeting at the level of cellular receptors has become one of the major goals of current adenovectorology. A good level of receptor selectivity would assure no toxicity and high efficacy and it would render secondary any other type of selectivity engineering. Unfortunately, changing the virus particle biodistribution and receptor interactions has proven very elusive, mostly because the basic knowledge from virologists does not apply to such an artifact as a bloodborne adenovirus. On the other hand, selectivity at the level of virus gene expression and replication has been attained much more successfully. The need to reach more tumor cells in a tumor mass prompted the use of replication-competent vectors contrary to the trend in noncancer gene therapy (5). Thus, the replicative vector has taken a preeminent role over the therapeutic payload gene of gene therapy. The replicating virus itself is the payload now that destroys the tumor cell and the exogenous transgenes have become secondary arms that help the virus in this goal. The adenovectors for cancer gene therapy are now designed with an oncolytic goal in mind, closer to the old concept of virus therapy of cancer than to the modern concept of gene therapy. Most of the vector development has been based on adenovirus serotype 5 and, unless otherwise specified, this chapter deals

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with this human serotype. A great deal of work has also focused on nongenetic modifications of the vector. Such transient modifications that do not pass to the vector progeny are described elsewhere (6).

2. Biodistribution Selectivity Selective accumulation of the injected dose at the tumor sites would be ideal for tumor targeting. Such a selective biodistribution of the virus particle must be distinguished from selective transduction. Biodistribution relates to the fate of the virus particles independent from gene expression. Despite this, reporter-gene expression has been often taken as synonymous with biodistribution. The biodistribution determines which cell targets are susceptible to transduction, but, on the contrary, transduction does not determine the tissue or organ biodistribution. In other words, certain modifications of the capsid that affect interaction with receptors or blood factors may affect the transduction of the virus but may not change its biodistribution. Biodistribution of adenovirus particles after intravenous administration can be measured by quantitative polymerase chain reaction (PCR) of the virus genome or labeling the capsid with fluorescent tags or isotopes that can be imaged. This kind of analysis reveals that adenovirus is enriched quickly in the liver and the spleen of mice. From the spleen, adenovirus escapes promptly and eventually most of the injected dose is collected in the liver. This biodistribution is a consequence of the low level of interaction of adenovirus with endothelial cells and the larger size of endothelial fenestrations in the spleen and the liver compared with other organs (Fig. 1); a size large enough to allow the extravasation of the 100-nmdiameter adenovirus capsid. The biodistribution, and in particular the adenovector hepatotropism, may be species specific. Studies in rabbits have shown that the size of the sinusoidal fenestrations determines the biodistribution (7). A size of sinusoidal fenestrations of 120 nm allows liver extravasation, in contrast to a pore size of 105 nm. The consequence of this biodistribution is a rapid decrease of the viremia, with a mean time of 2 minutes in mouse and 15 minutes in humans. Why does the virus accumulate in the liver but not in the spleen? One factor could be that the flow through the interstitial fluid in the liver is much slower than the flow through the spleen. The virus accumulates and remains floating in the space of Disse between the endothelial cells and the hepatocytes (8, 9). In addition to this passive uptake, there is an active uptake mediated by cellular interactions that seems to be more efficient in the liver than in the spleen. This active or

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CAR Hepatocyte NFkB

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Fig. 1. Emerging picture of cells and receptors that affect the biodistribution of adenovirus in the bloodstream. In the bloodstream, adenovirus is decorated with blood factors that bind to fiber knob sites overlapping with Coxsackie and adenovirus receptor (CAR)-binding sites and provide a bridge for virus binding to heparan sulfate proteoglycans (HSPG, black triangle) (see note added to proofs). These interactions then become responsible for biodistribution and cellular uptake instead of CAR. Furthermore, CAR is barely accessible because it is a regulatory component of the tight junctions of polarized epithelial cells. In liver sinusoids, the virus is mainly taken up by Kupffer cells through HSPGs, scavenger receptors, and other unknown interactions. In response, Kupffer cells produce primarily tumor necrosis factor (TNF) to attract neutrophils that adsorb the adenovirus capsid via complement receptor 1 (CR1) and Toll-like receptors (TLR). When antibodies against adenovirus are present, virus can also be opsonized using immunoglobulin receptors (FcgR) on monocytes and neutrophils. Other blood cells such as erythrocytes and platelets also bind to adenovirus. Platelets with virus (binding via von Willebrand factor [vWF]) are phagocytosed by Kupffer cells. Pro-inflammatory chemokines such as interleukin (IL)-6 and TNF can induce the expression of integrins and contribute to adenovirus uptake by other cells such as endothelial cells. Extravasated virus also accumulates in the space of Disse and binds to hepatocytes. As a result, little virus is available to reach tumors. Initial entry in tumors is also mediated mostly by HSPG. After replication, the intratumoral amount of virus increases and overcomes the lower amount of blood factors present. The depolarized epithelial tumor cells expose CAR and integrins, and therefore classic CAR-mediated entry can occur. Overexpressed fiber opens tight junctions to facilitate the spread of progeny virus.


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receptor-mediated capture may lead to transduction events, such as in the case of hepatocytes, or not, such as in the case of macrophages (Kupffer cells in the liver). The elimination of Kupffer cells or macrophages, in general, to some extent decreases the virus uptake by liver and increases the persistence of the virus in blood. However, even without macrophage uptake, the virus resides mainly in the liver and is cleared very rapidly from blood. How can this biodistribution be avoided? One possibility is to inhibit the flexibility of the fiber. The fiber must bend to allow the capsid to pass through the liver and spleen fenestrations. Another hypothetical possibility is the enlargement of the capsid size, using long protruding peptide chains. As in any preclinical model of cancer, it is always important to bear in mind that animal models may differ considerably from each other and from human disease. This is particularly worrisome for biodistribution of adenovectors. There are fundamental differences in the toxicology of Ad5 between rats, mice, pigs, and rabbits following intravenous administration (10–12). In mice, pigs, and rabbits, but not in rats, 10e12 viral particles (vp)/kg is well tolerated. Nonhuman primates seem as resistant as mice. This may be explained by Ad5 erythrocyte binding in rats but not in rabbit or mouse (12). Importantly, human erythrocytes show erythrocyte hemagglutination with Ad5 similar to rats. The interaction of Ad5 with human erythrocytes can greatly diminish the bioavailability of the injected vector dose (13). These erythrocyte interactions seem to be Coxsackie and adenovirus receptor (CAR) dependent, and they could be avoided with CAR-ablating mutations (14). Although the extrapolation from mice to humans must be taken with caution, the hepatotropism found in mice agrees with the transaminitis found after intravenous injection of adenovectors in humans or the hepatitis found after dissemination of adenovirus infections (15). The few data on biodistribution of adenovirus in humans from necropsies after virus administration also point out the tropisms toward liver and spleen (16). Accordingly, in humans, the liver is also the main target of transgene expression after systemic administration (17). In addition to this interaction with erythrocytes, other interactions with blood cells can affect the biodistribution. Among different serotypes, this seems more relevant for serotypes 4 and 11, in which binding of up to 5% of the injected dose to platelets has been detected. For serotype 5, this does not seem to be a significant fate (0.1% of the injected dose). The fate of the platelet-bound virus is also the liver Kupffer cells (18). The capsid proteins responsible for interaction with platelets seem to be the fiber and the penton base. After the identification of the specific binding sites, this problem may be addressed using a treatment of antibodies against platelets before the administration of the vector. This should increase tumor transduction and decrease liver transduction.

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The interaction of the adenovirus capsid with blood factors also affects biodistribution (19). The interaction of the fiber with mainly factor X, but also factor IX and C4BP, allows the virus to dock at cell surface heparin sulfates (see note added to proofs). In vivo, many cells, such as hepatocytes, macrophages (Kupffer cells in the liver), and likely tumor cells (our unpublished data), capture the vector mostly through these indirect interactions. Binding to complement C3b and to antibodies against the capsid can also affect the biodistribution because this binding promotes the opsonization by neutrophils (20). The capsid sites involved in these interactions are being mapped to improve the tumor-selective biodistribution and transduction of adenovectors. So far, these interactions have been partially ablated using a combination of three knob mutations: one extending the HI-loop with 12 amino acids after residue 547 using an heterologous mock-binding peptide such as SKCDCRGECFCD, a second that deletes FG loop residues TAYT (489–492), and a third that changes tyrosine 477 to alanine (21).

3. Transductional Selectivity To design tumor-targeting modifications in the vector, it is convenient to separate the two steps of gene transfer and expression that are fused into the broad concept of transduction. Gene transfer from the cell surface to the nucleus is highly dependent on the capsid composition of the vector. Gene expression depends on the activity of the promoter. Therefore, a more restricted use of the term transduction may be adopted that refers only to the first step, that is, to the transfer of DNA in a way that can lead to gene expression. This definition would include the steps of cell binding, entry, and intracellular trafficking that allows the virus DNA to reach the nucleus and initiate gene expression. However, it would not include the transcription of the vector genes, a step that may be modulated through the use of tumor-selective promoters (transcriptional targeting). Although not straightforward, this restricted definition of transduction is useful to separate tumor-targeting strategies based on cellular interactions from those based on promoter activation. This is important because gene expression is also used to study the transductional selectivity. For this, a vector with a reporter gene driven by promoter that can yield ubiquitous gene expression should be used. Usually a vector with luciferase under the cytomegalovirus (CMV) promoter has been used. The use of tumor-selective promoters would render an estimation of a transductional selectivity that is the result of the sum of transduction and transcription selectivity and it would not allow us to study separately these two events


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unless the experiment contains a nontranscriptionally selective vector control with the same capsid composition. On the other hand, transduction requires cell entry but entry and transduction are different endpoints. For example, the adenovector enters macrophages with a degradation fate not leading to transduction. The biodistribution of the vector particle determines the cells susceptible to transduction. The factors that determine the transduction of these target cells in vivo are becoming clear. The initial attempt to avoid liver transduction by eliminating CAR and integrin interactions failed. As mentioned above, in vivo blood factors bind to adenovirus capsid and promote interactions with different receptors. Using the triple mutation that affects resides Y477 and TAYT 489/492 and that extends the HI-loop with the RGE mock-binding peptide (named YT-RGE), Shayakmetov et al. reduced the virus load in the liver 50-fold, and, at a transductional level, this detargeting is around 100-fold (our unpublished results) (21). This triple mutation however does not preclude completely the transduction of the liver, and a residual interaction with blood factors that promotes hepatocyte infection remains. This can be deduced through in vivo experiments using warfarin, an inhibitor of vitamin K-dependent coagulation factors (FII, FVII, FIX, FX, and protein C), where the level of transduction detargeting is even more dramatic (300-fold, and our unpublished results). In the future, a detailed study of the capsid binding sites to these factors may allow the long awaited liver-detargeted adenovector (22). Considering the knob as a principal binding site for these factors, a more radical approach would be to get rid of the knob (see note added to proofs). The knob is necessary for the fiber trimerization that ensures its insertion at the penton base. Therefore, this approach requires a trimerization domain that substitutes the missing knob. This domain has been taken from the Moloney murine leukemia virus envelope glycoprotein, the T4 fibritin protein, the neck region peptide of human lung surfactant protein D and the sigma 1 reovirus protein (23–26). It is expected that deknobbed adenovectors will detarget the liver more efficiently than the triple YT-RGE vectors but this needs to be confirmed. The inefficient translation of deknobbed fiber messenger RNA (mRNA) into protein may also frustrate this approach (27). As long as liver detargeting is achieved, the opportunity for tumor targeting arises. In vitro, genetic tumor targeting has been achieved using peptides and serotype switching (28). The insertion of tumor-targeting peptides is limited by the low affinity of these ligands and by structural constraints imposed by the fiber that preclude the viable insertion or presentation of many peptides. The RGD motif is exceptional and it may allow the targeting of many integrin-overexpressing tumors. Another fiber modification that has proven very useful to target tumor cells in vitro is the use of the serotype 35 knob. This knob binds to

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CD46, a receptor essential for tumor cell survival and upregulated in tumor cells. The knob replacement would be the most versatile solution for liver detargeting and tumor targeting if complex ligands such as single-chain antibodies could be incorporated. The recent use of stable single-chain antibodies (scFv) in an optimized deknobbed vector is encouraging (29). An alternative to substituting the knob with tumor-targeting ligands would be to use deknobbed adenoviruses with tumor-targeting ligands in other exposed capsid proteins. The insertion and presentation of large targeting ligands such as single-chain antibodies into protein IX is a remarkable step forward (30). The hypervariable loops of the adenovirus hexon, on the other hand, can only accommodate small targeting peptides such as RGD (31). However, these approaches may not be compatible with the highly orchestrated entry and trafficking of the capsid from the membrane to the nucleus. The fiber falls off soon after entry to release the capsid into the endosome. In contrast neither protein IX nor hexon release from the capsid before reaching the nucleus. Therefore, the receptor-targeted capsid via pIX or hexon sticks to the receptor on the endosome membrane, and efficient trafficking is only possible when low-affinity ligands are used (32).

4. Transcriptional Selectivity Once the adenovector has reached the nucleus of the transduced cell, gene expression can proceed. As mentioned, it is convenient to separate the concept of transcription from the concept of transduction. As we have seen, the selectivity at the level of vector biodistribution and subsequent entry into cells that can express the vector’s genes need to be determined in vivo. The genetic modifications intended to modify these steps affect the capsid composition of the vector. However, transcriptional selectivity is independent from capsid composition and depends on regulatory DNA sequences. Hence, there are two fundamental differences in the experimental approach to tackle scientifically transductional or transcriptional tumor targeting. Because transductional targeting affects the capsid, it can be applied to any kind of vector, replicative or not. In other words, results can be extrapolated from a luciferase vector to an oncolytic adenovirus, for example, as long as they have the same capsid composition. Transcriptional targeting results cannot be extrapolated from one vector to another with a different virus DNA content. The second difference is that in vitro transcriptional targeting results are much more relevant than in vitro transductional targeting results.


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This second advantage led to the situation that, at the level of gene expression, the tumor targeting achieved so far is remarkable. Many different promoters have been used in nonreplicative adenovectors (and see Chapter “Expression Targeting for Tumor Specificity”) (33). As a general consideration, the use of promoters active in a broad range of tumor types would be ideal to apply one vector to different tumor types. Promoters such as hTERT, E2F1, or HIF are very attractive in this regard. In replication-defective adenovectors, one can afford the insertion of long regulatory DNA sequences, assuring a tighter regulation and a lower degree of interference from virus DNA. The current trend toward replication-competent adenovectors imposes new challenges. The size of the promoter to be inserted is a major limitation, considering a size limit of 38 kb for the final vector. With this caveat, artificially designed modular promoters may offer an advantage. To diminish the effect of viral enhancers, tumor-specific promoters regulated by transcriptional activators and repressors are preferred. In addition, insulators have been inserted next to the promoters to block those enhancers. The presence of cryptic transcription initiation sites, for example, present in the inverted terminal repeats (ITRs), can be blocked using polyadenylation signals upstream of the tumor-selective promoter. Another strategy used to eliminate the interference form the E1a enhancers that overlap with the left-end packaging signal has been the relocation of the packaging signal to the right end of the genome. From the point of view of the gene or genes to be controlled, E1a is a key gene because it is the first gene to be expressed and controls the rest of the early virus genes. It is important to bear in mind that replacing the E3 promoter can interfere with control of E2, and insertion of promoters in E4 can reduce replication in permissive cells (34, 35). Double regulation of two early transcription units improves the tumor selectivity of replication. However, direct or inverted repeats should be avoided to maintain the stability of the genome (35). For example, viruses such as CV716, 733, and 740 with E1a and E1b under the same promoter are unstable (36). Alternatively, an internal ribosome entry site (IRES) has been used to link E1a and E1b to obtain a remarkable selectivity index (105-fold better replication in tumor permissive cells than in nonpermissive cells). The use of two different promoters for the same goal has the limitation that it restricts replication to tumor cells that coexpress both promoters. The control of E4 has also been considered alone or in combination with E1. However, E4 modifications have been associated with lower genome stability. In summary, taking into account stability issues and size limitations, currently, the preferred design for transcriptional targeting is an insulated modular short synthetic promoter that controls an E1a-IRES-E1b unit.

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5. MutationRescue Selectivity Adenovirus expresses several proteins that modulate the cell cycle, protein translation, and cell death of the infected cells to promote its replication. These proteins have functional similarities with cellular proteins that are commonly altered in tumor cells. A mutation or deletion of such virus proteins can be rescued in tumor cells. This functional complementation has been used to design conditionally replicating adenoviruses with oncolytic properties. The first deletion used with this goal in mind was the deletion of E1b-55K (37). Adenovirus blocks early apoptosis of infected cells using E1B-55K in concert with E4orf6 to degrade p53 via ubiquitin-mediated proteolysis. An adenovirus without E1b-55k depends on the absence of cellular p53 activity, a trait found in 50% of tumors. However, E1b-55K has other functions and the oncolytic application of such a mutant (Onyx-015) has revealed that those other functions are important to maintain the replication potency in tumor cells. For example, during the late phase, E1B-55K forms a different complex with E4orf6 to shut off host mRNA nuclear export. E1b-55K also facilitates the nuclear localization of transcription factor YB1 that activates the E2 late promoter (38). Many tumors cannot complement efficiently the defect in RNA transport and the nuclear translocation of YB1. A better design for p53-defective tumors requires the dissection of p53 binding from these functions. The interaction of E1a with pRB has also been used to design tumor-selective replication-competent adenovectors. Binding of E1a proteins to pRB results in the release of E2F from the E2F–pRb complexes. Free E2F activates the transcription of cell cycle progression genes. Given the universal activation of the pRB pathway in tumors, the deletion of such a virus function can be complemented in all tumor cells. Two conserved regions in E1a, known as CR1 and CR2, bind pRB. E1a also binds p300 through CR1 and amino-terminal residues to stimulate E2F transcriptional activity. Taking into account these interactions, different mutations have been used to achieve tumor-selective replication. CR2 mutants preserve the oncolytic potency in tumor cells and are attenuated in arrested normal cells (39, 40). In contrast, the sole mutation of CR1 renders adenovirus barely selective and attenuates replication in tumor cells. Accordingly, the CR1 mutation may increase the selectivity of certain CR2 mutants, but the preservation of the potency in tumor cells should be carefully studied (41). The most selective combination of mutations seems to be the amino terminal and CR2 double-deletion mutation, such as in the dl4-25/121-128 adenovirus. In combination with these E1a mutations, the deletion of E4-orf6/7 has been proposed


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to increase tumor-selective replication because this virus protein also binds pRB. Current data, however, does not favor the use of this deletion (35). Adenovirus inhibits apoptosis at the early phase of adenovirus infection. The main virus protein that mediates this inhibition is the bcl-2 homolog, E1b-19K. Apoptosis resistance is a common trait in cancer and therefore E1B-19Kd deletion mutants can be rescued in cancer cells (42). The phenotype of E1b-19K mutants is cellular toxicity (cyt phenotype) and degradation of the cellular genome (deg phenotype) with the typical ladder pattern associated with apoptosis. This cyt and deg phenotype results in a great reduction of virus yield. In tumor cell lines in which apoptosis is inhibited, E1b-19K mutants grow to wild-type levels; even an oncolytic potency increase compared with adenoviruses has been described with E1b-19K. This oncolytic potency increase has been associated with an earlier release of virions from the infected cells causing an accelerated cell-to-cell spread (43). Protein translation is also enhanced by adenovirus and in tumor cells in a way that mutant adenoviruses unable to promote protein translation can be rescued in tumor cells. Adenovirus encodes virus-associated (VA)-I, a VA-II RNA at the late phase that acts as a competitive inhibitor of the activation of protein kinase R (PKR) by other double-stranded RNAs produced during infection. PKR is a protein kinase induced by interferon (IFN) that phosphorylates the eIF-2-alpha translation factor and leads to protein synthesis shutoff in infected cells (44). VA-RNAs counteract this IFN response to adenovirus, and VA-RNA mutants are blocked by IFN. However, tumor cells present a truncated IFN pathway (45). Hence, in tumor cells, the deletion of VA-RNAs can be rescued (46, 47). This same rational has been applied to design oncolytic herpes and influenza viruses (48, 49) and, in general terms, viruses sensitive to IFN have been used as oncolytic viruses. The tumor-selective replication of viruses unable to block the IFN pathway correlates with an enhanced initiation of protein translation found in tumor cells. Several pathways, including Ras and PI3K, contribute to protein translation in tumor cells and there is a lack of correlation between virus replication and each of these pathways considered separately.

6. Mutations for Efficacy Different viruses are under study as oncolytic agents (see Chapters 29 to 32). With regard to oncolytic potency, small RNA viruses that replicate in the cytoplasm have a clear advantage over large DNA viruses that replicate in the nucleus. A fast life

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cycle produces a rapid cytopathic effect in cultured monolayers or in tumor models. For example, as low as ten infectious units of an oncolytic picornavirus can spread in a few days through a 1-cm-diameter tumor. The spread of adenovirus is far from such a speed. Adenovectors replicate in the nucleus of infected cells, with a life cycle of approximately 40 h. A time course of synchronic wild-type adenovirus progeny production in 293 cells follows a sigmoid curve with a lag period of 6 h and a plateau at 36 h. The slope of this sigmoid curve at the time of maximal increase of production (50% of total virus production) gives an idea of the speed of replication. A comparison of this one-step growth curve between replication-defective adenovectors (E1-deleted or gutless), oncolytic adenoviruses, and wild-type adenovirus can reveal replication speed differences that otherwise become indistinguishable in a burst assay (total yield per infected cell at a late time point). The size of a plaque in a regular plaque assay can also reveal differences in the duration of the virus life cycle. Because the plaque is the consequence of multiple rounds of infection, in this assay, the differences are multiplied. Large plaques reveal the presence of adenoviruses that spread faster. The one-step growth curve and the plaque assay also differ in an important detail: to estimate the virus yield in a given time point of a grow curve, the researcher freezes and thaws the infected cells to “extract” the virus. During a plaque assay, the infected cell releases the virus naturally. A classic one-step growth curve using crude cell extracts is appropriate to measure the kinetics of the virus life cycle up to formation of mature virions in the nucleus. However, the most inefficient step in the adenovirus life cycle seems to be the release of virions. Therefore, to measure this release step, it is necessary to measure virus yields in the supernatant of the infected cells. This supernatant kinetics curve will take longer than the crude lysate curve to reach a plateau. In summary, time-course supernatant release assays and plaque-size assays can help us to predict the oncolytic potency of adenovirus mutants. Another appealing method to compare the oncolytic potency of different adenovectors is competition (50). Tumor cells are co-transfected with virus DNA or co-infected with viruses and the best growers take over after several passages. This method can also be used to obtain the most potent viruses of a pool of mutants. After initial major efforts toward obtaining selective oncolytic adenoviruses, it is now becoming increasingly important to enhance the potency of the adenovirus. Each step of the virus life cycle is amenable to optimization for this purpose. At the level of cell entry, the capsid can be modified to enhance the infectivity toward tumor cells. The RGD motif has been used in this regard (51). Note that the concepts of enhanced potency and tumor targeting may or may not overlap. One can design vectors that are very infectious in tumor cells as well as in normal cells and then


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design selectivity properties at a post-entry step. Because of the paucity of CAR in tumors, there is ample room to improve the potency of adenovectors by inserting ligands that bind to tumor cells (28). However, most of the mutations that have rendered fast-spreading viruses affect the release of virus from the cell. We have already mentioned that E1b-19K deletion mutants are more cytopathic than wild-type adenovirus in certain tumor cell lines. A premature apoptotic cell death may decrease the final virus yield (burst size) but it may also accelerate the release of virions into the supernatant. This accelerated release translates into an increased oncolytic potency. This suggests that faster release and not virus yield is a better parameter to predict oncolytic potency. A second mutation that has been used to accelerate the release of adenovirus is the overexpression of adenovirus death protein (ADP). ADP is the only E3 protein expressed from the major late promoter. It is an integral membrane protein localized in the Golgi apparatus, endoplasmic reticulum, and nuclear membrane. ADP contributes to the release of virions from infected cells and its overexpression enhances this release and the oncolytic activity of adenovirus (52). Conversely, the absence of ADP in E3-deleted replicating adenoviruses has been associated with a lower potency (53, 54). ADP overexpression has been achieved by either deleting other E3 open-reading frames or inserting a second copy of the major late promoter upstream of ADP. The mechanisms of cell killing by ADP are not yet elucidated. On one hand, ADP can block the action of E1b-19K thus mimicking the caspase-dependent pro-apoptotic phenotype of E1b-19K mutants. However, ADP also promotes caspase-independent apoptosis, and, contrary to E1b-19K, ADP is expressed late in infection (55). ADP also binds to MAD2B, a component of the spindle assembly checkpoint protein that counteracts adenovirusmediated lysis (56). Despite the uncertain mechanism, the combination of ADP overexpression with selective replication greatly increases the prospects of adenovirus therapy (57). Growth competition assays have revealed another important genomic region to take into account to increase oncolytic potency. When tumor-selective promoters are inserted upstream of E1a, the packaging elements V, VI, and VII should not be eliminated (50). These studies also indicated that the orientation of expression cassettes in the genome may affect the propagation of the virus and confirmed the importance of the E3 region. When repeated passaging in tumor cells was used to select the more potent adenoviruses after random mutagenesis, several interesting mutations were selected (58). For example, a mutation that truncates the c-terminus of the i-leader protein or mutations that truncate E4-ORF1 or E4-ORF2 were selected. How these mutations accelerate propagation remains to be studied, but the use of these mutations in oncolytic adenovectors is warranted.

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Random mutagenesis and serial propagation in plaque assays has also been applied to an E3-deleted, and therefore small plaque phenotype, adenovirus mutant (59). In this case, there is a strong pressure to select mutations that compensate for E3 loss. Accordingly, most selected “large plaque” (relative to the small plaque parental virus) mutants contained deletions affecting E1b-19K, suggesting the antagonistic function of ADP and this protein. In practical terms, this means that when E3 is deleted from the backbone of the oncolytic virus (for example, to insert transgenes), then the deletion of E1b-19K is highly recommended. Interestingly, a c-terminal i-leader truncating mutation was also found, and, in this case, its effect was additive to the presence of E3. A comparison of plaque size with the kinetics and yield of virus for each mutant also reveals that the spread phenotype does not correlate with virus replication. With regard to oncolytic potential, spread potency seems a better marker than virus yield.

7. Transgenes: Immunocentrics and Virocentrics

Another approach to increase the potency of an oncolytic adenovirus is to arm it with transgenes that favor the spread or cytotoxicity of the virus. There are two opposite strategies to gain antitumor activity. Those that do not believe that the virus can reach all disseminated tumor cells and replicate in them until no tumor cells are left are strong proponents of the idea of viruses as sparks of immune responses. In such a scenario, stimulation of the immune system is advantageous. It is only with the help of the immune system that a complete therapy may occur. This view has been defined as immunocentric. On the other hand, others see the immune system as a foe of the virus that neutralizes the therapeutic spread of the virus. This view has been defined as virocentric. There is experimental and clinical evidence in favor of each view. In preclinical studies, virotherapy in combination with immunostimulation has improved the oncolytic outcome for many different oncolytic viruses, although, for adenovirus, the species specificity has precluded a proper evaluation of this strategy. On the other hand, the use of immune inhibitors has also improved the outcome of preclinical virotherapy. In a clinical setting, immunocentric proponents can claim reports of occasional cures observed with subcutaneous tumor lysates. On the other hand, virocentric proponents can reference better responses in immunosuppressed patients or even complete cures in patients getting virus infections after bone-marrow transplantation. Immunocentric proponents have armed adenovirus with GM-CSF or interleukins. Virocentric proponents try to use the limited space to insert genes that increase virus spread, such as


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p53, prodrug-converting genes, fusogenic genes, or stroma-digesting genes. From the patient and environment standpoints, an immunocentric approach is safer than arming the virus with destruction weapons, particularly if enhancing potency mutations are also used. However, the virus capsid per se is a formidable inflammatory and danger signal and the addition of immune stimulation genes may become superfluous. Given the low predictive value of preclinical immunotherapy, conclusions are difficult to reach. Cancer immunotherapy is evolving continuously with drugs that are much simpler than a virus, and the competition for pharmaceutical development is going to be difficult. Because chemotherapy is often immunosuppressive, concomitant chemotherapy and virotherapy can be considered as virocentric. As in any kind of tumor therapy, because every tumor and patient is different, there will be cases in which immunostimulation may be much better than immune-suppression, or conversely. Certain parameters may be used to predict the best application of viruses, such as the amount of stroma in tumors, vascularization, location of metastases, immune status of the patient, immune characteristics of the tumors, etc. Finally, a general scheme can be used that exploits both strategies sequentially: initially the patient has less probability to be immune to the vector and transient immune-suppression or concomitant chemotherapy combined with a systemic oncolytic vector armed for maximal spread can offer the best therapeutic opportunity. After a few weeks, facing a lack of response, the virus now armed with immune-stimulatory genes can be used intratumorally, trying to elicit an immune response.

8. Conclusion The many years of high expectations laid on adenovirus as a vector or oncolytic agent have fostered a myriad of approaches to troubleshoot the selectivity and efficacy limitations encountered during this journey. Still, major obstacles remain. Nevertheless, the amount of information gathered on adenovirus entry, replication, and release allows for the current design of better adenovectors against cancer. This added value is a shared contribution of many research groups. However, the long history has a clear drawback: an optimized antitumoral adenovirus uses intellectual property from many sources. Without a clear patent position, no company will back such an optimal design. The bet is not small because the costs for good laboratory practices (GLP) preclinical studies and good manufacturing practices (GMP) clinical virus is approximately half a million US dollars for a first test in 21 patients. There is a risk then that such an optimization process becomes only an academic exercise. Without optimization at the

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level of selectivity and potency, the possibility of success is dim. With adenovirus, as with other viruses, virotherapy and gene therapy for cancer need to carefully approach clinical development to show more than merely lack of toxicity. Note added to proofs: FX binds to Ad5 hexon and has a major role in hepatotropism (Waddington et al. 2008. Cell 132: 397–409).

Acknowledgments I thank the collaborative effort of the Virus Therapy Group at the Institut Català d?Oncologia involved in the author’s results mentioned herein. Special thanks to Manel Cascallo and Juan Fueyo for their close collaboration. Thanks to Cristina Balague for critical reading of the manuscript. The author is supported by Bio2005-08682-C03-01 from the Ministerio de Ciencia y Tecnología of the Government of Spain, the EU 6th FP research contract 18700 (Theradpox, RA), and the Network of Cooperative Research on Cancer (C03-10), Instituto de Salud Carlos III of the Ministerio de Sanidad y Consumo, Government of Spain. References 1. Leopold PL, Crystal RG. (2007) Intracellular trafficking of adenovirus: many means to many ends. Adv Drug Deliv Rev 59:810–21. 2. Jiang H, Gomez-Manzano C, Aoki H, et al. (2007) Examination of the therapeutic potential of Delta-24-RGD in brain tumor stem cells: role of autophagic cell death. J Natl Cancer Inst 99:1410–4. 3. Ito H, Aoki H, Kuhnel F, et al. (2006) Autophagic cell death of malignant glioma cells induced by a conditionally replicating adenovirus. J Natl Cancer Inst 98:625–36. 4. Volpers C, Kochanek S. (2004) Adenoviral vectors for gene transfer and therapy. J Gene Med 6 Suppl 1:S164–71. 5. Alemany R. (2007) Cancer selective adenoviruses. Mol Aspects Med 28:42–58. 6. Kreppel F, Kochanek S. (2008) Modification of adenovirus gene transfer vectors with synthetic polymers: a scientific review and technical guide. Mol Ther 16:16–29. 7. Lievens J, Snoeys J, Vekemans K, et al. (2004) The size of sinusoidal fenestrae is a critical determinant of hepatocyte transduction after adenoviral gene transfer. Gene Ther 11:1523–31. 8. Bernt KM, Ni S, Gaggar A, Li ZY, Shayakhmetov DM, Lieber A. (2003) The effect

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Designing Adenoviral Vectors for Tumor-Specific Targeting 14. Nicol CG, Graham D, Miller WH, et al. (2004) Effect of adenovirus serotype 5 fiber and penton modifications on in vivo tropism in rats. Mol Ther 10:344–54. 15. Steiner I, Aebi C, Ridolfi Luthy A, Wagner B, Leibundgut K. (2008) Fatal adenovirus hepatitis during maintenance therapy for childhood acute lymphoblastic leukemia. Pediatr Blood Cancer 50(3):647–9. 16. Hamid O, Varterasian ML, Wadler S, et al. (2003) Phase II trial of intravenous CI-1042 in patients with metastatic colorectal cancer. J Clin Oncol 21:1498–504. 17. Connelly S. (1999) Adenoviral vectors for liver-directed gene therapy. Curr Opin Mol Ther 1:565–72. 18. Stone D, Liu Y, Shayakhmetov D, Li ZY, Ni S, Lieber A. (2007) Adenovirus-platelet interaction in blood causes virus sequestration to the reticuloendothelial system of the liver. J Virol 81:4866–71. 19. Baker AH, McVey JH, Waddington SN, Di Paolo NC, Shayakhmetov DM. (2007) The influence of blood on in vivo adenovirus bio-distribution and transduction. Mol Ther 15:1410–6. 20. Cotter MJ, Zaiss AK, Muruve DA. (2005) Neutrophils interact with adenovirus vectors via Fc receptors and complement receptor 1. J Virol 79:14622–31. 21. Shayakhmetov DM, Gaggar A, Ni S, Li ZY, Lieber A. (2005) Adenovirus binding to blood factors results in liver cell infection and hepatotoxicity. J Virol 79:7478–91. 22. Parker AL, Waddington SN, Nicol CG, et al. (2006) Multiple vitamin K-dependent coagulation zymogens promote adenovirusmediated gene delivery to hepatocytes. Blood 108:2554–61. 23. van Beusechem VW, van Rijswijk AL, van Es HH, Haisma HJ, Pinedo HM, Gerritsen WR. (2000) Recombinant adenovirus vectors with knobless fibers for targeted gene transfer. Gene Ther 7:1940–6. 24. Krasnykh V, Belousova N, Korokhov N, Mikheeva G, Curiel DT. (2001) Genetic targeting of an adenovirus vector via replacement of the fiber protein with the phage T4 fibritin. J Virol 75:4176–83. 25. Magnusson MK, Hong SS, Boulanger P, Lindholm L. (2001) Genetic retargeting of adenovirus: novel strategy employing “deknobbing” of the fiber. J Virol 75:7280–9. 26. Mercier GT, Campbell JA, Chappell JD, Stehle T, Dermody TS, Barry MA. (2004) A chimeric adenovirus vector encoding reovirus attachment protein sigma1 targets cells

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Chapter 3 Analysis of HSV Oncolytic Virotherapy in Organotypic Cultures Giulia Fulci and Brent Passer Summary Tumor-selective replication-competent viral vectors, such as oncolytic herpes simplex virus (HSV) type I (HSV-1), represent an attractive strategy for tumor-based therapies because these viruses can replicate and spread in situ exhibiting cytopathic effects through direct oncolytic activity. These lytic viruses offer a distinct advantage over other forms of cancer therapies in that they are self-perpetuating and can spread not only in the tumor itself, but also to distant micrometastases. Translational studies aimed at identifying novel virotherapies for human cancers are incumbent upon the appropriate experimental models. While animal models are the preferred choice for efficacy studies of HSV virotherapy, we have developed a novel complementary approach toward assessing the effectiveness of oncolytic HSV therapy in both brain and prostate cancers. This experimental model takes advantage of previously published work in which human prostate cancer biopsies and rodent brain slices can be easily maintained ex vivo. The advantage of these systems is that the three-dimensional structure remains intact. Thus, all of the factors that may affect viral entry and replication, such as cell–cell and cell–matrix interactions, and interstitial fluid within this three-dimensional milieu remain preserved. Moreover, with respect to the brain, this system offers the advantage of direct access to brain cells, such as microglia and astrocytes, and circumvents the problems associated with the presence of the blood–brain barrier. Key words: Brain tumors, HSV, oncolytic virotherapy, organotypic cultures, prostate cancer.

1. Introduction The discovery of tumor-selective replication-competent viral vectors, designated also as oncolytic viruses (OVs), represents an attractive strategy for cancer therapies. OVs can replicate in situ, spread their progeny throughout the neoplastic mass, and reach isolated migrating cancerous cells as well as distal metastases (1).


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Several virus strains are being used as cancer therapeutic OVs (1). Herpes simplex virus (HSV)-derived OVs (HSV-OVs) were first designated to treat brain tumors, but have been found to also be very efficient in other cancer types (2–4). The advantage of using HSV is that it can be engineered with large genetic inserts and its replication can be effectively inhibited with pharmaceuticals (1). Although OVs can kill tumor cells grown in vitro with high efficiency, they present reduced replication when applied in vivo, which results in a low therapeutic efficiency in clinical trials. This suggests that physiological aspects of the tumor and host decrease the OV’s therapeutic potential, and preclinical research aimed at understanding and overcoming these physiologic aspects is currently ongoing. Solid tumors are formed of both cancerous and noncancerous cells. The noncancerous cells include blood vessel endothelial cells, hematopoietic cells from circulating blood, and local and peripheral cells of the innate and adaptive immune system. Together, these cells form the tumor stroma, a complex and changing structure that can influence tumor formation, progression, and response to therapy. Moreover, the compact structure of solid cancers is hold together by a tight extracellular protein matrix that does not allow efficient spread of drugs, including OVs. Recent discoveries have demonstrated that disruption of the extracellular matrix and/or suppression of innate immunity from the cancer stroma can increase the efficacy of cancer virotherapy (5–18). In vivo models are the most reliable means by which to evaluate efficacy for experimental cancer treatments. However, in vivo studies can present a number of limitations in understanding the physiological mechanisms that influence the outcome of a specific therapy. Indeed, studies performed in vivo can be time-consuming, cost-ineffective, and present difficulties in drug delivery. Moreover, they may also require complex surgeries that limit the total number of animals analyzed, thus limiting statistical analyses and parallel comparisons. Thus, to understand how physiological aspects of tumor influence the outcome of cancer virotherapy, we have developed a novel complementary approach using prostate cancer biopsies and rodent brain slices cultured ex vivo (19–24). The advantage of these organ cultures is that all factors that may affect viral entry and replication, such as cell– cell and cell–matrix interactions, and interstitial fluid within an intact three-dimensional milieu remain preserved. Therefore, we have been able to evaluate the role of phaogocytic microglia in brain tumor virotherapy (14) and to analyze the efficacy and cancer selectivity of HSV-OV in human prostate tumors (Passer et al., unpublished).

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2. Materials 2.1. Preparation of Organotypic Brain Slices

1. Five to 7 days postnatal mice or rats. 2. Vibratome with magnifier lens (Leika VT100S, Bannockburn, IL) (see Note 1). 3. Dissecting microscope (Nikon SMZ 1000, Melville, NY) (see Note 1). 4. Surgery tools (see Note 2): feather blades (Ted Pella, Redding, CA), knife, small spatula (4 × 150 mm), large scissors, and small dissecting scissors (Fine Science Tools Inc., Foster City, CA), the finest brushes provided by any art supply store. 5. Pasteur pipettes. 6. Super Glue (Staples). 7. Gey’s balanced salt solution pH 7.2 (Gey’s BSS) (SigmaAldrich, St. Louis, MO) enriched with 36 mM D-glucose (Fisher Scientific, Pittsburgh, PA) and with a penicillin/streptomycin (P/S) solution (Invitrogen, Carlsbad, CA) diluted 1:1,000. 8. Millicell cell culture plate insert, hydrophilic polytetrafluoroethylene (PTFE) filter 0.4 μm (Millipore, Billerica, MA). 9. Culture medium (see Note 3): 50 mL minimum essential medium (MEM, Cellgro, Herndon VA), 25 mL heat-inactivated fetal bovine serum (FBS; see Note 4) (Hyclone, Logan, UT), 25 mL Hank’s balanced salt solution (Hank’s BSS, Cellgro), 0.5 mL glutamine (Cellgro), 100 µL P/S solution (Invitrogen), 650 mg D-glucose (Fisher Scientific).

2.2. Tumor Establishment in Organotypic Brain Slices

1. Trypsin-EDTA (Cellgro). 2. Cell culture medium: Dulbecco’s modified essential medium (DMEM, Cellgro) supplemented with 10% heat-inactivated FBS (Hyclone) and 1% P/S solution (Invitrogen). 3. Sterile phosphate buffer solution (PBS, Cellgro). 4. Cell counter. 5. 10-cm cell culture dishes. 6. Dissecting microscope (Nikon).

2.3. Prostate Organ Culture

1. Prostate organ culture complete media (POC-CM): RPMI1640 (Cellgro) supplemented with 10% heat-inactivated FBS (Hyclone), 1 mM sodium pyruvate (Cellgro), 1 mM nonessential amino acids (Invitrogen), 1 mM P/S/glutamine (Invitrogen), Ultrafoam (collagen sponge) (Davol Inc., Cranston, RI).

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2. Prostate biopsies (Massachusetts General Hospital [MGH] Pathology Department; Institutional Review Board [IRB] approved). 2.4. Oncolytic HSV Strains and Other Pharmaceuticals

1. Oncolytic HSV strain hrR3 (25): derived from HSV Kos; inactivation of the ICP6 gene by Escherichia coli LacZ gene insertion. Dr. Sandra Weller at the University of Connecticut provides the virus. 2. Oncolytic HSV strain G47 (26): derived from the HSV backbone F; ICP6 gene inactivation by E. coli LacZ gene insertion, deletion of both γ34.5 alleles and of the α47 gene. MediGene Inc. (San Diego, CA) provides working preparations of this virus. 3. Dr. Robert Lee at the Ohio State University provides clodronate liposomes (14).

2.5. Cryostat Sectioning of Prostate Tissue

1. OCT freezing media (Tissue-Tek, Torrance, CA). 2. 2-Methyl butane (Sigma-Aldrich). 3. Cryomolds (Tissue-Tek). 4. Dry ice.

2.6. Staining of Brain and Prostate Organ Cultures

1. Glass coverslips (round glass coverslips that fit in a 24-well cell culture plate). 2. 24-Well cell culture plates. 3. Protein block (DAKO-Cytomation, Glostrup, Denmark). 4. Antibody diluent solution (DAKO-Cytomation). 5. Phosphate buffer (PBS, Sigma-Aldrich) (see Note 5). 6. Primary antibodies (see Note 6): mouse anti-rat CD68, mouse anti-rat CD163, rat anti-mouse CD68, and mouse anti-HSV-gC (AbD Serotec, Oxford, UK), mouse antiHSV-ICP4 (US Biological, Swampscott, MA). 7. Secondary antibody (see Note 6): FITC-conjugated antimouse IgG (Jackson Immunoresearch, West Grove, PA). 8. Vectashield® medium containing 4´ ,6-diamidino-2-phenylindole (DAPI) (Vector Laboratories, Burlingame, CA). 9. Acetone, ethanol (EtOH), and xylene (these do not have to be extremely pure, the cheapest brand from your local pharmacy will be good). 10. Isolectin GS-IB4 from Griffonia simplicifolia, Alexa Fluor® 488 conjugate (Molecular Probes, Carlsbad, CA). 11. X-gal, potassium ferricyanide, potassium ferrocyanide, and 1 M MgCl2 solution (Sigma-Aldrich) (see Note 7). 12. Hematoxylin 2 (Richard Allan Scientific, Kalamazoo, MI).


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1. African green monkey kidney cells (Vero, American Type Culture Collection [ATCC], Manassas, VA). 2. DMEM tissue culture medium (Cellgro) supplemented with 10% heat-inactivated FBS (Hyclone) and 1% P/S solution (Invitrogen). 3. Sterile glucose-enriched PBS (Sigma-Aldrich). 4. Trypsin–EDTA (Cellgro). 5. 6-Well tissue culture plates. 6. Cell counter. 7. Human anti-HSV IgG (Gamunex, Talecris Biotherapeutics, Research Triangle Park, NC).

3. Methods 3.1. Preparation and Culture of Organotypic Brain Slices

1. Animals are killed and plunged into a 70% EtOH solution, made with Millipore distilled water, and immediately transferred to a prechilled Petri dish maintained on ice. 2. The animals are decapitated and the skull is removed by making two horizontal cuts with small dissecting scissors; the cuts should start on each side of the head at the level just above the ears. 3. The cranial nerves are cut from the cerebrum with a small spatula, which is also used to disrupt the dura mater and to ease the brain out of the cranium. 4. The entire cerebrum is immerged in prechilled Gey’s BSS in a sterile Petri dish. 5. The sagittal surface of the brain is glued with Super Glue to a chilled metal base in the chamber of a vibratome and the chamber is filled with cold Gey’s BSS. Gey’s BSS in the vibratome chamber should be changed after each animal. 6. Coronal sections at a thickness of 300 µm are cut from the posterior pole (remember to keep the blade and the block of tissue always moist), transferred to a separate 60-mm dish with ice-cold Gey’s BSS, and incubated at 4°C for up to 3 h. 7. The slices are then transferred to the surface of the Millicell culture plate insert by means of a Pasteur pipette filled with one drop of Gey’s BSS, which serves to keep the tissue moistened. 8. The Millicell culture plate insert is placed in a 24-well tissue culture dish on top of a layer of culture medium. The medium should not cover the membrane so that the explant remains well exposed to the air.

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9. After the brain slice is placed on the Millicell insert, the drop of Gey’s BSS is vacuum-aspirated off, and the slice is incubated at 36°C in a chamber with 5% CO2-enriched air. 10. Brain slice cultures are fed using sterile procedures under a laminar flow hood by changing half of the growing medium every 2–3 days. 11. Three weeks later, the organotypic brain culture has matured to the point at which experimental manipulations can be performed. 3.2. Tumor Establishment, Drug Treatment, and Virus Infection in Organotypic Brain Slices

1. 10 mL of warmed sterile PBS are added to the bottom part of a 10-cm culture dish. 2. Trypsinized brain tumor cells are resuspended in 10% FBS supplemented with the P/S solution at a final density of 100,000 cells/mL (see Note 8). 3. A drop of cell suspension is gently placed (~20 μL or ~2,000 cells) on the upper lid of a 10-cm cell culture dish. About 25 drops can be placed in one dish. 4. The bottom part of the dish, containing sterile PBS, is carefully covered with the upper lid where the cell drops are placed, and the hanging drops are allowed to grow for 3–4 days to allow the cells to aggregate in spheres (27). 5. Under a dissecting microscope, one sphere is removed (pipette setting: 1.2 μL) and placed onto the brain slice surface without damaging the slice. 6. The sphere will adhere to the brain slice through interaction with surrounding astrocytes and small tumor structures will be formed. Depending on the tumor model, this process may take only 8 h or as long as 1 week; this has to be established empirically with different cancer cells. 7. Phagocytic microglial cells can be depleted with the use of clodronate liposomes. One milliliter of liposome solution (0.2 mg/mL in feeding medium) is placed below the insert holding the brain slice and on top of the brain slice. The slice is incubated with the liposome solution for 24 h, after which it is rinsed twice and fresh medium is put under the slice for further growth. Treatment with liposomes is performed before tumor establishment to avoid the disruption of the tumor. The tumor growth is not altered by this treatment. 8. OV infection of the tumor is performed by placing a sterile plastic ring (cut from the top edge of a 200-μL pipette tip) on top of the tumor and by filling it with 40–50 μL of OV suspension in sterile PBS (see Note 9). The brain slice is then incubated for 3 h at 37°C, after which the virus is rinsed out with several changes of feeding medium.


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All procedures described below should be performed in a laminar flow hood. To maintain the structural integrity of the tissue, prostate biopsies must be manipulated immediately after receiving. 1. Human prostate cancer tissues are obtained from the MGH Pathology Department (IRB-approved) after radical prostatectomies. 2. Prostate tissues are placed into a 10-cm Petri dish containing RPMI media supplemented with a P/S/glutamine solution and cut into 2–4 mm3 fragments. 3. Tissue fragments are placed in a 48-well plate containing 250 μL/well of POC-CM and inoculated with 5 × 104–5 × 106 plaque-forming units (pfu) of G47Δ for 1–2 h at 37°C in a humidified incubator with 5% CO2-enriched air. 4. During inoculation, the collagen sponge is cut into a ~10 × 10-mm square under sterile conditions and placed into 2 mL of POCCM. Tumor fragments are then transferred to the semi-submersed sponge and incubated at 37°C in a 5% CO2-humidified incubator for up to 2 weeks. 5. At various time points after infection, tissues are removed and placed in OCT in a cryomold. Tissues are then allowed to slowly freeze in a 2-methyl butane bath prechilled in dry ice, and can either be stored at – 80°C or immediately cut with a cryostat in 5 μM sections.

3.4. Staining of Brain and Prostate Organ Cultures (13, 14)

All staining procedures are performed on tissues that were dried overnight, fixed in ice-cold acetone, and stored at – 20°C or –80°C (see Note 10).

3.4.1. Immunohistochemistry Staining of HSV and Microglia (see Notes 11 and 12)

The incubation times required for the brain slices are in bold; the other incubation times refer to the prostate slices. If only one time is indicated, it refers to both the prostate and brain slices. 1. The brain and prostate slices are rehydrated in 300 μL PBS for 30 min at room temperature (RT). 2. Endogenous protein background is blocked with 300 μL serum-free protein block for 10 min or 2 h at RT. 3. Sections are then incubated with the primary antibody appropriately diluted in 300 μL of antibody diluent solution overnight or for 48 h at 4°C. 4. After three washes in 500 μL PBS at RT for 30 min each, the slices are incubated with the secondary antibody for 45 min at RT or 24 h at 4°C. 5. After three washes in 500 μL PBS at RT for 30 min each, the slices are mounted with Vectashield® medium containing DAPI for nuclear counterstaining.

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3.4.2. Staining of Oncolytic HSV β-Galactosidase Activity

Because both hrR3 and G47 contain the E. coli LacZ gene, X-gal staining on tissue sections can be performed to evaluate viral spread. 1. Rehydrated tissues are covered with a freshly prepared X-gal solution containing: 0.5 mL of X-gal stock in DMSO, 0.05 mL of 1 M MgCl2, 1 mL of 125 mM potassium ferricyanide, 1 mL of 125 mM potassium ferrocyanide, and 1X PBS and 22.45 mLs of 1X PBS (total volume 25 mL). 2. Tissues are then incubated for 2–4 h (until neat blue HSV colonies appear) at 37°C in the dark, and excessive X-gal solution is rinsed off by submerging the tissues in clean PBS two to three times. 3. The tissues are counterstained through a 2 min incubation in hematoxylin. 4. Excess of hematoxylin is rinsed off with large volumes of deionized water. The tissues are submerged in fresh clean water for 5 s; this is repeated several times until the rinsing water remains colorless. 5. The stained tissues are dehydrated in increasing concentrations of ethanol (2 min at 75% EtOH, 2 min at 95% EtOH, and 2 min at 100% EtOH) and mounted in xylene.

3.4.3. Live Microglia Immunofluorescence

Isolectin binds specifically to microglia cells and fluorescent conjugated forms of this agent can be used for live microglia staining. 1. The slices are submerged in an isolectin solution (5 μL/mL) in culture medium. 2. Next, the slices are washed three times with 500 μL PBS for 30 min each and can be analyzed under an inverted fluorescence microscope.

3.5. Virus Extraction from Brain and Prostate Organ Cultures Slices (12, 14)

1. The tissues are resuspended in an appropriate volume of DMEM supplemented with a P/S solution diluted 1:100. 2. The tissues are then manually homogenized by pipetting them up and down. 3. The homogenized tissues are frozen in a dry ice–EtOH bath and thawed at 37°C three consecutive times. 4. After the final thaw, the freeze-thawed material is sonicated for 10 s in an ice bath to ensure release of the virus. 5. The cellular debris from the tissue lysates is pelleted through centrifugation at 14,000g for 10 min at 4°C. 6. Virus-containing supernatants are collected in new tubes and stored at – 80°C. 7. Virus titers can then be determined as described next.


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1. Vero cells are plated at 400,000 cells/well in complete DMEM in 6-well plates and allowed to grow overnight. 2. The following day, serial dilutions (1:10) of the OV stock are made in glucose-enriched PBS. 3. OV inoculum (0.7 mL) is added to duplicate wells and incubated for 1 ½ h at 37°C. During this incubation, the plates must be rocked every 15 min to avoid drying of the cells. 4. OV inoculum is removed and 2 mL of DMEM media containing 1% FCS, 1% P/S solution, and human anti-HSV IgG is added to each well. 5. The virus is allowed to grow for 4 days. The presence of antiviral IgG in the culture medium will not allow the spread of OV particles to distant cells. Therefore, only the cells adjacent to the initially infected cell will be infected by the virus, thus allowing the formation of distinct virus plaques that will correspond to the initial pfu of the OV stock. 6. After staining of virus β-galactosidase activity as described in Subheading 3.4.2, the OV plaques can be easily counted (see Note 13).

4. Notes 1. The vibratome and dissecting microscope are placed under the laminar hoods during preparation and sectioning. 2. All tools for preparing the organ cultures must be sterile. The instruments are sterilized with a high-temperature heater with glass beads after dissecting each animal, and cooled with icecold sterile phosphate buffer (PBS). All surfaces of the slicing apparatus that come in contact with the tissue, as well as the blades and the fine brushes, are sterilized with 70% alcohol. 3. To avoid excessive autofluorescence during the imaging experiments, all media reagents must be without phenol red. The medium is sterilized through a 0.2-μm filter before use (SCGVU02RE, Millipore). 4. The serum (FBS) is stored at –20°C. Before using, it is thawed at room temperature, and incubated in a water bath at 56°C for 30 min for complement inactivation. 5. Dissolve 1 packet of the buffer powder in 1 L of deionized water for a 1X ready-to-use solution, and store at room temperature. 6. The optimal dilution of primary and secondary antibodies will have to be determined empirically for each model. In all cases, we have used a 1:100 dilution for the primary antibody and a 1:200 dilution for the secondary antibody.

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7. Prepare X-gal stock solutions by dissolving 50 mg/mL X-gal in DMSO and store in the dark at – 20°C; prepare 10 mL of 125 mM potassium ferricyanide (0.41 g in 10 mL 1X PBS) and 10 mL of 125 mM potassium ferrocyanide (0.53 g in 10 mL 1X PBS) and store at room temperature. 8. Different tumor cells require different culture treatments. In our experiments, we have used mainly U87 human glioblastoma and D74 rat glioma cells transduced with the HSV receptor HveC (D74-HveC (12)). Both of these cell lines are detached from the culture dish through trypsinization. Briefly, cells are washed with prewarmed sterile PBS, incubated with trypsin–EDTA at 37°C until all cells are detached from the culture dish, and resuspended in complete growth medium. The serum of the medium inactivates completely the trypsin enzymatic activity. 9. The quantity of virus to be used depends on the OV strain, tumor model, and experiment. We have used 10 plaqueforming units (pfu) of hrR3 in the D74-HveC rat glioma model. 10. The tissues are dried overnight and submerged in a bath with ice-cold acetone for 10 min. Then, the tissues are air-dried for 30 min at RT before storage. It is important to use large volumes of acetone to avoid evaporation of the acetone during the fixation procedure. To avoid melting of the 24-well cell culture plate, the organotypic brain slices are fixed on a glass cover slip. Then, they are placed and stored in 24-well cell culture plates. Because of the thickness of the organotypic brain slices, staining of these tissues requires prolonged incubation time to allow the reagents to penetrate the whole tissue in a homogeneous way. Indeed, even though these slices are too thick for high-resolution IHC, they are too thin to be cut into thinner, 2- to 6-μm slices. The same problems were not present with the prostate organ cultures, which were easily cut into 5-μm slices (see Subheading 3.3) before staining. 11. Staining of the tumor implanted on the ex vivo brain slices is very difficult. Indeed, because of its fragile structure, the tumor tends to easily detach from the brain slice. To avoid this problem, the staining and washing passages must be reduced to a minimum. Thus bright light staining with DAB detection is not recommended, and fluorescent staining is preferred. 12. Microscopy imaging of 300-μm-thick tissues is very challenging. Good cellular resolution can be achieved only at high magnifications (40 × 10 or higher). A broad view of the tumor tissue and surrounding brain performed at low magnification has very poor resolution. The best way to image the tumor is on unstained slices under bright light microscopy (14). The tumor is thick enough to give good contrast


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and under these conditions the tissue is not disrupted. DAPI staining of the tumor tissue at low magnification gives a dark-black contrast because of autofluorescence caused by the thickness of the tumor (14). 13. Before staining the cells for β-galactosidase activity, the cells must fixed to the cell culture dish. This is performed by incubating the cells for 5 min at 4°C within the following freshly prepared fixing solution: 2.7 mL of 37% formaldehyde plus 0.4 mL 25% gluteraldehyde in 50 mL of 1X PBS. There is no need to rinse the cells before fixation or before adding the LacZ staining solution.

Acknowledgments This work was supported by a US Department of Defense (DOD) grant W81XWH-05-1-0367 to Brent J. Passer.

References 1. Fulci G, Chiocca EA. (2003) Oncolytic viruses for the therapy of brain tumors and other solid malignancies: a review. Front Biosci 8:e346–60. 2. Varghese S, Rabkin SD, Nielsen GP, MacGarvey U, Liu R, Martuza RL. (2007) Systemic therapy of spontaneous prostate cancer in transgenic mice with oncolytic herpes simplex viruses. Cancer Res 67:9371–9. 3. Varghese S, Rabkin SD, Nielsen PG, Wang W, Martuza RL. (2006) Systemic oncolytic herpes virus therapy of poorly immunogenic prostate cancer metastatic to lung. Clin Cancer Res 12:2919–27. 4. Liu R, Varghese S, Rabkin SD. (2005) Oncolytic herpes simplex virus vector therapy of breast cancer in C3(1)/SV40 T-antigen transgenic mice. Cancer Res 65:1532–40. 5. McKee TD, Grandi P, Mok W, et al. (2006) Degradation of fibrillar collagen in a human melanoma xenograft improves the efficacy of an oncolytic herpes simplex virus vector. Cancer Res 66:2509–13. 6. Ganesh S, Gonzalez Edick M, Idamakanti N, et al. (2007) Relaxin-expressing, fiber chimeric oncolytic adenovirus prolongs survival of tumor-bearing mice. Cancer Res 67:4399–407.

7. Ikeda K, Ichikawa T, Wakimoto H, et al. (1999) Oncolytic virus therapy of multiple tumors in the brain requires suppression of innate and elicited antiviral responses. Nat Med 5:881–7. 8. Ikeda K, Wakimoto H, Ichikawa T, et al. (2000) Complement depletion facilitates the infection of multiple brain tumors by an intravascular, replication-conditional herpes simplex virus mutant. J Virol 74:4765–75. 9. Hirasawa K, Nishikawa SG, Norman KL, et al. (2003) Systemic reovirus therapy of metastatic cancer in immune-competent mice. Cancer Res 63:348–53. 10. Balachandran S, Barber GN. (2004) Defective translational control facilitates vesicular stomatitis virus oncolysis. Cancer Cell 5:51–65. 11. Balachandran S, Thomas E, Barber GN. (2004) A FADD-dependent innate immune mechanism in mammalian cells. Nature 432401–5. 12. Wakimoto H, Fulci G, Tyminski E, Chiocca EA. (2004) Altered expression of antiviral cytokine mRNAs associated with cyclophosphamide’s enhancement of viral oncolysis. Gene Ther 11:214–23. 13. Fulci G, Breymann L, Gianni D, et al. (2006) Cyclophosphamide enhances glioma virotherapy by inhibiting innate immune responses. Proc Natl Acad Sci USA 103:12873–8.

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14. Fulci G, Dmitrieva N, Gianni D, et al. (2007) Depletion of peripheral macrophages and brain microglia increases brain tumor titers of oncolytic viruses. Cancer Res 67:9398–406. 15. Lamfers ML, Fulci G, Gianni D, et al. (2006) Cyclophosphamide increases transgene expression mediated by an oncolytic adenovirus in glioma-bearing mice monitored by bioluminescence imaging. Mol Ther 14:779–88. 16. Iankov ID, Pandey M, Harvey M, Griesmann GE, Federspiel MJ, Russell SJ. (2006) Immunoglobulin g antibody-mediated enhancement of measles virus infection can bypass the protective antiviral immune response. J Virol 80:8530–40. 17. Power AT, Wang J, Falls TJ, et al. (2007) Carrier cell-based delivery of an oncolytic virus circumvents antiviral immunity. Mol Ther 15:123–30. 18. Li H, Zeng Z, Fu X, Zhang X. (2007) Coadministration of a herpes simplex virus-2 based oncolytic virus and cyclophosphamide produces a synergistic antitumor effect and enhances tumor-specific immune responses. Cancer Res 67:7850–5. 19. Nevalainen MT, Harkonen PL, Valve EM, Ping W, Nurmi M, Martikainen PM. (1993) Hormone regulation of human prostate in organ culture. Cancer Res 53:5199–207. 20. Shinmura Y, Kosugi I, Kaneta M, Tsutsui Y. (1999) Migration of virus-infected neuronal cells in cerebral slice cultures of developing mouse brains after in vitro infection with murine cytomegalovirus. Acta Neuropathol (Berlin) 98:590–6.

21. Jung S, Ackerley C, Ivanchuk S, Mondal S, Becker LE, Rutka JT. (2001) Tracking the invasiveness of human astrocytoma cells by using green fluorescent protein in an organotypical brain slice model. J Neurosurg 94:80–9. 22. de Bouard S, Christov C, Guillamo JS, et al. (2002) Invasion of human glioma biopsy specimens in cultures of rodent brain slices: a quantitative analysis. J Neurosurg 97: 169–76. 23. Markovic DS, Glass R, Synowitz M, Rooijen N, Kettenmann H. (2005) Microglia stimulate the invasiveness of glioma cells by increasing the activity of metalloprotease-2. J Neuropathol Exp Neurol 64:754–62. 24. Belmadani A, Tran PB, Ren D, Miller RJ. (2006) Chemokines regulate the migration of neural progenitors to sites of neuroinflammation. J Neurosci 26:3182–91. 25. Goldstein DJ, Weller SK. (1988) An ICP6::lacZ insertional mutagen is used to demonstrate that the UL52 gene of herpes simplex virus type 1 is required for virus growth and DNA synthesis. J Virol 62:2970–7. 26. Todo T, Martuza RL, Rabkin SD, Johnson PA. (2001) Oncolytic herpes simplex virus vector with enhanced MHC class I presentation and tumor cell killing. Proc Natl Acad Sci, USA 98:6396–401. 27. Del Duca D, Werbowetski T, Del Maestro RF. (2004) Spheroid preparation from hanging drops: characterization of a model of brain tumor invasion. J Neurooncol 67:295–303.

Chapter 4 Use of Minicircle Plasmids for Gene Therapy Peter Mayrhofer, Martin Schleef, and Wolfgang Jechlinger Summary A large number of cancer gene therapy clinical trials are currently being performed that are attempting to evaluate novel approaches to eliminate tumor cells by the introduction of genetic material into patients. One of the most important objectives in gene therapy is the development of highly safe and efficient vector systems for gene transfer in eukaryotic cells. Currently, viral and nonviral vector systems are used, both having their advantages and limitations. Minicircles are novel supercoiled minimal expression cassettes, derived from conventional plasmid DNA by site-specific recombination in vivo in Escherichia coli for the use in nonviral gene therapy and vaccination. Minicircle DNA lacks the bacterial backbone sequence consisting of an antibiotic resistance gene, an origin of replication, and inflammatory sequences intrinsic to bacterial DNA. In addition to their improved safety profile, minicircles have been shown to greatly increase the efficiency of transgene expression in various in vitro and in vivo studies. In this chapter, we describe the production, purification, and application of minicircle DNA and discuss the rationale of the improved gene transfer efficiencies compared to conventional plasmid DNA. Key words: Bacterial backbone sequences, biosafety, minicircle DNA, nonviral gene therapy, persistent transgene expression, plasmid, site-specific recombination.

1. Introduction Cancer gene therapy aims to eliminate tumor cells, while avoiding adverse effects on healthy tissues, through the introduction of genetic material (DNA and RNA) into the cells of a patient. Strategies of cancer gene therapy may be broadly subdivided into (i) the application of genes inhibiting tumor growth-like tumor suppressor genes (1), genes stimulating apoptosis (2), or genes blocking angiogenesis (3); (ii) turning off genes in tumor cells


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with antisense oligonucleotides or small interfering RNA (siRNA) (e.g., genes inhibiting apoptosis) (4); and (iii) cancer vaccination or immunotherapy, which aims to induce a cellular immune response against specific tumor antigens like HER2/NEU in breast cancer therapy (5), thus, enabling the immune system to recognize and kill tumor cells (6, 7). In the broadest sense, vaccines delivering genetic material that prevent but do not treat cancer, for example, a human papilloma virus (HPV) DNA vaccine preventing cervical carcinoma, may as well be attributed to the field of cancer gene therapy (8). Plasmid DNA is commonly used for gene therapy, with naked plasmid DNA trials already contributing 18% of all clinical trials in mid-2007 (9). Most possibly, the steady increase in the use of nonviral vector systems can be attributed to safety concerns associated with viral vector technologies (10–12), leading to the decrease in viral-based trials (9, 13). Although considered safe and easy to produce in large-scale quantities (14), one disadvantage of nonviral vectors is the low gene transfer efficiency when compared with viral vector systems (9). Therefore a major research focus is the improved delivery and optimization of the plasmid DNA itself in terms of (1) enhancing the efficiency of plasmid uptake into the cells and the nucleus, (2) enhancing gene expression, (3) extending the time of gene expression, and (4) designing the desired immune response, which should be enhanced in vaccination strategies, but avoided in the case of therapeutic gene delivery. In addition to the improvement of gene transfer efficiencies, biosafety issues have to be addressed adequately. Conventional plasmid vectors can be subdivided into a bacterial backbone and a transcription unit. The transcription unit usually carries the target gene or sequence along with necessary regulatory elements (15). The bacterial backbone includes elements like an antibiotic resistance gene, an origin of replication, unmethylated CpG motifs, and potentially cryptic expression signals (15). Some of these sequences are required for the production of plasmid DNA, but each of them can cause serious biological safety problems. Among other things, regulatory agencies therefore recommend totally avoiding the use of antibiotic resistance markers (16–18). In addition to the dissemination of antibiotic resistance genes via horizontal gene transfer, the expression of bacterial genes in patients represents another serious problem. For instance, it has been demonstrated that intramuscular injection of plasmid DNA not encoding any eukaryotic promoter element into rabbits leads to expression of bacterial sequences and therefore to an unintended immune response (19). Additionally, the production of antibiotic resistance genes from cryptic upstream eukaryotic expression signals results in an altered gene expression

Use of Minicircle Plasmids for Gene Therapy

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profile in the mammalian cell itself (20, 21). More importantly, the bacterial backbone DNA has been shown to be responsible for silencing the expression of the encoded antigen (22). Bacterial DNA contains unmethylated CpG motifs, which activate the innate immune system of the host by binding to the Toll-like receptor 9 (TLR9) of antigen-presenting cells (23, 24). Because unmethylated CpG motifs are much less frequent in vertebrate DNA, it seems that these sequences act as an alarm signal for the immune system during bacterial infections. After binding to TLR9, a cascade of immunostimulatory events leads to the activation of diverse eukaryotic transcription factors, which, in turn, upregulate the expression of cytokines and chemokines. Finally, the binding of CpG motifs to TLR9 leads to the maturation, differentiation, and proliferation of natural killer (NK) cells, macrophages, and T cells, which themselves secrete cytokines that direct the immune system into a TH1-dominated response (23, 24). This adjuvant effect is certainly valuable for vaccination purposes, where the induction of an immune response is necessary for the protective effect. However, there are different optimal immunogenic CpG motif hexamer sequences for different species because TLR9 also differs markedly between species (25, 26). Further, different CpGs and flanking sequences have been shown to activate different types of immune cells and cytokines when co-applied as adjuvants on oligonucleotides (23). Thus, even if the CpG adjuvant effect is beneficial for vaccination purposes, the incorporation of defined amounts of species-specific CpG motifs stimulating the desired immune response should be preferred in rational vaccine design. For therapeutic purposes in gene therapy, for which an immune stimulation is not required, those sequence motifs are clearly counterproductive. Summing up, the bacterial backbone of conventional plasmid DNA (1) constitutes a significant portion of the plasmid DNA without a therapeutic effect (“junk” DNA), leading to an decreased bioavailability because of the increased size of the plasmid DNA; (2) leads to inefficient and short-lived expression of the transgene in eukaryotic cells; (3) consists of sequences associated with biosafety concerns; and (4) contains motifs eliciting an inflammatory response in the host. It seems obvious that the removal of bacterial backbone DNA can greatly improve the safety and efficiency of plasmid DNA used for gene therapy and vaccination. An in vitro system (MIDGE) based on excision of the therapeutic expression cassette (minimal expression unit) by endonuclease digestion is described in Chapter 2.6 the MIDGEvector system. Here we describe the production, purification, and application of supercoiled minimal plasmids, also known as minicircle DNA, devoid of bacterial backbone sequences for application in nonviral gene transfer.

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2. Production of Minicircle DNA by Site-Specific Recombination In Vivo

Minicircles are the result of an in vivo site-specific recombination process. The parental plasmid (PP) carries the eukaryotic expression cassette flanked by two recognition sites of a sitespecific recombinase (Fig.1a). The in vivo expression of the respective recombinase results in the excision of the interjacent DNA sequences, dividing the parental plasmid into two supercoiled molecules: (1) a replicative miniplasmid carrying the undesired backbone sequences, and (2) a minicircle carrying the therapeutic expression unit (Fig.1). The recombinases used so far for minicircle DNA production were either derived from the λ integrase (tyrosine recombinase) family (the integrase of bacteriophage lambda, the Cre recombinase from bacteriophage P1, and the FLP recombinase of the yeast plasmid 2-µm circle) or the serine recombinase family (the integrase of Streptomyces bacteriophage Phi31 or the ParA resolvase from the multimer resolution system of the broad host range plasmid RK2 or RP4) (27–31). There are differences in the recombination events mediated by the respective recombinases that are of high importance for the in vivo production of minicircle DNA in Escherichia coli (32). The genetic crossover between the recombination sites of the Cre recombinase (lox) and of the FLP recombinase (FRT) regenerates a site of identical or highly similar sequences and thus the mediated recombination event is bidirectional and fully reversible, giving rise to intramolecular and intermolecular recombination events (32, 33). This bidirectionality of the Cre and FLP recombinases, in addition to leading to the production of monomeric minicircle molecules, results in the unwanted formation of dimeric, trimeric, etc. minicircles, parental plasmids, miniplasmids, and mixed concatemers (27, 31). Thus, Bigger and coworkers constructed a parental plasmid with one lox site being mutated to prefer unidirectional recombination events (27). However, even if the generation of monomeric minicircle molecules was improved, a high amount of concatamerization was still observed (27). The lambda integrase on the other site is supposed to be able to favor unidirectional recombination events between attB and attP sites in the absence of the Xis protein (32). The resulting molecules carrying attL and attR sites are recombined mainly in the presence of Xis. However, it was demonstrated that about 30% of the minicircles produced with the lambda integrase system were present as dimers in a Xisnegative environment (34). The additional application of the multimer resolution system of the broad host range plasmid RK4 allowed efficient resolution of the dimeric minicircles originally produced by the lambda integrase (34). The integrase


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Fig. 1. Schematic illustration of the production of minicircle DNA via site-specific recombination. As an example, minicircle production using the ParA resolvase system is illustrated (Adapted from Mayrhofer et al., submitted). The parental plasmid contains the parA resolvase gene under transcriptional control of the arabinose promoter/operator system, whereas the transcription unit is flanked by the corresponding resolution sites. After transformation of the parental plasmid into a bacterial strain, the recombination process can be induced by the addition of arabinose. The ParA resolvase catalyzes site-specific recombination, resulting in a replicative miniplasmid and a minicircle, which carries the transcription unit. (a) Schematic drawing of a parental plasmid (PP) and the products of the recombination process (minicircle [MC] and miniplasmid [MP]). CMV, major immediate early (IE) enhancer containing promoter of the human cytomegalovirus; luc, luciferase reporter gene; SV40 polyA, polyadenylation signal derived from the simian virus 40; bla, ampicillin resistance gene; MB1, origin of replication derived from pUC19; parA, parA resolvase gene; P, arabinose-inducible promoter; araC, repressor/inducer of the PBAD promoter; res, resolution sites. (b) Undigested DNA preparations of the parental plasmid before and after induction of recombination analyzed by agarose gel electrophoresis: lane 1, 1-kb size marker; lane 2, without addition of 0.5% l-arabinose, only parental plasmid is visible; lane 3, the parental plasmids are completely divided into minicircles and miniplasmids 60 min after induction, mainly supercoiled forms of MC and MP are visible beside slight traces of open circular (oc) forms of MC and MP. (c) The recombination products (see d, lane 3) were digested with NsiI; different concentrations of digested and undigested samples were analyzed by agarose gel electrophoresis. Lane 1, 1-kb size marker; lanes 2–7, the minicircles and miniplasmids were linearized (MC: 4,040 bp, MP: 4,353 bp), no parental plasmid was detected although 700 ng of the recombination products was applied (NsiI fragments of parental plasmid: 4.8 and 3.6 kb). The gel photos were kindly provided by PlasmidFactory GmbH Co. KG (Bielefeld, Germany).

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of bacteriophage Phi31, a member of the large serine recombinase family, has been shown to be strictly unidirectional. This enzyme mediates recombination events between an attP and an attB site, resulting in recombination products containing attL and attR sites (35, 36). The reverse reaction has neither been observed in E. coli nor in vitro (35, 36). Thus, minicircle production can be driven to a high percentage without reversible reactions and without the generation of concatemers (28). Likewise, the ParA resolvase of the multimer resolution system of the broad host range plasmids RK2 and RP4, belonging to the small serine recombinase family of recombinases, is only able to mediate intramolecular and not intermolecular recombination events between the corresponding directly repeated resolution sites and is thus not able to revert excision events (37–39). The ParA resolvase therefore was able to direct the recombination event in one direction to completion and no multimers or other concatamerization events were observed during minicircle production (30). To accomplish an efficient recombination process, the stringent control of repression and expression of the site-specific recombinase in E. coli is of high importance. In the first place, the expression system should efficiently silence gene expression before induction to avoid premature recombination leading to the displacement of the parental plasmid by the miniplasmid. The temperature-sensitive lambda cI857/pR promoter (29, 31) and the PBAD/araC arabinose expression system (27, 28, 30) have been shown to be able to inhibit background expression of the recombinases in the uninduced state. On the other hand, a high ratio of recombinase to plasmid is necessary to yield high recombination rates. Approaches integrating a single copy of a respective recombinase gene under control of the lambda or arabinose expression system in the chromosome of E. coli to obtain minicircle production strains led to an overall low recombination efficiency of 50–90% at maximum (27, 29, 31, 40, 41). Transformation of a low copy number plasmid expressing the ParA resolvase in addition to a compatible high copy number plasmid containing the corresponding resolution sites in E. coli led to a recombination efficiency of only 50% (30). However, the integration of recombinase expression systems into the same plasmid that contains the corresponding recombination sites led to a very high recombination efficiency of 97% in the case of the Phi31 integrase (28) and to a complete recombination when the ParA resolvase system was applied (30) according to agarose gel electrophoresis (AGE). The residual amount of parental plasmid 60 min after ParA-mediated recombination was undetectable using AGE (Fig. 1) and a recombination efficiency of 99.57% has been determined by quantitative real-time PCR (42).


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3. Purification of Minicircle DNA The in vivo recombination process in E. coli generates three plasmid species: (1) minicircles, (2) miniplasmids, and (3) residual amounts of parental plasmids. Thus, the molecule of interest, the therapeutically valuable minicircle, has to be separated from the other plasmids after recombination. So far, in all of the studies described below, the miniplasmids and thus the parental plasmids as well have been linearized by restriction digestion, and the minicircle was subsequently purified via ultracentrifugation in cesium chloride. The major disadvantages of this method are the relatively low yield of minicircle DNA, the high costs of enzymes, and the labor intensity, which make the method unsuitable for large-scale preparations of minicircle DNA. Chen and coworkers developed a method that degrades the remaining miniplasmid DNA and parental plasmid DNA (pDNA) in vivo via the coexpression of an restriction enzyme (43). The homing endonuclease used in this procedure (I-SceI) is expressed together with the Phi31 recombinase as a bicistronic messenger RNA (mRNA) (43). After induction, minicircles and miniplasmids are generated and the expressed endonuclease recognizes an 18-bp sequence that is not present in the E. coli genome and the transcription unit, but is incorporated into the backbone sequence of the parental plasmid DNA (43). Miniplasmids as well as parental plasmids are degraded by linearization and the activity of E. coli exonucleases. However, it has been reported that, when using this system even 240 min after induction of the recombinase and the endonuclease, 3% of miniplasmids and parental plasmids are still detectable (43). Another approach is based on the affinity purification of the minicircle DNA. In this case, a short recognition sequence is integrated into the parental plasmid at a position that is located on the minicircle after the recombination process. This system of course assumes highly efficient recombination, because only plasmids carrying the recognition sequence can be separated from plasmids without the recognition sequence (e.g., minicircle and parental plasmid cannot be separated). In principle there are a few affinity purification systems for DNA under development either based on DNA–DNA interaction, like the triple helix system (44), or on protein–DNA interaction (45, 46). Currently, none of the mentioned systems have been reported to have been used to separate a mixture of minicircle and miniplasmid DNA. So far the only system described for this purposed is based on the interaction of a direct tandem repeat of modified lactose operator sites (lacOs sites) serving as the recognition sequence and the repressor of the lactose operon (LacI) (30, 42, 47). Plasmid

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DNA containing these recognition sequences can be reversibly bound to a solid support carrying LacI, thereby enabling the separation from DNA without LacI (42). This system allows the preparation of highly pure minicircle DNA and is scaleable for industrial production (42 and Chapter 5.4. Production of plasmid-DNA as pharmaceutical).

4. Minicircle Vectors as Improved Molecules for In Vitro and In Vivo Gene Transfer

Darquet and coworkers (1997) were the first to show that minicircle DNA leads to much higher transgene expression levels in cell culture experiments than the parental plasmid or other conventional control plasmids encoding the same transgene (29). The same molarity of the reporter gene expression cassette was used in the comparative experiments in five different cell lines, whereby the lower amount of minicircle DNA was compensated by the addition of unrelated pDNA (stuffer DNA) to avoid an amount-associated effect on the gene transfer efficiencies. The expression levels caused by the minicircle DNA-encoded transgene cassette were at least three to ten times higher than those achieved with parental plasmid or conventional plasmid DNA, depending on the cell lines (Table 1) (29). The results of this first study were confirmed by other groups using the same application scheme. Not surprisingly, transfecting the same amount of minicircle DNA and parental plasmid (weight-to-weight basis) led to a even more significant increase (8.8-fold up to 40-fold) in the expression of the encoded transgene, because in this case more reporter gene cassettes were administered (Table 1) (27, 29, 40). A vast increase in minicircle reporter gene activity (up to 152-fold) over parental plasmid was observed when the same molarity of the expression cassette was used (mole-to-mole comparison) but no stuffer DNA was added to the minicircle DNA samples (27). These cell culture experiments were performed with Lipofectamine and, thus, the lower amount of minicircle DNA used in the molar-to-molar comparison studies also led to the use of a lower amount of the transfection reagent. Lipofectamine is applied in a certain ratio to the DNA to be transfected and also exhibits cytotoxic effects at certain ratios (27). These experiments highlighted the benefit of minicircle DNA for transfections, because lower amounts of DNA containing the same number of expression cassettes need less transfection reagent. By maximizing the amount of the transcriptional units per weight of DNA but minimizing the amount of the transfection reagent, the expression levels could be increased greatly by reducing the cytotoxicity of the lipoplexes.

Transgeneb

Luc

Luc

Luc

Luc

Luc

Luc

ß-gal

ß-gal

Luc

Luc

Luc

Luc

Luc

Luc

hAAt

Systema

λ Int/att

λ Int/att

λ Int/att

λ Int/att

λ Int/att

λ Int/att

λ Int/att

λ Int/att

λ Int/att

λ Int/att

λ Int/att

Cre/lox

Cre/lox

Cre/lox

ϕC31 Int/att RSV

CMV

CMV

CMV

SV40

CMV

SV40

CMV

CMV

CMV

CMV

SV40

SV40

SV40

SV40

PRc

C57BL/6 mice

HeLa

HeLa

HeLa

TU182

C57BL6 mice

C57BL6 mice

lacZ mice

NIH 3T3/RSM

RSM

NIH 3T3

HSM

RSM

H460

3LL/NIH3T3

Cell lined/animals

HD/8.5

LF/0.24

LF/0.5

LF/0.24

IT/10

IM/10

IM/10

IM/10

LF/0.65

LF/0.55

LF/0.55

LF/0.46

LF/0.46

LF/0.46

LF/0.46

m:m

m:m

w:w

m:m + st

w:w

w:w

w:w

w:w

m:m + st

m:m + st

m:m + st

m:m + st

m:m + st

m:m + st

m:m + st

560

152

8.8

4.5

40

50

32

13

3

5.5

3

>10

10

7

4

TF-method (route)e/µg MC Ratio MC:PPf Expression MC > PPg

(continued)

(28)

(27)

(27)

(27)

(40)

(40)

(40)

(40)

(40)

(40)

(40)

(29)

(29)

(29)

(29)

References

Table 1 Comparison of the expression level of transgenes encoded on minicircle DNA (MC) or parental plasmids (PP) in various in vitro and in vivo systems

Use of Minicircle Plasmids for Gene Therapy 95

hFIX

IFNγ

IFNγ

IFNγ

ß-gal

luc (+HSV amplicon)

ϕC31 Int/att

ϕC31 Int/att

ϕC31 Int/att

ϕC31 Int/att

Cre/lox

ϕC31 Int/att CAG

CMV

CMV

CMV

CMV

RSV

PRc

MRC9

MF-1 mice

293/NIH3T3/ NPC

293/NIH3T3/ NPC

293/NiH3T3/ NPC

C57BL/6 mice

Cell lined/animals

HSV virions/106 TU

HD/0.5

LF/0.14

LF/0.14

LF/1

HD/16.2

tu:tu (m:m)

w:w

m:m

m:m + st

w:w

m:m

20

10

30

5.5/10

21/40 (NPC)

45

TF-method (route)e/µg MC Ratio MC:PPf Expression MC > PPg

(60)

(51)

(48)

(48)

(48)

(28)

References

a Site-specific recombination system used for the production of MCs. λ, bacteriophage lambda; Int, Integrase; att, attachment site of bacteriophage integrases; Cre, bacteriophage P1 derived recombinase; lox, recognition site of Cre recombinase; ϕC31, bacteriophage ϕC31. b Luc, luciferase gene; ß-gal, ß-galactosidase gene; hAAt, human α1-antitrypsin; hFIX, human serum factor IX; IFNγ, interferon γ; HSV, herpes simplex virus. c PR, eukaryotic promoter systems. CMV, major immediate early (IE) enhancer containing promoter of the human cytomegalovirus; SV40, early promoters of the simian virus 40; RSV, early promoters of the Rous sarcoma virus; CAG, hybrid CMV enhancer—modified chicken ß-actin promoter. d 3LL, mouse Lewis lung carcinoma; NIH3T3, murine fibroblast; H460, human non-small cell lung carcinoma; RSM; rabbit aortic smooth muscle; HSM, human aortic smooth muscle; TU182, human and neck carcinoma cells implanted in nude mice; 293, human embryonic kidney; NPC, nasopharyngeal carcinoma; MRC9, human fibroblast. e TF, transfection method, LF, lipofection; IM, intramuscular; IT, intratumoral; HD, hydrodynamic; TU, transducing units. f M:m, molar:molar: the same molarity of the transgene encoded on MC and PP is applied (e.g., 1 µg PP 11.3 kb compared with 0.14 µg of 1.6 kb MC); m:m + st, molar:molar + stuffer DNA: stuffer DNA is added to the MC samples (e.g., 1 µg of 11.3 kb PP compared with 0.14 µg of 1.6 kb MC + 0.86 µg unrelated pDNA [total 1 µg]); w:w, weight:weight: same amount (e.g., 1 µg) of PP and MC is applied. g X-fold expression of transgene encoded on MC when compared with the expression level of the same transgene encoded on the PPv.

Transgeneb

Systema

Table 1 (continued)

96 Mayrhofer, Schleef, and Jechlinger


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The first in vivo studies using minicircle DNA in comparison with unrecombined plasmid was performed by Scherman’s group in 1999, demonstrating that the effect of minicircle DNA application in vivo was even more profound than in vitro (40). The injection of minicircle DNA into mouse cranial tibial muscle or human head and neck carcinoma cells led to 13–50 times more reporter gene expression than with parental plasmids or larger plasmids in weight-to-weight comparison experiments (Table 1) (40). However, one of the most impressive results obtained to identify minicircle DNA as powerful nonviral vector for gene therapy purposes was described by the group of Mark Kay (28). They compared the application of equimolar amounts of (1) purified linear expression cassettes (derived by the excision of the expression cassette with endonucleases and subsequent purification), (2) a mixture of linearized expression cassette and the linearized bacterial backbone, (3) uncut parental plasmid, and (4) minicircle DNA encoding the respective expression cassette. The expression cassette consisted of a gene encoding human α1-antitrypsin (AAT) under control of the Rous sarcoma virus (RSV) promoter and the bovine growth factor polyadenylation signal (28). Three weeks after hydrodynamic injection, the serum concentrations of AAT obtained from mice that received the linearized purified expression cassette were three times higher than those obtained from mice that received the two fragments, and 20- to 43-fold higher than those of mice that received the parental plasmid (28). The serum AAT levels from mice that received minicircle DNA were 10–13 times higher than those from mice that received the purified expression cassette and thus were 200- to 560-fold higher when compared with serum concentrations reached after the injection of parental plasmid (28). Although initially the serum concentrations were comparably high in the four different groups, the serum reporter level decreased dramatically and rapidly in the three non-minicircle groups, especially in the parental plasmid group (28). These results demonstrated that supercoiled minicircle DNA is by far the most efficient form of an expression cassette to elicit persistent and high-level transgene expression (28) (Table 1). Based on these results, a nonviral cancer gene therapy approach was performed by Wu and coworkers to investigate the antitumor effect of interferon (IFN)-γ gene transfer on human nasopharyngeal carcinoma by using IFNγ–minicircle DNA (48). Expression levels of IFNγ were 11–14 times higher after intratumoral administration of IFNγ-minicircles in nasopharyngeal carcinoma (NPC) CNE-2 and C666-1 cell line-xenografted mice compared with the IFNγ expression measured after injection of the IFNγ–parental plasmids. Using the minicircle approach, IFNγ could be detected in tumors until day 21 after infection compared with day 7 when parental plasmids were applied (48). Further, minicircle IFNγ treatment achieved tumor inhibition

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rates of 77.5% and 83% in CNE-2 and C666-1 xenografted mice (parental plasmid: 32.4% and 58.5%, respectively) (48). Minicircle DNA treatment additionally increased the median survival of human NPC-xenografted mice up to 77 days after DNA injection, whereas mice treated with IFNγ-expressing parental plasmids survived 47 days, and the control group that did not receive an IFNγ expression cassette survived 38 days (48). Further studies supported the idea that minicircle DNA improves gene transfer efficiency (49, 50). Park and coworkers demonstrated that the application of minicircle DNA led to an efficient delivery of an adiponectin gene to diet-induced obese mice, whereas parental plasmids had no effect on the adiponectin levels (49). The sufficient blood level of adiponectin in the minicircle-treated mice normalized the parameters related to insulin resistance, thus, this treatment is proposed as a potential gene therapy strategy in cases of type 2 diabetes (49). Currently, additional features are being incorporated in minicircles, e.g., to improve their nuclear uptake via the attachment of nuclear localization signal peptides (51) or to improve their persistence in cells by introducing a scaffold/matrix attachment region (S/MAR) enabling stable replication and maintenance in dividing eukaryotic cells (31).

5. Rationale of the Improved Expression of Transgenes by Minicircle DNA 5.1. Size and Structure

One of the most obvious differences between minicircle DNA and conventional or parental plasmids is their size. It has been proposed that smaller plasmids like minicircles have better bioavailability characteristics than larger ones, endowing smaller plasmids with an advantage to overcome obstacles on their way to gene expression. Before a plasmid-encoded gene can be expressed, it needs to pass several barriers. It has to diffuse into the tissue, enter through the cell membrane, escape the endo/lysosome in case of internalization via endocytosis, diffuse into the cytoplasm, and, finally, cross the nuclear membrane. That the size of plasmid DNA might have an influence on the above-mentioned processes has been discussed in different reports (40, 41, 52). Another important feature of minicircle DNA produced by in vivo recombination is that the supercoiled structure remains unchanged. It has been proposed that the supercoiled structure of plasmids is superior to linear plasmid structures in terms of transfection efficiencies (29, 53, 54); this was thought to be caused by the higher intranuclear concentration of the


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supercoiled molecules (54). Chen and coworkers demonstrated that mice transfected with minicircles encoding an AAT expression cassette exhibited 10–13 times more serum AAT levels when compared with mice transfected with linearized purified expression cassettes (22). 5.2. Transgene Silencing by Bacterial Backbone DNA

In addition to the advantages of minicircle DNA in terms of bioavailability because of their size and physiochemical properties, the removal of the bacterial backbone leads to higher transfection efficiencies, which can be ascribed to the intrinsic nature of bacterial DNA. The removal of the bacterial backbone DNA also leads to the reduction or elimination of bacterial unmethylated CpG motifs. These motifs are characterized by CpG dinucleotides flanked by two 5′ purines and two 3′ pyrimidines, which have been shown to be present much more frequently in bacterial than in mammalian DNA. These unmethylated motifs have been shown to induce inflammatory responses when applied in vivo (23, 24). These inflammatory reactions are clearly unwanted side effects for the delivery of plasmids expressing therapeutic proteins, but may be of value as adjuvants for vaccine applications. In addition, the inflammatory responses have been proposed to lead to short-lived transgene expression caused by the presumed necrosis- or apoptosis-mediated cell death of transduced cells mediated by a cytotoxic immune response (55–58). However, recent studies point out that the inflammatory response may not be the main mechanism of transgene silencing by bacterial backbone DNA (22, 59, 60). Chen and coworkers have demonstrated that the covalent linkage of bacterial backbone DNA to a eukaryotic expression cassette leads to transgene silencing in vivo, whereas the co-application of bacterial backbone DNA not covalently connected to the expression cassettes had only a minor effect on transgene expression (22, 28). Further, they were able to demonstrate that in vivo (i) minicircle DNA, (ii) a mixture of bacterial backbone DNA and expression cassettes both linearized, and (iii) parental plasmid including the bacterial backbone DNA, each containing the same number of expression cassettes, persisted in transfected cell for an equal time without the loss of cells harboring bacterial DNA (28). Likewise, Suzuki and coworkers reported that they detected the same amount of parental vectors containing a bacterial backbone and minicircle vectors in infected cells, but detected much lower gene expression levels when using the parental vector (60). Thus, the elimination of transduced cells by a cytotoxic response does not seem to be the likely mechanism of transgene silencing. The silencing of viral promoters has also been proposed to be the consequence of an inflammatory response not directly linked to a destruction of transduced cells. The production of IFNγ and other cytokines was thought to directly inhibit transcription driven by viral

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promoters (61, 62). However, Chen et al. (2004) tested three different viral and four different mammalian promoters, which were silenced in the mouse liver only when covalently connected with the expression cassette. Taken together, the results indicated that gene silencing is not a consequence of direct inhibition of the promoters, but a result of events that first occur at the bacterial backbone sequences and then spread to the promoter structures of the expression cassette. Therefore, the molecular mechanisms of transcriptional silencing driven by bacterial sequences in mammalian cells were investigated (59, 60). Transcription of genes in eukaryotic organisms is highly dependent on the higher-order chromatin structure (63, 64). The basic units of chromatin are histones, which associate with DNA to form two main structures, whereby (1) euchromatin is accessible to DNA binding proteins, which are necessary for a variety of fundamental nuclear processes including transcription, and (2) heterochromatin is densely packed and mostly inaccessible for trans-acting factors and transcriptionally silent (64). The hyperacetylation or methylation pattern of core histones, especially H3 and H4, can be used to distinguish these two chromatin forms. Whereas the methylation of H3 at lysine 4 (H3K4me) is involved in euchromatin formation and has been shown to recruit chromatin-remodeling enzymes that lead to a relaxed chromatin structure, the methylation of the same core histone at lysine 9 (H3K9me) is associated with heterochromatic structures and the recruitment of the heterochromatin protein HP1 leading to gene silencing (64). It has been shown that bacterial sequences like, e.g., the ampicillin-resistance gene, were associated with a high ratio of H3K9me to H3K4me (heterochromatin indicator) immediately after transfection, whereas the eukaryotic elements vice versa showed much lower ratios (euchromatin indicator) (60). While the ratio of H3K9me to H3K4me was persisting over time in the bacterial sequences, the H3K9me to H3K4me ratio in the eukaryotic sequence elements increased gradually until it reached the value observed for the bacterial sequences (60). These data indicated that the bacterial sequences are rapidly recognized by the mammalian cells and packed into a heterochromatic structure being immediately silenced. Then the inactive chromatin is hypothesized to spread into the surrounding eukaryotic sequence elements leading to the downregulation of the eukaryotic promoter. Riu and coworkers investigated other euchromatin- and heterochromatin-associated histone methylation patterns and proteins involved in heterochromatin formation accompanying histone structures complexed with minicircles and parental plasmids, respectively (59). In agreement with the data presented by Suzuki and coworkers, silencing of the transgene from the parental plasmids is accompanied by an increase


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in heterochromatin-associated factors and methylation patterns, whereas the pattern of histone modifications associated with minicircles devoid of surrounding bacterial sequences was consistent with an euchromatin structure (59, 60). These data comply with the observation that the bacterial backbone sequence has to be covalently bound to the transgene expression cassette to silence gene expression because the heterochromatin structure can spread from the bacterial sequences into the eukaryotic expression cassette (22, 28). Which factors or patterns actually are involved in the remodeling of chromatin to turn euchromatin into a heterochromatin structure at bacterial sequences is, so far, unknown. However, many different backbone elements were investigated for their ability to diminish or abolish transcription (22). It may be concluded that the silencing mechanism is not sequence specific, but inherent to prokaryotic DNA, thus suggesting that the CpG islands common to all tested backbone elements are a logical candidate factor for transgene silencing (22, 59). However, other typical prokaryotic sequence elements (e.g., characteristic GC to AT ratios) may also be involved in the recognition of prokaryotic DNA in mammalian cells inducing a defense mechanism that results in heterochromatin formation (59, 65).

6. Conclusion Minicircles are supercoiled minimized plasmids devoid of bacterial backbone sequences and encoding only the therapeutic gene of interest together with regulatory sequences. They have been shown in various in vitro and in vivo studies to greatly improve transgene expression, both in terms of expressed levels and persistent expression. The elimination of the bacterial backbone results in the reduction or even elimination of immunostimulatory CpG motifs, which are clearly unwanted for therapeutic gene delivery in patients. Furthermore, minicircles are devoid of sequences attributed with safety concerns such as antibioticresistance genes. Recently it has been proposed that bacterial backbone sequences silence transgene expression, therefore accounting for the unsatisfactory transgene expression levels of conventional plasmid DNA. Minicircles lacking these undesired sequences therefore offer a new concept in highly efficient and safe nonviral gene transfer. New approaches are under way to efficiently produce and purify minicircles at large scale, which will be a prerequisite for their future application in clinical studies.

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Chapter 5 Transposable Elements as Plasmid-Based Vectors for Long-Term Gene Transfer into Tumors John R. Ohlfest, Zoltán Ivics, and Zsuzsanna Izsvák Summary A primary limitation to using nonviral vectors for cancer gene therapy is transient expression of the therapeutic gene. Even when the ultimate goal is tumor cell death, a minimum threshold of gene expression is required to kill tumor cells by direct or indirect mechanisms. It has been shown that transposable elements can significantly enhance the duration of gene expression when plasmid DNA vectors are used to transfect tumor or tumor-associated stroma. Much like a retrovirus, transposon-based plasmid vectors achieve integration into the genome, and thereby sustain transgene expression, which is especially important in actively mitotic cells such as tumor cells. Herein we briefly discuss the different transposons available for gene therapy applications, and provide a detailed protocol for nonviral transposon-based gene delivery to solid experimental tumors in mice. Key words: Antiangiogenic, cancer gene therapy, convection-enhanced delivery (CED), glioma, glioblastoma, immunotherapy, interferon, nonviral vectors, polyethylenimine (PEI), Sleeping Beauty, transposon.

1. Introduction 1.1. Transposable Elements as Nonviral Delivery Vectors in Gene Therapy

Currently, both viral and nonviral methods are used for therapeutic gene delivery, including cancer therapy. The widely used retroviral and lentiviral vectors are efficient in stable gene transfer, but their capacity for cargo is limited (1), their stability/quality may be compromised by incorporation of mutations by reverse transcriptase (2), and their use raises serious safety concerns. Preferential integration of retroviral and lentiviral vectors into expressed genes (3, 4) poses the risk of inadvertent oncogene activation and resulting development of cancer (5). In contrast to integrating


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viral vectors, unintegrating viruses, such as adenovirus, may need to be repeatedly administered in vivo to achieve sustained expression (especially in mitotic cells, where extrachromosomal viral genomes may be lost). However, repeated administration might provoke an undesired immune response. A further problem with viral approaches is that vector production is laborious and expensive. Nonviral vectors are usually less limited in terms of cargo, their large-scale manufacture is simple, and they are considerably safer than viral approaches. However, their greatest limitation in clinical gene transfer is their relative inefficiency in providing long-term therapeutic transgene expression. Nonetheless, if the same clinical efficacy could be achieved by using nonviral therapy as with a viral vector, nonviral vectors might be preferentially used. The establishment of nonviral, plasmid-based, integrating vectors based on naturally occurring and engineered transposable elements generated considerable interest in developing them as efficient and safe vectors for human gene therapy (6, 7). Transposable elements can be considered as natural, nonviral delivery vehicles, capable of efficient genomic insertion. Thus, transposons can address one of the main problems of nonviral technologies: stable genomic integration provides long-term expression of therapeutic genes. However, nonviral gene delivery approaches still have problems concerning efficacy. Since the resurrection of Sleeping Beauty, the first transposable element shown to be active in mammalian cells, other alternative systems appeared on the palette of available tools capable of gene insertion in human cells. The toolbox of currently available transposons include Sleeping Beauty (8), Frog Prince (FP) (9), mariner and Minos from the Tc1/mariner family (reviewed in (6, 7)), piggyBac (10), and Tol2 (11), representatives of two other transposon families. In addition, PhiC31 (12), a site-specific recombinase system has been tested as a delivery vector of therapeutic genes. All of the transposons mentioned above have a simple structure and require only a single protein, a transposase/ recombinase, to catalyze integration. In its natural form, a transposon typically consists of a gene encoding the transposase, which is flanked by terminal inverted repeats (IRs) containing binding sites for the transposase. Importantly, the transposase gene can be physically separated from the IRs and the transposase can be supplied to initiate the transposition reaction in trans. Thus, in principle, the transposase can mobilize any sequence (e.g., a therapeutic gene), as long as it retains the IRs (Fig. 1). The transposase gene can be located on the same DNA molecule as the transposon (one-component system), or supplied on another DNA molecule (two-component system) (Fig. 2), or supplied in the form of messenger RNA (mRNA). In one-component systems, only one DNA molecule should be delivered, whereas in the two-component systems, to ensure transposon-mediated

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Fig. 1. Mechanism of SB-mediated chromosomal integration. SB transposons are flanked by terminal inverted repeat (IR) elements that serve as transposase binding sites and mark the end of the integrating unit. The target TA dinucleotide where integrations occurs is duplicated upon integration.

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Fig. 2. Suggested plasmid DNA vector configurations. The gene encoding the Sleeping Beauty (SB) transposase enzyme to facilitate integration can be supplied on a separate plasmid (trans) or in the plasmid backbone (cis). For trans delivery, a cytomegalovirus (CMV) promoter is used to regulate SB expression and expression of the reporter gene/therapeutic gene. For cis delivery, use of a weaker promoter such as phosphoglycerate kinase (PGK) to regulate SB expression to avoid overexpression inhibition is recommended.

integration, both molecules should be present in the same cell. Both vector systems have to be optimized for efficient cellular delivery and genomic insertion in different tissues (13). Proof-of-principle studies to use transposons in gene therapy are documented for Sleeping Beauty (14), PhiC31 (15), and Tol2 (11), and Sleeping Beauty was demonstrated to be a promising vector in cancer gene therapy in several glioblastoma models

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(16, 17). Importantly, transposons do not replicate themselves, and they can only be mobilized if the transposase is present. One of the key parameters concerning the usefulness of these elements is their overall activity. Importantly, hyperactive Sleeping Beauty systems (unpublished), piggyBac (18), and Tol2 (11) have integration frequencies approaching the efficacy of viral vectors. Because of their versatility and simplicity, transposons have the potential to be the “next-generation” delivery vectors in gene therapy. Learning the lessons from the viral approaches, each of the transposon vector systems should be carefully evaluated for efficacy, safety, and therapeutic potential. Vectors that are efficient at integrating foreign DNA into chromosomes have good potential for providing lifelong gene expression. However, the integration profile of retroviral vectors raises a serious safety issue. To minimize the risk of oncogenesis after random vector integration, a technology had been developed that allows for site-specific integration into the target cell genome based on the phage PhiC31 integrase (12). Unfortunately, genome stability is still a concern with PhiC31, because its activity was clearly associated with recombination events between and within chromosomes (19, 20). From the transposon vector systems discussed above, the integration profile of Sleeping Beauty is closest to random (21, 22) and, in contrast to retroviral vectors, it does not prefer to integrate into transcriptionally active regions in the genome. Similarly to retroviruses, although to a lesser degree, piggyBac also shows nonrandom integration site distribution in human cells, including a higher preference for integrations in regions surrounding transcriptional start sites and within long terminal repeat elements (10). To minimize the risk of oncogenesis after random insertional integration, transcriptional shielding of transposon-borne transgene cassettes as well as targeting integration to specific locations in the human genome is desirable. Indeed, it was recently shown that incorporation of chromatin insulators into Sleeping Beauty vectors significantly diminished their ability to transcriptionally transactivate nearby promoters (23). Furthermore, to improve biosafety of integrating transposon vectors, proof-of-concept studies of modifying the integration profile of Sleeping Beauty (24, 25) and piggyBac (26) transposons have been performed. These studies demonstrated the potential feasibility to target transposon integration to relatively “safe” locations in the genome. A further concern of using certain transposon vectors can be a potential interference with endogenous elements in the human genome. Thus, in principle, an exogenous source of transposase could mobilize some of the endogenous human transposons, possessing high similarity to the vector. Sequences similar to mariner or piggyBac (derived from fly and moth genomes, respectively) are well represented in the human genome (27, 28). Despite the

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huge evolutionary distances, the possibility of cross-mobilization as a potential safety issue needs to be addressed before any clinical application. Advantageously, Sleeping Beauty and Frog Prince (originated from fish and frog genomes, respectively) have no similarity to any endogenous element in the human genome. 1.2. Background on the Use of Transposons for Cancer Gene Therapy

The absolute level and duration of transgene expression are important factors that influence the efficacy of cancer gene therapy (reviewed in (29)). For this reason, viral vectors have been preferentially used in cancer gene therapy applications because they have historically obtained superior gene transfer to tumor cells relative to nonviral vectors. There are many reasons to account for the short-lived expression of plasmid DNA, including degradation after entering the cell and loss of the extrachromosomal plasmid upon cell division. Cancer gene therapy strategies such as antiangiogenesis and interfering RNA vectors, however, will theoretically have less therapeutic impact if gene expression is transient because these approaches rely on sustained inhibition of their target(s) (30). Transposon-based plasmid gene transfer has shown superior therapeutic efficacy relative to the identical gene delivered on a standard plasmid DNA molecule. Sleeping Beauty (SB)-mediated transposition (Fig. 1) was found to be required for long-term expression (1 month or longer) of a luciferase reporter gene after in vivo transfection of U87 glioma tumors in mice. Without transposition, detectable gene expression persisted for less than 15 days (17). Moreover, chromosomal integration of the transposon in vivo was documented by cloning the insertion sites from the glioma cell genomic DNA. No insertions were recovered after standard plasmid gene transfer (17). In a therapeutic application, SB-mediated plasmid gene transfer of antiangiogenic genes significantly inhibited tumor growth, and extended survival in a mouse model of high-grade glioma, whereas the nonintegrating plasmids had only marginal effects (30). In another study, it was shown that SB-mediated delivery of the interferon gamma (IFN-γ) gene inhibited glioma growth by achieving sustained expression. The standard IFN-γ plasmid molecule had no therapeutic effect and gene expression was undetectable at 2 weeks (16). Taken together, these studies have demonstrated that transposon-mediated integration from a plasmid DNA molecule significantly increases the efficacy of cancer gene therapy. In all of the aforementioned studies (16, 17, 30), transfection of plasmid DNA into experimental tumors was achieved by intratumoral injection of polyethylenimine (PEI)/DNA complexes. Therefore, this method to achieve gene transfer will be described in this chapter (PEI/DNA complex administration has also been described in dogs and in humans (31, 32)). Nonetheless, alternative nonviral transfection reagents could

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be used, including intravenous injection and tumor-targeted transfection complexes. The essential requirement for effective transposon-based cancer gene therapy is to express the transposase enzyme in the same cells transfected with the transposon DNA substrate, by whatever means necessary (see Note 2 if long-term expression is not achieved). Accordingly, even viral vectors have been used to deliver the SB transposon components in vivo (33, 34).

2. Materials 2.1. Transfection of Subcutaneous Murine Tumors

The materials listed below are sufficient to transfect ten subcutaneous tumors of 5–10 mm in diameter growing in adult mice with 10 μg of total plasmid DNA vector(s). For larger numbers of mice or greater amounts of DNA, the materials should be increased accordingly. Co-transfection can be achieved as long as the total amount of DNA and PEI remains proportional (three plasmids at 3.3 μg/plasmid could be transfected instead of a single plasmid DNA vector at 10 μg dose). All materials listed below should be sterile. 1. 14 μL of 150 mM linear 22-kDa polyethylenimine (PEI; can be purchased from several commercial sources). 2. 100 μg of endotoxin-free/low plasmid DNA. 3. 500 μL of 5% glucose. 4. 1.5-mL microcentrifuge tubes. 5. Microcentrifuge. 6. Ten ½-cc syringes fitted with 28- or 30-gauge needles.

3. Methods 3.1. Transfection of Subcutaneous Murine Tumors

This method is appropriate to transfect ten subcutaneous tumors by intratumorally injecting a 50 μL volume of DNA/PEI complexes into each tumor. An efficient way to prepare these complexes is to generate a “master mix” solution with a final volume of 500 μL. This can be accomplished as follows (see Note 1 if the transfection fails).

3.1.1. PEI/DNA Complex Preparation

1. Label two microcentrifuge tubes as “DNA” and “PEI.” 2. Pipette 100 μg of plasmid DNA into the “DNA” tube.


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3. Pipette the amount of 5% glucose needed to bring the final volume of solution in “DNA” tube to 250 μL (if the plasmid concentration was 1 μg/μL, you would have added 100 μL of plasmid and 150 μL of glucose). 4. Pipette 14 μL of PEI and 236 μL of 5% glucose into the “PEI” tube. 5. Vortex or shake both tubes briefly (make sure the solutions are well mixed). 6. Centrifuge for 10 s at 1,000g (to ensure all the solution is at the bottom of the tubes after vortexing). 7. Pipette the entire 250 μL solution in the “PEI” tube into the “DNA” tube (important: do not reverse the order, always add PEI to DNA). 8. Vortex and centrifuge again as in steps 5 and 6. 9. Incubate the complexes for 15 min, they are then ready to be injected. 3.1.2. Intratumoral Injection

Intratumoral injection is a simple method to directly target the tumor and stromal cells for transfection. This basic procedure may require modification as described in the notes.

3.1.3. Anesthetize Mice

Sterilize the skin covering the tumor area with an appropriate method (it is also recommended to remove the hair above the tumor with a shaver). Tumors between 5 and 10 mm in diameter are typically good candidates for this procedure, so inspect the tumors and arrange the mice in groups containing similar sizes and appearances of the tumors for best results (larger tumors with necrotic cores may be harder to transfect). Load 50 μL of the DNA/PEI complex master mix into a ½-cc syringe and grasp the tumor gently with your fingers or tweezers (whatever gives you better control). Ideally, you will inject four different locations (with an even distance in between each injection) in the tumor mass to achieve wider transfection (12–13 μL/site). Inject the tumor in two to four quadrants by making only a single needle puncture through the skin aimed at the center of the tumor (Fig. 3a). Specifically, inject 12–25 μL per site into the tumor center slowly over 30–60 s, then withdraw the needle slightly and redirect it into a different part of the tumor and inject another 10–25 μL. Do not completely remove the needle from the skin and create a second puncture, because this will create a reflux point when the next portion of the tumor is injected. After the entire 50 μL has been delivered, leave the needle in the tumor mass for at least 30 s before slowly withdrawing it to avoid reflux. Measure reporter gene expression at the desired time points (see ref. (17) for

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a

Single Needle Puncture Through Skin

Intratumoral Needle Position(s)

4

1

2

3

Tumor Mass

b

Tubing Needle / Catheter

Micropump

Tumor Mass

Fig. 3. Intratumoral injection schematic. (a) Intratumoral bolus injection with a syringe can be accomplished by pulling the needle back (but not out of the skin) and redirecting the needle tip to inject different portions of the tumor mass. (b) Convection-enhanced delivery is an alternative method to bolus injection but requires a micropump and tubing.

examples of useful reporter assays for transposition). If no therapeutic effect is observed, consider the remarks in Note 3.

4. Notes 1. Poor efficiency of initial gene transfer: Many important factors influence the transfection of tumor cells using intratumoral injection. These include the DNA quality, charge of the DNA/PEI complex, variations in tumor necrosis and intestinal pressure, tumor size, injection method, and cell types that comprise the tumor. Always use high-quality DNA with an OD 260/280 ratio of 1.8 or greater and that is free of endotoxin.


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If you encounter significant reflux using bolus injection or experience very high resistance during the injection, you may consider using convection-enhanced delivery (CED). CED is a slow, continuous delivery of infusate that establishes a pressure gradient in the tumor to achieve a more widespread distribution (reviewed in (35)). For instance, you could attach the syringe to an electronic micropump and deliver the infusate as shown in Fig. 3b at a flow rate of 1–5 μL/min (see ref. (30) for detailed protocol). If you observe that a white precipitate is formed when DNA and PEI are mixed, this can be addressed by lowering the final concentration of DNA (it is not recommended to exceed 0.5 μg/μL). You may also reduce precipitation by reducing the amount of salt that is delivered with DNA (go from 1X Tris/ EDTA [TE] buffer to 0.1X, or DNA in water may be best). In addition to precipitation, the ratio of nitrogen atoms in the PEI to phosphate in the DNA backbone (e.g., the N/P ratio) influences the net complex charge and can be optimized. In the protocol described, the N/P ratio is 7, which is typically a good place to start for in vivo testing. The following formula can be used to determine the N/P ratio for empirical optimization: μl of150 mM linear 22 kDa PEI =

(μg of DNA × 3) × N/P ratio 150

2. Good initial gene transfer but not long-term expression: This problem may occur if the transfected cells die or because of lack of transposition, only the later will be addressed here. For certain transposon systems, a phenomenon termed overexpression inhibition can occur wherein too much transposase enzyme can reduce transposition rates. This is true of SB, for instance (7). This problem can be avoided by using weaker genetic promoters to regulate SB expression if the SB gene is delivered in cis (one-component system), or reducing the amount of SB-encoding plasmid delivered in trans (twocomponent system). A second consideration when delivering the SB gene in cis is the orientation of the SB expression cassette relative to the expression cassette within the transposon IR/direct repeats (DRs). If the two expression cassettes are transcribed in an opposing fashion, there is a potential for several problems including “collision” of transcriptional machinery and interfering RNA effects. This configuration is not recommended. In addition, the number of base pairs that separate the genetic promoter that regulates SB expression from the IR/DR where SB must bind to facilitate transposition is probably important. The best results have been obtained in our laboratory when the distance from any portion of transcribed DNA relative to the IR/DR is at least 200 bp. The recommended

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starting conditions using SB are as follows: for trans delivery of SB, use a 1:20 ratio of a cytomegalovirus (CMV)-regulated SBencoding plasmid to transposon substrate. For cis delivery, use a weak promoter such as phosphoglycerate kinase (PGK) (with cis configuration, every cell has a chance to undergo transposition). The suggested vector configurations are shown in Fig. 2. 3. Good gene transfer, long-term expression, but not therapeutic effect: In this situation, it is important to consider the mechanisms and theoretical appeal of the therapeutic gene product being expressed in the tumor. For instance, if the gene product is directly and rapidly cytotoxic to the tumor cells, you probably will not obtain any therapeutic benefit from SB-mediated integration and long-term expression because the transfected cells die before this could occur. If using this approach, you may also wish to conduct the experiments using murine tumors in immunocompetent mice, because a secondary antitumor immune response may occur after the initial tumor cell death. In cancer gene therapy it is difficult to transfect the majority of tumor cells, and so the use of therapeutic genes that are secreted or prime an immune response is strongly recommended to achieve best results. Top candidate genes would be those that are secreted, potent inhibitors of angiogenesis and/or immunostimulatory cytokines. A related class of genes that has great theoretical appeal is co-stimulatory molecules, which are often transmembrane proteins, but facilitate the priming of an innate or adaptive antitumor immune response via direct interaction with tumor-infiltrating leukocytes. These types of transgene are more likely to maximize the therapeutic effect of long-term expression because of sustained inhibition of angiogenesis or enhanced duration of immune stimulation relative to conventional plasmid vectors (reviewed in (29)).

References 1. Lundstrom, K. (2003) Latest development in viral vectors for gene therapy. Trends Biotechnol 21, 117–22. 2. Pathak, V.K. and H.M. Temin (1990) Broad spectrum of in vivo forward mutations, hypermutations, and mutational hotspots in a retroviral shuttle vector after a single replication cycle: substitutions, frameshifts, and hypermutations. Proc Natl Acad Sci USA 87, 6019–23.

3. Schroder, A.R., P. Shinn, H. Chen, C. Berry, J.R. Ecker, and F. Bushman (2002) HIV-1 integration in the human genome favors active genes and local hotspots. Cell 110, 521–9. 4. Bushman, F.D. (2003) Targeting survival: integration site selection by retroviruses and LTR-retrotransposons. Cell 115, 135–8. 5. Hacein-Bey-Abina, S., C. Von Kalle, M. Schmidt, et al. (2003) LMO2-associated clonal


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19. Liu, J., I. Jeppesen, K. Nielsen, and T.G. Jensen (2006) Phi c31 integrase induces chromosomal aberrations in primary human fibroblasts. Gene Ther 13, 1188–90. 20. Ehrhardt, A., J.A. Engler, H. Xu, A.M. Cherry, and M.A. Kay (2006) Molecular analysis of chromosomal rearrangements in mammalian cells after phiC31-mediated integration. Hum Gene Ther 17, 1077–94. 21. Vigdal, T.J., C.D. Kaufman, Z. Izsvak, D.F. Voytas, and Z. Ivics (2002) Common physical properties of DNA affecting target site selection of sleeping beauty and other Tc1/mariner transposable elements. J Mol Biol 323, 441–52. 22. Yant, S.R., X. Wu, Y. Huang, B. Garrison, S.M. Burgess, and M.A. Kay (2005) Highresolution genome-wide mapping of transposon integration in mammals. Mol Cell Biol 25, 2085–94. 23. Walisko, O., A. Schorn, F. Rolfs, A. Devaraj, C. Miskey, Z. Izsvak, and Z. Ivics (2008) Transcriptional activities of the Sleeping Beauty transposon and shielding its genetic cargo with insulators. Mol Ther 16, 359–69. 24. Yant, S.R., Y. Huang, B. Akache, and M.A. Kay (2007) Site-directed transposon integration in human cells. Nucleic Acids Res 35, e50. 25. Ivics, Z., A. Katzer, E.E. Stuwe, D. Fiedler, S. Knespel, and Z. Izsvak (2007) Targeted Sleeping Beauty transposition in human cells. Mol Ther 15, 1137–44. 26. Maragathavally, K.J., J.M. Kaminski, and C.J. Coates (2006) Chimeric Mos1 and piggyBac transposases result in site-directed integration. FASEB J 20, 1880–2. 27. Laha, T., A. Loukas, S. Wattanasatitarpa, et al. (2007) The bandit, a new DNA transposon from a hookworm-possible horizontal genetic transfer between host and parasite. PLoS Negl Trop Dis 1, e35. 28. Lander, E.S., L.M. Linton, B. Birren, et al. (2001) Initial sequencing and analysis of the human genome. Nature 409, 860–921. 29. Ohlfest, J.R., A.B. Freese, and D.A. Largaespada (2005) Nonviral vectors for cancer gene therapy: prospects for integrating vectors and combination therapies. Curr Gene Ther 5, 629–41. 30. Ohlfest, J.R., Z.L. Demorest, Y. Motooka, et al. (2005) Combinatorial antiangiogenic gene therapy by nonviral gene transfer using the sleeping beauty transposon causes tumor regression and improves survival in mice bearing intracranial human glioblastoma. Mol Ther 12, 778–88. 31. Oh, S., G.E. Pluhar, E.A. McNeil, et al. (2007) Efficacy of nonviral gene transfer in the canine brain. J Neurosurg 107, 136–44.

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32. Ohana, P., P. Schachter, B. Ayesh, et al. (2005) Regulatory sequences of H19 and IGF2 genes in DNA-based therapy of colorectal rat liver metastases. J Gene Med 7, 366–74. 33. Yant, S.R., A. Ehrhardt, J.G. Mikkelsen, L. Meuse, T. Pham, and M.A. Kay (2002) Transposition from a gutless adeno-transposon vector stabilizes transgene expression in vivo. Nat Biotechnol 20, 999–1005. 34. Bowers, W.J., M.A. Mastrangelo, D.F. Howard, H.A. Southerland, K.A. Maguire-

Zeiss, and H.J. Federoff (2006) Neuronal precursor-restricted transduction via in utero CNS gene delivery of a novel bipartite HSV amplicon/transposase hybrid vector. Mol Ther 13, 580–8. 35. Raghavan , R. , M.L. Brady, M.I. RodriguezPonce , A. Hartlep , C. Pedain , and J.H. Sampson (2006) Convection-enhanced delivery of therapeutics for brain disease, and its optimization . Neurosurg Focus 20 , E12 .

Chapter 6 Designing Plasmid Vectors Oleg Tolmachov Summary Nonviral gene therapy vectors are commonly based on recombinant bacterial plasmids or their derivatives. The plasmids are propagated in bacteria, so, in addition to their therapeutic cargo, they necessarily contain a bacterial replication origin and a selection marker, usually a gene conferring antibiotic resistance. Structural and maintenance plasmid stability in bacteria is required for the plasmid DNA production and can be achieved by carefully choosing a combination of the therapeutic DNA sequences, replication origin, selection marker, and bacterial strain. The use of appropriate promoters, other regulatory elements, and mammalian maintenance devices ensures that the therapeutic gene or genes are adequately expressed in target human cells. Optimal immune response to the plasmid vectors can be modulated via inclusion or exclusion of DNA sequences containing immunostimulatory CpG sequence motifs. DNA fragments facilitating construction of plasmid vectors should also be considered for inclusion in the design of plasmid vectors. Techniques relying on site-specific or homologous recombination are preferred for construction of large plasmids (>15 kb), while digestion of DNA by restriction enzymes with subsequent ligation of the resulting DNA fragments continues to be the mainstream approach for generation of small- and medium-size plasmids. Rapid selection of a desired recombinant plasmid against a background of other plasmids continues to be a challenge. In this chapter, the emphasis is placed on efficient and flexible versions of DNA cloning protocols using selection of recombinant plasmids by restriction endonucleases directly in the ligation mixture. Key words: Longevity of gene expression, modulation of immune response in cancer gene therapy, plasmid manipulation, plasmid stability, restriction endonuclease digestion of ligation mixture, transgene expression.

1. Introduction Nonviral gene therapy protocols often use plasmid DNA or its artificial derivatives for gene delivery. In addition to their therapeutic DNA load, these plasmids necessarily contain a bacterial replication origin and a selection marker, usually a gene conferring Wolfgang Walther and Ulrike S. Stein (eds.), Methods in Molecular Biology, Gene Therapy of Cancer, vol. 542 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-561-9_6

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antibiotic resistance. Adequate expression of the therapeutic gene or genes is ensured by the use of appropriate promoters and other regulatory elements. In cancer gene therapy, the transient expression of a therapeutic gene is often sufficient to trigger the destruction of malignant cells. However, if long-term expression is desired, the plasmid vectors should include genetic elements for either episomal maintenance or chromosomal integration of the transgene expression cassette. Recombinant plasmids are assembled via molecular cloning in bacteria. Cloning protocols for construction of recombinant plasmids are presently so abundant that it is often difficult to choose a version that is the most appropriate for a particular constellation of the DNA cloning circumstances. Common difficulties include selection of a desired recombinant plasmid against a background of other plasmids and structural/maintenance plasmid instability. Techniques relying on site-specific or homologous recombinations are preferred for construction of large plasmids (>15 kb), whereas digestion of DNA by restriction enzymes with subsequent ligation of the resulting DNA fragments continues to be the mainstream approach for generation of small- and medium-size plasmids. In this chapter, the emphasis is placed on efficient and flexible versions of DNA cloning strategies using direct selection of recombinant plasmids by restriction endonucleases in the ligation mixture.

2. Choice of a Bacterial Plasmid Replicon

The standard bacterial replicon used in artificial plasmids is the ColE1-type origin of replication from the plasmid pMB1, which evolved via plasmids pBR322 and pUC8/9 (1–3). This replication origin is associated with multicopy plasmid lifestyle, where plasmids are distributed fairly randomly during cell division. Multicopy status ensures high plasmid DNA yield, which is often desirable. In fact, the majority of modern plasmid vectors contain a mutated version of the pMB1 replicon, providing higher than wild-type plasmid copy number. However, it is not uncommon for a high copy number status to exacerbate a detrimental effect of certain plasmid elements on the bacterial growth, resulting in selective advantage of spontaneous plasmid deletion mutants, which manifests itself as structural plasmid instability. In addition, random, uncontrolled distribution of plasmids during bacterial cell division produces plasmidless cells, which, given some selective advantage, can outgrow cells with plasmids. Both structural and maintenance instability of recombinant plasmids bearing therapeutic gene modules often interfere with plasmid propagation in bacteria. Thus, plasmid replicons alternative to pMB1 that

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possess efficient partitioning mechanisms should often be considered in plasmid design. An array of stable replicons was used for propagation of recombinant plasmids. The most well known replicons are derived from plasmids R6K, RSF1010, F-factor, and bacteriophage P1 (4–6). The stable replicons tend to be the low copy number replicons, which give a low yield of plasmid DNA. In fact, both partitioning mechanisms and a low copy number mode of replication contribute to structural and maintenance stability.

3. Choice of a Plasmid Marker A bacterial plasmid selection marker is a necessary tool to control the maintenance of plasmids by elimination of plasmidless cells. Traditionally, antibiotic-resistance genes are used as bacterial selection markers. These genes can be large (e.g. the tetracyclineresistance cassette includes the 1,206-bp gene tetA and the 624-bp gene tetR) or short (e.g. the blasticidin-resistance gene bsr is 429 bp). The size of the bacterial marker gene can be used to modulate the CpG-motif load of the plasmid vector. A common marker is the ampicillin-resistance gene coding for β-lactamase. When culturing ampicillin-resistant bacteria in liquid culture, it is important to realize that secreted β-lactamase destroys added ampicillin within several hours and subsequent bacterial growth occurs without selection, with ensuing reduction of the DNA yield of maintenance instability-prone plasmids. Genes for resistance to the ionic heavy metals such as mercury (Hg2+) can be used as an alternative to antibiotic-resistance markers. This type of bacterial resistance is normally determined not by a single gene but by a gene cluster, which can increase the plasmid vector size and its CpG-motif load. However, if a minimal size of the plasmid selection gene is required, a combination of plasmidborne suppressor genes and amber, ochre, or opal mutated chromosome-resident resistance genes looks particularly attractive as a plasmid selection system. The plasmid-borne tyrosine ambersuppressor gene supF (103 bp in size) was previously successfully used for selection of plasmid–bacteriophage recombinants (7).

4. Choice of Transgene Promoter, Polyadenylation Site, and Expression Enhancements Elements

The level of transgene expression is very important in gene therapy. Most often, the transgene expression needs to be maximized. An appropriate transcription promoter and a polyadenilation site

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are strictly necessary elements of the standard eukaryotic transcription unit served by RNA polymerase II. Although the promoter is a directional element, polyadenilation sites are known to function in both alternative orientations. Promoters vary in their strength and ability to maintain durable expression in vivo. As a rule of thumb, strong and ubiquitous promoters (e.g. viral cytomegalovirus [CMV] promoter) have a tendency to shut down early, whereas weaker tissue-specific promoters can provide longer expression. Nearly all actively translated messenger RNA (mRNA) are known to contain a sequence similar to 5′-GCCACC-3′ immediately before the translation start. This sequence is called a “Kozak consensus sequence” and is known to boost mRNA translation. Thus, a Kozak consensus sequence or its extended version 5′-GCCGCCACC-3′ should be incorporated immediately before the start codon of the therapeutic transgene if a suitable sequence is not already present. Transcriptional enhancers are often used to enhance the therapeutic transgene expression. Many of them are tissue-specific and can be exploited to drive predominantly tumour-specific gene expression. The so-called post-translational enhancer element from Woodchuck Hepatitis Virus (WPRE) can be used to improve stability of the transgenic mRNA and thus to increase longevity of the transgene expression. This element can be placed between the coding sequence and the polyadenilation sequence. However, the potential oncogenic activity of WPRE should also be considered (8). Evaluation of this activity is complicated by the vastly different tumorigenesis mechanisms in humans and experimental rodents.

5. Modulation of Immune Response via Vector Design and Use of the Minimized DNA Vectors

DNA extracted from Escherichia coli is immunogenic to humans because of the activation of the innate immune response by the unmethylated 5¢-CG-3¢ sequences within the so-called bacterial CpG sequence motifs. Some immune response to incoming DNA might be beneficial in some forms of cancer gene therapy (9). Clearly, immune response can be enhanced by the increased size of the CpG motif-containing bacterial sequences. Alternatively, if the immune response needs to be minimized, CpG-depleted or CpG-ablated plasmid backbones can be used to decrease the CpG-motif load (10) or bacterial sequences can be removed from


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the plasmid DNA to produce minimized DNA vectors. Known minimized DNA vectors include minicircle DNA vectors (11) and MIDGE vectors.

6. Elements for Mammalian Episomal Maintenance and Chromosomal Integration

7. Choice of Bacterial Strain for Plasmid Manipulation and Propagation

Synthetic plasmid-based vectors can be custom-made to pass a series of barriers presented by target cells and immune system, however, these vectors often provide only transient maintenance of a transgene within a cell (12, 13). Therefore, plasmid gene vectors capable of persistent episomal maintenance or chromosomal integration are preferable in many therapeutic settings. Epstein-Barr virus’s oriP-EBNA1 mammalian episomal replication module is a classic DNA element to consider for inclusion into the therapeutic plasmids. Scaffold/matrix attachment region (S/MAR) is another DNA element that can be included in the plasmid vectors to ensure episomal maintenance (14). A fair frequency of chromosomal integration in target human cells can be achieved by inclusion of attB site for Streptomyces bacteriophage φC31 within therapeutic plasmids and use of site-specific φC31 integrase (15). The artificial mammalian transposon Sleeping Beauty can also be used to achieve stable chromosomal integration (reviewed in Chapter 5).

Bacterial strains used to establish and propagate recombinant plasmids constitute an important choice for the cancer gene therapist. Strains harbouring recA mutations, which result in homologous recombination defect, are known to be beneficial for structural plasmid stability (16). The critical step in an artificial plasmid’s life is when the artificial plasmid is first established after the transformation of bacteria. Recombinant plasmids often exhibit idiosyncratic behaviour during this period, being unequivocally stable in one recA strain and demonstrating marked instability in another recA strain. HB101-based E. coli strain Stbl3TM and JM109-based E. coli strains Stbl2TM and Stbl4TM (Invitrogen, Carlsbad, CA) are often used in situations where recombinant plasmid stability is an issue. A specialized E. coli strain GT116 was developed by InvivoGen (San Diego, CA) to maintain DNA constructs coding for short hairpin RNA (shRNA), a common experimental anticancer agent (17).

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8. Cloning Strategies Based on Bacterial Colony Tests, Positive Selection of Recombinant Plasmids In Vivo, and Directional Insertion of DNA Fragments

The assembly of recombinant plasmids is technically well established. However, the process is marred by multiple pitfalls, which slow down the construction. Plasmid generation mediated by restriction enzymes and T4 DNA ligase continue to be the mainstream approach to plasmid engineering. Generation of DNA fragments by polymerase chain reaction (PCR) or annealing of complementary single-stranded oligonucleotides is also common. In addition to plasmid instability, the difficulties include reduced efficiency of blunt DNA ends’ ligation compared with efficiency of cohesive ends’ ligation and the background of bacterial clones harbouring the insertless self-ligated vector. Phosphatase treatment is commonly used to block self-ligation of vector DNA. However, a trace quantity of incompletely removed phosphatase can dephosphorylate the insert DNA and, therefore, ruin the insert–vector ligation. Thus, calf intestinal phosphatase (CIP, Invitrogen) and Antarctic phosphatase (New England Biolabs, Ipswitch, MA) are preferable to bacterial alkaline phosphatase (BAP, Invitrogen) because of their efficient and complete inactivation by heat. The use of phosphatase can reduce the overall efficiency of cloning because dephosphorylated vector termini have a 50% diminished activity in the intermolecular joining during ligation, and survival of the formed ligation products relies on the repair of their phosphodiester bonds in vivo. This reduction in cloning efficiency is particularly critical in experiments relying on blunt DNA ends’ ligation. Thus, specialized cloning vectors were developed that allow circumvention of the use of phosphatase by simplifying the selection of recombinant plasmid clones with antibiotic-resistance marker inactivation (1, 2), colony colour tests (3, 18), or in vivo positive selection set-ups (19–21). The ability of Vaccinia virus topoisomerase I to mediate extra-efficient DNA ligation reaction is exploited in specialized TOPO® cloning vectors supplied by Invitrogen (22). However, in complex recombinant DNA assemblies, it is not possible to use specialized cloning vectors repeatedly because one round of genetic manipulation inactivates the insert selection machinery of the cloning vector. In such situations, the most powerful screening technique is colony PCR, which can be tailored to identify clones with an insert and/or a specific orientation of the insert (23). Intramolecular ligation can be also blocked by a “directional placing” technique, relying on an insertion using a pair of incompatible cohesive ends that block intramolecular ligation. Directional placing is limited by the choice of restriction sites and occasional insert orientation-dependent instability of recombinant plasmids. In general, all of the above approaches are not always sufficient for rapid selection of insert-containing plasmids.


9. Cloning Strategies Based on Selection of Recombinant Plasmids by Restriction Endonucleases Directly in the Ligation Mixture

9.1. Selection Strategy Based on the Abrogation of a Restriction Site After Intermolecular Ligation of Compatible Cohesive DNA Ends

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A very useful method of positive selection of the desired recombinant plasmids consists of restriction enzyme digestion of the ligation mixture containing wanted and unwanted ligation products. Some particular applications of this approach were published in the technical literature (24–26), however, the general description of the method is presented here for the first time. The tactic exploits the fact that E. coli cells are poorly transformed by linear DNA (27). This allows the inactivation of the self-ligated vector DNA and other unwanted circular plasmid species by their DNA cleavage with an appropriate restriction enzyme. This selection approach requires that the vector, the desired insert DNA, and their junctions do not contain any internal sites for the restriction enzyme used to digest the ligation mixture. However, with complete plasmid DNA sequences presently standardly available, the restriction enzymes required for plasmid selection can be predicted by computer analysis of the sequences and taken into account in plasmid design. Quite often, preliminary insertion of a short linker containing restriction sites tailored for selection of recombinants by restriction digestion can be an attractive strategy expediting the overall DNA manipulation. Considering the not so uncommon difficulties in finding the desired recombinant plasmid clone in the library of bacterial transformants, this selection approach is highly beneficial, resulting in 30–100% of bacterial transformants harbouring the desired recombinant plasmid (O. Tolmachov, unpublished observations). I present here four major variations of this strategy, which may provide a timesaving algorithm for efficient generation of plasmid vectors for cancer gene therapy. Selection strategy 1 exploits the reconstitution of the cloning site in the self-ligated vector DNA and the abrogation of this restriction site in the recombinant plasmid after ligation of the complementary sticky ends produced by certain pairs of restriction endonucleases. For example, if the vector plasmid contains a unique cloning XhoI site and the insert DNA fragment is excisable by SalI with no internal XhoI sites, then the XhoI-digested vector plasmid is first ligated to the SalI-excised insert DNA. After inactivation of ligase, the ligation mixture is digested with XhoI and used to transform competent bacterial cells. Thus, the final digestion with XhoI linearizes only the unwanted circular products of intramolecular ligation of vector DNA containing a reconstituted XhoI site, making them incapable of replication in bacteria. So, only the circular products of intramolecular ligation, which contain the XhoI-uncleavable SalI–XhoI junctions, are selected for establishment in the bacterial cells. This approach

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is very effective because of the high ligation efficiency of cohesive DNA ends and it works very well for the cohesive ends’ families XhoI/SalI, SpeI/XbaI/NheI/AvrII, and BamHI/BglII/BclI. If the insert is generated by PCR, the appropriate flanking recognition sites can be engineered into the PCR product through the PCR primers. If the insert is produced by annealing of homologous oligonucleotides, the end sequences can be designed to create cohesive ends that form uncleavable sites after ligation to the cut vector DNA. 9.2. Selection Strategy Based on the Abrogation of a Restriction Site After Intermolecular Ligation of Blunt DNA Ends Produced by Different Restriction Endonucleases

Selection strategy 2 relies on the reconstitution of the cloning site for a blunt-end-generating restriction enzyme in the self-ligated vector DNA and the abrogation of the sites in vector–insert intermolecular ligation reactions. Selection of the insert-containing plasmids is accomplished by the digestion of the ligation mixture with the blunt-end-producing restriction enzyme that was used initially to cleave the vector plasmid. Again, linearized DNA species are unable to produce plasmids after transformation of bacteria. For example, a SmaI-digested vector is ligated to an EcoRV-excised insert and the resultant ligation mixture is subjected to SmaI treatment to select intermolecular vector–insert ligation products. Selection strategy 2 can be applied quite broadly because there is a wide choice of restriction endonucleases producing blunt DNA ends with all the blunt ends being compatible. Vector linearization with octanucleotide-recognizing enzymes (e.g. SrfI) instead of hexanucleotide recognizing restriction endonucleases increases the spectrum of amenable (e.g. SrfIsite-free) inserts because, on average, a specific sequence of eight nucleotides occurs only once in 65,536 bp of DNA sequence with a random distribution of A, T, C, and G. In addition to digestion by blunt ends producing enzymes, the insert DNA fragments with blunt ends can be prepared by enzymatic blunt ending of endonuclease-produced DNA fragments with cohesive ends (e.g. by Klenow fragment of E. coli polymerase I), annealing of complementary oligonucleotides, filling in of the staggered ends of Taq-PCR amplicons or PCR with blunt-end-producing proofreading polymerases such as Pfx (Invitrogen). A thorough computer sequence analysis must be performed to ensure that the desired recombinant plasmid does not contain recognition sites for the endonuclease that is used to digest the ligation mixture.

9.3. Selection Strategy Based on the Formation of a Re-Cleavable Site After Intramolecular Ligation of Blunt-Ended Cohesive DNA Ends

Selection strategy 3 relies on formation of a re-cleavable site after self-ligation of blunt-ended single-stranded vector overhangs produced by some restriction endonucleases (Table 1). The same restriction site is extremely unlikely to appear at new vector– insert junctions. For example, ligation of filled in HindIII overhangs produces a recognition sequence for NheI that spans both ligated DNA ends and that can be used to linearize unwanted


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Table 1 Candidate restriction endonucleases to digest the ligation mixture produced after ligation of the blunt-ended vector to the insert DNA. The Klenow fragment of Escherichia coli DNA polymerase I can be used to fill in 5’ overhangs or, alternatively, to trim 3’ overhangs. Octanucleotide-recognizing enzymes, which have broader applicability, are shown in bold Enzyme to cleave a unique site in the vector DNA

Type of the procedure to blunt-end the vector termini

Candidate enzymes to digest the ligation mixture

Acc65I

Filling in

SnaBI

AclI

Filling in

MluI

AflII

Filling in

PacI

AgeI

Filling in

EagI

AlwNI

Trimming

PvuII

ApaLI

Filling in

SphI

AscI

Filling in

BssHII

BamHI

Filling in

ClaIa

BclI

Filling in

ClaIa

BglII

Filling in

ClaIa

BsiWI

Filling in

SnaBI

BspEI

Filling in

EagI

BspHI

Filling in

NsiI

BsrGI

Filling in

SnaBI

BssHII

Filling in

BssHII

BstBI

Filling in

NruI

ClaIa

Filling in

NruI

DraIII

Trimming

PmlI

EagI

Filling in

EagI, NgoMIV, FseI

EcoRI

Filling in

AseI, XmnI

HindIII

Filling in

NheI

KasI

Filling in

BssHII

MfeI

Filling in

AseI

MluI

Filling in

BssHII

NarI

Filling in

AscI, BssHII (continued)

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Table 1 (continued) Enzyme to cleave a unique site in the vector DNA

Type of the procedure to blunt-end the vector termini

Candidate enzymes to digest the ligation mixture

NcoI

Filling in

NsiI

NgoMIV

Filling in

EagI, NgoMIV

NheI

Filling in

NheI

NotI

Filling in

EagI, FseI, NgoMIV

PciI

Filling in

NsiI

PspOMI

Filling in

FseI, NgoMIV

PspXI

Filling in

PvuI

SalI

Filling in

PvuI

XhoI or PaeR7I

Filling in

PvuI

XmaI

Filling in

EagI

a

Occasionally the vector DNA should be isolated from Dam− host, such as E. coli DM1 to ensure ClaI cleavage of the Dam-methylated vector DNA. If the insert DNA is extracted from a Dam+ bacterial host, some or all of its ClaI sites could be completely protected by Dam methylation from ClaI cleavage during ClaI digestion of the ligation mixture.

self-ligated vector DNA. As in the selection strategy 2 above, blunt-ended insert fragments can be generated by a variety of techniques. Ligation of the blunt-ended overhangs produced by a number of restriction enzymes generates rare 8-bp-long recognition sites of PacI, FseI, and AscI (Table 1) that are unlikely to be present inside the insert DNA, the new junctions, and the outlying regions of the vector. 9.4. Selection Strategy Using the Inactivation of All ReplicationCompetent DNA Species in the Ligation Mixture by Digestion with Restriction Endonucleases

Often the self-ligated vector is not the only unwanted replicationcompetent DNA species present in the ligation mixture. Multiple background-generating DNA species can arise in plasmid DNA fragment exchange cloning approaches, which are often used in the DNA assembly experiments. A typical background reduction strategy involves mechanical removal of all unwanted DNA fragments before ligation by gel purification of the wanted DNA fragments. DNA fragment isolation is particularly critical when the bacterial selection marker on a competing replicon-containing DNA fragment coincides with the bacterial selection marker of the vector plasmid. However, DNA fragment purification is often cumbersome. In addition, gel extraction can introduce ligase inhibitors and/or damage the DNA ends.


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Selection strategy 4 can circumvent isolation of the vector and insert DNA fragments and, thus, can simplify the cloning experiment. The strategy relies on the presence of certain restriction sites in the unwanted plasmid DNA fragments and their absence in the desired insert, which can be easily established by computer analysis of the plasmid DNA sequences. Digestion of the ligation mixture using carefully selected restriction endonucleases linearizes all the circular DNA species containing the unwanted DNA fragments, thus making them incapable of replication in bacteria. This selection strategy is often applicable in situations where the excised unwanted vector fragment is, in fact, a segment of the multi-cloning site and, so, contains an array of restriction sites that are not found elsewhere in the vector.

10. Cloning Strategies Relying on Site-Specific or Homologous Recombination

DNA cloning relying on site-specific recombination for joining of DNA fragments can be a convenient way to generate plasmid vectors, particularly when repetitive insertions are performed into the same vector (as, e.g., in Gateway® insertion technology by Invitrogen). This technique requires a specialized acceptor plasmid containing recombination site(s) such as the att sites of the bacteriophage λ Int-mediated recombination system or the loxP site(s) of the bacteriophage P1 Cre-mediated recombination system. The important advantage of molecular cloning methods based on site-specific recombination is the ability to manipulate large plasmids (>15 kb), which are difficult to modify by more conventional means. This advantage is shared by the plasmid manipulation protocols, which are based on homologous recombination. Homologous “recombineering” is normally performed in vivo in bacteria using a swarm of DNA structures including a plasmid coding for a recombination function such as RecET, a target plasmid, a donor PCR amplicon, or a donor plasmid (28).

11. Conclusion Four lines of consideration dictate the design of plasmid gene vectors for cancer gene therapy. Firstly, the plasmid vector must contain the DNA elements providing for the structural and maintenance stability of the vector during propagation in bacteria. Secondly, the therapeutic transgene must be efficiently expressed in target cells necessitating inclusion of a number of DNA modules

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within the vector. Thirdly, immune response to the bacterial CpG motifs should be taken into account and the plasmid vector sequences should be chosen to allow modulation of this response to an optimal level for specific cancer therapy settings. Fourthly, use of DNA elements assisting efficient construction of new plasmid gene vectors should be considered. Examples of such construction-facilitating sequences include site-specific recombination sites and polylinker sequences tailored for selection of recombinant plasmids by the restriction endonuclease digestion of the ligation mixture. References 1. Bolivar, F., Rodriguez, R. L., Betlach, M. C., and Boyer, H. W. (1977) Construction and characterization of new cloning vehicles. I. Ampicillin-resistant derivatives of the plasmid pMB9. Gene. 2, 75–93. 2. Bolivar, F., Rodriguez, R. L., Greene, P. J., Betlach, M. C., Heyneker, H. L., and Boyer, H. W. (1977) Construction and characterization of new cloning vehicles. II. A multipurpose cloning system. Gene. 2, 95–113. 3. Vieira, J., and Messing, J. (1982) The pUC plasmids, an M13mp7-derived system for insertion mutagenesis and sequencing with synthetic universal primers. Gene. 19, 259–268. 4. Chistoserdov, A. Y., and Tsygankov, Y. D. (1986) Broad host range vectors derived from an RSF1010::Tn1 plasmid. Plasmid. 16, 161–167. 5. Sarovich, D. S., and Pemberton, J. M. (2007) pPSX: a novel vector for the cloning and heterologous expression of antitumor antibiotic gene clusters. Plasmid. 57, 306–313. 6. Wu, F., Levchenko, I., and Filutowicz, M. (1995) A DNA segment conferring stable maintenance on R6K gamma-origin core replicons. J. Bacteriol. 177, 6338–6345. 7. Seed, B. (1983) Purification of genomic sequences from bacteriophage libraries by recombination and selection in vivo. Nucleic Acids Res. 11, 2427–2445. 8. Kingsman, S. M., Mitrophanous, K., and Olsen, J. C. (2005) Potential oncogene activity of the woodchuck hepatitis post-transcriptional regulatory element (WPRE). Gene Ther. 12, 3–4. 9. Storni, T., Ruedl, C., Schwarz, K., Schwendener, R. A., Renner, W. A., and Bachmann, M. F. (2004) Nonmethylated CG motifs packaged into virus-like particles induce protective cytotoxic T cell responses in the absence of systemic side effects. J. Immunol. 172, 1777–1785. 10. Yew, N. S., Zhao, H., Przybylska, M., Wu, I. H., Tousignant, J. D., Scheule, R. K., et al. (2002)

11.

12.

13.

14.

15.

16.

17.

CpG-depleted plasmid DNA vectors with enhanced safety and long-term gene expression in vivo. Mol. Ther. 5, 731–738. Bigger, B. W., Tolmachov, O., Collombet, J. M., Fragkos, M., Palaszewski, I., and Coutelle, C. (2001) An araC-controlled bacterial cre expression system to produce DNA minicircle vectors for nuclear and mitochondrial gene therapy. J. Biol. Chem. 276, 23018–23027. Vaysse, L., Gregory, L. G., Harbottle, R. P., Perouzel, E., Tolmachov, O., and Coutelle, C. (2006) Nuclear-targeted minicircle to enhance gene transfer with non-viral vectors in vitro and in vivo. J. Gene Med. 8, 754–763. Tolmachov, O., and Coutelle, C. (2007) Covalent attachment of multifunctional chimeric terminal proteins to 5’ DNA ends: a potential new strategy for assembly of synthetic therapeutic gene vectors. Med. Hypotheses. 68, 328–331. Lufino, M. M., Manservigi, R., and WadeMartins, R. (2007) An S/MAR-based infectious episomal genomic DNA expression vector provides long-term regulated functional complementation of LDLR deficiency. Nucleic Acids Res. 35, e98. Calos, M. P. (2006) The phiC31 integrase system for gene therapy. Curr. Gene Ther. 6, 633–645. Tolmachov, O., Palaszewski, I., Bigger, B., and Coutelle, C. (2006) RecET driven chromosomal gene targeting to generate a RecA deficient Escherichia coli strain for Cre mediated production of minicircle DNA. BMC Biotechnol. 6, 17. Gu, W., Putral, L., Hengst, K., Minto, K., Saunders, N. A., Leggatt, G., et al. (2006) Inhibition of cervical cancer cell growth in vitro and in vivo with lentiviral-vector delivered short hairpin RNA targeting human papillomavirus E6 and E7 oncogenes. Cancer Gene Ther. 13, 1023–1032.

Designing Plasmid Vectors 18. Chaffin, D. O., and Rubens, C. E. (1998) Blue/white screening of recombinant plasmids in Gram-positive bacteria by interruption of alkaline phosphatase gene (phoZ) expression. Gene. 219, 91–99. 19. Bernard, P. (1996) Positive selection of recombinant DNA by CcdB. Biotechniques. 21, 320–323. 20. Choi, Y. J., Wang, T. T., and Lee, B. H. (2002) Positive selection vectors. Crit. Rev. Biotechnol. 22, 225–244. 21. Gabant, P., Van Reeth, T., Dreze, P. L., Faelen, M., Szpirer, C., and Szpirer, J. (2000) New positive selection system based on the parD (kis/kid) system of the R1 plasmid. Biotechniques. 28, 784–788. 22. Heyman, J. A., Cornthwaite, J., Foncerrada, L., Gilmore, J. R., Gontang, E., Hartman, K. J., et al. (1999) Genome-scale cloning and expression of individual open reading frames using topoisomerase I-mediated ligation. Genome Res. 9, 383–392. 23. Al-Allaf, F. A., Tolmachov, O., Themis, M., and Coutelle, C. (2005) Coupled analysis

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of bacterial transformants and ligation mixture by duplex PCR enables detection of fatal instability of a nascent recombinant plasmid. J. Biochem. Biophys. Methods. 64, 142–146. Fromme, T., and Klingenspor, M. (2007) Rapid single step subcloning procedure by combined action of type II and type IIs endonucleases with ligase. J Biol Eng. 1, 7. Gupta, S., Arora, K., Sampath, A., Khurana, S., Singh, S. S., Gupta, A., et al. (1999) Simplified gene-fragment phage display system for epitope mapping. Biotechniques. 27, 328–330, 332–324. Worthington, M. T., Pelo, J., and Lo, R. Q. (2001) Cloning of random oligonucleotides to create single-insert plasmid libraries. Anal. Biochem. 294, 169–175. Cosloy, S. D., and Oishi, M. (1973) Genetic transformation in Escherichia coli K12. Proc. Natl. Acad. Sci. USA. 70, 84–87. Sawitzke, J. A., Thomason, L. C., Costantino, N., Bubunenko, M., Datta, S., and Court, D. L. (2007) Recombineering: in vivo genetic engineering in E. coli, S. enterica, and beyond. Methods Enzymol. 421, 171–199.

Chapter 7 Development of Bacterial Vectors for Tumor-Targeted Gene Therapy Li-Jun Jia and Zi-Chun Hua Summary Gene therapy holds great promise for the treatment of cancer. The success of the strategy relies on effective gene transfer into tumor microenvironments. Although a variety of gene delivery vehicles, such as viral vectors, has been developed, most of them suffer from some limitations, including inadequate tumor targeting, inefficient gene transfer, and potential toxicity. This situation suggests that it is necessary to develop novel vectors for effective tumor-targeted gene transfer. The discovery of tumor-targeting bacteria has spurred interest in the use of these bacteria as gene transfer vectors. In this review, we focus on the current status of the development of bacterial vectors for cancer gene therapy and highlight some of the directions that the field may take. Key words: Administration routes, anti-angiogenesis, bacterial vector, cancer gene therapy, cytokine, gene expression, prodrug therapy, RNA interference, tumor targeting.

1. Introduction Gene therapy, as an alternative treatment of human cancer to conventional surgery, radiotherapy, and chemotherapy, holds great promise (1). The success of gene therapy relies on effective gene transfer into tumor microenvironments. Ideal gene delivery vectors should (1) be safe; (2) selectively target tumors; (3) be motile and disseminate within tumors in large areas; (4) exert long-term efficacy; (5) have a large capacity to contain multiple therapeutic genes; and (6) be controllable and eradicable after delivery (2–4). In the past 2 decades, a variety of gene delivery vehicles has been developed. However, vectors that meet all or most of the requirements are few. Most of


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them have some drawbacks such as inadequate tumor targeting, inefficient gene transfer, limited delivery capacity, or potential risk to patients (2–4). This unsatisfactory situation suggests that it is necessary to develop novel vectors for effective tumor-targeted gene transfer.

2. Bacterial Vectors and Their Advantages

2.1. Tumor-Targeting Potential

It has been found that some obligate or facultative anaerobic bacteria, such as Salmonella, Bifidobacterium, and Clostridium, can preferentially replicate in hypoxic and necrotic areas of tumors and suppress their growth (5–8). The discovery of this kind of bacteria (so-called tumor-targeting bacteria [TTBac]), has spurred interest in the use of these bacteria as antitumor agents and gene transfer vectors for tumor-targeted therapy (Fig. 1) (9–14). Compared with conventional gene delivery carriers, such as oncolytic viruses, bacteria possess a series of unique advantages as follows. TTBac preferentially or exclusively replicate in tumors, which results in the ratio of bacterial titer between tumor and normal tissues of more than 1,000–10,000:1 (9, 11). Preferential localization of the bacteria within tumors is mainly attributable to the unique tumor microenvironments, which contains hypoxic and

Fig. 1. Bacteria-mediated cancer therapy.

Bacterial Vector-Mediated Cancer Gene Therapy

133

necrotic regions, has plenty of nutrition required for bacteria replication, and inhibits bacterial clearance because of the immuneescaped environment in tumors (9, 12). 2.2. Significant Efficacy on Cancer Cells in Hypoxic Regions

Solid tumors contain hypoxic regions resulting from inadequate blood supply. Cancer cells in these areas are resistant to traditional chemotherapies and radiotherapies (15, 16). However, TTBac appear to preferentially replicate in, and destroy, hypoxic regions via induction of tumor necrosis (17) and stimulation of antitumor immunity (18, 19) as well as inhibition of tumor angiogenesis (20, 21).

2.3. Self-Replication Within Tumors and Long-Term Anticancer Effect

TTBac are capable of self-replicating in a supportive tumor microenvironments. After a single infection, bacteria can selectively localize within tumors as live biological agents for a long period (22). Self-replication and long-term persistence of TTBac within a tumor can amplify the expression of therapeutic genes, simplify the treatment procedure, and reduce the potential toxicity.

2.4. Effective Dissemination Within Tumors

Cancer cells are generally surrounded by an extracellular matrix (ECM) within some kinds of tumor (23). This barrier significantly limits the dissemination of therapeutic agents within the tumor. Bacteria can secret specific enzymes to degrade the ECM, which enables the highly motile bacteria to spread throughout the tumor (24).

2.5. Large Delivery Capacity

Viral vectors have small genome sizes that limit their delivery capacity for therapeutic genes (2, 3, 25). In contrast, bacteria have much larger genomes and can carry multiple genes to attack the tumor, thus improving efficacy (26, 27).

2.6. Safety

Unlike some viral vehicles that can carry a risk of genome integration, bacterial vectors show very little risk during self-replication. Moreover, bacterial vectors for cancer therapy are essentially nontoxic or can be made genetically attenuated (28, 29). In addition, bacteria are generally sensitive to antibiotics and can be cleared from the body if a significant adverse effect is encountered (29–31). The safety of TTBac, such as Clostridium and Salmonella, has been demonstrated in clinical trials (29, 32–34).

3. BacteriaMediated Cancer Gene Therapy

As introduced above, TTBac are very promising candidates as gene therapy vectors. Three genera, Salmonella, Bifidobacteria, and Clostridia, have been tested for this purpose (Table 1). Salmonella, as facultative anaerobes, can proliferate in both

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Table 1 Frequently used tumor-targeting bacteria Bacteria

Anaerobic features

Safety

Tumor localization

Gene Oncolysis delivery

Vaccine vector

Salmonella Facultative anaerobe

Engineering attenuated; high safety

Preferentially in tumor; intracellular or extracellular

Significant

Prokaryotic or eukaryotic expression

Available

Clostridia

Engineering attenuated; high safety

Exclusively in tumor; extracellular

Significant

Prokaryotic expression

N/A

Nontoxic; used in dairy industry

Exclusively in tumor; extracellular

Minor

Prokaryotic expression

N/A

Obligate anaerobe

BifidoObligate bacteria anaerobe

aerobic and anaerobic conditions, which enables Salmonella to colonize both large and small tumors as well as micrometastases (35, 36). Clostridia and Bifidobacteria, as obligate anaerobes, can exclusively grow in hypoxic regions of large tumors (37, 38). Bacteria-mediated cancer gene therapy targets either tumor cells directly or tumor vascular endothelial cells and other stromal cells indirectly (Table 2). 3.1. Bacteria-Directed Enzyme Prodrug Therapy

Prodrug-converting enzymes, such as cytosine deaminase (CD), nitroreductase (NTR), or thymidine kinase of herpes simplex virus (HSV-TK), have been found in bacteria, fungi, and viruses, but not in mammalian cells (39, 40). The enzymes can convert nontoxic prodrugs into toxic agents. For example, CD converts the nontoxic prodrug 5-fluorocytosine (5-FC) to the cytotoxic antimetabolite 5-fluorouridine (5-FU), which is further metabolized into inhibitors of RNA and DNA biosynthesis, thus with the potential of killing tumor cells (39, 40). Direct systemic administration of larger doses of activated cytotoxic agents (such as 5-FU) is currently used in the clinical treatment of a variety of cancers, such as colorectal, stomach, head and neck, and breast carcinomas. However, the treatment is often coupled with significant adverse effects because of normal cell killing. This should be preventable by tumor-targeted delivery of prodrug-converting enzymes followed by the administration of nontoxic prodrugs. The selective activation of prodrugs in tumors by exogenous enzymes can be achieved via several approaches, including conventional gene-directed enzyme prodrug therapy (GDEPT), virusdirected enzyme prodrug therapy (VDEPT), antibody-directed enzyme prodrug therapy (ADEPT) (40), and bacteria-directed enzyme prodrug therapy (BDEPT) developed recently (refer to text below).


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Table 2 Bacterial vector-mediated cancer gene therapies Therapeutic genes

Bacterial strains

Models Results

Reference

Clostridium beijerinckii

In vitro

Expression of CD and conversion of 5-FC to 5-FU

(41)

C. acetobutylicum

Rat

Tumor-specific prodrug conversion

(43, 44)

Prodrug-converting enzyme Cytosine deaminase

Salmonella typhimu- Mice rium

Tumor-selective expression of CD and (47) conversion of 5-FC to 5-FU

S. typhimurium

Mice

CD expression, 5-FC to 5-FU conver- (48) sion and tumor suppression

S. typhimurium

Human

Bacterial colonization in tumor and 5-FC to 5-FU conversion were demonstrated in two of three refractory cancer patients

(32)

Bifidobacterium longum

In vitro

Expression of CD and conversion of 5-FC to 5-FU

(49)

B. longum

Rat

Tumor-selective expression of CD, conversion of 5-FC to 5-FU and tumor suppression

(50)

B. longum

In vitro

A mutant CD gene with a mutation at the active site generated significantly enhanced enzymatic activity

(51)

HSV-TK

S. typhimurium

Mice

TK expressed, tumor suppressed, and survival prolonged

(46)

Nitroreductase

C. beijerinckii

Mice

Tumor-selective expression of Escherichia coli nitroreductase

(42)

C. sporogenes

Mice

(45) Sustained antitumor effects was achieved by repeated cycles of Clostridium-directed enzyme prodrug therapy

Angiogenesis inhibitors Thrombospondin-1

S. choleraesuis

Mice

TSP1 expression, angiogenesis inhibition, tumor growth suppression, and survival prolongation

(55)

Endostatin

S. choleraesuis

Mice

Endostatin expression, angiogenesis inhibition, tumor growth suppression, and survival prolongation

(56)

B. adolescentis

Mice

Endostatin expression, angiogenesis inhi- (57) bition, and tumor growth suppression

B. longum

Mice

Tumor growth suppression and survival prolongation

(58) (continued)

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Table 2 (continued) Therapeutic genes

Bacterial strains

Models Results

Reference

TNF-alpha

C. acetobutylicum

In vitro Tumor-specific delivery of TNF-alpha and by recombinant Clostridium. Sigin rat nificantly improved tumor specificity via radio-induced promoters

(66–67)

Interleukin-2

C. acetobutylicum

In vitro

Secretory production of rat interleukin-2

(69)

LIGHT

S. typhimurium

Mice

Inhibition of tumor growth and metastases, induction of inflammatory cell infiltration within tumor

(70)

Catenin β-1

E. coli

Mice

(74) E. coli encoding short hairpin RNA (shRNA) against catenin β-1 induce significant gene silencing in human colon cancer xenografts in mice

STAT3

S. typhimurium

Mice

S. typhimurium expressing Stat3siRNAs significantly inhibited tumor growth, tissue metastasis, and extended animal survival

Cytokines

siRNA

(75)

Clostridia, Salmonella, and Bifidobacteria have been used to selectively deliver prodrug-converting enzymes into tumors (32, 41–51). The expression of enzymes after treatment with recombinant bacteria was detected in tumors, rather than in normal tissues. The enzymes effectively converted the prodrugs to cytotoxic agents, and induced significant antitumor effects in a variety of models without significant adverse effects (41–51). Moreover, repeated treatments of BDEPT induced sustained antitumor effects (45). Based on the efficacy and safety of BDEPT observed in preclinical studies, a pilot clinical trial of Salmonella typhimurium VNP20009 expressing Escherichia coli cytosine deaminase (TAPET-CD) was performed in refractory cancer patients (32). Three patients received intratumoral injections of three doses (3 × 106 to 3 × 107 cfu/m2) of TAPET-CD, followed by 5-FC treatment. Two of the three patients had bacteria accumulation in the tumor and conversion of 5-FC to 5-FU as a result of CD


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expression. During the treatment, no significant adverse effects were observed (32). The results demonstrate that Salmonellamediated CD delivery into tumor can be safe and functional. In this very preliminary trial, although protein expression and prodrug conversion were observed, TAPET-CD did not induce observable anticancer effects (32). Similar to its parental strain, VNP20009 (29), poor tumor colonization of TAPET-CD (103–106 cfu/g tumor) in cancer patients was demonstrated (32), which might result in inadequate prodrug conversion and toxic drug production, thus limiting the efficacy. In addition, the failure may also result from the essential limitations of BDEPT. The present TTBac is mainly distributed in hypoxic regions that are poorly vascularized or avascularized. Thus, the delivery of prodrugs into all tumor areas is a barrier, and further genetic modification is needed to better optimize the targeting by the bacteria. 3.2. Bacteria-Mediated Anti-Angiogenic Therapy

It is well known that angiogenesis is essential for the growth and metastases of tumors. Angiogenesis inhibition delays tumor growth and metastasis (52, 53). Anti-angiogenic therapies using endogenous inhibitors, therapeutic antibodies, and low-dose metronomic chemotherapeutics are being intensively evaluated in preclinical and clinical studies (53, 54). The treatments inhibit tumor angiogenesis via “direct” pathways by killing differentiated endothelial cells or circulating endothelial progenitor cells, and via “indirect” pathways by increasing endogenous inhibitors of angiogenesis (such as thrombospondin 1 [TSP1]) and/or decreasing endogenous promoters of angiogenesis, such as vascular endothelial growth factor (VEGF) (53, 54). Recently, Bifidobacteria- and Salmonella-mediated endostatin and TSP-1 gene therapy were reported (55–58). After systemic infection, recombinant bacteria specifically targeted and preferentially replicated within tumors. Expression of endostatin and thrombospondin-1 within tumors inhibited tumor angiogenesis, delayed tumor growth, and prolonged animal survival in primary and metastatic tumor models (55–58). The studies demonstrated that bacterial vectors are suitable to deliver antiangiogenic genes for cancer therapies. Although acting as anti-angiogenic gene carriers, the bacterial vector alone, especially for Salmonella, displayed anti-angiogenic effects. An in vitro study showed that S. typhimurium directly induced the apoptosis of endothelial cells in a dose- and time-dependent manner, and the majority of the apoptotic effect appeared to be lipopolysaccharide (LPS)mediated (Thamm DH, 2005). We found that S. typhimurium significantly reduced the tumor microvascular density (MVD) and the tumor VEGF level (20). The study by Lee et al. also found that S. choleraesuis suppressed endothelial cell proliferation and reduced tumor MVD (21). In addition, bacterial

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infection-induced cytokines, such as interferon (IFN)-γ, interferon-inducible protein (IP)-10, and interleukin (IL)-18, have potential anti-angiogenic activity (21, 59, 60). Thus, Salmonella-directed anti-angiogenic gene therapy exerts a synergic anti-angiogenesis efficacy induced by both specific therapeutic genes and bacterial functions. Combination therapies with TTBac and anti-angiogenic or antivascular agents further support the rationale of bacteriadirected anti-angiogenic gene therapy. We found that the combination of S. typhimurium VNP20009 and purified endostatin protein or cyclophosphamide used at regimens targeting angiogenesis significantly reduced tumor microvascular density (MVD), and enhanced tumor necrosis and anticancer effects when compared with bacteria treatment alone (20, 61). The combination of S. typhimurium VNP20009 with a single compound isolated from Chinese herbal medicine with an anticancer activity also displayed synergistic antitumor efficacy, significantly reduced tumor microvascular density and VEGF expression, and increased tumor necrosis (unpublished data). Dang et al. performed combination therapy with Clostridium novyi and antivascular microtubule destabilizers that radically disrupted the preexisting tumor microvasculature. Microtubule destabilizers reduced blood flow to tumors, and enlarged the hypoxic and necrotic regions, which favored C. novyi-NT proliferation and exertion of antitumor effects (62). 3.3. BacteriaMediated Cytokine Gene Therapy

The induction of an anticancer immune response has been a long-sought goal (63). It has been found that immunoregulatory cytokines, such as granulocyte-macrophage colony-stimulating factor (GM-CSF), interleukin (IL), tumor necrosis factor-α (TNF-α), and interferon (IFN), activate antitumor immunization in both preclinical and clinical studies (64, 65). Clinical studies with cytokines, such as IL-12, showed that systemic administration of cytokines at therapeutic doses were highly toxic (65). Therefore, novel tumor-targeted delivery systems for cytokine therapies need to be developed. TTBac represent an innovative tool to selectively deliver cytokines into tumors. Previous studies showed that C. acetobutylicum- or Salmonella-mediated delivery of a TNF-α gene induced tumor-specific protein expression (66–68), and C. acetobutylicum encoding IL-2 induced secretory expression of the protein at high concentrations (69). Most recently, attenuated S. typhimurium was engineered to produce the human cytokine LIGHT, a TNF-family cytokine, for cancer therapy (70). LIGHT binds to both the lymphotoxin-β receptor (LTβR) and the herpes virus entry mediator (HVEM) receptor, and stimulates emigration of dendritic cells (DCs), T cells, B cells, and natural killer (NK) cells. LIGHT is also a growth factor for cells involved in antigen


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presentation and immune response initiation. Attenuated S. typhimurium-mediated LIGHT administration induced significant antitumor activity. The efficacy is mediated by receptor (LTβR and HVEM)-dependent activation of B cells, CD4 (+) T cells, CD8 (+) T cells, and DCs (70). In addition to LIGHT, Fas L and IL-18 have been tested in S. typhimurium-mediated delivery systems and significant antitumor effects were also obtained (70). During treatment, no systemic toxicity was observed. These studies demonstrate the feasibility of bacteria-mediated tumortargeted delivery of cytokines. Aside from acting as a tumor-targeted delivery system, the bacterial vector itself can exert immune-stimulatory effects. C. novyi-NT stimulates a severe inflammatory response within the tumor and the production of a series of inflammatory cytokines, therefore inducing specific antitumor cellular immune responses (18). S. choleraesuis causes a significant increase in neutrophils and CD4 (+) and CD8 (+) T cells in tumors (21). Recently, it was reported that attenuated S. typhimurium can invade melanoma cells and cause the presentation of antigenic determinants of bacterial origin, which makes them targets for anti-Salmonellaspecific T cells (19). The findings demonstrated the immunestimulatory effect of bacteria vectors, by inducing the production of inflammatory cytokines, the infiltration of immune cells into tumors, and the presentation of bacterial antigenic determinants by tumor cells. 3.4. BacteriaDelivered RNA Interference Therapy

RNA interference (RNAi) is a conserved mechanism from plants to humans for specific silencing of target genes, and becomes an important tool for cancer gene therapies (71). In mammals, a series of vectors such as viruses, liposomes, and nanoparticles containing small interfering RNA (siRNA)-encoding DNA have been exploited to mediate gene-specific silencing (72, 73). However, the delivery vectors have limitations as mentioned above. Recently, bacteria-mediated siRNA delivery into mammalian cells has been established. Xiang et al. reported for the first time that bacteria can be engineered to produce an siRNA targeting a mammalian gene to induce trans-species RNAi both in vitro and in vivo (74). In this study, nonpathogenic E. coli were engineered to transcribe siRNAs from a plasmid containing the invasin gene Inv and the Listeriolysin O gene HlyA, which encode two bacterial factors needed for successful transfer of the siRNAs into mammalian cells. The administration of E. coli encoding siRNA against catenin β-1 induced significant gene silencing in the intestinal epithelium and in human colon cancer xenografts in mice. Next, S. typhimurium was used to deliver Stat3-specific siRNA for antitumor therapy (75). Compared with bacterial treatment alone, Stat3 siRNA-expressed S. typhimurium inhibited tumor growth, reduced metastases, and extended animal survival more

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significantly. The findings demonstrate that attenuated bacteria could be used as siRNA delivery systems. The technical breakthrough of bacteria-based siRNA delivery in mammals not only broadens the application of bacterial vectors, but also has a profound impact on the discovery of novel anticancer targets. Conventionally, an ideal cancer target should be overexpressed in cancer cells but not in normal cells (76). Thus, its inhibition will have selective killing effects on cancer cells without toxicity to normal cells. This criterion makes many ubiquitously expressed molecules that are critical for cell growth and function “undrugable.” The highly tumor-targeted silencing of target genes by bacteria-delivered siRNA will make the ubiquitously expressed “undrugable” molecules “drugable.” This should broaden the spectrum of anticancer targets. The success in bacteria-mediated RNAi delivery also suggests that the bacteria could be used to deliver therapeutic microRNAs (miRNA). MiRNAs are small noncoding RNAs that can regulate gene expression (77, 78). Accumulating evidence highlights their effects in cancer initiation, progression, and prognosis (77, 78). Some miRNAs are found to produce tumor suppressor effects by silencing the expression of oncogenes, thus anti-oncogenic miRNAs may be delivered by TTBac to achieve anticancer effects.

4. Optimization of Bacteria-Mediated Delivery Systems

4.1. Bacterial Vectors 4.1.1. Screening for Tumor-Targeting Strains

To maximize the potency of bacteria-mediated cancer gene therapy, elements of systems including bacterial vectors, gene expression constructs, and administration routes need to be optimized (Fig. 2). Tumor-targeting potential is one of the most potent advantages of bacterial vectors. Based on the features of bacteria, several aspects should be kept in mind. (1) Not all bacterial strains have equal tumor-targeting potentials. Dang et al. systematically assessed anaerobic bacteria for their capacity to grow expansively within transplanted tumors. Among 26 different strains from three different genera including Bifidobacteria, Lactobacilli, and Clostridia, only two strains (C. novyi and C. sordellii) exhibited extensive spreading throughout the poorly vascularized regions of the tumors after the systemic injection of spores (17). (2) Tumor-targeting strains are not limited to the most frequently tested Salmonella, Bifidobacteria, and Clostridia. Some strains from other genera, including E. coli, Vibrrio chlolerae, and Listeria monocytogenes, also have tumor-targeting potential (79, 80). (3) The extent of preferential replication of different strains in the


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Fig. 2. Optimization of bacteria-mediated gene delivery systems.

same tumor model or a given strain in different tumor models varies significantly (9). These findings suggest that new tumortargeting strains should be further screened and a suitable tumor model for a given strain needs to be identified. 4.1.2. Attenuation of Virulent Tumor-Targeting Strains

Safety is one of the major concerns for the clinical application of the bacterial vectors. Although essentially nonpathogenic, bacteria have been used for anticancer therapy; the most progress has been made through using attenuated mutants of pathogenic

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bacteria. To attenuate virulent strains, two strategies based on forward genetic or reverse genetic techniques can be exploited. Using the forward genetic strategy, attenuated mutants can be selected naturally or delicately induced by chemical and physical mutagens (81). The genetic information involved in bacteria attenuation by this strategy then needs to be further identified. Although this strategy has been used to screen attenuated strains for the development of gene transfer vectors or vaccine strains, it has a main disadvantage of an unidentified genetic background, which poses a risk of virulence recovery in certain conditions (81). Reverse genetics is an approach to discovering the function of a known gene that proceeds in the opposite direction of forward genetic screening. The attenuation of the virulent strains by reverse genetics is based on the identification of virulent genes, which are then inactivated, resulting in significant attenuation of strains. This strategy is performed from genotype to phenotype and has been successfully applied to generate well-defined tumortargeting strains, such as genetically attenuated S. typhimurium and C. novyi (17, 82). Bacterial lipopolysaccharide (LPS) induces the production of TNF-α and other host-derived inflammatory cytokines, resulting in a severe inflammatory response (septic shock). S. typhimurium VNP20009 is a lipid A-modified (MsbB−), auxotrophic (PurI−) strain (30, 82). The disruption of the Salmonella msbB gene impairs the synthesis of lipid A, a key component of LPS. The parental strain of VNP20009 with the wild-type msbB gene killed all mice after infection because of an inflammatory response induced by LPS. Compared with the parental strain, the MsbB mutant strain reduces TNF-α induction and increases the LD50 by 10,000-fold. The auxotrophic MsbB mutant retained its ability to invade, preferentially replicate, and induce antitumor activities in a variety of tumor models (30, 82). Phase I clinical trials of VNP20009 in patients with metastatic melanoma demonstrated its safety with a maximum tolerated dose of 108 cfu/m2 (29). C. novyi-NT is another promising biologic agent for cancer therapy (17). The strain preferential grows within tumors and induced significant tumor necrosis by killing surrounding tumor cells. The parental strain of C. novyi-NT is highly toxic and kills all mice 16–18 h after the infection. The lethal toxicity of the wild-type bacteria results from the release of toxins from the bacteria germinating within the tumors. Since the lethal toxicity gene is carried in phage, bacterial spores are heated to inactivate the phage carrying the lethal toxin gene to attenuate the parental strain (17). Toxicological studies of this attenuated strain show that it is rapidly cleared from the circulation and normal tissues without clinical toxicity (31). Compared with strains screened by forward genetics (81), strains isolated from reverse genetic screens (82)have defined


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genetic backgrounds, which is important for the development of anticancer bacteria. The availability of bacterial entire genome sequences (26, 27), the identification of bacterial virulent genes (83), and the easy manipulation of genetic modification significantly facilitate the process of attenuating virulent strains by reverse genetics. 4.1.3. Enhancement of Selective Tumor Localization of Bacterial Vectors

The anticancer effect of bacteria is closely correlated with selective tumor localization. In phase I clinical trials of VNP20009 given to patients with metastatic melanoma, poor tumor localization of bacteria and no significant antitumor effects were observed (29). The finding indicates that improvement of tumor localization of bacteria is urgently required for clinical application of tumor-targeting bacteria. For this purpose, two strategies, either by creating a more ideal tumor microenvironment for bacteria replication or by enhancing the affinity of bacteria to cancer cells, can be exploited. It has been shown that the combination of bacteria and other treatments, such as chemotherapeutics (21, 84), traditional medicine components, angiogenesis inhibitors (61), or antivascular agents (62, 85), significantly improved selective tumor localization of bacteria. We have performed combination therapies with S. typhimurium plus cyclophosphamide or endostatin. The combinations have induced significant tumor angiogenesis suppression, enhanced tumor hypoxia and necrosis, and therefore facilitated bacterial proliferation (20, 61). Improved tumor targeting of bacteria is also demonstrated in combination therapies with C. novyi plus microtubule destabilizers (62), C. sporogenes plus combretastatin A-4 phosphate (85), or S. choleraesuis plus cisplatin (84). The enhancement of tumor localization of bacteria by combining other anticancer agents aiming to enhance tissue hypoxia and necrosis improves the clinical value of bacteria. The use of bacteria displaying tumor-targeting molecules represents another strategy to improve selective replication of bacteria in tumors. S. typhimurium VNP20009 expressing carcinoembryonic antigen (CEA)-specific single chain antibody fragments (scFv) on the cell surface is generated to increase tumor targeting of bacteria in vivo. The modified bacteria enhanced the localization in CEA-expressing tumors and therapeutic effects (86). The display of antibody fragments on the surface of bacteria represents a novel strategy for targeting specific antigen-expressing tumors. Similarly, bacteria displaying other tumor-affinity molecules, such as tumor-targeting peptides, receptors, and/or ligands, also hold great promise for improving the tumor-selective localization of bacteria (87, 88).

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4.2. Gene Expression Systems 4.2.1. Modes of Gene Expression

Therapeutic genes can be expressed by either prokaryotic or eukaryotic expression constructs. Prokaryotic expression of a gene occurs in recombinant bacteria and the expressed protein is subsequently secreted out of the bacteria to exert an anticancer effect. In contrast, eukaryotic expression of a gene depends on the effective invasion of bacterial vectors into cancer cells and thus the delivery of an expression plasmid from bacteria into cells for protein expression. Both protein expression systems have their own advantages and disadvantages as discussed below.

Prokaryotic Expression

Prokaryotic expression of genes can be achieved by recombinant bacteria carrying a prokaryotic expression plasmid or carrying a chromosomally inserted gene expression cassette. This strategy is suitable to both intracellular and extracellular bacteria. To accomplish the secretion of a therapeutic protein from bacteria into extracellular space, secretion signals need to be integrated into the DNA constructs. Efficient secretion expression of HSVTK by Salmonella, and IL-2 and TNF-α by Clostridium have been reported (8, 66, 69). However, the expression of mammalian genes may be limited because of the difference in codon preference between bacteria and mammalian cells. For example, the expression of GC-rich mammalian genes may be limited in C. novyi-NT carrying a high AT content genome (26). Moreover, the activity of some mammalian proteins expressed by bacteria may be impaired because of inadequate translational modifications in bacteria (89). In addition, prokaryotic expression cannot be applied to deliver therapeutic siRNA or microRNA because their synthesis and modification require eukaryotic cellular machinery (90).

Eukaryotic Expression

Eukaryotic expression of mammalian genes has native posttranslational modifications to achieve full biological activity. This strategy is suitable to intracellular bacteria such as Salmonella and Escherichia, but not extracellular bacteria such as Bifidobacteria and Clostridia. Salmonella has been used to deliver mammalian expression constructs encoding endostatin, TSP-1, and cytokine LIGHT for cancer therapy (55, 56, 70). The success of this strategy depends on the ability of recombinant bacteria to invade mammalian cells and to deliver DNA from bacteria into host cells for protein expression. Although the existence of TTBac within cancer cells in vivo has been observed, the percentage of infected cells is very low and most of the bacteria reside in the extracellular environment (11). Thus, improvement of bacterial invasion is important for effective gene transfer into host cells. Avogadri et al. found that only invasive S. typhimurium was able to infect tumor cells, whereas a noninvasive strain was unable to effectively invade tumor cells,


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suggesting that it is necessary to use a invasive strain for effective cell invasion (19). Yersinia invasin binds to β1 integrin receptors, which are located on the membrane of mammalian cells, thereby facilitating the Yersinia entry into the cells (91). It has been used to mediate cell invasion of other bacterial strains. E. coli expressing an invasin gene successfully invaded Hela, U2OS, and HepG2 cancer cells (92). Moreover, invasin-mediated cancer cell entry of E. coli has further been demonstrated in in vivo studies. To avoid the entry of invasinexpressing bacteria into normal cells and confine invasin-expressing bacteria to malignant cells, inducible expression of invasin genes by hypoxia, a hallmark of solid tumors, was exploited (92). The findings highlight the potential of using Yersinia invasin to improve the selective cancer cell invasion of recombinant bacteria. Another key factor affecting bacteria-mediated gene therapy is the release efficacy of DNA constructs into the cytoplasm of host cells. After cell invasion, Salmonella remains and divides within phagosomes, and transfers DNA from bacteria into cells (93, 94). The improvement of DNA transfer would enhance the efficacy of therapy. Listeriolysin from L. monocytogenes can induce phagosome lysis and cause bacteria to escape from phagosomes (95). The use of recombinant Salmonella expressing additional listeriolysin results in a higher level of gene expression (96). Listeriolysin has also been applied to facilitate the transfer of the siRNA-encoding plasmids from bacteria into host cells (74). These findings demonstrate that listeriolysin is a useful tool to enhance gene delivery by bacteria. It is reported that eukaryotic promoters (PCMV and PRSV) can direct protein synthesis in gram-negative bacteria using the lacZ and green fluorescent protein (GFP) genes as reporters (97). The GFP synthesized in E. coli can be transferred to mammalian cells where it remains detectable and stable (97). We have investigated whether a reporter gene controlled by a eukaryotic promoter could be expressed by tumor-targeting S. typhimurium; VNP20009 was electroporated with a pGL3-Control vector in which a reporter luciferase gene was controlled by an SV40 promoter. The recombinant Salmonella were cultured in LB medium and sampled for luciferase activity assays. Figure 3 shows that a significant level of luciferase was expressed by VNP20009. In addition, we found that the expression capability of an SV40 promoter in Salmonella varies significantly in different strains. In S. typhimurium strain C77, an SV40 promoter has very weak activity. This result raises the question regarding to what extent the gene under the control of a eukaryotic promoter was really expressed by mammalian cells but not in bacteria. The findings also suggest caution when interpreting data on the DNA delivery from bacteria to mammalian cells..

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Fig. 3. Eukaryotic promoter-directed gene expressions in tumor-targeting Salmonella typhimurium.

Double Expression System

In contrast to the single expression of a therapeutic gene, doubleexpression vectors containing both prokaryotic and eukaryotic expression cassettes in a plasmid could be designed. A double-expression vector might allow intrabacterial expression of a gene from a prokaryotic expression cassette and intracellular expression from a eukaryotic expression cassette, thus employing a wide range in bacteria-mediated gene therapy and greatly improving efficacy.

4.2.2. Inducible Expression of Therapeutic Genes

The ideal bacteria-mediated gene therapy requires high tumorspecific expression of genes with the minimum distribution of the gene products into normal tissues. Although TTBac can preferentially replicate in tumors, a small number of bacteria, especially from facultative anaerobes, may replicate in normal tissues, which poses the potential risk of toxicity to normal tissues (22). Thus, spatial and temporal control of gene expression is of importance for improving tumor specificity and reducing potential toxicity. To control gene expression, unique features of tumor microenvironment (such as hypoxia), anticancer treatments (such as radiotherapy and chemotherapy), and small molecular weight chemicals can be exploited as inducers. Radiation-induced gene expression has been demonstrated by using recA promoters in Clostridium (67). The recA gene, belonging to the SOS-repair system in bacteria, is induced by ionizing irradiation (98). The production of TNF-α is increased by ionizing irradiation of Clostridium containing TNF-α complementary DNA (cDNA) under control of the recA promoter (67). Repeated irradiation induced repeated gene activation and a significantly increased TNF-α production. These results demonstrated that spatial and temporal control of gene expression by a bacterial vector can be achieved using a radio-inducible promoter, which thus provides a new means of increasing the selectivity in cancer treatment.


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One hallmark of tumors, hypoxia, has been used to induce tumor-targeted gene expression in Salmonella. A hypoxia-inducible promoter (HIP-1) derived from a portion of the endogenous Salmonella pepT promoter drives reporter gene expression under hypoxia, but not under normoxia (99). Genetic engineering of regulatory elements within HIP-1 allowed the fine tuning of gene induction. Moreover, HIP-1 can drive hypoxia-mediated gene expression in bacteria that have colonized in tumors (99). In addition, the promoter of formate dehydrogenase (fdhF), one of the most strongly hypoxia-induced genes, can be used to induce gene expression (92). Some small molecules, such as L-arabinose, have been used to induce bacteria-mediated gene expression (100, 101). L-arabinose can activate the PBAD promoter from the arabinose operon of E. coli. One study used the L-arabinose and PBAD promoter system to control Salmonella-mediated gene expression. Administration of L-arabinose induced strong expression of reporters by the bacteria in tumors (101). As an inducer of gene expression, L-arabinose possesses some advantages, such as biocompatibility and effective distribution in the entire tumor. Eukaryotic inducible-expression systems have also been investigated for tumor-selective gene expression. The early growth response 1 (Egr1) gene is induced by irradiation and the inducibility is attributable to ten nucleotide motifs of consensus sequence CC (A/T)6GG (CArG elements) (102). These elements respond to ionizing radiation-induced reactive oxygen species (ROS) and H2O2. Further studies found that the Egr1 promoter is also induced by ROS-inducing chemotherapeutics, such as cisplatin, doxorubicin, cyclophosphamide, and 5-FU (103). Thus, the Egr1 promoter has been intensively used to control gene expression. To induce gene expression under hypoxia, promoters containing hypoxia-responsive elements with the consensus sequence (A/G) CGT (G/C) (G/C) bonded by hypoxia-inducible factor (HIF) have been developed (102). Thus, there is considerable potential for eukaryotic inducible promoters to be incorporated into eukaryotic gene expression systems delivered by bacteria. 4.3. Administration Routes for Bacterial Vectors

To increase the reliability of bacterial infection, TTBac are conventionally injected intravenously (i.v.), intraperitoneally (i.p.), or intratumorally (i.t.) into animals and humans for cancer therapy. Most recently, we have found that tumor-targeting S. typhimurium VNP20009 could be administrated orally as an effective anticancer agent (104). After oral administration, VNP20009 targeted and preferentially accumulated within tumors, leading to significant anticancer effects either as a single-agent therapy or as part of a combination therapy. No obvious toxicity is observed during the treatments. Comparative analysis of toxicity in tumor-

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bearing and tumor-free mice further revealed that orally administrated Salmonella had higher safety compared with conventional systemic delivery of bacteria (104). The findings indicate that oral administration of TTBac is effective and safe. The approach is also highly applicable to other tumor-targeting bacteria, especially those that naturally infect their host via the alimentary tract. Thus, this approach provides an alternative route in the application of bacteria as a potential antitumor agent. In addition, oral administration of tumor-targeting Salmonella can be used to deliver DNA vaccine to induce specific anticancer responses. It is well established that oral treatment of nontumor-targeting S. typhimurium containing DNA vaccine could induce specific anticancer immunity (105, 106). After oral infection of recombinant Salmonella, DNA encoding protective antigens would be released from the bacteria vector and enter Peyer’s patches in the small intestine. The DNA is subsequently transcribed, expressed, processed, and presented by antigenpresenting cells (such as macrophages and dendritic cells) in Peyer’s patches, and then a specific immune response is stimulated (105, 106). We have found that oral delivery of tumor-targeting S. typhimurium VNP20009 can induce a specific immune response against a hemagglutinin reporter gene (unpublished data). Using tumor-targeting Salmonella as a delivery vector for DNA vaccine, dual therapeutic effects mediated by both specific antitumor immunity and bacteria therapy should be induced. In addition, this strategy will not only be suitable for preventive vaccination, but also be valuable for therapeutic immunization. Different routes of bacterial administration have their own features and applications (Table 3). Intratumoral injection ensures the most direct intratumoral delivery of bacteria and avoids systemic toxicity. However, it has less efficiency in small tumors. Instead, systemic injection, i.v. and i.p., has the potential of

Table 3 Advantages and disadvantages of different routes of bacterial administration Parameters

i.v.

i.p.

i.t.

Orally

Stress

+++

+++

+++

+

Simplicity

+

+

+

+++

Potential toxicity

+++

+++

+

+

Access to metastases

+++

+++

−

+++

Suitability to bacterial strains

+++

+++

+++

++

Delivery of DNA vaccine

−

−

−

+++


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targeting both accessible tumors and small metastases, although systemic injection is often coupled with systemic toxicity. Compared with the above-mentioned routes, oral administration is safer, more convenient, and can target both large tumors and small metastases. The potential disadvantage of oral treatment is that a higher therapeutic dose is needed compared with other routes (104). However, this may not be a major concern in clinical application of the approach because of the low cost of bacteria preparation. In addition, oral infection is only suitable to alimentary tract-invasive strains, compared with the broad suitability of conventional routes to most tumor-targeting strains. However, this feature may suggest that oral treatment will be more effective on tumors in the alimentary tract, such as gastric and colorectal cancer, than other routes. The choice of appropriate administration routes will be based on a comprehensive consideration of a series of factors including (1) the tumor site, size, and metastasis; (2) the type of therapeutic gene; (3) potential toxicity and patient health status; (4) the therapeutic gene and bacterial vector used; and (5) the simplicity of the therapeutic procedures.

5. Concluding Remarks The unique advantages of tumor-targeting bacteria suggest that they can be potent gene-delivery vectors. Preclinical and clinical studies have heightened interest in their potential. Further studies in the following directions should be performed to enhance the applicability of tumor-targeting bacteria in the clinic. First, thorough elucidation of the mechanisms responsible for tumor targeting of bacteria and improvement of their tumor localization are essential for the clinical application of the bacterial vectors. Poor tumor targeting of TAPET-CD and its parental strain VNP20009 in cancer patients was observed, although those strains are very promising in a variety of tumors and animal models (29, 32). The reasons for the huge difference in tumor-targeting potential between animal models and humans largely remain unknown. Therefore, the ultimate understanding of the mechanisms will definitely contribute to the development of bacterial vectors. Second, a larger number of therapeutic genes needs to be screened. As a new member in the family of gene delivery carriers, bacterial vectors have only been investigated with a limited number of genes so far. It is well known that cancer cells carry a series of hallmarks such as unlimited growth, resistance to apoptosis, sustained angiogenesis, and immune evasion. Any genes that reverse those processes could be delivered by bacteria to delay tumor formation, growth, and metastasis. In addition,

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the success in bacteria-mediated siRNA delivery significantly broadens the potential anticancer targets. With the progress of genome-wide RNAi screening, more potential anticancer targets will be identified (107, 108). Finally, successful bacteria-mediated gene therapy relies on efficient gene delivery and expression in which many factors that play key roles may be optimized, such as the use of TTBac from different genera, the control of bacteria invasion and DNA delivery, and the modes of gene expression as well as the routes of bacteria administration. The comparison and optimization of these parameters could lead to the establishment of a formidable bacteria delivery system. In conclusion, as an important example of gene delivery systems, bacterial vectors hold great promise for tumor-targeted gene therapy. Further development and optimization of related elements in the system will definitely promote their application potential.

Acknowledgments The authors thank Dr. Brooks Low, Dr. John Pawelek, Dr. Yi Sun, and Dr. Zhen Wang for critical reading of the manuscript and editorial work. The authors are grateful to grants from the Chinese National Nature Sciences Foundation (30425009, 30500637) and the Jiangsu Provincial Nature Sciences Foundation (BK2007715). References 1. Cross D, Burmester JK. (2006) Gene therapy for cancer treatment: past, present and future. Clin Med Res. 4:218–27. 2. Palmer DH, Young LS, Mautner V. (2006) Cancer gene-therapy: clinical trials. Trends Biotechnol. 24:76–82. 3. Seth P. (2005) Vector-mediated cancer gene therapy: an overview. Cancer Biol Ther. 4:512–7. 4. Dong JY, Woraratanadharm J. (2005) Gene therapy vector design strategies for the treatment of cancer. Future Oncol. 1:361–73. 5. Möse JR, Möse G. (1964) Oncolysis by Clostridia. I. Activity of Clostridium Butyricum (M-55) and Other nonpathogenic Clostridia against the Ehrlich carcinoma. Cancer Res. 24:212–6 6. Gericke D and Engelbart K. (1964) Oncolysis by Clostridia. II. Experiments on a Tumor Spectrum

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Subsection C Nonviral Transfer Technologies in Cancer Gene Therapy

Chapter 8 Electroporative Gene Transfer Marco Schmeer Summary Membrane electroporation (MEP) uses short high-voltage pulses to render cell membranes transiently porous and therewith permeable to otherwise impermeable substances. This technique was first described, in vitro, by Neumann in 1982 (1). In vivo, this method is restricted to solid tissues accessible to the electrodes used to apply the electric field pulses. Electroporation of cell tissue gains increasing importance especially in clinical applications such as electrochemotherapy (ECT) of, e.g., skin tumors, and for gene therapy (2, 3, 4). The various applications of MEP include, in addition to the direct functional transfer of genes (electrotransfection, electrogenetransfer) and drugs, the release of proteins, and the electrotransfer of ionic dyes into cells (5). But, nevertheless, the mechanism of pore opening and resealing as well as the transfer, especially of DNA, is not yet completely understood. Key words: Electrogenetransfer, electrotransfection, membrane electroporation, membrane pores.

1. Introduction Biological cells, very much simplified, consist of a conductive medium (cytoplasm) surrounded by a dielectric surface layer, the lipid membrane. This cell membrane is a nonpermeable hydrophobic barrier controlling exchanges of molecules between the cytoplasm and the medium surrounding the cell. Therewith, most hydrophilic molecules, e.g., DNA, are unable to enter the cell without any external forces. When an electric field is applied to the lipid membrane of a biological cell, electric charges are accumulated at the membrane pole caps facing the electrodes. As shown in Fig. 1, this leads to a modulation of the membrane potential (ionic Maxwell–Wagner polarization) inducing membrane fields that are orders of magnitude higher than the external field. These strong fields Wolfgang Walther and Ulrike S. Stein (eds.), Methods in Molecular Biology, Gene Therapy of Cancer, vol. 542 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-561-9_8

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Fig. 1. Model cell of radius a and membrane thickness d: Distribution of charges during application of an external electric field; θ is the angle between the direction of the external electric field and a considered point on the membrane surface.

Fig. 2. Electroporative gene transfer: Because of an increased membrane potential, the applied electric field causes the creation of membrane pores enabling the transport of otherwise nonpermeating substances. Additionally, the electric field electrophoretically moves the DNA molecules into the cell.

cause global and local structural changes of the lipid membrane phase, resulting in electropores that are the structural basis for enhanced transmembrane transport of small ions and larger ionic or dipolar molecules (6) (see Fig. 2). For effective electroporative transmembrane transport, the electric field strength apparently has to exceed a minimum value (threshold). Electric field pulses, applied to densely packed pellets of Chinese hamster ovary (CHO) cells as simple tissue models as

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well as to mouse skin tumors have been shown to cause electric conductivity changes analyzable in terms of kinetic normal modes (7,8). Such analyses result in at least two different pore states: small, short-lived pores and larger long-lived pores that stay open for seconds or even minutes after termination of the electric field pulse. Especially the latter appear to cause effective transmembrane transport of otherwise nonpermeating substances. However, this electric field-induced formation of membrane pores is not sufficient for an effective DNA transfer. Plasmid DNA added immediately after field pulse application does not enter the cellular interior through the membrane pores that are still open at this moment like, e.g., macromolecular dyes do (9). Additionally, the efficacy of the electroporative gene transfer significantly depends of the applied pulse protocol, e.g., the pulse duration. Besides the formation of membrane pores, the electric field—according to the currently widely accepted model—can electrophoretically move the negatively charged DNA molecules toward and finally through the cellular membranes (6). This means that only tissue areas covered by the electric field become permeabilized and transfected. Additionally, the DNA has to be as near as possible to the cell membranes of the target tissue when the electric field is applied (see Note 1). The field pulse protocol has to be carefully adjusted to the biological system that has to be transfected depending on the range of variations that is possible with the available field pulse generator. Here, it is not enough to overcome the abovementioned minimum threshold value of the applied field strength because also very high field strengths and larger pulse durations may cause irreversible cell damage. Additionally, the unavoidable high voltage drop at the electrode surfaces may cause trouble in tissue because of a large number of dead cells in this area. In electrochemotherapy, cell damage is less critical because of apoptotic and immunologic cell removal. But, nevertheless, the electrically induced damage may cause negative side effects like necrosis, ulceration, and wound appearance (10). Especially in gene therapy, massive cell damage has to be avoided because a loss of viability causes less transfection efficacy.

2. Materials 1. (Plasmid) DNA, aqueous suspension (stable at 4°C for some weeks, stable at at most −20°C for years). For clinical applications, this DNA has to be produced under Good Manufacturing Practices (GMP) (see Chapter 25, “Production of Plasmid DNA as Pharmaceutical”).

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2. Electroporation device (e.g., Cliniporator, Igea, Carpi, Italy; CUY21, Sonidel, Dublin, Ireland; MedPulser, Inovio, San Diego, USA). 3. Electrodes (with—where applicable—conductive electrode paste). 4. Syringe with sterile cannula.

3. Methods In brief, a DNA suspension is injected into the target tissue and subsequently electric field pulses are applied using an electrode setup (plates or needles) connected to a pulse generator. Basically, there are two different types of electric field pulses that can be delivered by electric field pulsers: (a) Exponential pulses: The applied electric field strength E decays from a maximum value E0 to zero with a time constant tE depending on the individual conductivity of the system between the electrodes. (b) Rectangular pulses: The voltage and duration of the pulses can be set at the device and do not depend on the electric behavior of the tissue. Therefore only this system allows or at least makes it much easier to apply a clearly predefined pulse protocol. To give an overview, different pulse protocols for treatment of different tumor tissues described in the literature are collected in Table 1. For the electroporation procedure, one possible procedure is described in the following section. 3.1. Preparation of the Plasmid DNA

The concentration of the plasmid DNA (here, e.g., pCMV-GFP) is adjusted in water for injection (WFI; PlasmidFactory, Bielefeld, Germany) suspension to 1 mg/mL (see Note 1).

3.2. Preparation of the Target Tissue

Because the tumor has to be easily accessible for the electrode system, nude mice bearing subcutaneous tumors with a size of around 4 mm (e.g., A549, MDA-MB-435, Detroid562 from ATCC, Manassas, VA, USA) serve as an appropriate animal model for in vivo experiments (16). Because the DNA injection as well as the application of the electric field pulses do not cause much pain, a local anesthetic is sufficient. Alternatively, depending on the guidelines to be applied, animals are anesthetized with Hypnorm or Midazolam (17).

Antisense ODNs against PLK-1 and BCL-2

pEGFP-N1

p.DOM-AH1 p.BCL1

Mouse subcutaneous tumor

Rat subcutaneous tumor

Mouse, intravenously injected tumor cells

Intramuscular injection (quadriceps)

Injection on tumor surface

Systemic, injection into tail vain

Intramuscular injection (quadriceps)

pcD2/hIFN-α2

Mouse leukemia

Plates, silver (with contact paste)

Plates, stainless steel

Plates, stainless steel (with contact paste)

Needle pair, stainless steel




Intratumoral injection

pEGFP-C1 pMC1-DT-A pCEP4/TK



Needle pair, gold plated

pEGFP-N1 pNGVL-mIL2 pNGVL-mIL12 pNGVL-β-gal

LacZ

Mouse melanoma

Injection in carotid artery

Electrodes

Needle array, stainless steel

LacZ, MCP-1

Rat brain tumor

Gene delivery


Gene/plasmid

Cancer/tumor type

10 Trains of 1,000 rectangular pulses, 150 V/cm, 200 µs (1 Hz)

8 Rectangular pulses (a) 1,300 V/cm, 0.1 ms (1 Hz) (b) 600 V/cm, 5 ms (1 Hz)

5 Rectangular pulses, 400 V/cm, 10 ms (1 Hz)


2 × 3 Rectangular pulses, 167 V/cm, 50 ms


1 Rectangular pulse, 70–90 V/cm, 5 ms (1 Hz)

8 Rectangular pulses, 600 V/cm, 97 µs

Pulse protocol

Table 1 Overview of different pulse protocols for treatment of various tumor models described in the literature

(18)

(17)

(16)

(15)

(14)

(13)

(12)

(11)

Reference

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Additionally, the tumor as well as the surrounding tissue have to be accessible for appropriately placing the electrodes. The skin surface should be shaved (if necessary) and amicrobic. 3.3. DNA Injection

A single dose of 50 µg plasmid DNA is injected into the tumor tissue using a microliter syringe (see Note 2).

3.4. Application of the Electric Field Pulses

The electrodes coated with contact paste are placed at two opposite sides of the tumor after intratumoral injection of the DNA. Here, two parallel stainless steel plates (5 to 10 mm wide, 0.5 to 1 mm thick) placed on a nonconductive, preferably tweezers-like holder serve as an easy to handle electrode setup. The parallel plates are connected with the pulse generator. Subsequently, five consecutive rectangular pulses of a field strength E = 400 V/cm and a duration t = 10 ms are applied at a frequency f = 1 Hz (see Notes 3 and 4).

3.5. Response Monitoring

Either the protein encoded by the transferred genetic material itself or its effect on the size of the tumor are monitored by appropriate individual analytical methods. The green fluorescence caused by expression of the green fluorescent protein (GFP, encoded on the plasmid DNA) mentioned above can be visualized in the surgically removed tumor tissue using a fluorescence microscope (excitation filter of 450–490 nm, emission filter 520 nm).

4. Notes 1. The DNA concentration depends on the injection procedure. To ensure a homogenous distribution throughout the electroporated tumor tissue, it could be necessary to dilute the DNA suspension to obtain larger volumes to inject. If the plasmid is stored at −20°C, it is stable in WFI for years. The benefit of WFI is that there are no side effects of the other substances contained in, e.g., buffer solutions. Saline, for example, should also be an option. The presence of positive ions is crucial because they allow the necessary contact between the negatively charged DNA with the cell surface, which is also negatively charged (6). In contrast to field pulse application to suspended cells, effects of the injected medium on the conductivity and osmolarity should be of minor importance in vivo because electrolytes are present in the tissue anyhow. Another important aspect is the size of the plasmid DNA. Maucksch et al. recently showed that the transfection efficacy in terms of the amount of plasmids transferred into the cell is higher

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with smaller plasmids (19). In this context, please see also the Chapter 4 “Use of Minicircle Plasmids for Gene Therapy” concerning the development of minicircle DNA—certainly the minimal circular DNA structure that can be used here. 2. For treatment of tumors, a local intratumoral injection of the DNA is necessary because for a systemic injection the required DNA amount would be too large to obtain a sufficient DNA concentration in the direct environment of the tumor cells. Using needle electrodes, better results are described in the literature if the DNA suspension is injected via these needles, preferably during the incision (20). 3. It is important to know that the electric field strength E = U/l as the driving force for changes of the membrane integrity depends on the applied voltage U and the electrode distance l. Therefore (if this is not done automatically by the device), the distance l has to be carefully measured to adjust the corresponding voltage U at the field pulse generator. Application of electric field pulses to living material always goes along with irreversible damage of a certain amount of cells. Various electrode effects like redox reactions as well as an increased voltage drop near the electrode surfaces cause this damage of a high fraction of cells, especially in the direct vicinity of the electrodes (21, 22). These cells may cause necrosis, possible ulceration, and wound appearance (10). Especially in gene therapy, this effect has to be avoided because a loss of viability causes less efficacy of the DNA transfer. Therefore, the correct pulse protocol is crucial for an effective treatment. Depending on the tumor tissue, it is recommended to place needle electrodes near to but outside the tumor because the cut tissue in the immediate vicinity of the electrodes might be able cause metastases. Here, the fact that cells in the immediate vicinity of the electrodes are irreversibly damaged in most cases may be even beneficial. The advantage of plate electrodes in contrast to needle electrodes is a more homogenous field distribution between the electrodes whereas with needle electrodes the electric field is completely inhomogeneous. This effect can be overcome to a certain extent using needle arrays, e.g., consisting of parallel rows of several needles. Another option is a coating of the electrodes with a conductive gel. Here, the high potential drop at the electrode surfaces (because of the roughness of the material and the density of the field lines at needle electrodes) causing redox reactions, including pH effects, and solvation of metal ions from the anode electrode takes place inside the gel. Therefore, the cells are prevented from influences of these effects (23). Clamping the tumor between two plate electrodes for field pulse application to subcutaneous tumors leads to a more homogenous

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field within the tumor tissue between the electrode plates. However, here one has to take into account the high resistance of the outer skin layer decreasing the effective electric field applied to the (skin-coated) tumor tissue. A contact paste between the electrodes allows at least a conductive contact between the metal and the tissue. 4. As discussed, several pulse protocols lead to a sufficient transfection result. Electric field pulses that are too long or a field strength that is too high leads to irreversible damage of parts of the target tissue, which may cause negative side effects. On the other hand, one has to overcome a certain threshold value for pulse duration and field strength to reach an amount of transfected cells sufficient for the intended effect. The time constant of exponential shaped field pulses depends on the conductivity of the substance between the electrodes. However, on the other hand, this type of pulse causes the creation of membrane pores as well as the electrophoretic movement of the DNA. A comparable effect cannot be seen after application of a single field pulse of rectangular shape. Here, trains of pulses have to be used where the intensity and duration of the field is easier to control than for the exponential pulses (6). Another elegant approach is to combine a short (microsecond) rectangular pulse of a high field strength to open the required large membrane pores followed by a longer (millisecond) lasting pulse of lower field strength enabling the movement of the DNA into the cell. Unfortunately, up to now this type of pulse protocol is possible only with very few devices (e.g., Cliniporator). Additionally, measurements of the conductivity changes appearing during the field pulse application because of the creation of new conductive pathways—the membrane pores—can in principle be used to (re-)adjust the applied field pulse (24).

References 1. Neumann, E., Schaefer-Ridder, M., Wang, Y., Hofschneider, P.H. (1982) Gene transfer into mouse lyoma cells by electroporation in high electric fields. EMBO J. 1, 1982, 841–845. 2. Mir, L.M., Tounekti, O., Orlowski, S. (1996) Bleomycin: revival of an old drug. Gen. Pharmacol. 27, 745–748. 3. Heller, R., Jaroszeski, M.L., Glass, L.E., Messina, J.L., Rapaport, D.P., De Conti, R.C., Frenske, N.A., Gilbert, R.A., Mir, L.M., Reintgen, D.S. (1996) Phase I/II trial for the treatment of cutaneous and subcutaneous tumors using chemoelectrotherapy. Cancer, 77, 964–971.

4. Heller, R., Jaroszeski, M.L., Glass, L.E., Messina, J.L., Rapaport, D.P., De Conti, R.C., Frenske, N.A., Gilbert, R.A., Mir, L.M., Reintgen, D.S., (1996) Phase I/II trial for the treatment of cutanous and subcutantous tumors using chemoelectrotherapy, Cancer, 77, 964–971. 5. Neumann, E., Tönsing, K., Kakorin, S., Budde, P., Frey, J. (1998) Mechanism of electroporative dye uptake by mouse B cells. Biophys. J. 74, 1998, 98–108. 6. Neumann, E. (1991) Membrane electroporation and direct gene transfer. Bioelectrochem. Bioenerg. 28, 247–267.

Electroporation 7. Schmeer, M., Seipp, T., Pliquett, U., Kakorin, S., Neumann, E. (2004) Mechanism for the conductivity changes caused by membrane electroporation of CHO cell – pellets. Phys. Chem. Chem. Phys. 6, 5564–5574. 8. Pliquett, U., Elez, R., Piiper, A., Neumann, E. (2004) Electroporation of subcutaneous mouse tumor by rectangular and trapezium high voltage pulses. Bioelectrochemistry 62, 83–93. 9. Neumann, E., Boldt, E. (1990) Membrane electroporation: the dye method to determine the cell membrane conductivity. In Nicolau, C., Chapman, D. (eds.) Horizons in Membrane Technology, Progress in Clinical and Biological Research. Wiley-Liss, New York, pp. 69–83. 10. Miklavcic, D.,Semrov, D., Mekid, H., Mir, L.M. (2000) A validated model of in vivo electric field distribution in tissues for electrochemotherapy and for DNA electrotransfer for gene therapy. Biochim. Biophys. Acta 1523, 73–83. 11. Nishi, T., Kimio, Y., Yanashiro, S., Takeshima, H. Sato, K., Hamada, K., Kitamura, I., Yoshikura, T., Saya, H., Kuratsu, J., Ushio, Y. (1996) High-efficiency in vivo gene transfer using intaarterial plasmid DNA injection following in vivo electroporation. Cancer Res. 56, 1050–1055. 12. Rols, M.P., Delteil, C., Golzio, M., Dumond, P., Cros, S., Teissie, J. (1998) In vivo electrically mediated protein end gene transfer in murine and gene transfer in murine melanoma. Nat. Biotechnol. 16(2), 168–171. 13. Goto, T., Nishi, T., Tamura, T., Dev, S.B., Takeshima, H., Kochi, M., Yoshizato, K., Kuratsu, J.-I., Sakata, T., Hofmann, G.A., Ushio, Y. (2000) Highly effective electro-gene therapy of solid tumor by using an expression plasmid for the herpes simplex virus thymidine kinase gene. PNAS 97(1), 354–359. 14. Lohr, F., Lo, D.Y., Zaharoff, D.A., Hu, K., Zhang, X., Li, Y., Zhao, Y., Dewhirst, M.W., Yuan, F., Li, C.-Y. (2001) Effective tumor therapy with plasmid-encoded cytokines combined with in vivo electroporation. Cancer Res. 61, 3281–3284. 15. Zhang, G.-H., Tan, X.-F., Shen, D., Zhao, S.Y., Shi, Y.-Y., Jin, C.-K., Guo, Y.-H., Chen, K.-H., Tang, J. (2003) Gene expression and antitumor effect following im electroporation

16.

17.

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delivery of human interferon α2 gene. Acta Pharmacol. Sin. 24(9), 891–896. Elez, R., Piiper, A., Kronenberger, B., Kock, M., Brendel, M., Hermann, E., Pliquett, U., Neumann, E., Zeuzem, S. (2003) Tumor regression by combination antisense therapy against Plk1 and Bcl-2. Oncogene 22, 69–80. Cemazar, M., Wilson, I., Dachs, G.U., Tozer, G.M., Sersa, G. (2004) Direct visualization of electroporation-assisted in vivo gene delivery to tumors using intravital microscopy – spatial and time dependent distribution. BMC Cancer 4:81, www.biomedcentral. com/1471-2407/4/81. Buchan, S., Gronevik, E., Mathiesen, I., King, C.A., Stevenson, F.K., Rice, J. (2005) Electroporation as a “Prime/Boost” Strategy for Naked DNA Vaccination against Tumor Antigen1. J. Immunol. 174, 6292–6298. Maucksch, C., Hoffmann, F., Schleef, M., Aneja, M. K., Rosenecker, J., Rudolph, C. (2005) In vitro transfection efficiency of concatameric plasmid DNA using nonviral transfection methods. Abstract book of the 13th Annual Congress of the European Society of Gene Therapy, p. 80. Tjelle, T.E., Salte, R., Mathiesen, I., Kjeken, R. (2006) A novel electroporation device for gene delivery in large animals and humans. Vaccine 24, 4667–4670. Pliquett, U., Gift, E.A., Weaver, J.C. (1996) Determination of the electric field and anomalous heating caused by exponential pulses with aluminum electrodes in electroporation experiments. Bioelectrochem. Bioenerg. 39, 39–53. Gehl, J., Sorensen, T.H., Nielsen, K., Raskmark, P., Nielsen, S.L., Skovsgaard, T., Mir, L.M. (1999) In vivo electroporation of skeletal muscle: threshold, efficacy and relation to electric field distribution. Biochim Biophys. Acta 1428, 233–240. Neumann, E., Pliquett, U., Seipp, T., Schmeer, M.(2003) Biokompatible Elektroden für in vivo Elektroporation zur Wirkstoffzufuhr, Utility model, No. 203 02 861.9. Pliquett, U., Schmeer, M., Seipp, T., Neumann, E. (2002) Fast recovery process after electroporation. IFMBE Proc. 3(1), 98–99.

Chapter 9 Gene Gun Delivery Systems for Cancer Vaccine Approaches Kandan Aravindaram and Ning Sun Yang Summary Gene-based immunization with transgenic DNA vectors expressing tumor-associated antigens (TAA), cytokines, or chemokines, alone or in combination, provides an attractive approach to increase the cytotoxic T cell immunity against various cancer diseases. With this consideration, particle-mediated or gene gun technology has been developed as a nonviral method for gene transfer into various mammalian tissues. It has been shown to induce both humoral and cell-mediated immune responses in both small and large experimental animals. A broad range of somatic cell types, including primary cultures and established cell lines, has been successfully transfected ex vivo or in vitro by gene gun technology, either as suspension or adherent cultures. Here, we show that protocols and techniques for use in gene gun-mediated transgene delivery system for skin vaccination against melanoma using tumor-associated antigen (TAA) human gp100 and reporter gene assays as experimental systems. Key words: DNA vaccine, gene gun, gp100, melanoma, metastasis, mouse model, tumor invasion.

1. Introduction The general aim for cancer treatment has been complete regression of primary and metastatic tumors in the patients or in the animal model hosts. Under this consideration, antigen-specific cancer immunotherapy or vaccine approaches signify a striking strategy for cancer therapy. Gene-based immunization with plasmid DNA and/or other transgene vectors has been shown to offer great potential to modulate humoral and cellular immune responses that can help to protect against cancers in experimental models. Gene gun technology has provided a useful means for


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direct transfer of DNA that can result in useful transgenic protein expression from mammalian gene expression vectors. Epidermal and/or intradermal administration of DNA vaccines using a gene gun apparently has a random chance to deliver specific transgenes into professional antigen-presenting cells (APCs, e.g., Langerhans or dendritic cells) in vivo (1). The gene gun technology was first reported by our research laboratory in 1990 (2) for in vivo and in vitro gene transfer into mammalian somatic tissues. This technology was later extended to various ex vivo gene transfer systems, including excised tissue explants or clumps, organoid tissues placed in culture vessels, and various primary cultures derived from them (3); see Notes 1 and 2). The technique also allows multiple gene delivery to target tissues, and is desirable for assessing the combinational therapy of cytokine or other immune-modifier genes in skin tissues as a model system. Studies have shown that gene gun-mediated immunization can be more efficient than direct needle injection of DNA into muscle, because it elicits similar levels of antibody (Ab) and cellular responses with less DNA. One possible explanation for this result is that gene gun-mediated immunization can more efficiently deliver DNA intracellularly into target cells of test tissues than direct needle injection (4, 5). In vivo electroporation into muscle may provide a higher immune response for DNA vaccines, as compared with gene gun into skin, but the two systems use different APC systems, and may thus complement each other in vaccination approaches (20). Previously we reported that gene gun-mediated vaccination with interleukin (IL)-12 complementary DNA (cDNA) could achieve effective transgenic IL-12 expression in the skin surrounding implanted tumors, and this caused effective regression of established Renca, MethA, SA-1, or L5178Y tumors (6). It is important to note that both humoral and cell-mediated immune responses, as well as Th1 and Th2 T-cell activities, were effectively elicited by using system of gene gun-mediated vaccination of the skin tissue, demonstrated for both small and large experimental animals (7–10). A clinical study to treat melanoma or sarcoma patients with gene-based vaccination, using autologous tumor cells transfected with a GM-CSF cDNA expression vector by the gene gun approach was reported (11), and, recently, a very similar clinical trial protocol to treat melanoma patients with human gp100 and GM-CSF genes was reported ((12); see Notes 4 and 5). Furthermore, a combinational transgene strategy for enhancing the effects of a heterologous prime and boost DNA vaccine has been obtained in our recent studies of cancer vaccines and immunotherapy (13). We found that vaccination of mice with the RANTES gene followed by the human gp100 gene at a specific time point via delivery with gene gun significantly reduced

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Fig. 1. (a) The Helios gene gun device is now commercially available from Bio-Rad. (b) In vitro gene delivery into attached monolayer cells using the Helios gene gun. (c) In vivo gene transfection into murine epidermis. The abdomen hairs were removed with a depilatory cream before the delivery of DNA-coated gold particles. (d) In vivo experiment on treatment of melanoma tumors in experimental mice at an effective DNA vaccine dose. Representative figures of lung tumor nodules in (i) a control (DNA empty vector only) and (ii) a human gp100 cDNA-treated group.

the primary growth and metastases of a melanoma tumor, and substantially enhanced the survival rate of test mice (13). The Helios gene gun (Fig. 1a–c) (Bio-Rad, Hercules, CA) became commercially available to public users in late 1997 and today is still the only standardized and routinely applicable gene gun device that is available on the market. In principle, a helium gas shockwave is used to propel the high-density DNA-coated gold particles to a very high velocity, efficiently enough to penetrate cell membranes of targeted cells/tissues, resulting in a direct physical delivery of plasmid DNA, RNA, or other polymer molecules at high copy numbers in a dry form. In vivo gene transfer has been used most successfully for transfection of skin

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tissue. In this chapter, we describe the protocols and techniques for the particle-mediated gene transfer method for gene transfection into skin tissues in vivo. The very specific application of this technology to cancer gene therapy (transfer of tumor-associated antigen) (Fig. 1d) is herein discussed.

2. Materials 2.1. Instrumentation

1. The commercially available Helios gene gun from Bio-Rad. Instructions for the mechanical operation and/or manipulation of the gene gun and associated devices are given in the manufacturer’s manual. 2. Compressed helium of grade 4.5 or higher. 3. Hearing protection device.

2.2. Elemental Gold Particle Preparations

Microscopic gold particles can be purchased from Degussa (South Plainfield, NJ). Gold particles of 0.95-μm diameter (for in vitro transfection of various leukocytes, lymphocytes, and tumor cells) and 2-μm-diameter particles were found to be the best for in vivo gene transfection into skin (14), and 1- to 2-μm-diameter gold particles were found to be the best for adhesive monolayer cells or cell aggregates and tissue clumps in culture (see Notes 1 and 2). It is important that these gold particles be obtained as elemental gold, not as gold salt or colloidal gold. If necessary, the gold particles can be washed and cleaned by rinsing them in distilled water, 70% ethanol, and 100% ethanol in sequence before use, and they can also be sterilized in phenol or CHCl3 if necessary. It is important to examine each lot of newly purchased gold particle preparation microscopically, making sure that the lot, particle size, and form are correct and appropriate as desired for test systems.

2.3. DNA Vectors

A clean plasmid DNA, for example, human gp100 and a number of pro-inflammatory cytokines (tumor necrosis factor [TNF]-α, nuclear factor [NF]-κB, and granulocyte macrophage colony stimulating factor [GM-CSF] (27, 28), dissolved in TE buffer (10 mM Tris-HCl, pH 7.0, and 1 mM EDTA) or distilled water should be used for coating particles. Cocktails of different DNA vector systems or preparations can be mixed in desired molar ratios in aqueous solution and then effectively loaded onto gold particles as follows (see Note 6). For exploratory gene transfer experiments, convenient and sensitive reporter gene systems that have low endogenous


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activity background (e.g., green fluorescence protein and luciferase) are recommended for verification of transient gene expression systems. 2.4. Coating of DNA onto Gold Particles for Gene Transfer

1. 0.05 M spermidine or polyethylene glycol (PEG) in H2O. Use fresh spermidine made weekly from a free-base solution (Sigma, St. Louis, MO). 2. 1 mg/mL polyvinylpyrrolidone (PVP; Sigma). 3. 2.5 M CaCl2 in H2O. 4. 100% ethanol kept at −20°C.

2.5. Treatment and Care of Skin

1. Oster electric hair clippers with a no. 40 blade (Fisher, Pittsburgh, PA). 2. Tegaderm adhesive (3M, St. Paul, MN) or Scotch tape (3M) or Nair depilatory hair remover (Princeton, NJ).

2.6. Tissue Extraction Buffers and Enzyme Assay Systems 2.6.1. Luciferase Assay

1. Extraction buffer: 100 μL of cold 10% Triton X-100 in 9.9 mL of cold phosphate-buffered saline (PBS). 2. Luciferase assay substrate (Promega, Madison, WI), dissolved in luciferase assay buffer at a concentration of 13.33 mg/ mL. 3. Luminometer (Lumat LB 9507, Berthold Systems, Pittsburgh, PA).

2.6.2. hAAT Assay

1. Antibody dilution buffer: 5% fetal bovine serum (FBS) in PBS. 2. First antibody: Goat antihuman α1-antitrypsin (Sigma) diluted in 50 mM carbonate buffer at a concentration of 3 μg/mL. 3. Second antibody: Rabbit antihuman α1-antitrypsin (hAAT) (Sigma) at least 1,000× diluted in dilution buffer. 4. Third antibody: Goat anti-rabbit IgG (H + L), peroxidaseconjugated (Pierce, Rockford, IL) at least 1,000× diluted in dilution buffer. 5. Substrate solution: Add 10 μL H2O2 to 1 mL 3,3′,5,5′-tetramethylbenzidine (TMB; Pierce) or 2,2′-azinobis(3-ethylbenzothiazoline-6-sulfonic acid)-diammonium salt (ABTS; Pierce). 6. Blocking solution: 0.5% bovine serum albumin (BSA) and 0.01% sodium azide in PBS. 7. Wash buffer: 0.05% Tween-20 in PBS, prepared freshly.

2.6.3. X-Gal Assay

1. 5-Bromo-4-chloro-3-indolyl-8-D-galactoside (X-gal) substrate, dissolved in dimethyl formamide at a concentration of 40 mg/mL.

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2. Buffer solution: 44 mM HEPES, 15 mM NaCl, 1.3 mM MgCl2, 3 mM K+ ferricyanide, 3 mM K+ ferrocyanide, pH 7.4. 3. X-gal buffer: Add X-gal substrate to buffer solution, making a final solution of 1 mg/mL. 2.6.4. Cytokines and Other Transgenic Protein Products Assays

General extraction buffer: 9.5 mL PBS (0.2 g potassium chloride, 0.2 g potassium phosphate monobasic, 8.0 g sodium chloride, and 1.15 g sodium phosphate dibasic in 1 L distilled H2O), 2.4 mg serine protease inhibitor Pefabloc®SC (4-[2aminoethyl]-benzenesulfonylfluoride hydrochloride; Roche), 0.5% Triton X-100, pH 7.2.

3. Methods 3.1. Coating DNA onto Gold Particles

1. Prepare DNA solution at a concentration of approximately 1 μg/μL and store at 4°C. 2. One cartridge of the Helios gene gun device (for one transfection) contains approximately 0.5 mg of gold particles. 3. Assuming that 40 cartridges will be needed, weigh 21 g of gold particles into a 1.5-mL microcentrifuge tube. 4. Add 250 μL of 0.05 M spermidine or PEG to the tube with the gold particles (e.g., use 200–300 μL for 20–50 mg and 400–500 μL for 120 mg). 5. Vortex and sonicate for 3–5 s to break up gold clumps. 6. Add DNA at 2.5 μg DNA/mg gold particle to 21 mg of gold. 7. Add 250 μL of 2.5 M CaCl2 (or the same volume as spermidine) dropwise while vortexing the tube at a low speed. 8. Incubate at room temperature for 10 min. While waiting, prepare a culture tube with 3 mL of 100% ethanol. 9. Microcentrifuge for 3–5 s. Remove and discard the supernatant. Break up the pellet by flicking the tube. Add 0.5 mL of cold 100% ethanol dropwise while gently vortexing the tube and then add 0.5 mL more of 100% ethanol. Mix by inversion. Wash the pellet with cold ethanol three times. 10. Transfer particles into a culture tube containing 3 mL of 100% ethanol (7 mg gold particle/mL ethanol). Sonicate to disperse particles. Particle suspensions can be used immediately or stored at 4°C or −20°C under stringent desiccation for 2–3 h.


3.2. Preparation of DNA Cartridges

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1. Sonicate DNA-coated gold particle suspension for 2–3 s, vortex, add PVP at 0.01 mg/mL, sonicate again, and immediately load the suspension into the Tefzel tubing following the manufacturer’s instruction (Bio-Rad). 2. Discard unevenly coated ends or portions of the tubing and cut the tubing into 0.5-inch pieces. Cartridges may be stored desiccated in a tightly sealed container at 4°C for several weeks.

3.3. Animal Care and Skin Treatment

1. Experimental animals (including mouse, rat, hamster, and rabbit) usually do not need to be anesthetized for particlemediated gene transfer into the skin. However, larger animals such as dogs and pigs in general need to be anesthetized before treatment by using ketamine (10 mg/kg body weight for intramuscular injection or 5 mg/kg for intravenous injection) or pentothal (12.5 mg/kg body weight for intramuscular injection or 5 mg/kg for intravenous injection) under the guidance of a consulting veterinarian as required by the Animal Welfare Act and administered by the institution’s Animal Care and Use Committee. 2. Animal hair in the target area (mostly the abdominal area for cancer gene therapy or vaccine studies) should be removed with clippers and shearing blades. We also used Nair depilatory cream to remove hair from animals. If a depilatory is used to remove stubble, the animals should be anesthetized.

3.4. Epidermis and Dermis Gene Transfer (see Notes 1 and 4)

1. Allow the container with the DNA–gold cartridges to reach room temperature before opening. 2. Attach the regulator to the compressed helium tank. Connect the feed hose to the regulator and gene gun, and then plug in the device. 3. Load the prepared cartridges into the 12-chamber cartridge holder, following the manufacturer’s instructions. 4. Put on a hearing protection device. Insert the loaded cartridge holder into the barrel of the gene gun device. Open the helium tank valve and the regulator valve. 5. Adjust the discharge pressure to the desired setting (usually 250–400 psi). 6. Restrain a mouse or other testing small animal in a hand or with a steady setup. Hold the nozzle of the device against the target skin area and discharge the device. If several skin transfections are required, turn the cartridge holder clockwise and discharge at another target skin area (Fig. 1c).

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7. If desired, apply a semiocclusive skin dressing such as Tegaderm (3M) or Opsite (Smith and Nephew, Hull, UK) to the transfected epidermis. 8. To transfect dermal tissues, anesthetize a mouse, make an incision, and dissociate the full-thickness skin tissue of the target size (∼3.2 cm2) from the facies and muscle tissue using standard surgical procedures and tools. Flip the skin flap over, exposing the dermal tissue, and transfect the fibroblasts, muscle cells, and other stromal cell types. Moisten the dermal tissues with sterile saline before closing. Close the incision with sutures or wound clips. 9. Collect skin samples from target sites at a designated time point for transgene activities or sequences as follows. 3.5. Reporter Gene Expression Assay (see Note 3) 3.5.1. Tissue Extraction

1. Collect transfected skin tissues from test animals by excising a small portion of the targeted skin area. A piece as small as 1–2 mm2 can be enough for Luc or X-gal reporter gene expression assays. The transgenic expression products may also be stripped off the target sites of the skin with Scotch tape or duct tape. 2. Drop the freshly excised skin piece or tape into a tube with the appropriate buffer (0.2–0.5 mL) and keep on ice if performing the assay right away; otherwise freeze at −20°C. 3. Cut with scissors, grind, or homogenize the frozen skin or tape immediately before carrying out the assay. The samples should be kept on ice. 4. Sonicate and then centrifuge the sample at a high speed to separate the tissue or tape from the lysate. Crude tissue extracts are used for various reporter gene assays.

3.5.2. Luciferase Assay

1. After the lysate has been collected, prepare dilutions in PBS if necessary. 2. Add 100 μL Luc assay substrate and 20 μL extraction buffer to the tube, and vortex. 3. Read the relative light units in a luminometer and run a standard curve to quantify the results.

3.5.3. hAAT Assay

1. Add 100 μL of skin tissue extract or tape-stripped cell extract to a 96-well plate precoated with anti-hAAT antibody. 2. Incubate at 37°C for 1 h. Add 90 μL of the secondary antibody (rabbit anti-hAAT, at least 1,000× diluted), and incubate again for 1 h. 3. Wash 3–5 times with wash solution (300 μL/well). 4. Add 85 μL of the third antibody (anti-rabbit, at least 1,000× dilution) and incubate for another 1 h. Wash 3–5 times.


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5. Add 80 μL substrate solution. Allow color formation at room temperature for 1/2–1 h. 6. Measure transgenic hAAT levels by an enzyme-linked immunosorbent assay (ELISA) reader. 3.5.4. X-Gal Staining

1. Perform whole-mount tissue staining by placing the excised skin target into X-gal buffer. For best results, glue the skin tissue onto the surface of a 35-mm dish and keep the skin stretched out for better examination. The tissue can also be fixed in methanol/acetone (1:1) for 10 min and then stained with X-gal buffer. 2. Section the tissue into approximately 10-μm sections by cryostat microtome or paraffin sectioning. Place the slide in a cold 1.5% glutaraldehyde solution for 10 min, and wash in cold PBS 5X for 5 min. Then stain the tissue slides with X-gal buffer for 1 h. Avoid prolonged staining, because it can often result in a nonspecific greenish blue background that can develop in hair follicles of certain skin tissues.

3.5.5. Cytokine Assay

ELISA tests are usually used for quantification of cytokines. Test reagents may be purchased as a kit or antibody pair samples and used in a sandwich-style assay. The ELISAs can be run using skin crude tissue extract (with general extraction buffer) or tapestripped cell extract.

3.5.6. Mouse Lung Metastasis Assay

Mice are intravenously injected with test tumor cells (e.g., B16 melanoma) before vaccination, and 5 days after tumor inoculation, mice are vaccinated with human gp100 cDNA expression plasmid at the abdominal skin area as described above. Three weeks later, mice are killed and colonies of tumor metastasis in lung tissue are scored (Figs. 1d and 2).

Fig. 2. Lung metastases in mice intravenously injected with test melanoma cells, where mice were treated with human gp100 cDNA vaccine or empty vector (control) only. The number of lung metastatic nodules in mice was visually scored 3 weeks after tumor injection, mean ± SEM; P < 0.01.

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4. Notes 1. The gene gun method as a nonviral system for gene transfer can be applied to a broad range of tissues and cell types; a key feature of this technology is its applicability to in vivo gene transfer to various tissues and organs, especially the epidermal skin cell layers (2, 3, 15, 16). The latter case thus permits a powerful nucleic acid or other gene-based vaccine strategy. 2. Two key technical advantages were observed for the particle-mediated gene delivery method: (1) a very wide range for DNA load (1 ng–10 μg/dose/transfection site) can be delivered into a 3- to 5-cm2 surface area of targeted tissues, resulting in different efficacies of transgene expression, depending on target cells, tissues, or organ types as well as the in vitro, ex vitro, or in vivo experimental conditions; and (2) there is little or no restriction on the molecular size or form of testing DNA, at least at a molecular size 40 kb as double-stranded DNA (14, 17, 18). 3. Various transgenic proteins expressed by reporter transgenes (e.g., Luciferase, green fluorescence protein, and β-Gal), candidate therapeutic genes (e.g., granulocyte-macrophage colony-stimulating factor (19), or vaccine/antigen genes (e.g., influenza, foot and mouth disease viral proteins) (20) have been readily detected at 0.6–2 ng for cytokines and luciferases, 0.2–5 μg or higher for relatively stable proteins such as human growth hormone or human α1 anti-trypsin, and 10–200 pg for specific viral protein antigens (e.g., SARS, HIV antigens, N. S. Yang et al., 2008). 4. The high accessibility of the skin as an exposed tissue and the organization of the epidermis make the skin an excellent target for gene gun-mediated gene transfer, not only for gene-based vaccination against infectious diseases, but also for serving as a transgenic bioreactor for gene therapy approaches, including cytokine gene therapy for cancer and DNA cancer vaccines, using either in vivo or ex vivo gene delivery strategies (21, 22). 5. Gene gun technology has also been applied in clinical trials of human gene therapy as a cancer vaccine, as a hepatitis B DNA vaccine, and for HIV gene therapy approaches (23–26). 6. Other than DNA and RNA (13, 14), peptide nucleic acid (PNA) (P. L. Fuller, 2004) may also be effectively used as a vector for gene-based vaccination using gene gun delivery.


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Acknowledgments This work was supported by a grant (No. 96-2320-B-001–008) from the National Science Council and a grant (No. 95N-1003) from the National Science and Technology Program for Agricultural Biotechnology, Taiwan. References 1. Condon, C., Watkins, S.C., Celluzzi, C.M., Thompson, K., and Falo, L.D. Jr. (1996). DNA-based immunization by in vivo transfection of dendritic cells. Nat. Med. 2, 1122– 1128. 2. Yang, N.S., Burkholder, J., Roberts, B., Martinell, B., and McCabe, D. (1990). In vivo and in vitro gene transfer to mammalian somatic cells by particle bombardment. Proc. Natl. Acad. Sci. USA 87, 9568–9572. 3. Cheng, L., Ziegelhoffer, P., and Yang, N.S. (1993). In vivo promoter activity and transgenic expression in mammalian somatic tissues evaluated by using particle bombardment. Proc. Natl. Acad. Sci. USA 90, 4454–4459. 4. Pertmer, T.M., Eisenbraun, M.D., McCabe, D., Prayaga, S.K., Fuller, D.H., and Haynes, J.R. (1995). Gene gun-based nucleic acid immunization: elicitation of humoral and cytotoxic T lymphocyte responses following epidermal delivery of nanogram quantities of DNA. Vaccine 13, 1427–1430. 5. Feltquate, D.M., Heaney, S., Webster, R.G., and Robinson, H.L. (1997). Different T helper cell types and antibody isotypes generated by saline and gene gun DNA immunization. J. Immunol. 158, 2278–2284. 6. Rakhmilevich, A. L., Turner, J., Ford, M. J., McCabe, D., Sun, W. H., Sondel, P. M., et al. (1996). Gene gun-mediated skin transfection with interleukin 12 gene results in regression of established primary and metastatic murine tumors. Proc. Natl. Acad Sci USA 93, 6291– 6296. 7. Fuller, D.H., Corb, M.M., Barnett, S., steamer, K., and Haynes, J.R. (1997). Enhancement of immunodeficiency virus-specific immune responses in DNA-immunized rhesus macaques. Vaccine 15, 924–926. 8. Yamano, T., Kaneda, Y., Huang, S., Hiramatsu, S.H., and Hoon, D.S.B. (2006). Enhancement of immunity by a DNA melanoma vaccine against TRP2 with CCL21 as an adjuvant. Mol. Ther. 13, 194–202. 9. Yang, S.C., Batra, R.K., Hillinger, S., Reckamp, K.L., Strieter, R.M., Dubinett, S.M.

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et al. (2006). Intrapulmonary administration of CCL21 gene-modified dendritic cells reduced tumor burden in spontaneous murine bronchoalveolar cell carcinoma. Cancer Res. 66, 3205–3213. Lian, H., Jin, N., Li, X., Mi, Z., Zhang, J., Sun, L. et al. (2007). Induction of an effective anti-tumor immune response and tumor regression by combined administration of IL-18 and apoptin. Cancer Immunol. Immunother. 56, 181–192. Mahvi, D.M., Sondel, P.M., Yang, N.S., Albertini, M.R., Schiller, J.H., Hank, J., et al. (1997). Phase I/IB study of immunization with autologous tumor cells transfected with the GM-CSF gene by particle-mediated transfer in patients with melanoma or sarcoma. Hum. Gene Ther. 8, 875–891. Cassaday, R.D., Sondel, P.M., King, D.M., Macklin, M.D., Gan, J., Warner, T.F., et al. (2007). A phase I study of immunization using particle-mediated epidermal delivery of genes for gp100 and GM-CSF into uninvolved skin of melanoma patients. Clin. Cancer Res. 13, 540–549. Aravindaram, K., Yu, H.H., Lan, C.W., Wang, P.H., Chen, Y.H., Chen, H.M. et al. (2009). Transgenic expression of human gp100 and RANTES at specific time points for suppression of melanoma. Gene Ther. (under revision). Qiu, P., Ziegelhoffer, P., Sun, J., and Yang, N.S. (1996). Gene gun delivery of mRNA in situ result in efficient transgene expression and immunization. Gene Ther. 3, 262–268. Jiao, S., Cheng, L., Wolff, J., and Yang, N.S. (1993). Particle bombardment mediated gene transfer and expression in rat brain tissues. Bio/Technology 11, 497–502. Burkholder, J.K., Decker, J., and Yang, N.S. (1993). Transgene expression in lymphocyte and macrophage primary cultures after particle bombardment. J. Immunol. Methods 165, 149–156. Christou, P. (1994). Application to plants, in Particle Bombardment Technology for Gene Transfer (Yang, N.S. and Christou, P.,

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Aravindaram and Yang eds.), Oxford University Press, New York, pp. 71–99. Yang, N.S. and Ziegelhoffer, P. (1994). The particle bombardment system for mammalian gene transfer, in Particle Bombardment Technology for Gene Transfer (Yang, N.S. and Christou, P., eds.), Oxford University Press, New York, pp. 117–141. Rakhmilevich, A.L., Imboden, M., Hao, Z., Macklin, M.D., Roberts, T., Wright, K.M., et al. (2001). Effective particle-mediated vaccination against mouse melanoma by coadministration of plasmid DNA encoding gp100 and granulocyte-macrophage colony-stimulating factor. Clin. Cancer Res. 7, 952–961. Yang, N.S., Wang, J.H., Lin, K.F., Wang, C.Y., Kim, S.A., Yang, Y.L., et al. (2005). Comparative studies of the capsid precursor polypeptide P1 and the capsid protein VP1 cDNA vectors for DNA vaccination against foot-and-mouth disease virus. J. Gene Med. 7, 708–717. Guo, Z., Yang, N.S., Jiao, S., Sun, J., Cheng, L., Wolff, J.A., et al. (1996). Efficient and sustained transgene expression in mature rat oligodendrocytes in primary culture. J. Neurosci. Res. 43, 32–41. Sun, W.H., Burkholder, J.K., Sun, J., Culp, J., Turner, J., Lu, X.G., et al. (1995). In vivo cytokine gene transfer by gene gun reduces tumor growth in mice. Proc. Natl. Acad. Sci. USA 92, 2889–2893. Mahvi, D.M., Burkholder, J.K., Turner, J., Culp, J., Malter, J.S., Sondel, P.M., et al.

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(1996). Particle-mediated gene transfer of granulocyte-macrophage colony-stimulating factor cDNA to tumor cells: Implications for a clinically relevant tumor vaccine. Hum. Gene Ther. 7, 1535–1543. Tan, J., Yang, N.S., Turner, J.G., Niu, G.L., Maassab, H.F., Sun, J., et al. (1999). Interleukin-12 cDNA skin transfection potentiates human papillomavirus E6 DNA vaccineinduced antitumor immune response. Cancer Gene Ther. 6, 331–339. Woffendin, C., Yang, Z., Udaykumar, K., Xu, L., Yang, N., Sheehy, M.J., et al. (1994). Non viral and viral delivery of a Human immunodeficiency virus protective gene into primary human T cells. Proc. Natl. Acad. Sci. USA 91, 11581–11585. Hogge, G.S., Burkholder, J.K., Culp, J., Albertini, M.R., Dubielzig, R.R., Yang, N.S., et al. (1999). Preclinical development of human granulocyte-macrophage colonystimulating factor-transfected melanoma cell vaccine using established canine cell lines and normal dogs. Cancer Gene Ther. 6, 26–36. Staniforth, V., Chiu, L.T., and Yang, N.S. (2006) Caffeic acid suppresses UVB radiationinduced expression of interleukin-10 and activation of mitogen-activated protein kinases in mouse. Carcinogenesis 27, 1803–1811. Chiu , S.C. and Yang , N.S. (2007) Inhibition of tumor necrosis factor-alpha through selective blockade of Pre-mRNA splicing by shikonin . Mol. Pharmacol. 71 , 1640 – 1645 .

Chapter 10 Ultrasound-Mediated Gene Transfection Loreto B. Feril Jr. Summary Ultrasound-mediated gene transfection (sonotransfection) has been shown to be a promising physical method for gene therapy, especially for cancer gene therapy. The procedure being done in vitro uses several ultrasound exposure (sonication) setups. Although high transfection rates have been attained in some of these setups in vitro, replicating similar levels of transfection in vivo has been difficult. In vivosimulated setups offer hope for a more consistent outcome in vivo. Presented in this chapter are typical methods of sonotransfection in vitro, methods when using a novel in vivo-simulated in vitro sonication setup and also sonotransfection methods when doing in vivo experiments. Factors that could potentially influence the outcome of an ultrasound experiment are cited. Several advantages of sonotransfection are recognized, although a low transfection rate is still considered a disadvantage of this method. To improve the transfection rate and the efficiency of sonotransfection, several studies are currently being undertaken. Particularly promising are studies using engineered microbubbles to carry the therapeutic genes into a particular target tissue in the body, then using ultrasound to release or deliver the genes directly into target cells, e.g., cancer cells. Key words: Cancer therapy, HIFU, sonotransfection, ultrasound-mediated transfection.

1. Introduction The use of ultrasound in medicine has been evolving rapidly in recent years. Not only has the use of ultrasound for diagnosis advanced from two-dimensional (2D) to three-dimensional (3D) then to four-dimensional (4D) imaging, but also new medical applications of ultrasound have been introduced. This is exemplified by its use as a therapeutic tool to treat tumors and many other potential applications under investigation, including the use of ultrasound for gene therapy(1–3) (Fig. 1).


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G. Thermal ablation

B. Cell necrosis

H. Gene regulation

C. Apoptosis CANCER D. Cell-growth inhibition

I. Tumor growth inhibition J. Metastasis inhibition

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K. Angiogenesis inhibition L. Sonopermeabilization

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EFFECTIVE CANCER TREATMENT

Fig. 1. Therapeutic ultrasound. Several bioeffects of ultrasound have been observed. Thermal ablation (G) of tumors using high-intensity focused ultrasound (HIFU) is currently in clinical application. Other effects of ultrasound are under investigation for possible clinical applications. Most of these effects can be attained using a relatively low-intensity ultrasound. One of these is the ultrasound-mediated gene transfection or sonotransfection (F) for its potential application in cancer gene therapy.

1.1. Ultrasound in Therapy

The use of ultrasound in therapy is a new modality revived from an old concept of using ultrasound radiation to treat deep-seated target tissues, such as tumors. Recent developments in the application of ultrasound to extracorporeally target deeper targets has been made possible also because of advances in imaging techniques such as the use of diagnostic ultrasound itself. High-intensity focused ultrasound (HIFU) technologies are now in clinical use (1, 4). As the name implies, in HIFU, ultrasound is focused to concentrate the energy into small volumes to thermally destroy the diseased tissue. This modality has been applied in treating prostate cancers, breast tumors, fibroids, and abdominal tumors (5–8). Although the mechanism is mainly thermal ablation (Fig. 1G), other mechanisms do exist (2).

1.2. Bioeffects of Ultrasound

Studies on the bioeffects of ultrasound are leading us to a better understanding of the mechanisms underlying how ultrasound at different intensities, frequencies, duty factors (DF), pulse repetition frequencies (PRF), and microenvironmental conditions can affect living cells and tissues both in vitro and in vivo (9, 10). Consequently, understanding the mechanism of bioeffects has led to more ideas on how to use ultrasound for therapy for a wide array of illnesses (Fig. 1). Among the potential applications is the use of ultrasound for gene transfection with cancer gene therapy in mind.

Ultrasound-Mediated Gene Transfection

1.3. UltrasoundMediated Gene Transfection (Sonotransfection)

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The discovery that most human diseases, such as cancers, genetic disorders, and metabolic disorders, are somehow gene related has led to a concept of therapy, called gene therapy (11). Gene therapy is carried out by introducing recombinant genes (therapeutic genes) into somatic cells to alter the course of a disease process. Several strategies have been designed to introduce functional genes and to allow integration into the nucleus of target cells. Viral-mediated gene transfer is efficient for the task, but cytotoxicity and antigenicity are among the limiting factors in therapeutic application. This is the reason behind the search for alternative methods to deliver therapeutic genes into target cells; a search for a nonviral method that is safer than the viral method (12, 13). These methods include electrotransfection, a physical method, and liposome-mediated transfection, a chemical method (14). The use of ultrasound in therapy and also in gene transfection has been investigated both in vitro and in vivo (2, 15–18). Just like the other nonviral methods, poor transfection rates remain a problem, so combinations with other methods are being used to try to improve the outcomes (19, 20). An attempt to optimize the conditions for ultrasound-mediated gene transfection has been made on skeletal muscle cells, but so far the mechanism remains generally unknown. The leading belief, however, is that ultrasound increases DNA uptake by the cells. Previously, we showed that optimizing ultrasound-induced apoptosis is possible based on the concept that the bioeffects of ultrasound are mainly caused by mechanical damage on the cell membranes (2, 21, 22). Understanding the membrane damage induced by ultrasound and the physiology of cellular membrane repair has led to an optimal apoptosis induction on cells in vitro using pulse ultrasound (23). Pulsing, at certain pulse duration, induces membrane damage, and the period of “no radiation” allows the membrane to undergo some repair. This has been the basis of the optimized sonotransfection in vitro (3). Sonotransfection methods that are being applied in vitro and in vivo are presented in this chapter.

2. Materials

2.1. Cells and Cell Culture

Different types of cancer cell lines have been used for sonotransfection. Cells that grow as suspension in a liquid culture medium include: (1) U937 (human monohistiocytic lymphoma) cell line, and (2) Meth A (murine fibrosarcoma); whereas cells that grow at the bottom of a treated culture dish include: (1) HeLa (human epithelial cervix adenocarcinoma), (2) PC-3 (human

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epithelial prostate adenocarcinoma), and (3) T-24 (human epithelial urinary bladder transitional carcinoma). Most of these cell lines can be obtained from ATCC, Manassas, VA, USA and also from the Human Sciences Research Resources Bank Human Sciences Foundation, Tokyo, Japan. All of these cell lines can be maintained in RPMI 1640 medium (Invitrogen, Groningen, The Netherlands) supplemented 10% heat-inactivated fetal bovine serum (FBS) (Gibco, Carlsbad, CA) at 37°C in humidified air with 5% CO2. In most instances, use of cells in log-phase is ideal and cell viability must be good (e.g., over 95%) when used for ultrasound experiments. 2.2. Ultrasound Apparatus and Intensity Measurement

Several ultrasound devices have been used in sonotransfection experiments using several ultrasound exposure (sonication) setups, including those described in Fig. 2. Examples of these ultrasound devices are: 1. ITO Ultrasonic by ITO ultrasonics Tokyo, Japan. With a 1-MHz resonant frequency, this device has two settings. The first setting is continuous whereas the other is pulsed with a variable PRF ranging from 0.5 to 100 Hz at a fixed DF of 50%. The plane unfocused transducer with a diameter of 3 cm can be used and is ideal with a 35-mm culture dish in the sonication experiments. The spatial average temporal average intensity (ISATA) values at continuous wave are 0.054, 0.233, 0.634, and 0.865 W/cm2 for the intensity readings 0.5, 1.0, 1.5, and 2 W/cm2, respectively. 2. Sonicmaster ES-2 by OG Giken Co. Ltd, Okayama, Japan With a resonant frequency of 1 MHz and 100 Hz PRF, this device is equipped with a built-in digital timer, intensity regulator, and DF controller. The transducer with a diameter of 5 cm can be fixed and mounted for experiments involving setups A, B, and C in Fig. 2, using an ultrasound power meter (UPM-DT-10E, Ohmic Instrument Co., Easton, MD), ISATA of 0.3 and 0.5 W/cm2 at 10% DF were 0.081 and 0.105 W/ cm2, respectively. The above intensities are the main intensities used in an optimization study (3). 3. ES-1 by OG Giken Co. Ltd, Okayama, Japan With a resonant frequency of 1 MHz, the transducer with a diameter of 3 cm is horizontally directed to hit the sample contained in the lower cylindrical portion (1.9-cm long and 2 cm in inner diameter with both ends covered with 6-mmthick polyester Mylar films) of a specially designed inverted “T” glass tube at a distance 20 cm used in previous studies (10, 15, 22). The effective output was determined and it was found that the intensities 0.5 and 1 W/cm2 on the reading meter of the device have measured values of 0.312 and 0.692 W/cm2, respectively.


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b DW S

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Fig. 2. In vitro setup for sonication experiments. a. Ultrasound travels horizontally through degassed distilled water (DW) to hit the sample (S) positioned at a distance. At the far end is an ultrasound absorber (UA) that prevents reflection of ultrasound. b. A dish containing the sample is positioned directly on top of the transducer (T) after applying acoustic gel to avoid air in between the transducer and the dish. c. The sample in a tube (could be rotated during sonication) is positioned a few centimeters from the transducer. Ultrasound is directed upward to hit the sample. d. For small samples (such as those using 24- or 96-well plates) small transducers (e.g., 2–10 mm in diameter) can be dipped directly into the sample for sonication.

4. SonoPore KTAC-4000 by Nepa Gene, Chiba, Japan. This device is variable in transmission frequency (0.2–5 MHz), voltage (0–100 V), DF (1–100%), burst rate or PRF (0.5–100 Hz), irradiation duration (0.1 s– h), pulse type (rectangular or sine), and pulse frequency pattern. The pulse frequency patterns have four types: type 1 (low frequency to low), type 2 (high to low), type 3 (low–high–low), and type 4 (high–low– high). The “sweeping” of frequencies can also be varied as to the sweep width or frequency range (0.1–99.9%) and sweep interval (0.2–100 ms). This device is used in the in vivo-simulated in vitro setup.

3 Methods 3.1. Measurement of Cell Viability by Trypan Blue Dye Exclusion Test

To determine the best condition to attain optimal transfection rate, you also need to know the loss of viability caused by sonication. The Trypan blue dye exclusion test is a simple method to measure loss of cell viability.

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1. The Trypan blue exclusion test is performed by mixing 200 μL of cell suspension with an equal amount of 0.3% Trypan blue solution (Sigma, St. Louis, MO, USA) in phosphatebuffered saline (PBS). 2. After 5 min of incubation at room temperature, count the cells excluding Trypan blue (unstained) using a Burker Turk hemocytometer (EKDS, Tokyo, Japan) to estimate the number of viable cells immediately after sonication. The viability ratio after sonication is calculated as the number of viable cells after sonication divided by the number of viable cells before sonication. 3.2. Luciferase Assay

The luciferase assay is used to assess ultrasound-mediated DNA transfection of cells, particularly to determine the transfection efficiency. 1. Plasmid vectors pGL3-control with the luciferase gene (Clontech Laboratories, Palo Alto, CA, USA) may be used as reporter. You can prepare plasmids on a large-scale basis using an endotoxin-free plasmid preparation kit (EndoFree Plasmid Kit; Qiagen, Valencia, CA, USA) and this can be stored at −20°C until use. 2. Twenty-four hours after treatment (e.g., sonication), prepare the cell lysate and assay the luciferase expression with the luciferase assay kit (Promega, Madison, WI, USA) using a luminometer (Turner designed luminometer TD-20/20; Promega). 3. Wash the cells in a culture dish twice with PBS before adding 500 μL of lysis buffer. 4. After a 15-min incubation at room temperature, collect the cell lysate and resuspend by pipetting. 5. Mix 20 mL of the supernatant with 100 μL of luciferase assay reagent. 6. Measure the luciferase expression using a luminometer. Count the luminescence of each sample for 10 s. 7. A portion of the lysate can be used to determine the protein concentration with a protein assay kit (Bio-Rad Laboratories, Hercules, CA, USA). 8. Mix 20 μL of each lysate with 1 mL of the protein assay reagent and place at room temperature for 5 min before analyzing using a spectrophotometer (DU60; Beckman Coulter, Fullerton, CA, USA) at 595-nm absorbance. 9. The protein concentration is calculated according to a standard curve plotted using IgG as a standard protein reference. The enzyme activity is expressed in relative light units (RLU) per milligram of protein where the luminescence count of


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each sample is divided by its protein content in milligrams to standardize the luciferase activity in accordance with the protein concentration of the cell lysate. 3.3. Fluorescence Microscopy and Flow Cytometry

To determine the transfection rate of a sonotransfection experiment, fluorescence microscopy, visually counting the transfected cells, or flow cytometry may be performed. 1. Using a plasmid containing the green fluorescence protein gene, pEGFP-N1, as a reporter, add medium containing pEGFP-N1 (final concentration of 1–100 μg/mL) into the sample cells. 2. Sonicate the cells in the presence of the pEGFP-N1 and microbubbles (see Subheadings 2.6.1 and 2.6.2 for the sonotransfection procedures). 3. Incubate the sonicated cells for 24 h. 4. Visualize the GFP expression using a fluorescence microscope (e.g., Nikon Eclipse TE 300; Nikon Corporation, Tokyo, Japan) with a fluoresceinisothiocyanate (FITC) filter and a digital camera (Hamamatsu Photonics K. K., Hamamatsu, Japan). The digital camera can be connected directly to a computer to capture visualized images and stored as computer files. 5. Determine the transfection rate by counting the cells in a number of fields and dividing the number of cells with GFP by the total number of cells counted in the selected fields. 6. The transfection rate can also be measured by flow cytometry. To do this, harvest the cells by trypsinization, wash them with PBS, and suspend them in a PBS solution before flow cytometry (Epics XL; Beckman Coulter). Cells emitting signals greater than the control are considered positive for GFP (or FITC). Normalize the values by deducting the reading values of the control (a sample that was not sonicated but was also treated with the GFP gene). The transfection rate of each sample is calculated as the percentage of cells with FITC signals above that of the control, divided by the total number of cells counted for that particular sample.

3.4. In Vitro Sonotransfection (24) 3.4.1. General Method (see Note 1)

1. Harvest the cells by trypsin treatment for attached cells and by centrifugation for suspension cells. 2. Resuspend the cells (e.g., four million) in 3.5 mL degassed Eagle’s minimum essential medium (Nissui Pharmaceuticals Co., Ltd., Tokyo, Japan) in a container for sonication, e.g., a polystyrene tube (Corning, Inc., Corning, NY, USA). 3. Just before sonication, add reporter plasmid DNA (e.g., pBKCMV-luc) to the cell suspension at a concentration

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of 10 μg/mL along with the microbubbles (see Notes 2 and 3). 4. Position the tube or dish containing the sample in front of the transducer. For the A and C setups in Fig. 2, the degassed distilled water in the tank may be circulated and adjusted at various temperatures (e.g., by Coolnit CL-80F, Taitec Co., Ltd., Tokyo, Japan), if desired (see Note 4). 5. Expose the sample to ultrasound (e.g., 1.0 MHz, DF of 50%, 3.6 W/cm2 in ISATA for 20 s). If using a tube in setup C of Fig. 2, the tube may be rotated during sonication at 30 rpm for uniform exposure. 6. After sonication, change the culture medium with freshly prepared medium, then incubate for 24 h before assay. 3.4.2. Sonicating Attached Cells

1. For attached cell types (e.g., HeLa) (Fig. 3), plate about one million cells in a 35-mm culture dish and incubate for 24 h to

1. Prepare cells

Cells in freshly prepared culture medium Plasmid DNA

2. Add plasmid DNA

3. Add microbubbles Add microbubbles just before sonication 4. Sonicate

5. Change culture medium

6. Incubate for 24 hrs

Ultrasound

Change the culture medium before incubation

Assay for transfection: GFP imaging or flow cytometry

7. Assay for transfection

Luciferase assay ELISA

Fig. 3. Sonication in vitro. Cells in a culture dish can be sonicated directly without being detached. This is usually applicable if sonication is performed according to a setup similar to setup B of Fig. 2. In this case, the diameter of the ultrasound transducer should not be less than the diameter of the dish containing the cells.


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attain at least 60% confluence (above 90% confluence may not be advisable, see Note 5). 2. Add 10 μg of expression vector to 0.75 mL culture medium containing the cells, incubate for 30 min, then add 0.75 mL freshly prepared medium containing microbubbles before sonication. Without microbubbles, bubbling air into the medium or slightly shaking the medium before adding the medium into the sample will help facilitate cavitation formation during sonication (see Note 2). 3. Collect the treated cells, then add 2.0 mL culture medium and incubate overnight before the cells are assayed for luciferase expression. Typical results of an optimized sonotransfection on HeLa cells are shown in Fig. 4, showing that sonotransfection can be superior to other nonviral methods. The epitheloid human carcimona of the cervix, HeLa cells, can be grown in a special culture container, OptiCellTM (BioCrystal, Ltd., Sanyo Biomedical, Osaka, Japan) (Fig. 5). OptiCell is a sterile, sealed cell culture environment between two optically clear gas-permeable growth surfaces in a standard microtiter plate-sized plastic frame with specially designed ports for access to the contents. OptiCell is the ideal environment for cell growth, microscopy, treatment, selection, separation, harvest, storage, and shipping. Optically clear gas-permeable growth surfaces allow diffusion of oxygen and carbon dioxide for optimal cell growth and permit microscopic examination at any stage of any cell process. OptiCell is compatible for use with standard,

3000 Luciferase expression/ mg protein

3.5. In Vivo-Simulated In Vitro Sonotransfection

2000

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0 Sonotransfection

Electrotransfection

Lipofection

Fig. 4. Optimized sonotransfection. This is a typical result in a series of sonotransfection experiments showing that sonotransfection can be superior to electroporation and liposome-mediated transfection. The bars in the graph show the luciferase expression and the micrographs on top of each bar represent the images of cells expressing GFP (cells appearing white).

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phase-contrast, confocal, and high-resolution time-lapse video microscopes and takes up a fraction of the space of conventional cell culture devices. Access ports allow aseptic access to the interior and its contents. 3.5.1. Sonotransfection Using an In Vivo-Simulated Setup

1. Plate 1 million cells in the OptiCell and incubate overnight or until confluence reaches 50–80%. 2. For sonication, prepare 10 mL of culture medium. 3. Add 10.0 μg/mL of plasmid DNA (e.g., pEGFP-N1) to the medium then agitate by pipetting several times. 4. Take out the medium contained in the OptiCell by introducing 10 mL of air into the OptiCell before withdrawing the medium. Try to avoid bubbling inside the OptiCell by positioning the OptiCell in such a way that the air is introduced from above and the medium withdrawn from below. 5. Add microbubbles at 10 μL/mL to the freshly prepared medium containing the plasmid DNA. Mix gently before aspirating the medium with the syringe and inject the medium into the OptiCell. Avoid splashing the medium inside the OptiCell by positioning the inlet below. When all medium is delivered inside the OptiCell, turn the OptiCell upside down without removing the needle then gently withdraw all air from inside the OptiCell. 6. Position the OptiCell for sonication on top of a 2-cm-thick oil-filled ultrasound transmitting pad (ESTEK, Tokyo, Japan) placed on top of the ultrasound transducer, with side B where cells are attached directed downward and facing the transducer (Fig. 5). 7. Place the ultrasound-absorbing pad (ESTEK) on top of the OptiCell after putting transmission gels in the path of the ultrasound field (see Note 6).

Ultrasound absorber Cells Culture medium OptiCellTM ferilism.com

Ultrasound transducer

Transmitting oil

Fig. 5. In vivo-simulated setup. This is a novel in vitro setup that simulates in vivo conditions. An OptiCellTM containing cancer cells is sandwiched between transmitting oil simulating soft tissue in the body and an ultrasound absorber on the other side.


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8. Sonicate the sample. Typical ultrasound conditions using SonoPore KTAC-4000 ultrasound device are: transmission frequency, 1.011 MHz; burst rate or PRF, 0.5; DF, 25%; voltage, 25 V; sweep frequency type 1 at 12% for 100 ms; rectangular pulse shape; and sonication time, 10–60 s. 9. After sonication, gently wash surfaces of the OptiCell with water to remove the gel, then wipe gently to dry. Try to avoid making scratch marks on the OptiCell because they may interfere when performing microscopy. 10. Incubate the cells for 24 h before assay (e.g., fluorescence microscopy when using a GFP gene). 11. From the sonicated areas, randomly select ten or more sites (×100 or ×200 magnification), take pictures of them, and save the images as computer files. 12. The rate of transfection is measured by dividing the total number of transfected cells (fluorescent cells) by the total number of visible cells. The cells may be counted manually. 3.6. In Vivo Sonotransfection

1. Anesthetize Wistar rats with 1.2 μL/g rat weight using Somnopentyl (Kyoritsu Seiyaku Corp., Tokyo, Japan).

3.6.1. Intravisceral Sonotransfection (17)

2. Remove the abdomen hair from the rat using a hair remover (Kanebo Ltd., Tokyo, Japan). 3. Empty the rat’s bladder by transurethral catheterization before introducing a 400-μL medium containing 0.5 million cells, 100 μg/mL pBKCMV-luc, and microbubbles (or an echo contrast agent, e.g., Levovist) into the bladder using a 1-mL syringe through the catheter. 4. Apply ultrasound (1 MHz, 0.78 W/cm2, and 30% duty cycle) transabdominally for 60 s using an ultrasound apparatus (Sonicmaster ES-2). 5. Withdraw the cell suspension from the bladder and cultivate the cell suspension at 37°C. 6. The assay can then be performed after a 24-h incubation.

3.6.2. Sonotransfection in Muscle (25)

1. Anesthetize mice by halothane inhalation (5% induction, 2% maintenance) in NO2/O2 (70%/30%) supplied over a cylindrical ventilation cap held over the head. 2. Inject 50 μL of gene preparation (e.g., pMhp53.MB) into shaved hind legs, intramuscular (i.m.) into the tibialis anterior muscle of C57B1/6 mice. 3. After the i.m. injections, sonicate the muscle (e.g., 2 min, duty cycle of 50%, at intensity 2 W/cm2). 4. Kill the mice 48 h after the i.m. injection and immediately resect and freeze the treated muscles.

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5. To evaluate expression in the muscle, 4-μm-thick slices are taken every 250 μm throughout the muscle. 6. Protein expression can then be assayed. For example, p53 protein expression can be evaluated by staining with p53 (N-19) (antibody against p53; Santa Cruz Biotechnology, Santa Cruz, CA, USA), followed by staining with a peroxidase-conjugated secondary antibody. 7. Peroxidase activity can be visualized with 3-amino-9ethyl-carbazole (AEC). 8. The positive-stained area on the slices can be assessed quantitatively using the Leica Qwin image analyzer. A novel sonication setup for tumors in vivo is shown in Fig. 6. 3.7. Future Directions

There are clear advantages of using ultrasound in transfection. First, unlike the viral method, immune reaction to the treatment is unlikely. Second, the ability of ultrasound to penetrate deeper into soft tissues makes it a noninvasive method when applied in vivo or clinically. Third, ultrasound can be localized, unlike the viral method and liposome-mediated transfection, which have generally systemic effects. However, the relatively low transfection rate compared with the viral method remains a big challenge for the researchers of sonotransfection. Another challenge is the lack of specificity in targeting cancer cells. Several studies are trying to address these issues. One is the use of engineered microbubbles (Fig. 7) that would carry the genes and seek cancer cells through a ligand specific to that particular cancer cell line (29). A highly localized effect and a very efficient transfection of therapeutic genes is expected if sonication is done after these gene-carrying bubbles have localized on the target cells.

Ultrasound transducer Coupling gel Silicon rubber Tumor Skin

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Fig. 6. In vivo sonication of tumor target. This setup can be used when sonicating tumors in vivo.


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Ligand Microbubbles

Microjets formed by sonication deliver genes into cancer cells Cancer

Microbubbles attach to cancer cells but not to normal cells

Successful sonotransfection after membrane repair

Fig. 7. Targeted sonotransfection using microbubbles. Microbubbles can be engineered to carry therapeutic genes and at the same time carry a ligand that specifically latches onto a particular cell (e.g., cancer cells belonging to a cancer cell line). Conceptually, these engineered microbubbles will be injected intravenously to a particular cancer patient, allowing time for the microbubble to localize on the cancer cells before sonication. Sonication will release and deliver the genes into the target cells with minimal effects on other cells in the body.

4. Notes 1. Bioeffects of ultrasound, including sonotransfection, are affected or influenced by many factors. Among them are: (1) the gas content of the sample and the type of gasses (26, 27) in it, (2) the temperature (15, 28) of the sample during sonication, (3) the ambient pressure within the sample and at the place of sonication, (4) the presence of reflective structures along the sonication path, and (5) the density of the sample, including the cell density of the sample (9). 2. Gas or gases dissolved in a medium determine the potential for cavitation formation, the type of cavitations that may occur during sonication, and the cavitational activity. If cavitation formation is desired, saturating the medium with air may be performed. Other gasses commonly used for an enhanced cavitation are argon (Ar) and diatomic gases such as O2 and N2. Triatomic gases such as N2O and CO2 usually inhibit formation of cavitations because of their high solubility in water. It is therefore expected that sonication of samples directly from incubation (growth medium likely to contain CO2) will give a different result than when sonication of sample is performed after having adding fresh medium (likely to contain dissolved air).

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Ambient pressure can also influence cavitation formation and activity. Regulating the ambient pressure during all experiments might be difficult, but it is good practice to try keeping all experiments in more or less the same pressure environment. 3. Instead of saturating the medium with gases, cavitational activities can also be attained by using engineered microbubbles, commercially called echo-contrast agents. Microbubbles (or echo-contrast agents) should be added to the sample just before sonication. Examples of microbubbles that are available in the market as contrast agents in sonography are: (1) Levovist® (SHU 508 A, Schering AG, Berlin, Germany), (2) Optison® (Molecular Biosystems, Inc., San Diego, CA), and Sonazoid® (Daiichi Sankyo, Tokyo, Japan). 4. The temperature of the sample could be a factor that will influence the outcome of your ultrasound experiments. It is known that temperature affects cells and tissue. Cell membranes are particularly vulnerable to temperature changes and temperature changes can potentially affect transfection. In ultrasound experiments that involve facilitation of cavitation, temperature can also influence the formation of cavitations and their activity. With therapeutic significance in mind, it is therefore advisable to keep the temperature of the sample similar to body temperature (e.g., 37°C) during experiments. In in vivo experiments, wherein higher intensities may be required, heating of the tissue may result. This can be avoided by using pulsed ultrasound and choosing a pulse rate and duty factor that will yield an optimal result without a significant temperature increase. 5. At higher densities, cavitation is less likely to occur. If cavitation exists, at higher densities it may be less likely to undergo inertial cavitation, that is, gain enough energy during oscillation then undergo implosion. To attain cavitation and facilitate inertial cavitation, it might be necessary to increase the ultrasound intensities if reducing the density is not possible, such as in in vivo experiments. 6. Reflection of ultrasound may result in overlapping of ultrasound waves, called standing wave formation. This will lead to a widened spectrum of ultrasound waves, from very low pressures to very high pressures. The ultrasound field is hard to define in this instance and results from experiments might be less predictable. Reflection of ultrasound happens when there are reflective surfaces (materials of different density than the site being exposed, e.g., bone next to a soft tissue target) close to the exposure site. To avoid this, one may need to use specialized ultrasoundabsorbing pads positioned at the far end along the exposure path. In some in vitro experiments using very low-intensity ultrasound, standing wave formation may be needed to attain the desired biological and sonochemical effects.


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References 1. ter Haar GR. (2001) High intensity focused ultrasound for the treatment of tumors. Echocardiography 18:317–22. 2. Feril LB, Jr., Kondo T. (2004) Biological effects of low intensity ultrasound: the mechanism involved, and its implications on therapy and on biosafety of ultrasound. Journal of Radiation Research 45:479–89. 3. Feril LB, Jr., Ogawa R, Tachibana K, Kondo T. (2006) Optimized ultrasound-mediated gene transfection in cancer cells. Cancer Science 97:1111–4. 4. Wu F, Wang ZB, Chen WZ, et al. (2004) Extracorporeal high intensity focused ultrasound ablation in the treatment of 1038 patients with solid carcinomas in China: an overview. Ultrasonics Sonochemistry 11: 149–54. 5. Uchida T, Baba S, Irie A, et al. (2005) Transrectal high-intensity focused ultrasound in the treatment of localized prostate cancer: a multicenter study. Hinyokika Kiyo – Acta Urologica Japonica 51:651–8. 6. Chaussy C, Thuroff S, Rebillard X, Gelet A. (2005) Technology insight: high-intensity focused ultrasound for urologic cancers. Nature Clinical Practice Urology 2:191–8. 7. Wu F, ter Haar G, Chen WR. (2007) Highintensity focused ultrasound ablation of breast cancer. Expert Review of Anticancer Therapy 7:823–31. 8. Chan AH, Fujimoto VY, Moore DE, Martin RW, Vaezy S. (2002) An image-guided high intensity focused ultrasound device for uterine fibroids treatment. Medical Physics 29 :2611–20. 9. Feril LB, Jr., Kondo T. (2005) Major factors involved in the inhibition of ultrasoundinduced free radical production and cell killing by pre-sonication incubation or by high cell density. Ultrasonics Sonochemistry 12:353–7. 10. Feril LB, Jr., Kondo T, Takaya K, Riesz P. (2004) Enhanced ultrasound-induced apoptosis and cell lysis by a hypotonic medium. International Journal of Radiation Biology 80:165–75. 11. Vessey CJ, Norbury CJ, Hickson ID. (1999) Genetic disorders associated with cancer predisposition and genomic instability. Progress in Nucleic Acid Research & Molecular Biology 63:189–221. 12. Uesato M, Gunji Y, Tomonaga T, et al. (2004) Synergistic antitumor effect of antiangiogenic factor genes on colon 26 produced by

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chemical and biological effects of ultrasound. Ultrasound in Medicine & Biology 12:151–5. 28. Kondo T, Kano E. (1987) Enhancement of hyperthermic cell killing by non-thermal effect of ultrasound. International Journal of Radiation Biology & Related Studies in Physics, Chemistry & Medicine 51:157–66. 29. Feril LB, Jr., Kondo T, Tabuchi Y, et al. (2007) Biomolecular effects of low-intensity ultrasound: apoptosis, sonotransfection, gene expression. Japanese Journal of Applied Physics 46:4435–40.

Chapter 11 Nonviral Jet-Injection Technology for Intratumoral In Vivo Gene Transfer of Naked DNA Wolfgang Walther, Iduna Fichtner, Peter M. Schlag, and Ulrike S. Stein Summary The main challenges for application of gene therapy to patients are poor selectivity in vector targeting, insufficient gene transfer, and great difficulties in systemic treatment in association with safety concerns for particular vector systems. For success in gene therapy, safe, applicable, and efficient transfer technologies are required. Because of the complex nature of targeted vector delivery to the tumor, our strategy for gene therapy is focused on the development of local nonviral gene transfer. This approach of local interference with tumor growth and progression could contribute to better control of the disease. Transfer of naked DNA is an important alternative to liposomal or viral systems. Different physical procedures are used for improved delivery of naked DNA into the target cells or tissues in vitro and in vivo. Among the various nonviral gene delivery technologies, jet-injection is gaining increased attractiveness, because this technique allows gene transfer into different tissues with deep penetration of naked DNA by circumventing the disadvantages associated with, e.g., viral vectors. The jet-injection technology is based on jets of high velocity for penetration of the skin and underlaying tissues, associated with efficient transfection of the affected area. The jet-injection technology has been successfully applied for in vivo gene transfer in different tumor models. More importantly, the efficacy and safety of jet-injection gene transfer have recently been investigated in a phase I clinical trial. Key words: gene transfer, intratumoral, jet-injection, naked DNA, nonviral.

1. Introduction The transfer of naked DNA for nonviral gene therapy is an alternative to liposomal or viral gene transfer technologies and it is gaining increasing importance. Naked gene transfer is efficiently used in DNA vaccination, genetic immunization, gene immunotherapy approaches, and other gene therapy applications (1–4). The growing impact of naked gene transfer on gene therapy is reflected by the Wolfgang Walther and Ulrike S. Stein (eds.), Methods in Molecular Biology, Gene Therapy of Cancer, vol. 542 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-561-9_11

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fact that, currently, approximately 18% of all clinical gene therapy trials are based on naked DNA gene transfer (5). For the delivery of naked DNA into cells or tissues, a great variety of procedures is used in vitro and in vivo. One well-known and early described procedure for naked DNA transfer was the needle and syringe injection. This technique went through several adaptations, such as the hydrodynamics pressure method that, in vivo, applies comparatively large volumes by needle injection within a few seconds for improved gene transfer (1, 2). To augment the application of naked DNA, various physical methods, including ballistic transfer, in vivo electroporation, sonoporation, or jet-injection, have evolved into applicable techniques. They were used for in vitro and in vivo gene transfer during the last decade and have shown their effectiveness in different experimental approaches (5–11). 1.1. Nonviral Gene Transfer Technologies

The majority of the nonviral gene transfer technologies are used for gene immune therapy or DNA vaccination studies. For intradermal or intramuscular applications, the use of naked DNA has proven to be an efficient vaccine against different viral infections or as an anticancer vaccine in numerous animal models (13–17). In the early 1990s, it was shown that simple needle and syringe injection of naked DNA is sufficient for in vivo gene transfer (1). Although needle injection is useful for naked DNA transfection in muscle tissue, this technique was only partially efficient for other tissue types, including tumors. This was one decisive reason why numerous studies deal with modifications of this procedure for the improvement of transfer efficiencies (18–21). This resulted in the development of, e.g., a hydrodynamics-based technology to deliver large volumes of more than 1 mL of naked DNA-containing solutions. These large volumes are either injected directly into the tissue by organ perfusion or applied by intravenous injection within short times of only a few seconds (22, 23). Although the efficiency of this procedure has been shown in several in vivo studies, at the current stage, this technique seems rather restricted to the perfusion of specific organs or particular portions of the desired organ, as shown for the liver or the kidney (23). The gene gun, or ballistic gene transfer, is based on the acceleration of DNA-coated gold or tungsten microparticles for gene transfer into different tissues. However, because of the technical characteristics, this transfer of plasmid DNA mediates only limited penetration and therefore does not reach deeper areas of the targeted tissues. This explains why most studies using particle bombardment for nonviral gene transfer are aiming at DNA vaccination or immunostimulatory approaches by targeting antigen-presenting cells (APC) in dermal and subdermal areas (24). Currently, many studies favor combinations of different nonviral transfer technologies to significantly improve in vivo gene transfer efficiencies. In this context, needle injection of naked DNA is

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combined with in vivo electroporation or focused ultrasound; or needleless technologies, such as jet-injection in combination with electroporation are used (9, 25, 26). 1.2. Jet-Injection for In Vivo Gene Transfer

The technique of jet-injection was initially developed and reported almost 50 years ago as a novel method for injecting insulin in a needleless fashion, and is also used for local application of various drugs as well as for immunization (27, 28). Among other physical delivery systems, jet-injection has developed to be an applicable gene transfer technology (29). This technique allows gene transfer into different tissue types with deeper penetration of the applied naked DNA compared with ballistic transfer systems (gene gun). Because of technological improvements, transfer efficiencies can be achieved by jet-injection that are comparable to in vivo electroporation or particle bombardment by application of significantly smaller amounts of naked DNA (12). The jet-injection technology is based on the use of fluid jets of high velocity possessing the required energy to penetrate skin and underlaying tissues, leading to efficient transfection of the jet-injected tissue areas (28, 30). The required acceleration of fluid jets is accomplished by either spring-forced systems or by applying pressurized air (gas powered) to the liquid, which is forced through a nozzle of very small diameter (3 weeks) after jet-injection before being killed for tumor removal and further analyses of gene expression. Tumors were excised and shock frozen in liquid nitrogen for subsequent preparation of cryosections for histochemical analysis, reporter gene assays of LacZ or GFP expression, or for the expression of further, therapeutically relevant genes, such as human tumor necrosis factor alpha (TNF-α) or the cytosine deaminase (CD) suicide gene. 2.2. Analysis of Transgene Expression in Jet-Injected Tumors

Initially, it was important to determine the transgene expression level and intratumoral distribution after jet-injection. In the majority of the in vivo studies, this was done with different reporter genes, such as LacZ or GFP (39, 40). Direct staining of cryosections was performed using X-gal to localize LacZ expression in the jet-injected tumor tissues. The tissues were cryosectioned and fixed in 2% formaldehyde to detect the LacZ expression in the jet-injected tumor. The slides were incubated with X-gal solution at 37°C for development of blue staining of the LacZ-transfected areas. The slides were evaluated in a light microscope (Fig. 2a). Figure 2a shows a representative appearance of LacZ expression detected by X-gal staining, which is scattered over a broad area of the jet-injected tumor tissue, with different staining intensities. As early as 24 h after jet-injection, LacZ gene expression was detectable in the jet-injected tumor tissue, however, the strongest gene expression started 48 h after jet-injection. Similar expression characteristics have been detected in tumors that were jet-injected with the GFP-expressing pEGFP-N1 plasmid (38, 39). For this, the tissues were also cryosectioned and fixed in formaldehyde for the detection of GFP expression in a fluorescence microscope (Fig. 2b). The jet-injection of GFP-expressing plasmid leads to bright fluorescence in the tumors, which again covers broad areas of the tissue (Fig. 2b). In addition to the use of reporter gene-expressing plasmids, the expression of jet-injected human TNF-α- or CD suicide gene-expressing plasmid DNA was analyzed at different times after jet-injection in xenotransplanted human colon carcinoma models. A high level of the cytokine or CD expression was detectable in these tumors as soon as 24 h after gene transfer at messenger RNA (mRNA) and at protein levels (40, 41). For TNF-α, we observed that the level of cytokine expression


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Fig. 2. Transgene expression in jet-injected tumor tissues (xenotransplanted human colon cancer). (a) LacZ expression (blue) detected in tumor cryosections by X-gal staining 48 h after intratumoral jet-injection of the naked pCMVβ plasmid DNA. (b) Fluorescence microscopy for detection of GFP expression in a tumor cryosection jet-injected with pEGFP-N1 plasmid DNA 48 h after the application. (c) CD suicide gene expression (brown staining) detected by CD-specific immunohistochemistry in cryosections of jet-injected tumor tissue 28 days after jet-injection. Arrows indicate the area of transgene expression (see Color Plates).

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remained at almost the same expression level during the observation time of 120 h after jet-injection. In the experiments with CD gene transfer, even 28 days after jet-injection, strong CD expression was detected at the mRNA level, and, more importantly, at the protein level (Fig. 2c) (41). Therefore, jetinjection gene transfer also ensures the efficient expression of therapeutic genes such as TNF-α or CD for several days or even weeks. This represents a duration of transgene expression that is sufficient to achieve effective therapeutic intervention for tumor treatment. 2.3. Analysis of the Stability of JetInjected Naked DNA

The stability of DNA is decisive for efficient gene transfer and foreign gene expression. It is known that preservation of plasmid topology, particularly of supercoiled plasmid DNA, has an impact on gene transfer efficiency. The jet-injection technology is based on the use of high pressures to eject the DNA-containing solution through the nozzle of the jet-injector, which has a narrow diameter of less than 0.4 mm. In fact, these conditions might create physical stress for the circular plasmid molecules, which might result in alteration of the DNA topology or damage of the DNA. Analyses of jet-injected plasmid DNA clearly demonstrated that marginal alterations of the plasmid DNA are apparent. We reproducibly found that the increase in jet-injection pressure leads to an increase of damaged DNA, reflected by the appearance of altered (in regard to DNA topology) or degraded DNA. However, the proportion of damaged plasmid DNA is low. Earlier quantitative analyses of jet-injected DNA using capillary gel electrophoresis (CGE) revealed that a maximum loss of 20% of the covalently closed circular (ccc) supercoiled form of the plasmid DNA occurs at the highest pressure setting of 3.0 bar (31, 39). The CGE analysis further demonstrated that reduced jetinjection pressure also reduces the loss of the ccc form of plasmid DNA to less than 7%. In contrast, our in vivo studies in different tumor models have shown that jet-injection pressures of 2.8–3.0 bar significantly improve the gene transfer efficiency. Therefore, for effective jet-injection gene transfer, conditions are used that represent the best compromise of jet pressure and preservation of intact DNA topology.

2.4. Jet-Injection for Antitumoral Gene Therapy: Preclinical Studies

The efficiency of jet-injection for in vivo gene transfer has been demonstrated in numerous tumor models. Therefore, it was of interest to analyze whether this technology is applicable for mediating therapeutic effects in in vivo tumor models. Therapeutic in vivo experiments using the jet-injection transfer of the cytosine deaminase (CD) suicide gene demonstrated antitumor effects (41). In these in vivo studies, human colon carcinoma (patient-derived tumor model 5734)-bearing NMRI–nu/nu male


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mice were jet-injected with the CD gene-harboring vector plasmid. By this single application of five jet-injections, a total dose of 50 μ g of CD-expressing plasmid DNA was installed intratumorally. The tumor volumes of control and treatment groups were measured during the 24-day treatment with the prodrug 5-fluorocytosine (5-FC). Antitumor effects were seen in the CD gene-transduced tumors starting from day 4 of 5-FC treatment compared with the non-jetinjected control group. The growth inhibitory effect lasted for the entire observation time and showed significant growth inhibition of the jet-injected colon carcinomas compared with the nontransduced and the 5-fluorouracil (5-FU)-treated animals. Detailed analyses of the jet-injected tumor revealed strong CD expression for the entire observation time. More importantly, jet-injection CD gene transfer caused necrosis in the 5-FC-treated tumors, supporting the therapeutic effectiveness of this nonviral approach. The jet-injection technology was also successfully used for the intratumoral application of a heat-inducible TNF-α expressing vector (42). In vivo experiments in human HCT116 colon carcinoma tumors showed efficient TNF-α expression that was heat inducible. This led to elevated transgene expression levels in the heated tumors. The jet-injection of the naked vector DNA led to therapeutic effects, reflected by a significant reduction in tumor growth of the jet-injected animals when heat-induced TNF-α expression was combined with chemotherapeutic treatment. In a different approach for tumor gene therapy, the jet-injection was used for the application of short hairpin RNA (shRNA)expressing plasmid vectors to reverse mdr1-mediated multidrug resistance (MDR) in tumors in mice. MDR renders tumors resistant to chemotherapeutic drugs mediated by ABC transporters such as mdr1 (43). The application of anti-mdr1 shRNA-expressing vectors into mdr1-overexpressing MaTu/Adriamycin tumors led to the sustained downregulation of the mdr1-mRNA and the mdr1 gene product, P-glycoprotein. Jet-injection of mdr1 shRNA reversed the resistant tumors to the sensitive phenotype, which then responded to subsequent chemotherapy with reductions in tumor size and growth. All of these therapeutic in vivo studies demonstrated the effectiveness and applicability of jet-injection based gene transfer as a therapeutic option for cancer treatment.

3. Clinical Application of Jet-Injection for Intratumoral Gene Transfer

The preclinical studies demonstrated the feasibility of jet-injection gene transfer. Based on this, a phase I clinical trial was performed to evaluate the safety and efficiency of intratumoral jet-injection

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LacZ-reporter gene transfer into skin metastases of melanoma patients (46). The patients were treated with jet-injection into a single cutaneous lesion (Fig. 3a). In the study, clinical safety monitoring was performed; plasmid DNA distribution and LacZ mRNA expression was analyzed by real-time polymerase chain reaction (PCR) and reverse transcriptase polymerase chain reaction (RT-PCR), complemented by Western blotting, immunohistochemistry, and X-gal staining of expression of functional LacZ protein (Fig. 3b). Systemic plasmid clearance was monitored by real-time PCR of blood samples at different time points before and after jet-injection. The study revealed that jet-injection was safely

Fig. 3. Clinical local application of jet-injection in a melanoma patient. (a) The skin lesion of the patient is jet-injected with the naked LacZ-expressing pCMVβ plasmid DNA. The inset shows the area of the tumor shortly after the jet-injection. (b) Detection of LacZ expression in the jet-injected tissue by X-gal staining of the cryosectioned tumor. The arrows indicate the LacZ-expressing tumor areas (blue) (see Color Plates).


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performed with no side effects for the patients. Plasmid DNA, the LacZ mRNA, and protein expression were detected in all treated lesions. Interestingly, similar patterns of transgene expression in the tumor were seen in the preclinical studies. Systemic plasmid distribution was detected shortly after jet-injection, peaking 30 min after application, followed by rapid DNA clearance after several hours. This clinical gene transfer trial demonstrated that jet-injection of plasmid DNA leads to the efficient expression of the functional transgene. No side effects were experienced, indicating safety for this nonviral transfer technology. Short-term and low-level systemic leakiness of the plasmid DNA was accompanied by rapid clearance. The data from this clinical trial support the feasibility of jet-injection gene transfer for gene therapy of cancer, particularly if local control of the disease is anticipated by this approach.

4. Conclusions This chapter describes the use of needle-free, low-volume jetinjection technology for gene transfer into tumors. Jet-injection has been extensively tested for its feasibility for in vivo transfer of naked DNA. It was demonstrated that jet-injection is successfully used for nonviral in vivo gene transfer and that it represents an attractive alternative to other established physical transfer technologies (29, 39, 44, 45). To obtain an applicable jet-injection-based technology, recent development has led to the construction of the multiuse SwissInjector prototype. This injector requires only small amounts of naked plasmid DNA associated with a significant reduction of ejected volumes and improved accuracy and reliability of DNA application. The Swiss-Injector system used in the in vivo studies is capable of ejection of low volume jets for single or repeated naked DNA application into the targeted tissue. This is advantageous for in vivo and for clinical application to transduce larger tissue areas. Regarding the safety of the jet-injection technology, we and others observed no serious side effects in preclinical jetinjection studies or in clinical application (30). Our qualitative and quantitative analyses revealed no significant loss of intact plasmid DNA with respect to the potential physical DNA damage by jet-associated shearing forces. This is important for the preservation of the functional topology of the DNA and for transgene expression (31, 45). The efficient expression of the LacZ and GFP reporter genes and also of the therapeutic human TNF-α and CD suicide gene has been demonstrated. More importantly, single applications of

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small amounts of naked vector DNA were sufficient to generate efficient transgene expression and to achieve antitumoral therapeutic effects in different in vivo tumor models. The pattern of transgene expression in the preclinical and clinical studies indicated that broad tumor areas are affected, providing a sufficiently high level and duration of transgene expression to exert a therapeutic effect. Taken together, the nonviral jet-injection gene transfer of naked DNA has the potential for clinical application in therapeutic settings, particularly if local gene therapy approaches are anticipated in cancer treatment.

Acknowledgments This work was kindly supported by the EMS Medical Systems SA, Nyon, Switzerland, and grants from the H.W. & J. Hector Foundation, Mannheim, Germany, and the Deutsche Forschungsgemeinschaft, Bonn, Germany. Support from A. Menne and T. Nellessen in the establishment of the jet-injection technology is appreciated. Help in the performance of the animal studies by M. Lemm and the excellent technical assistance of D. Kobelt, J. Aumann, and L. Malcherek are gratefully acknowledged. References 1. Wolff J. A., Malone R. W., Williams P., Chong G., Acsadi A., Jani A., Felgner P. L. (1990) Direct gene transfer into mouse muscle in vivo. Science 247: 1465–1458. 2. Herweijer H., Wolff J. A. (2003) Progress and prospects: naked DNA transfer and therapy. Gene Ther 10: 453–458. 3. Niidome T, Huang L. (2002) Gene therapy progress and prospects: nonviral vectors. Gene Ther 9: 1647–1652. 4. Romano G. (2007) Current development of nonviral-mediated gene transfer. Drug News Perspect 20: 227–231. 5. Edelstein M. L., Abedi M. R., Wixon J. (2007) Gene therapy clinical trials worldwide to 2007 – an update. J. Gene Med 9: 833–842. 6. Klinman D. M., Conover J., Leiden J. M., Rosenberg S. A., Sechler J. M. G. (1999) Safe and effective regulation of hematocrit by gene gun administration of an erythropoietinencoding DNA plasmid. Hum Gene Ther 10: 659–665. 7. Sikes M. L., O’Malley B. W., Finegold M. J., Ledley F. D. (1994). In vivo gene transfer into rabbit thyroid follicular cells by direct DNA injection. Hum Gene Ther 6: 837–844.

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transfection with interleukin 12 gene results in regression of established primary and metastatic murine tumors. Proc Natl Acad Sci USA 93: 6291–6296. Liu M. A., Ulmer J. B. (2000) Gene based vaccines. Mol Ther 1: 497–500. Macklin M. D., McCabe D., McGregor M. W., Neumann V., Meyer T., Callan R., Hinshaw V. S. Swain W. F. (1998) Immunization of pigs with a particle-mediated DNA vaccine to influenza A virus protects against challenge with homologous virus. J Virol 72: 1491–1496. Turner J. G., Tan J., Crucian B. E., Sullivan D. M., Ballester O. F., Dalton W. S., Yang N. S., Burkholder J. K., Yu H. (1998) Broadened clinical utility of gene gun-mediated granulocyte-macrophage colony-stimulating factor cDNA-based tumor cell vaccines as demonstrated with a mouse myeloma model. Hum Gene Ther 9: 1121–1130. Davis H. L., Demeneix B. A., Quantin B., Coulombe J., Whalen R. G. (1993) Plasmid DNA is superior to viral vectors for direct gene transfer into adult mouse skeletal muscle. Hum Gene Ther 4: 733–740. Heinzerling L., Burg G., Dummer R., Maier T., Oberholzer P. A., Schultz J., Elzaouk L., Pavlovic J., Moelling K. (2005) Intratumoral injection of DNA encoding human interleukin 12 into patients with metastatic melanoma: clinical efficacy. Hum Gene Ther 16: 35–48. Zhang G., Vargo D., Budker V., Armstromg N., Knechtle S., Wolff J. A. (1997) Expression of naked plasmid DNA injected into afferent and efferent vessels of rodent and dog livers. Hum Gene Ther 8: 1763–1772. Budker V., Zhang G., Danko I., Williams P., Wolff J. A. (1997) The efficient expression of intravascularly delivered DNA in rat muscle. Gene Ther 5: 272–276. Liu F., Song Y. K., Liu D. (1999) Hydrodynamics-based transfection in animals by systemic administration of plasmid DNA. Gene Ther 6: 1258–1266. Zhang G., Song Y. K., Liu D. (2000) Longterm expression of human alpha1-antitrypsin gene in mouse liver achieved by intravenous administration of plasmid DNA using hydrodynamics-based procedure. Gene Ther 7: 1344–1349. Pertmer T. M., Eisenbraun M. D., McCabe D., Prayaga S. K., Fuller D. H., Haynes J. R. (1995) Gene gun-based nucleic acid immunization: elicitation of humoral and cytotoxic T-lymphocyte responses following epidermal delivery of nanogram quantities of DNA. Vaccine 13: 1427–1430.

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25. Wells D. J. (2004) Gene therapy progress and prospects: electroporation and other physical methods. Gene Ther 11: 1363–1369. 26. McCreery T. P., Sweitzer R. H., Unger E. C., Sullivan S. (2004) DNA delivery to cells in vivo by ultrasound. Methods Mol Biol 245: 293–298. 27. Weller C., Linder M. (1966) Jet injection of insulin vs the syringe-and-needle method. JAMA 195: 844–847. 28. Baxter J., Mitragotri S. (2006) Needle-free liquid jet injections: mechanisms and applications. Expert Rev Med Devices 3: 565–574. 29. Mitragotri S. (2006) Current status and future prospects of needle-free liquid jet injectors. Nat Rev Drug Discov 5: 543–548. 30. Furth P. A., Kerr D., Wall R. (1995) Gene transfer by jet injection into differentiated tissues of living animals and in organ culture. Mol Biotechnol 4: 121–127. 31. Walther W., Stein U., Fichtner I., Voss K., Schmidt T., Schleef M., Nellessen T., Schlag P. M. (2002) Intratumoral low volume jetinjection for efficient nonviral gene transfer. Mol Biotechnol 21: 105–115. 32. Walther W., Stein U., Siegel R., Fichtner I., Schlag P. M. (2005) Use of the nuclease inhibitor aurintricarboxylic acid (ATA) for improved non-viral intratumoral in vivo gene transfer by jet-injection. J Gene Med 7: 477–485. 33. Heansler J., Verdelet C., Sanchez V., Girerd-Chambaz Y., Bonnin A., Trannoy E., Krishnan S., Meulien P. (1999) Intradermal DNA immunization by using jet-injectors in mice and monkeys. Vaccine 17: 628–638. 34. Choi A. H., Smiley K., Basu M., McNeal M. M., Shao M. Bean J. A., Clements J. D., Stout R. R., Ward R. L. (2007) Protection of mice against rotavirus challenge following intradermal DNA immunization by Biojector needlefree injection. Vaccine 25: 3215–3218. 35. Seigne J., Turner J., Diaz J., Hackney J., PowSang J., Helal M., Lockhart J., Yu H. (1999) Feasibility study of gene gun mediated immunotherapy for renal cell carcinoma. J Urol 162: 1259–1263. 36. Yamashita Y., Shimada M., Hasegawa H., Minagawa R., Rikimaru T., Hamatsu T., Tanaka S., Shirabe K., Miyazaki J., Sugimachi K. (2001) Electroporation-mediated interleukin-12 gene therapy for hepatocellular carcinoma in the mice model. Cancer Res 61: 1005–1012. 37. Hui K. M., Chia T. F. (1997) Eradication of tumor growth via biolistic transformation with allogeneic MHC genes. Gene Ther 4: 762–767.

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38. Walther W., Stein U., Fichtner I., Schlag P. M. (2001) In vivo gene transfer of naked DNA into xenotransplanted colon carcinoma by jetinjection. Langenbeck’s Archives Surg 2001, 30: 69–72. 39. Walther W., Stein U., Fichtner I., Malcherek L., Lemm M., Schlag P. M. (2001) Nonviral in vivo gene delivery into tumors using a novel low volume jet-injection technology. Gene Ther 8: 173–180. 40. Walther W., Stein U., Fichtner I., Schlag P. M. (2004) Low-volume jet-injection for efficient in vivo gene transfer. Mol Biotechnol 28: 121–128. 41. Walther W., Stein U., Fichtner I., Aumann J. , Arlt F., Schlag P. M. (2005) Nonviral jet-injection gene transfer for efficient in vivo cytosine deaminase suicide gene therapy of colon carcinoma. Mol Ther 12, 1176–1184 . 42. Walther W., Arlt F., Stein U., Fichtner I., Schlag P. M. (2007) Heat-inducible in vivo gene therapy of colon carcinoma by human

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mdr1 promoter regulated TNF-α expression. Mol Cancer Ther 6: 235–243. Stein U., Stege A., Walther W., Lage H. (2008) Complete in vivo reversal of the multidrug resistance (MDR) phenotype in a breast cancer model by jet-injection of anti-MDR1 short hairpin RNA-encoding plasmid DNA. Mol Ther 16: 178–186. Ren S., Li M., Smith J. M., DeTolla L. J., Furth P. A. (2002) Low-volume jet injection for intradermal immunization in rabbits. BMC Biotechnol 23: 10. Cartier R., Ren S. V., Walther W., Stein U., Lewis A., Schlag P. M., Li M., Furth P. A. (2000) In vivo gene transfer by low-volume jet injection. Anal Biochem 282: 262–265. Walther W., Siegel R., Kobelt D., Knösel T., Dietel M., Bembenek A., Aumann J., Schleef M., Baier R., Stein U., Schlag P.M. (2008) Novel jet-injection technology for nonviral intratumoral gene transfer in patients with melanoma and breast cancer. Clin Cancer Res 14: 7545–7553.

Subsection D Experimental Studies in Cancer Gene Therapy

Chapter 12 Methods for Constructing and Evaluating Antitumor DNA Vaccines Brian M. Olson and Douglas G. McNeel Summary An antitumor DNA vaccine is a bacterial DNA plasmid that encodes the complementary DNA (cDNA) of a tumor antigen. When injected into recipients, antitumor DNA vaccines have been shown to elicit both humoral and cellular immunity against the encoded tumor antigen. These vaccines represent a relatively new immunotherapeutic technique being investigated as a means to deliver a target antigen and elicit or augment antitumor antigen-specific immune responses. One of the primary advantages of DNA vaccines as opposed to some other methods of antigen delivery is that they can be easily constructed, purified, and delivered to recipients. In this review we describe this process, detailing the procedures used to construct, purify, deliver, and evaluate the efficacy of DNA vaccines. We begin by describing the process of molecularly constructing the vaccine, from selecting a bacterial plasmid to form the backbone of the vaccine, cloning the antigen cDNA into this plasmid, and confirming the sequence and orientation of the completed vaccine. This is then followed by a series of experiments that can be used to ensure that the antigen encoded by the vaccine is transcribed and translated after being taken up by eukaryotic cells. We then describe large-scale purification procedures that can be used to obtain sufficient quantities of plasmid DNA to conduct in vivo immunization experiments. Finally, we provide an immunization protocol that can be used to evaluate the immunological efficacy of the constructed DNA vaccine. By following these protocols, it is possible to construct, purify, deliver, and evaluate the efficacy of antitumor DNA vaccines. Key words: DNA vaccine, immunization, immunotherapy, plasmid purification, tumor antigen.

1. Introduction The development of DNA vaccines designed to elicit or augment antitumor immune responses represents a new, potentially powerful weapon in the arsenal of tumor immunologists. A DNA vaccine is comprised of a bacterial DNA plasmid backbone into Wolfgang Walther and Ulrike S. Stein (eds.), Methods in Molecular Biology, Gene Therapy of Cancer, vol. 542 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-561-9_12

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which is inserted the nucleic acid coding sequence of the targeted tumor antigen. When this vaccine is injected into a recipient, it can be directly taken up by antigen-presenting cells (APCs), causing the encoded antigen to be transcribed and translated by the host’s cellular machinery. This results in antigen-derived peptides being processed through the endogenous antigen presentation pathways, ultimately being presented on the surface of APCs bound to major histocompatibility complex (MHC) class I and class II complexes (1, 2). These two methods of antigen presentation can thus lead to the activation of not only CD4+ T cells (by either direct presentation or cross-priming), but can also lead to the activation, maturation, and proliferation of cytotoxic antigenspecific CD8+ T cells. The ability of DNA vaccines to potentially augment both CD4+ and CD8+ immune responses has led them to be evaluated in various tumor models, where they have demonstrated the ability to generate protective antitumor immunity in vivo (3–5). Some of these preclinical experiments have already been translated into human studies, with vaccines against various tumor antigens currently being tested in early stage clinical trials for toxicity as well as immunological and antitumor efficacy (6–10). There are several advantages to DNA vaccines as opposed to other methods of antigen delivery. One of the most significant advantages of DNA vaccines is that after being taken up by antigen-presenting cells, the target antigen is expressed by the host’s own cellular machinery. This causes the antigen to be naturally folded and processed by the endogenous antigen-presenting pathway, and most importantly results in the development of both humoral and cellular immunity. And, as opposed to viral vaccines, which can also induce the natural expression of the antigen by host APCs, DNA vaccines have not been shown to become incorporated into the host genome, preventing insertional mutagenesis as well as the genomic insertion and expression of the target antigen, which in some cases may act as an oncogene (11). Furthermore, cells that take up the DNA vaccine can continue to drive expression of the encoded antigen, potentially leading to an extended immune response (12). DNA vaccines can also be used to target multiple antigens, as well as to incorporate costimulatory molecules and other adjuvants into the vaccine construct to enhance its immunological efficacy—in fact, the bacterial DNA plasmid itself might contain hypomethylated CpG repeats, a TLR9 agonist, that serve as an intrinsic adjuvant and contribute to the immunological efficacy of the vaccine (13–17). Despite these advantages, DNA vaccines share a disadvantage with all other antigen-specific vaccines in that they are targeting a specific antigen, whereas nonspecific tumor vaccines, such as whole cell vaccines, can potentially elicit responses to any and all antigens that may be displayed by these vaccines. This

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highlights the importance of preclinical in vitro studies evaluating whether cytotoxic T cells specific for the antigen targeted by the DNA vaccine exist within the immunorepertoire of patients, and if these cells can recognize and lyse tumor cells. However, this antigen specificity can also be advantageous, because nonspecific vaccines may also elicit responses against irrelevant or deleterious antigens that may not contribute to antitumor immunity. Another potential disadvantage to DNA vaccines is that some methods of delivering this naked DNA to live recipients simply injects the DNA outside of the host’s APCs, leading to very low transfection efficiencies and potentially decreasing the efficacy of the immune response that is generated. However, by using these simple naked DNA plasmids as a vaccine, it is possible to easily construct, modify, and deliver these DNA vaccines to recipients, a process that we describe in detail in this review. To create a DNA vaccine, cDNA of the tumor antigen to be targeted is amplified and inserted into a bacterial DNA plasmid backbone under the expression of a eukaryotic transcription promoter. The sequence and orientation of this vaccine is then confirmed using a variety of methods to ensure that the cloning process produced the desired vaccine construct. After constructing this vaccine, it is essential to verify that the antigen is expressed when it is taken up by eukaryotic APCs, because this is required to activate an antigen-specific immune response. To confirm the expression of the antigen, the vaccine is used to transfect eukaryotic cell lines in vitro, and RNA and protein samples are collected to evaluate the antigen transcription and translation, respectively. After the construction and confirmation of the DNA vaccine, it is then necessary to purify large quantities of the vaccine, which are required for immunization studies. Once the vaccine has been constructed and purified, immunization studies can be conducted, evaluating the immunological and antitumor efficacy of the vaccine in vivo. This review will cover this process from beginning to end; from the creation of a DNA vaccine to immunization procedures that can be used in preclinical rodent studies, and will outline the methods required to create, purify, deliver, and evaluate the efficacy of antitumor DNA vaccines.

2. Materials 2.1. Constructing DNA Vaccines 2.1.1. Cloning Target Antigen cDNA into an Immunization Vector

1. Bacterial DNA plasmid, such as pNGVL-1 (National Gene Vector Laboratory, Indianapolis, IN) or pCDNA (Invitrogen, Carlsbad, CA) (see Note 1). 2. Target antigen gene-specific 3′ and 5′ flanking DNA primers (Sigma-Aldrich, St. Louis, MO) (see Note 2).

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3. Antigen template DNA (see Note 3). 4. QIAGEN Taq polymerase chain reaction (PCR) kit: store at −20°C (Qiagen, Valencia, CA). 5. dNTP mix: store at −20°C (Promega, Madison, WI). 6. Electrophoresis-grade agarose (Invitrogen). 7. Tris Borate EDTA (10× solution, National Diagnostics, Atlanta, GA). 8. Ethidium bromide (EtBr): this is a potent carcinogen—use care when handling. (Thermo Fisher, Waltham, MA). 9. 6X Blue/Orange DNA loading dye: store at −20°C (Promega). 10. Bio-Rad Sub-Cell hand-case gel system (Bio-Rad, Hercules, CA). 11. 1-kb DNA ladder: store at −20°C (Promega). 12. QIAquick Gel Extraction kit (Qiagen). 13. Plasmid- and antigen-specific restriction enzymes, and appropriate buffers (New England Biolabs, Ipswitch, MA). 14. T4 DNA Ligation kit: store at −20°C (New England Biolabs). 15. QIAGEN EZ Competent Cells: store at −80°C (Qiagen). 16. Luria broth (LB) bacterial growth media: 1% (w/v) Tryptone, 1% (w/v) NaCl, 0.5% (w/v) yeast extract; sterilized by autoclave (Sigma-Aldrich). 17. LB-Agar plates: 1.5% (w/v) agar dissolved into LB media (autoclave to mix and sterilize), and subsequently mixed with the appropriate antibiotic. 18. Antibiotic for plasmid selection: the antibiotic depends on the bacterial plasmid selected in step 2.1.1.1. 2.1.2. Confirmation of the Vaccine Construct

1. Materials for an agarose gel (steps 2.1.1.6–2.1.1.11).

Production and Purification of Transduced Bacterial Plasmid DNA

1. QIAGEN Taq polymerase kit: store at −20°C (Qiagen).

Gene- and PlasmidSpecific PCR

2. QIAprep Spin Miniprep Kit (Qiagen).

2. Internal gene-specific 3′ and 5′ primers to be used for determining correct insertion as well as antigen transcription (Sigma-Aldrich). 3. Plasmid-specific primers for sites upstream and downstream of inserted antigen cDNA (Sigma-Aldrich). 4. Materials for an agarose gel (steps 2.1.1.6–2.1.1.11).

Restriction Digestion

1. Plasmid-specific and antigen-specific restriction enzymes and buffers that can be used to determine the presence and orientation of construct (see Note 4). 2. Materials for an agarose gel (steps 2.1.1.6–2.1.1.11).

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1. Big Dye v3.1 kit: store at −20°C (Applied Biosystems, Foster Center, CA). 2. Gene- and plasmid-specific primers. 3. Sequencing alignment software.

2.2. Confirming Expression of the Target Antigen 2.2.1. Transfection of Cell Lines

1. Chinese hamster ovary (CHO) cells. 2. RPMI 1640 media (Sigma-Aldrich) supplemented with 10% fetal bovine serum (FBS, HyClone, Ogden, UT) and 1% penicillin-streptomycin (RPMI/FBS): store at 4°C. 3. 60-mm flasks (Thermo Fisher). 4. QIAGEN Effectene transfection kit: store at 4°C (Qiagen). 5. 1X PBS: diluted from 10X PBS stock in sterile H2O.

2.2.2. Confirming Transcription of the Target Antigen

1. QIAGEN RNeasy Plus Mini RNA purification kit (Qiagen). 2. β -Mercaptoethanol. 3. QIAGEN One-Step Reverse-Transcription (RT)-PCR kit: store at −20°C (Qiagen). 4. Gene-specific primers. 5. Materials for an agarose gel (steps 2.1.1.6–2.1.1.11).

2.2.3. Confirming Translation of the Target Antigen

1. Protein purification buffer: 50 mM Tris-HCl, pH 7.5, 150 mM NaCl, 1% Triton X-100.

Protein Preparation

2. Laemmli’s loading buffer, 5X solution: 65 mM Tris-HCl, pH 6.8, 20% glycerol, 2% SDS, 5% β -mercaptoethanol, 0.1% bromophenol blue. Store at 4°C. 3. Prestained molecular weight Kaleidoscope markers (Bio-Rad). Store at −20°C. 4. Ultraviolet cuvettes (International Crystal Laboratories, Garfield, NJ). 5. Bradford reagent: store at 4°C (Sigma-Aldrich). 6. Bovine serum albumin (BSA, Roche Applied Science, Indianapolis, IN).

SDS-PAGE

1. Bio-Rad Mini-Protean 3 Cell (Bio-Rad). 2. Resolving gel buffer: 1.5 M Tris-HCl, pH 8.8, 0.1% sodium dodecyl sulfate (SDS). 3. Stacking gel buffer: 0.5 M Tris-HCl, pH 6.8, 0.1% SDS. 4. Thirty percent acrylamide/bis solution (37.5:1 with 2.6% C). This is a potent neurotoxin when unpolymerized. Store at 4°C (Bio-Rad). 5. Ammonium persulfate (APS): prepare 10% solution in H2O, and store at 4°C (Bio-Rad).

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6. N, N, N, N′-Tetramethyl-ethylenediamine (TEMED): store at 4°C (Bio-Rad). 7. Water-saturated isobutanol: mix equal volumes of isobutanol and water, and allow it to separate into layers (top layer containing isobutanol). 8. SDS-polyacrylamide gel electrophoresis (PAGE) running buffer (5X): 125 mM Tris-HCl, 960 mM glycine, 0.5% (w/v) SDS. Western Blotting for Antigen Protein Expression

1. Bio-Rad Mini Trans-Blot Electrophoretic Transfer Cell (Bio-Rad). 2. Western transfer buffer: 25 mM Tris-HCl, 190 mM glycine, 20% (v/v) methanol, 0.05% (w/v) SDS. 3. Nitrocellulose membrane (Millipore, Bedford, MA). 4. 3MM Chromatography paper (Whatman, Maidstone, UK). 5. Tris-buffered saline with Tween (TBST, 10X): 1.4 M NaCl, 3 mM KCl, 250 mM Tris-HCl, pH 7.4, 1% Tween-20. 6. Ponceau-S stain (Sigma-Aldrich). 7. Blocking buffer: TBST supplemented with 5% (w/v) bovine serum albumin (BSA) (Sigma-Aldrich). 8. Antibody buffer: TBST supplemented with 3% (w/v) BSA. 9. Antigen-specific primary antibody. 10. Secondary alkaline phosphatase-labeled antibody (BectonDickinson Biosciences, San Jose, CA) (see Note 5). 11. Color development solution: 7.5 mM Tris-HCl, 100 mM Trizma base, 100 mM NaCl, 10 mM MgCl2. Filter-sterilize and store at room temperature. 12. 5-bromo-4-chloro-3-indolyl-phosphate (BCIP), resuspended to a concentration of 50 mg/mL in 100% dimethylformamide (DMF). Store at 4°C (Roche Applied Science). 13. Nitroblue tetrazolium salt (NBT), resuspended to a concentration of 50 mg/mL in 70% DMF. Store at 4°C (Roche Applied Science).

2.3. Large-Scale Production and Purification of DNA Vaccine

1. LB media (as above).

2.3.1. GigaPrep Purifications

4. Gene- and plasmid-specific PCR materials (as in step 2.1.2.1).

2. Endo-free GigaPrep Kit (Qiagen). 3. UV cuvettes (International Crystal Laboratories). 5. Restriction enzymes (as in step 2.1.2.2). 6. DNA sequencing materials (as in step 2.1.2.3).


2.4. Evaluating the Immunological Efficacy of DNA Vaccines 2.4.1. Intradermal Immunization

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1. 10X Phosphate buffered saline (PBS, Sigma-Aldrich). 2. Isoflurane (Halocarbon, River Edge, NJ). 3. Isoflurane vaporizer. 4. 1-cc insulin syringes (Thermo Fisher).

2.4.2. Evaluation of Antigen-Specific Immune Response

1. 21-Gauge 1.5-inch needles (Becton-Dickinson Biosciences).

Rodent Euthanasia and Sera/Spleen Harvest

4. 15-mL Conical tube (Becton-Dickinson Biosciences).

2. 1- and 5-mL Syringes (Becton-Dickinson Biosciences). 3. 1.5-mL Microcentrifuge tubes (Thermo Fisher). 5. 50-mL Conical tube (Becton-Dickinson Biosciences). 6. Hanks’ Balanced Salt Solution (HBSS, Invitrogen). 7. Microsieve set (Thermo Fisher). 8. Pasteur pipets (Sigma-Aldrich). 9. Histopaque 1083: store at 4°C (Sigma-Aldrich). 10. RPMI/FBS (Sigma-Aldrich, HyClone): store at 4°C. 11. Freezing medium: 90% FBS (Hyclone) and 10% dimethyl sulfoxide (DMSO, Sigma-Aldrich). Do not mix until use, and cool FBS and DMSO to 4°C before use. 12. 2-mL Cryovials (Corning, Corning, NY). 13. Mr. Frosty freezing container (Sigma-Aldrich).

Antibody Enzyme-Linked Immunosorbent Assay

1. Immulon 4 HBX 96-well plates (Thermo Fisher). 2. Carbonate buffer: 4 mM Na2CO3 and 9 mM NaHCO3 mixed in distilled water, pH 9.6. Store at 4°C. 3. Purified protein of interest. 4. Nonspecific protein (such as duck ovalbumin; Thermo Fisher). 5. PBS supplemented with 1% BSA (Sigma-Aldrich). 6. PBS supplemented with 0.1% Tween-20. 7. Horseradish peroxidase (HRP)-labeled antibody. 8. Tetra-methyl benzidine (TMB) microwell peroxidase substrate system (KPL Inc, Gaithersburg, MA). 9. 1 N Hydrochloric acid (HCl, Thermo Fisher).

Antigen-Specific Interferon-GammaSecreting Responses

1. Mouse interferon-gamma (IFNγ) ELISPOT Development Module (R&D Systems, Minneapolis, MN): resuspend capture antibody in PBS and detection antibody in PBS/1% BSA, store at 4°C. 2. 1X PBS and 1X PBS + 0.05% Tween-20. 3. Multiscreen HA 0.45-µm nitrocellulose 96-well plates (Thermo Fisher).

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4. Purified protein of interest (various sources; e.g. Invitrogen, Sigma-Aldrich, etc.). 5. Nonspecific protein (such as duck ovalbumin; Thermo Fisher). 6. Concanavalin A (ConA, Calbiochem, San Diego, CA). 7. Tumor cell lines: one line expressing the antigen of interest, and another line that does not express this antigen (ATCC, Manassas, VA). 8. AP-labeled streptavidin (GE Healthcare). 9. Color development solution. 10. BCIP, as in step 2.2.3.3. 11. NBT, as in step 2.2.3.3. Antigen-Specific Cytotoxic Immune Responses

1. Mouse granzyme B ELISPOT Development Module (R&D Systems): resuspend capture antibody in PBS and detection antibody in PBS/1% BSA, store at 4°C. 2. 1X PBS and 1X PBS + 0.05% Tween-20 (Sigma-Aldrich). 3. Multiscreen HA 0.45-µm nitrocellulose 96-well plates (Thermo Fisher). 4. Purified protein of interest (various sources; e.g., Invitrogen, Sigma-Aldrich). 5. Nonspecific protein (such as duck ovalbumin; Thermo Fisher). 6. Concanavalin A (Calbiochem). 7. Tumor cell lines: one line expressing the antigen of interest, and another line that does not express this antigen (ATCC). 8. AP-labeled streptavidin (GE Healthcare). 9. Color development solution (as above). 10. BCIP, as in step 2.2.3.3. 11. NBT, as in step 2.2.3.3.

3. Methods 3.1. Constructing DNA Vaccine

This section describes the process of constructing a DNA vaccine targeting a tumor protein antigen. These vaccines are created by first amplifying the cDNA of the desired antigen using primers that incorporate restriction sites just upstream and downstream of the coding region of the antigen. These restriction sites are specific for sites located within the DNA plasmid expression


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vector, which serves as the backbone for the DNA vaccine. The amplified antigen is digested along with the plasmid, and these digested fragments can be ligated together to form the complete DNA vaccine. The finished vaccine is then used to transduce bacterial cells, which produce plasmid DNA (pDNA) that can be analyzed to determine the success of the cloning process using PCR, restriction enzyme digestions, and DNA sequencing to ensure the proper sequence and orientation of the inserted gene. 3.1.1. Cloning Target Antigen cDNA into the Immunization Vector

1. After identifying the antigen that is to be encoded by the DNA vaccine, gene-specific 3′ and 5′ flanking DNA primers are designed, incorporating restriction enzyme sites just upstream and downstream of the 5′ and 3′ ends of the flanking primers appropriate for cloning into the vector of choice. 2. Amplify the cDNA of the antigen of interest using flanking primers and a DNA template. These instructions assume the use of the Qiagen Taq polymerase kit. In this protocol, 10 µL of the 10X PCR buffer is mixed with 2 µL of each 10 µM dNTP stock, 0.1 µM of each flanking primer, 0.5 µL Taq DNA polymerase, 15,000 × g for 30 min at 4°C. Wash once with 10 mL of 70% endotoxinfree ethanol, and centrifuge again at >15,000 × g for 15 min at 4°C. 13. Remove the ethanol wash, and allow the DNA pellet to dry for approximately 15 min. Resuspend the DNA in 1 mL endotoxin-free TE, saving a 20-µL sample for analysis of purification efficiency. 3.3.2. Analysis of Purified DNA

1. Analyze the concentration of the purified DNA and protein contamination by measuring the optical absorption at 260 and 280 nm. 2. Analyze the samples taken during the GigaPrep purification (bacterial lysate, lysate column flow-through, wash flowthrough, eluted DNA, and precipitated DNA) by mixing samples with 6X Blue/Orange loading dye, and separating on a 1% TBE agarose gel (see Note 23). 3. Analyze the purified DNA using restriction digestion (see Note 24), PCR analysis, and DNA sequencing, as in step 3.1.2.

3.4. Evaluating the Immunological Efficacy of DNA Vaccines

3.4.1. Intradermal Rodent Immunizations

The last section of this protocol describes a technique that can be used to immunize rodents with the vaccine. Although there are several methods that can be used to deliver the vaccine to the recipients, this protocol describes how to inject the vaccine intradermally into rodents (see Note 25). Furthermore, this protocol also describes examples of three assays that can be used to evaluate whether immunization with the DNA vaccine can elicit both an antibody response (as determined by enzyme-linked immunosorbent assays) and a cellular immune response (as determined by IFNγ and granzyme ELISPOT assays [see Note 26]). 1. This protocol assumes that rodents will be immunized intradermally with 100 µg plasmid DNA using hypodermic needles. 2. Purified plasmid DNA is diluted to 1 µg/µL in sterilized 1X PBS (see Note 27). For each animal to be immunized, add 100 µL of this solution to a 1.5-mL microcentrifuge tube, adding an extra 100 µL to the total to account for syringe dead-volume. This solution is mixed and kept on ice until the rodents are anesthetized (see Note 28). 3. Anesthetize the rodents using isoflurane and an isofluorane vaporizer.


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4. Remove the rodents from the vaporizer and maintain sedation using a nose cone. Turn the animals so that their right ears are exposed, and carefully insert the needle (bevel-side up) into the ear pinna intradermally (see Note 29). 5. Slowly inject 100 µL of the DNA vaccine into the ear, watching to ensure that there is not leakage through injection hole (see Note 30). 6. When the vaccine injection is finished, slowly remove the needle, rotating it to prevent any additional leakage during removal. 3.4.2. Analyzing the Immunological Efficacy of DNA Vaccine Rodent Euthanasia and Spleen/Sera Harvest

1. Animals should be humanely killed according to the institution’s protocol. 2. Sera can be collected by cardiac puncture. Use a 22-gauge needle attached to a 1-mL syringe. Insert the needle into the chest cavity at a 45-degree angle and collect blood. Deposit the collected blood into a 1.5-mL microcentrifuge tube. 3. Collect the spleens using a sterile scissors and tweezers. Insert the spleens into a sterile 50-mL Falcon tube filled with 10 mL cold HBSS. Keep the tubes on ice until processing. 4. Process the spleens into splenocytes by passing the spleens through a microsieve using 10 mL HBSS and the plunger of a 5-mL syringe (see Note 31). Collect the spleen cells and put into a 15-mL Falcon tube. 5. Gently underlay the cells with 3 mL Histopaque 1083. Centrifuge the samples at 400 × g for 40 min, turning the centrifuge brake off to prevent disruption of buffy coats. 6. Collect the buffy coat using a Pasteur pipette, and insert it into a fresh 15-mL Falcon tube (see Note 32). Fill the remaining volume of the tube with HBSS, and centrifuge the cells for 10 min at 180 × g. Remove the media, and resuspend the processed splenocytes in RPMI/FBS. Use fresh cells to set up immunological assays in steps 3.4.2.2–3.4.2.4, and freeze the remaining cells. 7. Freeze the excess splenocytes by centrifuging samples for 10 min at 180 × g, and resuspending the cell pellets in a solution of 90% FBS and 10% DMSO at a concentration of 20 × 106 cells/mL. Place 1 mL into a cryovial, and freeze the samples in Mr. Frosty overnight in a −80°C freezer. The next day, remove the samples and freeze them in a liquid nitrogen Dewar. 8. Process the blood into sera by centrifuging the microcentrifuge tubes containing blood for 5 min at ~1,500 × g. Collect the sera and place into a fresh 1.5-mL microcentrifuge tube. Freeze the sera until analysis.

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Antigen-Specific Antibody Enzyme-Linked Immunosorbent Assay

1. This experiment assumes the availability of purified protein and antibody reagents for the antigen of interest (see Note 33). Coat a 96-well Immulon plate with 50 µL of the antigen of interest (diluted in Carbonate buffer) or with 50 µL of Carbonate buffer alone. Let the plate sit overnight at 4°C. 2. Remove the coating solution, and block the plates by addition of 100 µL PBS/1% BSA. Let the samples rotate at room temperature for at least 1 h, and then remove the blocking solution and wash the plate three times with PBS/0.1% Tween-20. 3. Add 50 µL of each sera sample, diluted from 1:25 to 1:200 in PBS/1% BSA. Samples of each dilution should be added to two wells: a well coated with the antigen of interest and a well coated with buffer alone. Let the samples rotate at room temperature for 1 h, then remove the sera and wash the plate three times with PBS/0.1% Tween-20. 4. Add an HRP-labeled secondary antibody specific for the antigen of interest, diluted in PBS/1% BSA. Rotate the plate at room temperature for 1 h, and then wash the plate four times with PBS/0.1% Tween-20. 5. Mix TMB development solution by mixing 4 mL TMB peroxidase substrate with 4 mL peroxidase substrate solution B. Add 75 µL of this solution to each well of the 96-well plate, and monitor color development by measuring the optical density at 650 nm. 6. When samples reach the desired level of development, quench the reaction by addition of 75 µL of 1 N HCl. Measure the optical density of the plates at 450 nm. 7. Analyze the responses by calculating the delta OD values (OD450[antigen] − OD450[buffer alone]), and comparing immunized animals with control-immunized animals.

Antigen-Specific IFNγ Secretion

1. This protocol assumes the use of the R&D Systems mouse IFNγ ELISPOT kits. The day before the splenocytes are to be processed, coat a 96-well nitrocellulose plate with 100 µL of the IFNγ capture antibody diluted in PBS. Let the plate sit at 4°C overnight. 2. In a sterile fashion, remove the antigen solution and add 300 µL of RPMI/FBS. Let the plates block at room temperature for at least 2 h. 3. Remove the blocking solution in a sterile fashion, wash the plates three times with 300 µL RPMI/FBS, and add 100 µL of splenocytes (at a concentration of 2 × 106 cells/mL, or 2 × 105 cells/well). Then add 100 µL of one of four antigen mixtures: 100 µL of media alone, 100 µL containing a nonspecific protein, 100 µL of the specific protein antigen of interest, or 100 µL of a 10 µg/mL ConA positive control. Alternatively,


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add 100 µL of one of two tumor cell lines: a tumor cell line that expresses the antigen of interest, and another tumor cell line that does not express the antigen. Let the samples incubate undisturbed in a 37°C/5% CO2 incubator for 48 h (see Note 34). 4. Remove the cells and wash the plates four times with 300 µL PBS/0.05% Tween-20. Add 100 µL of the secondary biotinlabeled detection antibody (diluted in PBS/1% BSA) to the plates, and let the samples sit overnight at 4°C. 5. Remove the supernatant, and wash the plates four times with 300 µL PBS/0.05% Tween-20. Add 100 µL of alkaline phosphatase-labeled streptavidin (diluted in PBS/1% BSA) and let the samples rotate for 1 h at room temperature. 6. Remove the streptavidin and wash the plates four times with 300 µL PBS/0.05% Tween-20, and once with 300 µL PBS. 7. Prepare the alkaline phosphatase development solution by mixing 150 µL NBT and 75 µL BCIP with 25 mL color development solution, and add 100 µL of this solution to the 96-well plates. Monitor the development, and when the desired development is reached, quench the reactions by washing thoroughly with distilled water. 8. Quantify the number of spots (each representing a cell secreting IFNγ) using an automated ELISPOT plate reader (see Note 35). Analyze the responses by subtracting the results for the media-alone samples from the nonspecific, specific, or positive-control samples, and compare the number of IFNγsecreting cells in the samples stimulated with the nonspecific protein versus the specific protein. Also, compare the antigenimmunized animals with controls to determine the immunological efficacy of the vaccination. Finally, by comparing IFNγ-secreting cells in cultures incubated with tumor cells that express or do not express the antigen of interest, it is possible to determine whether the immunized splenocytes can recognize tumor cells. Antigen-Specific Cytotoxic Immune Responses

1. This protocol assumes the use of the R&D Systems mouse granzyme B ELISPOT kit. The day before the splenocytes are to be processed, coat a 96-well nitrocellulose plate with 100 µL of the granzyme capture antibody diluted in PBS. Let the plate sit at 4°C overnight. 2. In a sterile fashion, remove the antigen solution and add 300 µL of RPMI/FBS. Let the plates block at room temperature for at least 2 h. 3. Remove the blocking solution in a sterile fashion, wash the plates three times with 300 µL RPMI/FBS, and add 100 µL of splenocytes (at a concentration of 2 × 106 cells/mL, or 2

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× 105 cells/well). Then add 100 µL of antigen mixtures: 100 µL of media alone, 100 µL containing a nonspecific protein, 100 µL of the specific antigen of interest, or 100 µL of a Concanavalin A positive control. Alternatively, add 100 µL of one of two tumor cell lines: a tumor cell line that expresses the antigen of interest, and another tumor cell line that does not express the antigen. Let the samples incubate undisturbed in a 37°C/5% CO2 incubator for 48 h. 4. Remove the cells and wash the plates four times with 300 µL PBS/0.05% Tween-20. Add 100 µL of the secondary biotinlabeled detection antibody (diluted in PBS/1% BSA) to the plates, and let the samples sit overnight at 4°C. 5. Remove the supernatant, and wash the plates four times with 300 µL PBS/0.05% Tween-20. Add 100 µL of alkaline phosphatase-labeled streptavidin (diluted in PBS/1% BSA) and let the samples rotate for 1 h at room temperature. 6. Remove the streptavidin and wash the plates four times with 300 µL PBS/0.05% Tween-20 and once with 300 µL PBS. 7. Prepare the alkaline phosphatase development solution by mixing 150 µL NBT and 75 µL BCIP with 25 mL color development solution, and add 100 µL of this solution to the 96-well plates. Monitor the development, and when the desired development is reached, quench the reactions by washing thoroughly with distilled water. 8. Quantify the number of spots (each representing a cell secreting granzyme B) using an automated ELISPOT plate reader. Analyze the responses by subtracting the results for the mediaalone samples from the nonspecific, specific, or positive-control samples, and compare the number of granzyme-secreting cells in the samples stimulated with the nonspecific protein versus the specific protein. Also, compare the antigen-immunized animals with controls to determine the immunological efficacy of vaccination. Finally, by comparing the number of granzyme B-secreting cells in cultures incubated with tumor cells that express or do not express the antigen of interest, it is possible to determine whether vaccination induced potentially cytotoxic, antigen-specific immune responses.

4. Notes 1. There are several factors that should go into deciding which bacterial plasmid should be used as the immunization vector for a DNA vaccine. The plasmid should contain a eukaryotic promoter that can drive expression of the antigen once it is taken


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up by host antigen-presenting cells. These promoters are usually ubiquitous, such as the cytomegalovirus (CMV) immediate-early promoter; however, it is also possible to incorporate cell-specific promoters, such as a dendritic cell (DC)-specific promoter, that will cause the antigen to be expressed only by antigen-presenting cells (18, 19). The plasmid should also contain appropriate restriction enzyme digestion sites that can be used to insert the cDNA of the antigen of interest into the plasmid backbone (these restriction sites will be incorporated into the antigen-specific gene flanking primers during the cloning process—see Note 2). The plasmid also has to contain an antibiotic-resistance gene, allowing bacterial cells that take up the plasmid to be selected based on their antibiotic resistance during the cloning and purification process. Other DNA elements can also be added to the plasmid to enhance the transcription and translation of the antigen, such as an Intron A, Kozak, or poly-adenylation (poly-A) sequences (18). Immunomodulatory sequences can also be added to the DNA plasmid, such as CpG repeats, which are hypomethylated cytosine–guanine repeats often encoded within the plasmid (15). CpG motifs are TLR9 agonists, a receptor that is expressed only by DCs and B cells (15–18, 20). By binding to TLR9 and helping to activate these professional APCs, CpG motifs can serve to activate an innate immune response at the injection site that can contribute to the desired antigen-specific immune response (15–18, 20). Finally, plasmids can also be selected to incorporate more than one antigen of interest. Through the use of an internal ribosomal entry site (IRES), other antigens can be inserted into the vaccine and their expression can be driven off the same promoter. These could potentially encode other tumor antigens or other immunomodulatory antigens such as cytokines that can promote a Th1-type immune response (interleukin [IL]-2, IFNγ, or granulocyte-macrophage colony stimulating factor [GM-CSF]) (18, 21). 2. When designing these primers, ensure that the predicted annealing temperature is at least 50°C to prevent nonspecific amplification (~20–25 base pairs). In addition, it is crucial to ensure that an AUG start codon is included at the 5′ end of the gene to ensure initiation of translation. To further enhance translation, a Kozak sequence can be added at the 5′ end of the start codon, and a poly-A tail can be added to the 3′ end of the coding region. These primers should also incorporate restriction enzyme sites on the flanking ends of the antigen cDNA coding region of the primers. These restriction sites should be unique sites located within the plasmid backbone, but not found in the antigen cDNA. Also, if possible, these enzymes should be compatible for use in simultaneous double restriction enzyme digestions.

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3. The source of template DNA used for amplification will depend on the antigen of interest. Libraries containing the cDNA from different species or tissues can be obtained from various commercial sources, such as ATCC or BioChain (Hayword, CA). Alternatively, if one has access to tissues or cell lines known to express the antigen of interest, a genomic DNA purification can be conducted (QIAGEN DNeasy Blood and Tissue Kit, Qiagen) and the purified DNA can serve as the template DNA for the antigen cDNA amplification PCR in step 3.1.1.2. 4. Preferably, at least one of these enzymes should be for a restriction site found only within the antigen cDNA. When this enzyme is used, it will allow for the determination of whether the purified plasmid DNA contains the antigen of interest. Alternatively, plasmid-specific restriction enzymes can also be used, and the size of the digested fragments can be used to determine whether the plasmid DNA contains the antigen of interest. 5. This antibody should either be specific for the species class the primary antibody was raised against, or against a streptavidin tag conjugated to the primary antibody. This antibody should also be conjugated to alkaline phosphatase. Alternatively, horseradish peroxidase (HRP)-labeled secondary antibodies can also be used, which will require the use of another development reagent such as ECL+ (Thermo Fisher) during Western blotting. 6. When cutting the DNA band out of the gel, try to minimize the gel slice by removing extra agarose. This will result in a higher DNA concentration after the extraction. 7. If the restriction sites incorporated into the flanking primers cannot be used in a simultaneous double digestion, refer to the manufacturer’s instructions for conducting sequential digestions. 8. Ligation reactions should also be conducted in which only plasmid DNA or only antigen cDNA is added. These two reactions will serve as negative controls, ensuring that the bacterial plasmid does not re-ligate to itself. Furthermore, also conduct a ligation reaction in which undigested plasmid is added. This serves as a positive control for the transformation reaction. 9. The success of the cloning and ligation reactions can be initially determined by comparing the number of colonies on the negative control plates (insert or vector alone) with those of the experimental plates. If the negative control plates have no or very few colonies, whereas the experimental plates have many more colonies, the cloning and transformation process was likely successful.


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10. Several alternative cell lines can be used for this transfection procedure, such as COS7 cells, which may require other growth medium conditions. An attractive alternative would be to identify a cell line that is derived from the same species for which the DNA vaccine is intended, one that does not naturally express the antigen of interest, and to use this line for the transfection reactions. In addition, during the transfection reaction, taking time points at 0, 24, and 48 h will usually provide an adequate timeframe to obtain results regarding the translation of the antigen of interest. Additional time points (earlier, later, or more frequent) can be included if more detailed expression kinetics are desired. 11. This protein purification procedure assumes that the target antigen is a cytoplasmic protein. If the antigen of interest is a nuclear or membrane protein, alternative purification methods may be required. Several kits are available, in particular, the CelLytic Nu-CLEAR and CelLytic MEM protein extraction kits (Sigma-Aldrich), which can be used to easily purify nuclear or membrane proteins. 12. If the protein concentration of the CHO lysate is too low to run 50 µg of protein on SDS-PAGE, several alternatives can be considered. First, larger flasks can be used to grow the CHO cells (such as 150-mm flasks), which will provide more cells and thus more protein. Alternatively, the lysis conditions can be enhanced by sonicating the CHO lysates instead of vortexing the lysates in step 3.2.3.1. Finally, if the protein concentration remains too low, the CHO cell pellets can be directly resuspended in 1X Laemmli’s loading buffer. This will make the determination of protein concentration impossible, but will result in loading as much protein as possible into the SDS gel. 13. It is crucial that all air bubbles are removed from between the SDS gel and the nitrocellulose membrane, because any remaining bubbles may disrupt the transfer of the proteins from the gel to the membrane. Consider using a pipette to roll the air bubbles out. Alternatively, the entire gel cassette can be assembled within a tray of Western transfer buffer, with the buffer level covering the entire cassette, which often reduces the presence of air bubbles. 14. If possible, run this reaction in a cold room at 4°C. Alternatively, consider adding an ice pack to the open space within the transfer tank to cool the buffer during the transfer, and add a stir bar to distribute the buffer throughout the tank. 15. Ensure that the corner cut (made just before the transfer) is indicated on the blot for orientation purposes.

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16. If multiple exposures are desired, chemiluminescent reagents can be used. If this is the case, the secondary antibody used during the Western blotting should be conjugated to HRP, and ECL+ reagent (GE Healthcare) along with autoradiography film (Thermo Fisher) can be used to develop the blot. 17. If no protein expression is detected by the Western blot, there are several potential interpretations. First, it could be that the time points selected do not cover the period when the antigen is expressed (either too early or too late). This can be addressed by taking additional CHO time points earlier, later, and more frequently (every 6–12 h). Second, the protein may become degraded during the lysis of the CHO cells—this can be addressed by treating the protein lysis buffer with protease inhibitors. Another possible interpretation is that the protein is degraded by cells soon after it is translated, preventing any protein accumulation and thus a lack of signal. This can be addressed by treating the CHO cells with a proteasome inhibitor for a day before the transfection to prevent any protein degradation. Last, it could be that the antigen is simply not translated. To address this, ensure that the antigen cDNA is properly inserted downstream of the plasmid promoter and that an AUG start codon is included. If so, consider incorporating other elements into the DNA to enhance the translation or stability of the antigen messenger RNA (mRNA), such as a Kozak sequence or a poly-A tail. 18. Several alternative purification methods can be used to isolate large quantities of plasmid DNA. One of the most commonly used alternatives is cesium chloride (CsCl) gradient purification. In this method, bacterial cells are lysed using an alkaline lysis solution, protein is precipitated with sodium acetate and removed by centrifugation, and DNA is precipitated with polyethylene glycol. DNA is then resuspended in a CsCl/EtBr solution, and subjected to ultracentrifugation overnight. The EtBr-containing DNA band is visualized using ultraviolet light, and removed using a 5-cc syringe. Residual EtBr is then removed by passing the sample over an ion exchange column, and DNA is eluted, precipitated, and analyzed as in step 3.3.2. However, EtBr-free purification methods are preferred if the final DNA is to be used for live experiments. Alternatively, several other companies (such as Promega or others) produce purification kits for plasmid DNA that can be used instead of Qiagen’s GigaPrep kit. There are also several commercial companies (such as Aldevron and others) that offer large-scale DNA purification services.


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19. The bacterial growth stage is one of the most important steps in the purification process that will determine the final yield of DNA, and several variables can be addressed to optimize the bacterial growth. For example, different types of growth medium will affect the rate of bacteria growth. Other types of media, such as CircleGrow (MP Biomedicals, Solon, OH), Terrific Broth (Becton-Dickenson Biosciences), etc., can potentially enhance the rate of growth. Regardless of the media used, it is important to ensure that bacteria have reached plateau-stage growth (at least 16 h). In addition, it is important to ensure proper aeration of the flasks during growth. Two-liter Pyrex flasks containing extra-deep baffles will enhance the agitation and aeration of the bacteria during cell growth, and can improve the overall DNA yield. However, although it is important to ensure optimal bacterial growth, the plasmid purification protocol is optimized for a 7.5-g bacterial pellet, and using too much bacteria in this protocol may result in increased levels of genomic DNA and decreased levels of the purified vaccine. 20. In the lysis and neutralization of the bacteria, it is important to ensure that the solutions are well mixed with the bacterial samples. If the bacterial pellet is not completely lysed, the overall yield of DNA will substantially decrease. To help in this process, consider using the QIAGEN LyseBlue reagent, which causes the lysate to turn blue when the lysis solution is mixed in, and white when the neutralization solution is mixed in. However, although sufficient lysis and neutralization is important, it is also important not to mix the bacterial lysates too vigorously, because this will result in excessive lysis and the release of genomic DNA, which will be purified along with the plasmid DNA and contaminate the final product. By mixing the samples vigorously 10–12 times after addition of lysis and neutralization buffers, the samples should be sufficiently mixed to prevent excessive lysis. It is also important to note that after the bacteria have been lysed, the samples should never be vortexed, which can result in shearing of plasmid DNA. 21. Sometimes, when large amounts of bacterial debris are being filtered, QIAfilter mega/giga cartridges can become clogged, requiring the use of an additional cartridge (which will decrease the overall yield of DNA). Alternatively, consider transferring lysates to 250-mL centrifuge tubes, centrifuge for 30 min at ~30,000 × g, remove supernatant, and transfer to fresh 250-mL tube, and spin for an additional 15 min at ~30,000 × g. 22. Endotoxins, or lipopolysaccharides, are bacterial membrane components that are present in the cell membrane

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of all Gram-negative bacteria. These endotoxins strongly influence the transfection of DNA into primary cells, which will decrease the amount of DNA taken up by host cells when the DNA vaccine is injected into recipients. Furthermore, these endotoxins can cause fever, the activation of the complement cascade, and toxic shock. As such, it is very important that these endotoxins are removed from the DNA before the vaccine is injected into live recipients. If using other methods of purification, endotoxins can be removed using commercial endotoxin removal solutions (Sigma-Aldrich, Promega, etc.) 23. If a GigaPrep does not result in high yields of DNA, this gel will be instrumental in determining where the problem lies. The expected pattern would be that a significant amount of DNA is identified in the bacterial lysate and the precipitated DNA, with low levels of DNA found in the flow-through and eluted DNA samples. The most common problem that is found is that low levels of DNA are detected in the bacterial lysate; if this is the case, the bacterial growth conditions should be optimized (see Note 19). 24. These restriction digestions, in addition to determining whether the expected DNA vaccine was purified, can be used to determine whether there are significant levels of genomic DNA contamination. If the digested samples show a high molecular weight smear, this indicates genomic DNA that is being digested multiple times. If this occurs, the lysate may have been treated too roughly (too much mixing in buffer P2 and P3, or vortexing). 25. Although intradermal injections are a common form of DNA immunization, there are several alternative modes and routes of DNA delivery, which are variables that should be considered for each specific application. Another common method that uses hypodermic needles is intramuscular injections, which are prepared in the same fashion as intradermal injections, but the needle is inserted directly into the muscle. This type of injection has been shown to induce DNA expression primarily in skeletal muscle, whereas intradermal injections usually induce expression in keratinocytes (12, 22, 23). However, although intramuscular and intradermal injections are both common methods of vaccine delivery, they both have been shown to have low levels of DNA transfection into target cells (which is unsurprising, because the DNA is simply being introduced outside of cells), and, as such, large amounts of DNA are required for each immunization. To overcome this issue, biolistic injection methods have been developed. These methods rely on the use of gene guns, helium-powered injection devices


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that deliver the DNA vaccine (coated onto microscopic gold beads) directly into the epidermis of the patients, a region rich with Langerhans cells. By directly injecting the DNAcovered beads into the target cells, gene gun injections require far less DNA per immunization (100 to 1,000 times less) to elicit similar levels of responses compared with hypodermic injections (24, 25). Another method that is being investigated to increase the levels of vaccine uptake is the use of in vivo electroporation. This technique involves injecting recipients with the DNA plasmid, and immediately after the injection, an electrical field can be applied using various methods, such as a DermaVax delivery system (Cyto Pulse, Glen Burnie, MD). This causes the cell’s plasma membrane to become more permeable, allowing more of the injected plasmid DNA to be taken up by the host cells. 26. To evaluate the immunological efficacy of DNA vaccines, there are many factors that should go into choosing which assays to use. For example, whether the vaccine is targeting an intracellular protein as opposed to a secreted or transmembrane protein can affect which assays are chosen to evaluate the immune response. If the vaccine is targeting an intracellular protein, detecting the presence of a cellular immune response may be more relevant. Alternatively, if a plasma membrane or secreted protein is being targeted, the detection of antibody responses can potentially be indicative of potential therapeutic benefit. However, in most cases, researchers evaluate the presence of both humoral and cellular immune responses, because there are several assays that can be easily conducted to evaluate the presence of these two branches of the adaptive immune response. To evaluate the presence of an antibody response, ELISA or immunoblots (as described in Step 3.2.3, substituting purified antigen for CHO protein lysates and sera for the primary antibody) are usually used to identify antigen-specific antibody responses in the sera of vaccine recipients. To detect cellular immune responses, there are many different assays that can be conducted. Although ELISPOT assays are becoming more commonly used, several other techniques are used to detect the frequency and functionality of antigen-specific immune responses, such as tetramer staining, T cell proliferation assays, cytotoxicity assays, or measuring the expression of various immunomodulatory cytokines (either cytokine release or intracellular cytokine expression) (26, 27). In all of these assays, it is essential to focus not only on the presence of an immune response, but on whether or not this immune response is specific for tumor cells. To evaluate antitumor immune responses, one of the most useful reagents is

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a syngeneic tumor cell line that either expresses or does not express the antigen of interest. These tumor cell lines can be used in conjunction with ELISPOT assays (as described in this review) to determine whether the recipient generated splenocytes that can specifically recognize these tumor cells. In an even more direct measurement of antitumor reactivity, these tumor cells can be used as target cells in cytotoxicity assays, helping to determine whether splenocytes from immunized recipients can recognize and lyse these cells. These studies can also be conducted in the presence of MHC class I blocking antibodies, the results of which can help determine whether the potential antitumor reactivity is mediated by CD8+ T cells. Ultimately, autologous tumor cells or tumor cell lines may be most useful in directly evaluating the antitumor efficacy of DNA vaccination in vivo. In these studies, rodents can be challenged with antigen-expressing tumor cells before or after DNA immunization (to determine the therapeutic and protective antitumor efficacy of the vaccine, respectively). Tumor growth curves can then be measured to determine whether immunizing rodents with the DNA vaccine can lead to a decrease or delay in tumor growth. However, these experiments are highly application specific; they will depend not only on the type of tumor being studied, but also the rodent model, route of tumor administration (subcutaneous, transgene driven, etc.), the type of antigen being targeted, and so forth. Although the methods described in this review focus on evaluating whether or not the DNA vaccine elicits an immune response in vivo, there are several reviews that can be used to help determine assays that may be helpful in evaluating the presence of an antitumor immune response (26–31). 27. Although this protocol assumes the use of DNA diluted in PBS, studies have evaluated the dilution buffers used in the administration of DNA vaccines, focusing on variables such as injection volume, tonicity of injection fluid, type of solute, etc. These studies have shown that injecting DNA in hypotonic solutions (such as water) resulted in 95% lower gene expression compared with buffered solutions (32). In addition, although Wolff and colleagues investigated various solutions and buffers on gene expression in the muscle, they found that no vehicle resulted in a significant increase in gene expression when compared with plasmid diluted in saline (33). 28. Although this protocol does not assume the use of any additional adjuvants, there are several adjuvants that have been given along with immunizations with DNA vaccines. One of the most common adjuvants is GM-CSF, which activates the production of white blood cells. Research has shown that


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GM-CSF can enhance the immunogenicity of DNA vaccines by inducing dendritic cell activation and migration as well as the expansion of B and T cells, which can facilitate the development of an adaptive immune response (34, 35). Research has also evaluated the use of other immunomodulatory cytokines, such as IL-2, IL-12, and IL-4, which have been shown to enhance immune responses or skew them toward either Th1- or Th2-type responses (35). These adjuvants are important, especially in the context of hypodermic injections, to help facilitate uptake of the DNA vaccine by the host’s antigen-presenting cells. 29. This can be facilitated by using a finger behind the ear to push the center of the ear pinna outward, exposing a small flap of skin in the center. This flap can be used as an injection site, inserting the needle bevel-side up into the flap until it is approximately 0.5 cm into the skin. When inserting the needle into the ear, ensure that the needle does not poke through the ear, which will cause the vaccine to leak. If this occurs, gently remove needle, and select a different insertion site in the ear for the injection. 30. Although the ear pinna is an easily accessible site for intradermal injections (and has been well characterized as an injection site (36)), there are other alternative sites for intradermal injections, such as the footpad or flank. 31. Be very gentle in processing cells—the more harshly they are processed, the worse the recovery will be. 32. The buffy coat is the cloudy, opaque layer located below the red HBSS and above the clear Histopaque layer, and contains the desired splenocyte population. 33. Several alternative methods can be used to detect antibody responses. If no purified antigen source is available, capture ELISAs can be conducted. In this protocol, 96-well plates are coated with an antibody specific for the antigen of interest, which is followed by incubation with a cell line known to express the protein of interest. During this incubation, the antigen will bind to the primary antibody, and, after washing off any nonspecific interactions, a secondary antibody (that is specific for a different epitope on the antigen than the primary antibody) can be added to the ELISA, followed by HRP detection. Alternatively, a Western blot can be conducted. In this case, a protein lysate can be prepared from a cell line that expresses the antigen of interest. This lysate can be separated using SDS-PAGE and analyzed on a Western blot using antigen-specific antibodies. 34. A potential variable of these ELISPOT assays that can be examined is the incubation time. Mouse ELISPOTs are

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usually conducted as 40- to 48-h assays, but this should be confirmed for each particular application. In addition, it is crucial that the plates are incubated in an incubator that will not be disturbed. Large amounts of jostling during the incubation can result in poorly defined spots and higher levels of background. 35. Alternatively, spots can be manually counted by placing the plate under a light microscope and counting individual spots.

Acknowledgments This work is supported by Department of Defense (DOD) grant W81XWH-07–1–0088 to BMO and by National Institutes of Health (NIH) grant K23 RR16489 to DGM. References 1. Liu , M. A. (2003) DNA vaccines: a review. J Intern Med 253, 402–10. 2. van der Bruggen, P. and Van den Eynde, B. J. (2006) Curr Opin Immunol 18, 98–104. 3. Ciernik, I. F., Berzofsky, J. A., and Carbone, D. P. (1996) Processing and presentation of tumor antigens and vaccination strategies. J Immunol 156, 2369–75. 4. Pilon, S. A., Piechocki, M. P., and Wei, W. Z. (2001) Vaccination with cytoplasmic ErbB-2 DNA protects mice from mammary tumor growth without anti-ErbB-2 antibody. J Immunol 167, 3201–6. 5. Roos, A. K., Pavlenko, M., Charo, J., Egevad, L., and Pisa, P. (2005) Induction of PSAspecific CTLs and anti-tumor immunity by a genetic prostate cancer vaccine. Prostate 62, 217–23. 6. Cassaday, R. D., Sondel, P. M., King, D. M., Macklin, M. D., Gan, J., Warner, T. F., Zuleger, C. L., Bridges, A. J., Schalch, H. G., Kim, K. M., Hank, J. A., Mahvi, D. M., and Albertini, M. R. (2007) A phase I study of immunization using particle-mediated epidermal delivery of genes for gp100 and GM-CSF into uninvolved skin of melanoma patients. Clin Cancer Res 13, 540–9. 7. Liu, M. A. and Ulmer, J. B. (2005) Human clinical trials of plasmid DNA vaccines. Adv Genet 55, 25–40. 8. Pavlenko, M., Roos, A. K., Lundqvist, A., Palmborg, A., Miller, A. M., Ozenci, V.,

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Bergman, B., Egevad, L., Hellstrom, M., Kiessling, R., Masucci, G., Wersall, P., Nilsson, S., and Pisa, P. (2004) A phase I trial of DNA vaccination with a plasmid expressing prostate-specific antigen in patients with hormone-refractory prostate cancer. Br J Cancer 91, 688–94. Triozzi, P. L., Aldrich, W., Allen, K. O., Carlisle, R. R., LoBuglio, A. F., and Conry, R. M. (2005) Phase I study of a plasmid DNA vaccine encoding MART-1 in patients with resected melanoma at risk for relapse. J Immunother (1997) 28, 382–8. Zlotocha, S., Staab, M. J., Horvath, D., Straus, J., Dobratz, J., Oliver, K., Wasielewski, S., Alberti, D., Liu, G., Wilding, G., Eickhoff, J., and McNeel, D. G. (2005) Phase I study of a plasmid DNA vaccine encoding MART-1 in patients with resected melanoma at risk for relapse. Clin Genitourin Cancer 4, 215–8. Nichols, W. W., Ledwith, B. J., Manam, S. V., and Troilo, P. J. (1995) Potential DNA vaccine integration into host cell genome. Ann N Y Acad Sci 772, 30–9. Wolff, J. A., Malone, R. W., Williams, P., Chong, W., Acsadi, G., Jani, A., and Felgner, P. L. (1990) Direct gene transfer into mouse muscle in vivo. Science 247, 1465–8. Disis, M. L., Shiota, F. M., McNeel, D. G., and Knutson, K. L. (2003) Direct gene transfer into mouse muscle in vivo. Immunobiology 207, 179–86.

Methods for Constructing and Evaluating Antitumor DNA Vaccines 14. McNeel, D. G., Schiffman, K., and Disis, M. L. (1999) Immunization with recombinant human granulocyte-macrophage colonystimulating factor as a vaccine adjuvant elicits both a cellular and humoral response to recombinant human granulocyte-macrophage colony-stimulating factor. Blood 93, 2653–9. 15. Hemmi, H., Takeuchi, O., Kawai, T., Kaisho, T., Sato, S., Sanjo, H., Matsumoto, M., Hoshino, K., Wagner, H., Takeda, K., and Akira, S. (2000) A Toll-like receptor recognizes bacterial DNA. Nature 408, 740–5. 16. Klinman, D. M., Yamshchikov, G., and Ishigatsubo, Y. (1997) Contribution of CpG motifs to the immunogenicity of DNA vaccines. J Immunol 158, 3635–9. 17. Krieg, A. M., Yi, A. K., Schorr, J., and Davis, H. L. (1998) The role of CpG dinucleotides in DNA vaccines. Trends Microbiol 6, 23–7. 18. Garmory, H. S., Brown, K. A., and Titball, R. W. (2003) DNA vaccines: improving expression of antigens. Genet Vaccines Ther 1, 2. 19. Weeratna, R. D., Wu, T., Efler, S. M., Zhang, L., and Davis, H. L. (2001) Designing gene therapy vectors: avoiding immune responses by using tissue-specific promoters. Gene Ther 8, 1872–8. 20. Hornung, V., Rothenfusser, S., Britsch, S., Krug, A., Jahrsdorfer, B., Giese, T., Endres, S., and Hartmann, G. (2002) Quantitative expression of toll-like receptor 1–10 mRNA in cellular subsets of human peripheral blood mononuclear cells and sensitivity to CpG oligodeoxynucleotides. J Immunol 168, 4531–7. 21. Liu, M. A., Wahren, B., and Karlsson Hedestam, G. B. (2006) DNA vaccines: recent developments and future possibilities. Hum Gene Ther 17, 1051–61. 22. Hengge, U. R., Walker, P. S., and Vogel, J. C. (1996) Expression of naked DNA in human, pig, and mouse skin. J Clin Invest 97, 2911–6. 23. Raz, E., Carson, D. A., Parker, S. E., Parr, T. B., Abai, A. M., Aichinger, G., Gromkowski, S. H., Singh, M., Lew, D., Yankauckas, M. A., and et al. (1994) Expression of naked DNA in human, pig, and mouse skin. Proc Natl Acad Sci USA 91, 9519–23. 24. Fynan, E. F., Webster, R. G., Fuller, D. H., Haynes, J. R., Santoro, J. C., and Robinson, H. L. (1993) DNA vaccines: protective immunizations by parenteral, mucosal, and gene-gun inoculations. Proc Natl Acad Sci USA 90, 11478–82. 25. Pertmer, T. M., Eisenbraun, M. D., McCabe, D., Prayaga, S. K., Fuller, D. H., and Haynes,

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Chapter 13 Immunity of Lentiviral Vector-Modified Dendritic Cells Shuhong Han and Lung-Ji Chang Summary Innovative approaches to induce a strong immune response are key to the success of immunotherapy. Dendritic cells (DCs) are professional antigen-presenting cells (APCs) equipped with co-stimulatory, adhesion, and major histocompatibility complex (MHC) molecules needed for initiation and reactivation of the immune response. DCs are able to initiate and stimulate both innate and adaptive immune responses and, by secretion of cytokines, chemokines, and expression of regulatory molecules, to shape the adaptive immune response toward a long-lasting memory immunity. DCs from the peripheral blood of immune-compromised patients, however, often display an immature phenotype with defective functions. This emphasizes the importance and potential of engineering antigen-specific DCs in vitro. A state-of-theart approach to overcome the prevailing immune dysfunction(s) in patients is to engineer DCs or DC progenitors to generate fully functional DCs for the modification of host immunity. Lentiviral vectors (LVs) are highly efficient gene transfer vehicles for engineering DC functions. Examples of lentiviral vectors encoding immune-modulatory genes and useful functional assays for the analysis of effector immune cell response are described in this chapter. Key words: Dendritic cell, immunotherapy, lentiviral vector, T cell.

1. Introduction Dendritic cells (DCs) are important immune cells in initiating and maintaining immunity (1). Efficient gene transfer into DCs has been difficult but lentiviral vectors (LVs) can transduce DCs at high efficiencies with little to no cytotoxicity (2–5). Modification of DCs with genetic vectors can potentially activate a strong protective immunity against infections and cancer (6–10). Many studies have attempted to express target antigenic genes rather than immune-modulatory genes in DCs, which may


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not overcome the preexisting intrinsic immune dysfunctions in patients. To establish an effective immune response, interactions between antigen-presenting cells and effector immune cells are required; these interactions involve multiple mediators, including major histocompatibility complex (MHC) molecules harboring antigenic peptides (signal 1); co-stimulatory molecules such as CD80, CD86, and intercellular adhesion molecule (ICAM)-1 (signal 2); and cytokines such as interleukin (IL)-2, IL-4, and IL-12 (signal 3) (11). Upregulation or downregulation of immune-modulatory gene expression in DCs can alter subsequent T cell responses. For example, production of IL-12 by DCs early in an immune response is critical for polarizing T cells toward Th1 function, which is essential for the clearance of intracellular pathogens. IL-10, on the other hand, suppresses IL-12 production from DCs and diminishes the commitment of Th1 differentiation (12, 13). Lentiviral transduction of DCs to deliver IL-12 gene or

Fig. 1. Immune analysis of DCs modified with LVs encoding immune-modulatory genes. DCs are generated from PBMCs as illustrated in the diagram and modified with LVs on day 5. After maturation, Ag-primed DCs and immune cells are cocultured for 10–20 days to generate Ag-specific effector cells. Examples of immune-modulatory genes, including co-stimulator genes (CD80, CD86, CD137L), cytokine genes (IL-12, IL-7), activation gene (ICAM-1), and chaperone gene (calnexin or CNX) are shown in monocistronic or multicistronic lentiviral constructs. DCs infected with LV-CD86 showed increased CD86 expression (from 49% to 76%, or mean fluorescence index from 28.8 to 71.4) as demonstrated by antiCD86 antibody staining and flow cytometry analysis.

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small interfering RNA gene targeting IL-10 has been shown to promote a strong Th1 response (4). Besides cytokine signaling, increased interaction of co-stimulatory molecules and adhesion molecules such as CD80, CD86, CD137, and ICAM-1 are also critical for the development of a long-lasting memory immunity (14–17) (see examples in Fig. 1). In addition, supraphysiological expression of a chaperone protein, calnexin, in DCs has been shown to overcome intrinsic T regulatory activity in cancer patients (18).. Extensive in vitro analysis of immune functions of DCs is necessary before the gene-modified cells are applied in human trials. Several experimental approaches for phenotyping and functional evaluation of human DCs and immune effector cells are described below.

2. Materials 2.1. Preparation of Peripheral Blood Mononuclear Cells and DCs

1. Ficoll-Hypaque solution (density 1.077 g/L) from GE Healthcare Bio-Sciences (Buckinghamshire, UK). 2. Complete RPMI 1640 medium (Gibco-BRL, Carlsbad, CA, USA) supplemented with 10% 56°C/30 min heat-inactivated fetal bovine serum (FBS) (HyClone, Logan, UT, USA) and supplemented with 100 μg/mL streptomycin and 100 IU/ mL penicillin (Gibco-BRL). 3. AIM-V medium (Gibco-BRL). 4. Recombinant human granulocyte macrophage colonystimulating factor (GM-CSF) and IL-4 (Biosource, Sunnyvale, CA, USA) are prepared as 1,000x stock and stored in aliquots at −80 ° C. 5. 6-well plates and 24-well low attachment plates (Costar, Corning, NY, USA).

2.2. Lentiviral Preparation and Transduction of Immature DCs and Maturation of Gene-Modified DCs

1. Lentivirus vectors encoding immune-modulatory genes, see Fig. 1 for examples. All lentiviral backbone plasmid components are available from National Institutes of Health, Bethesda, MD, USA (see Note 1). 2. Polybrene is dissolved in appropriate medium as a stock solution of 20 mg/mL (Sigma, St. Louis, MO, USA). 3. Superfect (available from Qiagen, Valencia, CA, USA) or other transfection reagents. 4. Recombinant human tumor necrosis factor (TNF)-α and interferon (IFN)-γ (R&D Systems, Minneapolis, MN, USA)

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and lipopolysaccharide (LPS, Sigma) are prepared as 1,000x stock solutions and stored in aliquots at −80°C. 5. Sterile 0.45-μm low protein-binding filters (Millipore, Billerica, MA, USA). 6. Genomic DNA preparation kit (Promega, Fitchburg, WI, USA). 2.3. Flow Cytometry Analysis

1. FACS buffer: phosphate-buffered saline (PBS) containing 2% FBS and 0.09% NaN3. 2. Fixation solution: FACS buffer containing 1% formaldehyde (Fisher Biotech, Springfield, NJ, USA). 3. Flow cytometry tubes (BD Biosciences, San Jose, CA, USA). 4. Normal mouse serum (Sigma) for blocking nonspecific antibody binding. 5. Fluorochrome-labeled antihuman CD11c, CD40, CD86, CD80, HLA-I, HLA-DR, CCR7, DC-LAMP, CD83, CD3, CD14, CD19, CD3, CD4, CD8 antibodies and isotype controls (BD Biosciences).

2.4. Assay for Antigen Uptake Function of Immature DCs

1. DQ-OVA is prepared as 1 mg/mL stock solution in PBS and stored at −20°C protected from light (Molecular Probes, Inc., Eugene, OR). 2. APC-conjugated antihuman CD11c antibody (BD Biosciences).

2.5. Allogeneic Mixed Lymphocyte Reaction Stimulated by DCs

1. Human AB serum (Valley Biomedical, Winchester, VA, USA). 2. Carboxyfluorescein succinimidyl ester (CFSE) (Molecular Probes, Inc., Eugene, OR) is dissolved in dimethyl sulfoxide (DMSO) as a 5 mM stock and stored in aliquots at −20°C. 3. 96-well U-shape plates (Costar).

2.6. Activation of Antigen-Specific T Cells by Lentiviral Vector-Modified DCs

2.7. Detection of Intracellular Cytokines by Flow Cytometry and CD107a Translocation Assay

1. Recombinant human IL-2, IL-7, and IL-15 (Gentaur, Aachen, Germany) are prepared in sterile PBS with 0.1% BSA and stored in aliquots at −80°C. 2. Anti-CD3 antibody for positive control of activated T cells (eBioscience, San Diego, CA, USA). 1. Fluorochrome-conjugated antihuman cytokine antibodies: anti-IL-2, anti-TNFα, anti-IFN-γ (BD Biosciences). 2. Golgiplug (Brefeldin A) or GolgiStop (Monensin) (BD Biosciences). 3. Fluorochrome-conjugated antihuman CD107a antibody (BD Biosciences).


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1. Propidium iodide (PI) is dissolved in PBS as 1,000x stock solution at 1 mg/mL and stored at −20°C protected from light (Sigma). 2. Calibrate beads: APC-conjugated beads as internal control (BD Biosciences).

2.9. Multimer Binding Analyses

1. MHC-peptide specific tetramers (Beckman Coulter, Fullerton, CA, USA) or pentamers (Proimmune, Bradenton, FL, USA) can be purchased from vendors, or requested from National Institutes of Health MHC Tetramer Core Facility (http:// www.niaid.nih.gov/reposit/tetramer/genguide.html).

3. Methods 3.1. Preparation of Dendritic Cells from Peripheral Blood

1. Place fresh heparinized blood into 15- or 50-mL conical centrifuge tubes and dilute with equal volume of PBS (see Note 2). 2. The blood/PBS mixture is gently layered over 1/3 to 1/2 volume of Ficoll-Hypaque solution and centrifuged at 600g, 18–20°C for 25 min. 3. Remove the upper layer that contains plasma and platelets and transfer the peripheral blood mononuclear cell (PBMC) layer to a new centrifuge tube; wash cells three times with two to three volumes of PBS, and centrifuge at 450g at room temperature for 10 min. 4. Resuspend cells in AIM-V and determine cell number and viability by trypan blue exclusion (see Note 3). 5. Plate PBMCs at 107 cells/2 mL/well in 6-well tissue culture plates and let adhere at 37°C for 2 h (see Note 4). 6. The nonadherent cells are gently removed and cryopreserved as source of immune cells. 7. The adherent cells are cultured for 5 days in AIM-V supplemented with 50 ng/mL GM-CSF and 25 ng/mL IL-4. The cells are fed with the same medium with growth factors every other day. 8. On day 5, immature cells are collected and plated in 24-well low attachment plates in AIM-V supplemented with 1 mg/mL LPS, 50 ng/mL TNFα, and 50 ng/mL IFN-γ for maturation (see Note 5). 9. Mature DCs are harvested 24–48 h later, and the phenotype of mature DCs is analyzed by flow cytometry.

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3.2. Preparation of Lentiviral Vectors Encoding Immune Modulatory Genes

1. Prepare high purity vector plasmids including packaging construct (pHP), envelope construct (pVSV-G), activating construct (pTat), and transducing construct (pTY) for DNA co-transfection (see Note 1, and Fig. 1 for illustration of lentiviral vectors). 2. Split 293T cells using trypsin and plate into 6-well plates at 90% confluency 16–24 h before DNA co-transfection. 3. Transfect 293T cells with all four plasmid DNA for vector production using Superfect per manufacturer’s instruction (Qiagen, Valencia, CA, USA). 4. Harvest virus supernatants at 12, 24, and 36 h after transfection, centrifuge at 1,000g for 20 min to remove cell debris, and filter through a sterile 0.45-μm filter; store vectors at −80°C in aliquots before use. Further concentration of vectors by centrifugation may be performed as described (19). 5. Titrate LVs by infecting TE671 at serial dilutions of 10−4, 10−5, and 10−6, as described (20). Harvest the infected cell genomic DNA using a genomic DNA preparation kit and perform quantitative PCR to determine the provirus copy number, which can be converted to infectious units.

3.3. Lentiviral Transduction of Dendritic Cells

1. Plate immature DCs into 24-well low attachment plates at 5 × 105 cells/well with 200 μL of AIM-V supplemented with 50 ng/mL GM-CSF, 25 ng/mL IL-4, and 5 μg/mL polybrene. 2. Add LVs at multiplicity of infection (MOI) of 20–40; one MOI equals one infectious unit per cell. 3. Incubate cells at 37°C for 2 h, with gentle tilting every 30 min. 4. Add 1 mL of AIM-V supplemented with 50 ng/mL GM-CSF and 25 ng/mL IL-4 and continue incubation for an additional 12 h. 5. Induce DC maturation by adding 1 μg/mL LPS, 50 ng/mL TNFα, and 50 ng/mL IFN-γ for 24–48 h (see Fig. 1 for illustration); mature DCs can be harvested and pulsed with peptides at 37°C, 5% CO2 for 2 h in a 24-well low attachment plate.

3.4. DC Phenotype Analysis

1. DCs are harvested and washed twice with cold FACS buffer and resuspended in cold FACS buffer at a concentration of 5 × 105 cells/mL. 2. Add 200 μL of suspended cells into flow tube for analysis. 3. To each tube, add mouse serum (10% final concentration) and incubate on ice for 30 min to block nonspecific antibody binding.


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4. Add 3 mL of cold FACS buffer and centrifuge at 400g for 5 min; discard supernatant and resuspend cells in 100 μL of FACS buffer; add specific fluorochrome-labeled antibodies and incubate on ice for 30 min. 5. Add 3 mL of cold FACS buffer and centrifuge at 400g for 5 min; discard supernatant and wash cells twice with cold FACS buffer. 6. Resuspend cells in 500 μL of cold fixation buffer and store at 4°C away from light for 1–24 h before FACS analysis. An example of DC phenotype analysis is shown in Fig. 2. 3.5. Analysis of Antigen Uptake Function of Immature DCs

1. Immature DCs are harvested and washed with AIM-V twice and resuspended in AIM-V at a concentration of 5 × 105 cells/ mL. 2. Add 200 μL of cells in a flow tube with specific amount of DQ-OVA (see Note 6). 3. Incubate at 37°C for 1 h and, at the same time, set up control groups and incubate at 4°C for 1 h. 4. Wash cells three times with cold FACS buffer and resuspend in 100 μL of cold FACS buffer. 5. Analyze DCs with APC-conjugated anti-CD11c antibody as described above (Subheading 3.4).

Fig. 2. DC phenotyping by surface marker staining and flow cytometry analysis. Mature DCs were generated as illustrated in Fig. 1 and stained with fluorochrome-labeled antibodies against different surface markers as indicated. The number represents percentage of positive cells.

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3.6. Allogeneic Mixed Lymphocyte Reaction for DC Functional Analysis

1. Wash PBMCs (as allogeneic responder cells) twice with PBS and resuspend cells in prewarmed (37°C) PBS containing 1 μM of CFSE at a density less than 2 × 107 cells/mL and incubate at 37°C for 15 min. 2. Centrifuge at 320g for 7 min and resuspend cells in prewarmed complete RPMI containing 10% FBS. 3. Incubate cells at 37°C for 30 min and wash once with complete RPMI containing 10% FBS. 4. Resuspend the CFSE-stained PBMC at 1 × 106 cells/mL in AIM-V containing 5% human AB serum and aliquot 100 μL into a U-bottom 96-well plate at 1 × 105 cells/well. 5. Add irradiated immature DCs or mature DCs (ratio to responder cells, 1:20) at a final volume of 200 μL/well. CFSE-labeled PBMCs without DCs are included in separated wells as control. 6. After 5 days, harvest cells and incubate with labeled antibodies against surface markers such as CD3, CD4, and CD8, and the intensity of CFSE is determined in gated CD3, CD4, or CD8 cell populations as analyzed by flow cytometry (see Fig. 3 example).

Fig. 3. Ag-specific CD8 T cell proliferation analysis by CFSE dye dilution method. DCs were pulsed with EBV LMP2 (late membrane protein 2) peptides and cocultured with autologous immune cells for 15 days. The expanded immune cells were labeled with CFSE and exposed to DCs prepulsed with control or EBV LMP2 peptides and cultured for 5 days. Agspecific CD8 T cells were detected by anti-CD8 antibody and CFSE density analyzed by flow cytometry. Positive control of nonspecific proliferation is shown as cells treated with anti-CD3/anti-CD28 antibodies.


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1. Thaw autologous nonadherent PBMCs in AIM-V medium and count cell number and determine viability by trypan blue exclusion. 2. LV-modified and/or peptide-loaded mature DCs in AIM-V are irradiated by 2,000 rad and washed twice with AIM-V. 3. Mix nonadherent PBMCs and irradiated DCs at a ratio of 20:1 and centrifuge at 320g for 7 min. Resuspend the mixed cells in AIM-V supplemented with 5% human AB serum (see Note 7). 4. Plate cells at 6 × 106 cells/2 mL/well in AIM-V in a 24-well plate (see Note 8) and on day 3, replace half of the medium with fresh medium containing 12.5 U/mL IL-2, 10 ng/mL IL-7, and 20 ng/mL IL-15, and feed the culture every other day with fresh cytokine-containing medium. 5. Split cells into two wells as necessary (see Note 9). 6. After incubation for 10–20 days, the cells can be analyzed for antigen-specific effector functions.

3.8. Analyses of Antigen-Specific Effector Functions 3.8.1. Proliferation Analysis by CFSE Dye Dilution

1. Prepare CFSE-labeled responder cells using DC-primed T cells. Antigen-specific T cells are washed with PBS twice and resuspended in prewarmed (37°C) PBS containing 1 μM CFSE (less than 2 × 107 cells/mL) and incubated at 37°C for 15 min ((21) (see Note 10). 2. Centrifuge at 320g for 7 min and resuspend cell pellets in prewarmed complete RPMI 1640 containing 10% FBS, and incubate for another 30 min. 3. Wash and resuspend cells in AIM-V containing 5% human AB serum and aliquot 100 μL/well at 1 × 106 cells/mL into a U-bottom 96-well plate (1 × 105 cells/well). 4. Prepare stimulators using mature DCs, monocytes, autologous Epstein-Barr virus (EBV)-transformed B lymphocytes (BLCLs) or Ag (antigen)-expressing cells. Irradiate the stimulators, DCs, and monocytes at 2,000 rad and BLCLs at 5,000 rad (see Note 11). 5. Wash stimulator cells twice with AIM-V and resuspend in AIM-V containing 5% human AB serum. The cell density for DCs/monocyets and BLCLs are 1 × 105 cells/mL and 2 × 105 cells/mL, respectively. 6. Aliquot 100 μL of irradiated stimulator cells into each well and mix well with responder T cells; the ratio for DCs/monocytes to T cells is 1:10, and for BLCLs to T cells is 1:5. Set up triplicates for each treatment. At the same time, set up a responderalone group without stimulators as a control for background proliferation, and responder T cells plus anti-CD3 antibody

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(1 μg/mL) as a positive control. The total volume of each well is 200 μL. 7. Incubate the plate at 37°C in a 5% CO2 incubator for 3 days; stain cells with antibodies for CD3, CD4, and CD8; and analyze by flow cytometry (see Note 12). 3.8.2. Analysis of Ag-Specific T Cell Response by Intracellular Cytokine Staining

1. Prepare responder cells (3 × 106 cells/mL) as described above (Subheading 3.8.1) without CFSE staining. 2. Prepare irradiated stimulator cells (DCs/monocytes, 3 × 105 cells/mL and BLCLs, 6 × 105 cells/mL) as described above (Subheading 3.8.1). 3. Add 100 μL of responder cells (3 × 105 cells/well) and 100 μL of irradiated stimulator cells in each well of a U-bottom 96-well plate; for DCs/monocytes, the ratio of T cells and stimulators is 10:1, and for BLCLs, the ratio is 5:1. Set up triplicates for each treatment. Responder cells alone without stimulators serve as a negative control for background cytokine secretion, and responder cells and stimulators plus anti-CD3 antibody (1 μg/mL) serves as a positive control. The total volume of each well is 200 μL. 4. Incubate at 37°C for 1 h, add GolgiPlug, Brefeldin A (BFA) to each well (0.1% v/v) and continue incubation for 5 h. 5. Collect cells, wash once with cold FACS buffer, and stain with fluorochrome-conjugated antihuman CD4, CD8, and CD3 antibodies. 6. Intracellular cytokine staining is performed using fluorochrome-labeled antihuman antibodies against IFN-γ, TNFα, and IL-2.

3.8.3. Non-Radioactive, CFSE-Based CTL Assay

1. Prepare CTLs as described in Subheading 3.8.1, without CFSE staining. 2. Prepare CFSE-labeled target cells (1 × 105 cells/mL) as described in Subheading 3.8.1 (22). 3. Aliquot 100 μL of target cells per well (1 × 104 cells/well) in a U-bottom 96-well plate, add CTLs to target cells at various ratios, for example, 0:1, 5:1, 10:1, and 20:1, centrifuge at 200g for 3 min, and incubate the plate at 37°C for 6 h. 4. Transfer cells to flow tubes and wash twice with cold FACS buffer. 5. Resuspend cells in 0.5 mL cold FACS buffer, add 1 μg/mL propidium iodide, and incubate at room temperature for 30 min. 6. Transfer cells on ice, spike in 2 × 104 calibration beads, mix well, and proceed to FACS analysis (see Note 13).


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7. Collect 10,000 events of the microbeads for each sample and register the CFSE-stained target cell number. Example of target cell killing by CMV–pp65-specific CTLs is shown in Fig. 4.

Fig. 4. A sensitive, nonradioactive, Ag-specific CTL assay. DCs were pulsed with CMV pp65 peptides and cocultured with autologous immune cells as described in Fig. 1. The CTLs were mixed with CFSE-labeled target cells (BLCLs), which were pulsed with pp65 peptides or control peptides (BLCL-mk) at different ratios (0:1, 5:1, and 10:1) and cultured for 5 h. The cells were spiked with control fluorochrome-labeled beads and analyzed by flow cytometry. The survival percentages were calculated based on the disappearance of target cells in relation to the control beads and shown as bar graphs.

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The result is presented as: %Survival=

3.8.4. CD107a Translocation Assay for CTL Function

RemainingCFSEstainedcells(withCTLs) × 100 RemainingCFSEstainedcells(withoutCTLs)

1. Prepare effector CTLs (3 × 106 cells/mL) as described in Subheading 3.8.1 without CFSE staining. 2. Prepare target cells as described in Subheading 3.8.1; for DCs and monocytes, the concentration is 3 × 105 cells/mL, and for BLCL, the concentration is 6 × 105 cells/mL. 3. Mix 100 μL of CTLs (3 × 105 cells/well) and 100 μL of irradiated target cells (10:1 for DCs and 5:1 for BLCL, for example) into a U-bottom 96-well plate; set up triplicates for each treatment. At the same time, set up CTLs alone without target cells as a background CD107a detection and CTLs and stimulators plus anti-CD3 antibody (1 μg/mL) as a positive control. 4. Add PE-labeled anti-CD107a antibody (refer to manufacture’s instructions) and incubate at 37°C, 5% CO2 for 1 h, and add GolgiPlug (0.1% v/v) for another 5 h (23) (see Note 14). 5. Wash cells and incubate with labeled antihuman CD8, CD4, and CD3 antibodies, followed by intracellular staining with antihuman cytokine antibodies against IFN-γ, TNFα, and IL-2.

3.8.5. Multimer Binding Analysis of Ag-Specific CTLs

1. Wash CTLs twice and resuspend in FACS buffer at a concentration of 5 × 105 cells/mL. 2. Add 200 μL of sample into a flow tube for each multimer assay. 3. To each tube, add appropriately titrated multimer; for MHC class I tetramer (e.g., from Beckman Coulter), incubate at room temperature for 30 min; for MHC class I pentamer (Proimmune), incubate at room temperature for 15 min; for MHC class II tetramer (Beckman Coulter), incubate at 37°C in 5% CO2 for 2 h (see Note 15). 4. Add 3 mL of cold FACS buffer, centrifuge at 400g for 5 min, discard supernatant, and resuspend cells in 100 μL of cold FACS buffer. 5. Add labeled antihuman CD4, CD8, and CD3 antibodies and incubate at 4°C for 30 min; dilute with 3 ml of FACS buffer and centrifuge at 400 g for 5 min (see Note 16). 6. Wash cells twice and resuspend in 500 μL of cold fixation FACS buffer for analysis.


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4. Notes 1. All vector plasmids are freely distributed from NIH AIDS Reference and Reagent Program (https://www.aidsreagent. org/Index.cfm) donated by the author; the plasmid DNA quality is important for high efficiency transfection and high titer vector production. 2. PBMCs from leukapheresis should be diluted with four volumes of PBS. 3. The washing step usually removes platelets. With excess platelet contamination, layer PBMCs (1–2 × 107 cells/mL) over 3 mL of FBS, centrifuge at 200g for 15 min, and discard the supernatant containing platelets. 4. If PBMCs are collected from granulocyte-specific colonystimulating factor (G-CSF)-mobilized peripheral blood, because of high monocyte content, fewer cells (~6–8 × 106 cells) should be plated into 6-well plates. 5. Highly adherent immature DCs can be treated with 5 mM EDTA in AIM-V at 37°C for 20 min to increase recovery. 6. The concentration of DQ-OVA should be titrated beforehand. A too-high concentration of DQ-OVA inhibits the uptake function of immature DCs. 7. When enriched T cells (for example, CD4+, CD8+, or CD3+ T cells) are used for coculture with DCs, the ratio of T cells to DCs should be adjusted to 10:1 (24). 8. During the initial 3–4 days of coculture, do not disturb the cells to ensure the quality of synapse formation of DCs and T cells. 9. The growth of T cells is dependent on appropriate cell density. If the cell number is low, cells should be plated into a U-bottom 96-well plate at 5 × 105 cells/well. 10. A high concentration of CFSE may affect T cell proliferation. An optimal concentration of CFSE may need to be determined for each experiment (25). 11. Avoid contaminating T cells in the stimulator populations to prevent a false CFSE-negative population. The stimulator groups should include both antigen-specific and antigen-nonspecific cells and a control group without responder T cells. 12. When BLCLs are used as stimulators, the surface marker CD3 can be monitored to distinguish T cells and B cells. 13. The size of the microbeads should match the size of the cells for analysis. The target cells and the beads should distribute within the same FSC and SSC (forward and side scatter) window.

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14. If FITC-labeled CD107a antibody is used, add 4 μM monensin as an inhibitor of protein secretion (23). 15. It is important to titrate each multimer before use. 16. Some human CD8 monoclonal antibody clones may interfere with the binding of the multimer to the T cell receptor (26).

References 1. Guermonprez P, Valladeau J, Zitvogel L, Thery C, Amigorena S. (2002) Antigen presentation and T cell stimulation by dendritic cells. Annu Rev Immunol 20:621–667. 2. Condon C, Watkins SC, Celluzzi CM, Thompson K, Falo LDJ. (1996) DNA-based immunization by in vivo transfection of dendritic cells. Nat Med 2:1122–1128. 3. Liu M. (1998) Transfected human dendritic cells as cancer vaccines. Nat Biotechnol 16:335–336. 4. Chen X, He J, Chang L-J. (2004) Alteration of T cell immunity by lentiviral transduction of human monocyte-derived dendritic cells. Retrovirology 1:37. 5. Wang B, He J, Liu C, Chang LJ. (2006) An effective cancer vaccine modality: Lentiviral modification of dendritic cells expressing multiple cancer-specific antigens. Vaccine 24:3477–3489. 6. Ludewig B, Ehl S, Karrer U, Odermatt B, Hengartner H, Zinkernagel RM. (1998) Dendritic cells efficiently induce protective antiviral immunity. J Virol 72:3812–3818. 7. Kirk CJ, Mule JJ. (2000) Gene-modified dendritic cells for use in tumor vaccines. Hum Gene Ther 11:797–806. 8. Jenne L, Schuler G, Steinkasserer A. (2001) Viral vectors for dendritic cell-based immunotherapy. Trends Immunol 22:102–107. 9. Murphy A, Westwood JA, Teng MW, Moeller M, Darcy PK, Kershaw MH. (2005) Gene modification strategies to induce tumor immunity. Immunity 22:403–414. 10. Breckpot K, Aerts JL, Thielemans K. (2007) Lentiviral vectors for cancer immunotherapy: transforming infectious particles into therapeutics. Gene Ther 14:847–862. 11. Lipscomb MF, Masten BJ. (2002) Dendritic cells: immune regulators in health and disease. Physiol Rev 82:97–130. 12. Mosmann TR, Coffman RL. (1989) TH1 and TH2 cells: different patterns of lymphokine secretion lead to different functional properties. Annu Rev Immunol 7:145–173.

13. Kalinski P, Hilkens CM, Wierenga EA, Kapsenberg ML. (1999) T-cell priming by type-1 and type-2 polarized dendritic cells: the concept of a third signal. Immunol Today 20:561–567. 14. Chambers CA. (2001) The expanding world of co-stimulation: the two-signal model revisited. Trends Immunol 22:217–223. 15. Chirathaworn C, Kohlmeier JE, Tibbetts SA, Rumsey LM, Chan MA, Benedict SH. (2002) Stimulation through intercellular adhesion molecule-1 provides a second signal for T cell activation. J Immunol 168:5530–5537. 16. Salomon B, Bluestone JA. (1998) LFA-1 interaction with ICAM-1 and ICAM-2 regulates Th2 cytokine production. J Immunol 161:5138–5142. 17. Sabatte J, Maggini J, Nahmod K, Amaral MM, Martinez D, Salamone G, Ceballos A, et al. (2007) Interplay of pathogens, cytokines and other stress signals in the regulation of dendritic cell function. Cytokine Growth Factor Rev 18:5–17. 18. Han S, Wang B, Cotter MJ, Yang LJ, Zucali J, Moreb JS, Chang LJ. (2008) Overcoming Immune Tolerance Against Multiple Myeloma With Lentiviral Calnexin-engineered Dendritic Cells. Mol Ther 16: 269–279. 19. Chang L-J, Zaiss A-K. (2001) Methods for the preparation and use of lentivirus vectors. In: Morgan J, ed. Gene Therapy Protocols, Vol. 2, 2nd ed. Humana Press, Totowa, NJ, 303–318. 20. Chang L-J, Zaiss A-K. (2001) Self inactivating lentiviral vectors in combination with a sensitive Cre/loxP reporter system. In: Walker J, ed. Methods in Molecular Medicine. Humana Press, Totowa, NJ, 367–382. 21. Lyons AB, Parish CR. (1994) Determination of lymphocyte division by flow cytometry. J Immunol Methods 171:131–137. 22. Jedema I, van der Werff NM, Barge RM, Willemze R, Falkenburg JH. (2004) New CFSEbased assay to determine susceptibility to lysis by cytotoxic T cells of leukemic precursor cells within a heterogeneous target cell population. Blood 103:2677–2682.

Immunity of Lentiviral Vector-Modified Dendritic Cells 23. Betts MR, Brenchley JM, Price DA, De Rosa SC, Douek DC, Roederer M, Koup RA. (2003) Sensitive and viable identification of antigen-specific CD8 + T cells by a flow cytometric assay for degranulation. J Immunol Methods 281:65–78. 24. Hopken UE, Lehmann I, Droese J, Lipp M, Schuler T, Rehm A. (2005) The ratio between dendritic cells and T cells determines the outcome of their encounter: proliferation versus deletion. Eur J Immunol 35:2851–2863.

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25. Mannering SI, Morris JS, Jensen KP, Purcell AW, Honeyman MC, van Endert PM, Harrison LC. (2003) A sensitive method for detecting proliferation of rare autoantigenspecific human T cells. J Immunol Methods 283:173–183. 26. Denkberg G, Cohen CJ, Reiter Y. (2001) Critical role for CD8 in binding of MHC tetramers to TCR: CD8 antibodies block specific binding of human tumor-specific MHC-peptide tetramers to TCR. J Immunol 167:270–276.

Chapter 14 Saporin Suicide Gene Therapy Natasa Zarovni, Riccardo Vago, and Maria Serena Fabbrini Summary New genes useful in suicide gene therapy are those encoding toxins such as plant ribosome-inactivating proteins (RIPs), which can irreversibly block protein synthesis, triggering apoptotic cell death. Plasmids expressing a cytosolic saporin (SAP) gene from common soapwort (Saponaria officinalis) are generated by placing the region encoding the mature plant toxin under the control of strong viral promoters and may be placed under tumor-specific promoters. The ability of the resulting constructs to inhibit protein synthesis is tested in cultured tumor cells co-transfected with a luciferase reporter gene. SAP expression driven by the cytomegalovirus (CMV) promoter (pCI-SAP) demonstrates that only 10 ng of plasmid DNA per 1.6 × 104 B16 melanoma cells drastically reduces luciferase reporter activity to 18% of that in control cells (1). Direct intratumoral injections are performed in an aggressive melanoma model. B16 melanoma-bearing mice injected with pCI-SAP complexed with lipofectamine or N-(2,3-dioleoyloxy-1-propyl) trimethylammonium methyl sulfate (DOTAP) show a noteworthy attenuation in tumor growth, and this effect is significantly augmented by repeated administrations of the DNA complexes. Here, we describe in detail this cost-effective and safe suicide gene approach. Key words: DOTAP, melanoma, nonviral vectors, polyethylenimine (PEI), ribosome-inactivating proteins, ricin, saporin, suicide gene therapy.

1. Introduction Bacterial and plant toxins have proven to be suitable candidates in cancer suicide gene therapies (2–4). Ribosome-inactivating proteins (RIPs) are potent irreversible inhibitors of protein synthesis that are found in different plant tissues (5) and in some bacteria (6). These N-glycosidases (EC3.2.2.22) are able to specifically depurinate a single adenine present in a stem-loop


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region of large ribosomal RNA (rRNA) subunits (A4324 in rat 28S) (7) causing translation block and apoptotic cell death. In addition to this specific depurination activity, some DNA fragmentation activity has been recently also reported (8). Possibly, as little as a single toxin molecule in the cytosol could be sufficient to kill a mammalian cell. The cell killing activity of a SAP mammalian codon-optimized gene was previously reported to be comparable to that of the Herpes simplex gene encoding thymidine kinase followed by ganciclovir treatment (9). This chapter describes in detail the nonviral gene delivery protocols established to check for optimal lipid vectors/DNA combination and the techniques used for the direct intratumoral injections in vivo in a mouse melanoma model. pCI-SAP in vivo gene delivery shows the great advantage of needing only a single delivery step and makes use of an irreversible enzymatic activity that is not dependent on the cell proliferative status.

2. Materials 2.1. Devices

1. Cell culture incubator at 37°C and 5% CO2. 2. Horizontal gel electrophoresis unit with power supply. 3. Polymerase chain reaction (PCR) Thermal Cycler. 4. Enzyme-linked immunosorbent assay (ELISA) plate reader. 5. Microplate luminometer. 6. Tissue homogenizer.

2.2. Cell Culture Basic Materials and Reagents (see Note 1)

1. Corning disposable sterile plastic pipettes and flasks (Sigma, St. Louis, MO). 2. RPMI 1640 medium (Euroclone, Celbio S.p.A, Milan, Italy). 3. High-glucose Dulbecco’s Modified Eagle’s Medium (DMEM) (Sigma). 4. Fetal bovine serum (Invitrogen, Carlsbad, CA). 5. 1X Phosphate-buffered saline (PBS) (GIBCO, Invitrogen). 6. 1X Trypsin–EDTA liquid: 0.25% trypsin; 1 mM EDTA • 4Na (GIBCOTM, Invitrogen). 7. 100X L-Glutamine: 200 mM glutamine liquid (GIBCOTM, Invitrogen). 8. 100X Penicillin–streptomycin solution: 10,000 U/mL penicillin and 10 mg/mL streptomycin in 0.9% NaCl (Sigma).

Saporin Suicide Gene Therapy

2.3. Plasmids and Molecular Cloning Reagents

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1. pCISfiRsrT is used as a backbone vector for the construction of the plasmids as well as a control (pMock) plasmid in experiments performed both in vitro and in vivo. The main features of this vector as well as generation of pCI-SAP are described below (see Subheading 3.1). pCISfiRsrTLuc (pCILuc) is used as a luciferase reporter gene construct and was provided by Lucia Monaco (DIBIT, San Raffaele, Milan). All the plasmids contain the ampicillin resistance gene allowing selective growth in Luria Broth (LB) media. 2. DH5α bacterial strain (F’, hsdR17, rk−mk+, recA1, endA1) is used in all transformations. 3. Bacteria growth medium: LB medium (Sigma). 4. Ampicillin (Sigma), used at 100 µg/mL as a selection agent. 5. All restriction enzymes and buffers are obtained from New England Biolabs, Beverly, MA. 6. Pfu TurboTM DNA polymerase with reaction buffer 10X (Stratagene, La Jolla, CA). 7. dNTPs (Sigma). 8. Thin-wall PCR tubes (Applied Biosystems, Foster City, CA). 9. Synthetic oligonucleotides for PCR reactions (ours were purchased from PRIMM srl, Milan, Italy). 10. Agarose electrophoresis grade (Sigma). 11. 10 mg/mL Ethidium bromide (Sigma) 12. Loading dye: 0.25% bromphenol blue and 40% (w/v) sucrose in water. 13. Running buffer: 1X TAE: 40 mM Tris acetate and 1 mM EDTA, pH 8.0. 14. EndoFreeTM Plasmid Giga Kit (Qiagen, Hilden, Germany). 15. Limulus amebocyte lysate (LAL) assay (Bio-Whittaker, Verviers, Belgium).

2.4. Cell Lines and Mouse Strains

1. B16F1 murine melanoma cell line (ATCC, Manassas, VA). 2. Vero African Green Monkey kidney epithelium cell line (ATCC). 3. C57BL/6 8-week-old female mice (Charles River Laboratories, Calco, Italy).

2.5. Nonviral Transfections Materials and Reagents

1. PEI 25: polyethylenimine (Sigma). 2. DOTAP: (N-[1-(2,3-dioleoyloxy)] -N,N,N-trimethylammonium propane methyl-sulfate (Sigma).

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3. LipofectamineTM 2000 Reagent (Invitrogen). 4. HEPES buffer (GIBCO, Invitrogen). 5. Apyrogenic 0.9% saline solution (S.A.L.F. S.p.A, Bergamo, Italy). 6. Falcon polystyrene 15-mL conical centrifuge tubes (BD Biosciences, Franklin Lakes, NJ). 7. 1.5-mL Microcentrifuge tubes (Eppendorf, Hamburg, Germany). 8. Costar cell culture plates, 48 flat-bottom wells (Sigma, St. Louis, MO). 9. RQ1 DNaseI, 1,000 U/µL and its 10X working buffer (Promega, Madison, WI). 10. 0.5 M EDTA, pH 8 (Sigma). 11. 1000 U/mL Heparin (Sigma). 2.6. Luciferase and Bradford Assays

1. Luciferase assay system (Promega). 2. Corning flat-bottom white polystyrene 96-well plates (Sigma). 3. Bovine serum albumin (BSA) (Sigma). 4. Lysis buffer: 25 mM Tris-HCl, pH 8; 2 mM DTT; 2 mM EDTA, pH 8; 10% glycerol; and 1% Triton X-100 (see Note 2). 5. Bradford reagent (Bio-Rad, Hercules, CA) (see Note 3).

2.7. MTT Assay

1. Costar cell culture plates, 96 flat-bottom wells (Sigma). 2. 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) (Calbiochem, EMD Biosciences Inc., San Diego, CA) as stock of 5 mg/mL in PBS. 3. Dimethylsulfoxide (DMSO) (Calbiochem, EMD Biosciences Inc.).

2.8. In Vivo Studies

1. Trypan blue (Sigma). 2. 1-cc Insulin syringe and a 27- or 30-gauge needle (BD Biosciences). 3. Ethanol for sterilization (Sigma). 4. 2,2,2-Tribromethanol (Sigma). 5. 2-Methyl-2 butanol (tertiary amyl alcohol) (Sigma). 6. Burker-Turk hemocytometer (Emergo, Landsmeer, The Netherlands). 7. Electronic digital caliper.


265

3. Methods 3.1. Plasmid DNA Preparation 3.1.1. Generation of Saporin-Encoding Constructs Suitable for Expression in Mammalian Cells

The pCI-SAP plasmid is prepared by inserting sequence-3 coding for a mature saporin seed isoform (12) into the mammalian expression vector pCISfiRsrT (10, 11) to allow saporin cytosolic expression (Fig. 1). In this way, saporin is directly expressed in the same compartment in which the target ribosomes are located. 1. pCISfiRsrT has the pCI plasmid backbone (Promega). A transcription terminator has been placed downstream from the polyadenylation signal to improve transcription efficiency. Because of the presence of unique SfiI and RsrII sites in these plasmids, our constructs can be used for in vitro amplification approaches that allow multiple cassette cloning or co-delivery of different genes (10, 11). 2. The DNA saporin sequence is obtained from the pET-11d-SAP3 plasmid (12) digested with NcoI and EcoRI and protruding ends filled in with Klenow DNA polymerase. 3. pCIsfiRsrT is linearized with SmaI and dephosphorylated by the calf intestinal phosphatase (CIP). SmaI site is not reconstructed after ligation. Clones are confirmed by several analytical cuttings (Fig. 1 and see Note 4).

Fig. 1. Schematic representation of pCI-SAP expression vector. The vector backbone derives from pCI and includes the RsrII and Sfi I sites, allowing cloning of multiple cassettes in a head-to-tail orientation (pCISfiIRsrT), the CMV promoter region with the immediate-early enhancer (IE), an intron (Intr), the T7-EEV primer binding within the T7 RNA polymerase promoter region (T7), the multiple cloning site, late SV40 polyadenylation signal (poly A), a transcription terminator, the phage F1 origin of replication (f1 Ori), and the B-lactamase resistance gene (AmpR). The arrow in black indicates saporin cDNA (SAP) in 5′–3′ orientation with some of the restriction sites used for the analytical cuttings shown.

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3.1.2. Preparation of Control Plasmid Coding for the Mutant Form SAP-KQ with Reduced Catalytic Activity

To confirm that the antitumor effects specifically rely on the expression of an active SAP gene within the cells/tumors, a SAP catalytic site mutant (SAP-KQ) that has a significantly reduced RIP activity (1) is used as a control. 1. The pCI-SAPKQ construct is prepared by in vitro oligonucleotide mismatch site-directed mutagenesis on the pCI-SAP plasmid template, using Pfu Turbo polymerase. 2. The oligonucleotides used are: forward 5′ CT ATT CAA ATG ACA GCT AAG GTA GCA CAG TTT AGG TAC ATT C 3′; reverse 5′ GAA TGT ACC TAA ACT GTG CTA CCT TAG CTG TCA TTT GAA TAG 3′ (mutated nucleotides are shown in bold). In this way, two mutations are inserted at the catalytic active site 176 Glu to Lys and 179 Arg to Gln. 3. PCR reaction conditions are: hot start at 94°C for 1 min, followed by 20 cycles at 95°C for 1 min, 55°C for 30 s, and 68°C for 10 min. The template is digested with DpnI.

3.1.3. Large-Scale Preparation of Plasmid DNA from Bacteria

All of the plasmids used in vivo are prepared using an EndoFreeTM Plasmid Giga Kit (see Note 5). 1. Plasmid DNA is first mixed with 200 µL of DH5α competent cells thawed in ice. The mixture is incubated on ice for 45 min, followed by heat shock for 2 min at 42°C, and than returned to the ice for 2–3 min. 2. The cell suspension is plated on LB agar plates containing ampicillin and left overnight at 37°C in a bacterial incubator. 3. A single colony is used to inoculate a starter culture of 12 mL of LB medium containing 100 µg/mL ampicillin and the bacterial culture is incubated for approximately 8 h at 37°C with vigorous shaking (∼300 rpm). 4. The starter culture is diluted 1:500 in 2.5 L of selective LB medium, distributed in five glass vessels of 500 mL each, and grown overnight at 37°C with vigorous shaking. 5. Bacterial cells are harvested by centrifugation (6,000 rpm in a Sorvall GSA rotor). Preparation of high-copy plasmid DNA was carried out according to the protocol suggested by the manufacturer. 6. Determine the yield by measuring the 260/280-nm absorbance using a spectrophotometer. Stocks of starter bacterial culture in 50% sterile glycerol are prepared and stored at −80°C. The purity of plasmid DNA and the relative amounts of linear and covalently closed circular forms is determined by agarose gel electrophoresis (see Note 6). 7. All plasmid preparations are tested for endotoxin contamination using the Limulus amebocyte lysate (LAL) assay (see Note 5).


3.2. Cell Culture

267

The cells used to optimize gene transfer formulations (Subheading 3.3), to test the activity of saporin-encoding constructs in vitro (Subheading 3.4), and to establish subcutaneous tumors (Subheading 3.5) are treated in a sterile environment under a cell culturing hood. Cells are grown in a cell culture incubator at 37°C, 5% CO2. Routinely, cell supernatants of semiconfluent cells are collected and tested for mycoplasma contamination. • Vero cells derived from African green monkey kidney epithelium, a well-characterized cellular model used to study toxin delivery (13–15), are grown in high-glucose Dulbecco’s Modified Eagle’s Medium (DMEM) supplemented with 5% FBS, 2 mM l-glutamine, 100 U/mL penicillin, and 100 µg/ mL streptomycin. • B16-F1 cells are maintained in RPMI 1640, supplemented with 5% FBS, 2 mM L-glutamine, 100 U/mL penicillin, and 100 µg/mL streptomycin (see Note 1). • Both Vero and B16 cells are grown in T75 flasks and passaged when approaching confluence, typically twice a week. 1. B16 cells are washed with PBS and detached by incubation in PBS (10 mL/flask) for 5 min. Vero cells are washed with PBS and detached by treatment with a 0.25%-EDTA trypsin solution (1.5 mL/flask). Flick the flask gently to facilitate cell detachment. 2. The cell suspension is transferred into a 15-mL Falcon tube and cells are pelleted by centrifugation (5 min at 300 g). 3. Cells are resuspended in fresh medium and divided into culture flasks with medium (11 mL/flask) in such a way that cells are diluted 1:10 every passage.

3.3. Optimization of DNA Complex Formulation for In Vitro and In Vivo Gene Transfer

Nonviral gene delivery approaches making use of synthetic cationic polymers and lipids are easy, reproducible, and less toxic and immunogenic as compared with viral vectors, thus, offering a valid alternative to use in both basic research laboratories and clinical settings (16). Binding of polycations (i.e., PEI) or lipids (i.e., DOTAP, lipofectamine) to plasmid DNA results in DNA compaction and neutralization of negative charges in the phosphate backbone of DNA. The resulting particles are reduced in size and are endowed with neutral or slightly positive charge, being, thus, more apt to interact with negatively charged plasma membrane (see Fig. 2). We present an example of the optimization strategy used for two transfecting agents subsequently used in in vitro (PEI) and in vivo experiments (DOTAP).

268


Fig. 2. In vitro optimization of relative N/P ratios for PEI/DNA (a) and DOTAP/DNA complexes (b). The N/P ratios chosen for their best performance are 9 for PEI/DNA complexes (corresponding to a complex charge of 1.5±) and 4.8 for DOTAP/ DNA complexes (corresponding to a complex charge of 4.8±), as determined by the agarose DNase I/heparin protection assays (see Subheading 3.3.2) and the transfection efficiency assays (lower panels). The graphs show the estimated luciferase activity in relative light units (RLU) per microgram of total proteins of cell lysates after transfection with the pCILuc complexes. M, molecular weight marker; DNA, uncomplexed/naked DNA.

3.3.1. Gel Retardation Assay for Complex Formation and Stability

Charge neutralization and plasmid DNA condensation are examined by evaluating the electrophoretic migration of the DNA complexes in agarose gels (17). DNA complexes are formed at different nitrogen-to-phosphate (N/P) ratios (see Table 1). The N/P ratio is calculated considering the number of protonable nitrogen residues in the amino groups of polycation/lipid and the anionic phosphates in the phosphate group of DNA molecules (see Note 7–8). 1. For each complex formation, prepare separately solution 1 with 2 μg of plasmid DNA (pCILuc) in 20 μL of apyrogenic 0.9% saline solution, then mix solution 2 containing the appropriate amounts of each vector (see Table 1) dissolved in


269

Table 1 Amounts of 100 mM PEI-25 and 5 mg/mL DOTAP used to formulate the DNA complexes at the indicated different nitrogen to phosphate (N/P) ratios. Please see Note 7 and 8 and Subheading 2.3.1 N/P ratio

100 mM PEI (mL)

N/P ratio

5 mg/mL DOTAP (mL)

0

0

0

0

1

0.8

0.5

0.43

3.6

2.7

1

0.86

5

4

1.4

1.2

9

7.2

2.8

2.4

15

12

4.8

4

20 μL of apyrogenic 0.9% saline solution (PEI), or HEPES buffer (DOTAP). 2. Plasmid DNA and PEI or DOTAP solutions are mixed and incubated for 20 min at room temperature. 3. Mix 20 μL of each solution with 4 μL of the loading dye and load into agarose gel wells (0.8% agarose in TAE buffer, containing 0.5 μg/mL ethidium bromide). Perform electrophoresis at 100 V for 40–60 min, and visualize DNA bands by UV illumination. Electroneutralized complexes are unable to migrate toward the anode in the agarose gel (see Fig. 2). 3.3.2. Nuclease Sensitivity Assays

Compacted DNA is more resistant to nuclease degradation and thus more stable both in the bloodstream and, once internalized, in the cytosol (Fig. 2 and see Note 9). 1. To check the integrity of plasmid DNA within the complexes, mix 2 µg of complexed DNA prepared as described above (see Subheading 3.3.1) with 2 µL of 1,000 U/µL DNase I and 5 µL DNase reaction buffer, to obtain a final enzyme concentration of 1 U/µg DNA in an overall reaction volume of 50 µL. 2. Incubate reaction mix for 10 min at 37°C. 3. Add 5 µL of 0.5 M EDTA, pH 8, to each mixture to stop the digestion. 4. Add 1,000 U/mL heparin to disrupt the complexes. 5. Load 20 µL of each mixture onto a 0.8% agarose gel.

3.3.3. Transient Transfections with Luciferase Reporter Gene

Complexes prepared using the pCILuc construct at different N/P ratios are used to transfect B16F1 cells (see Table 1 and Fig. 2). Experimental steps 1–2 and 4–8 are the same as described below

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in Subheading 3.4.1. DNA complexes are prepared as previously described (Subheading 3.3.1), using 0.8 μg of pCILuc for each transfection. For quantification of luciferase expression, see Subheading 3.4.2. The best performing delivery systems chosen upon in vitro optimization are next tested for their ability to deliver the reporter gene in vivo (see Subheading 3.5 and Note 10). 3.4. Expression of SAPEncoding Constructs In Vitro

To assess the ability of toxin-encoding plasmids to induce the production of bioactive proteins, transfection studies in vitro are performed, using a luciferase reporter assay (see Fig. 3).

3.4.1. Co-Transfection Assay to Test Expression of SAP-Encoding Constructs

1. In a 48-well tissue culture plate, seed ~1.6 × 104 cells/well in 200 μL of complete medium. For every transfection condition, cells are plated at least in triplicates. 2. Incubate the cells at 37°C in a CO2 incubator until the cells are 70–80% confluent. This will usually take 16–24 h.

Fig. 3. Cytotoxicity of pCI-SAP construct in vitro. Vero (a) or (b–d) B16 cells were co-transfected with increasing amounts of PCI-SAP and with a constant amount of PCILuc (0.4 μg/well). The results are presented as mean ± SD of four points per group. (a–c) By increasing the amounts of SAP-encoding construct, the protein synthesis progressively decreased as reflected by the reduction in luciferase activity, which is estimated in RLU per microgram of protein and is expressed as a percentage of the controls using PCILuc alone. (d) The MTT assay was performed 24 h after transfection of B16 cells with PEI/ PCI-SAP complexes to detect cell death as a response to transfection. Maximum mortality is observed already with 0.02 μg of plasmid used. **p < 0.01, *** p < 0.001 and correlates with the expected transfection efficiencies of PEI-25 (Reproduced from (1)).


271

Table 2 Cells are co-transfected with increasing amounts of pCISAP mixed with complementary decreasing amounts of the empty vector (pmock), and with a constant amount of luciferase-encoding plasmid (0.4 mg/well). The total amount of DNA used for each transfection is 0.8 mg/well Sample n°

pmock (mg)

pCI-SAP (mg)

pCILuc (mg) Total (mg)

1

0.4

0

0.4

0.8

2

0.3

0.1

0.4

0.8

3

0.2

0.2

0.4

0.8

4

0.1

0.3

0.4

0.8

5

0

0.4

0.4

0.8

3. The day after, prepare separately the following solutions (the amounts are per well): Solution 1: For each transfection, dilute 0.8 μg of plasmid DNA (see Table 2) in 10 µL of a sterile, apyrogen, isotonic solution of 0.9% sodium chloride. Solution 2: For each transfection, dilute 0.22 μL of 100 mM PEI in 10 µL of a sterile, apyrogen, isotonic solution of 0.9% sodium chloride. 4. Combine the two solutions, mix gently, and incubate at room temperature for 15 min to allow complex formation. 5. Add dropwise the DNA–PEI complexes to cells, without removing the medium. 6. Centrifuge the plate at 1,500 rpm for 2 min (see Note 11). 7. Incubate the cells overnight at 37°C in a CO2 incubator. It is not necessary to replace the medium, because no toxicity is observed. 8. Assay cell extracts for luciferase gene activity 24–48 h after the start of the transfection (see Note 12). 3.4.2. Luciferase Assay and Bradford Assay

Luciferase activities in the cell lysates are measured using a Luciferase Assay System and results are standardized for total protein content, measured with a Bradford protein assay method. 1. Put the plate on ice (see Note 13). 2. Remove the medium by aspiration and wash the cells with PBS. 3. Remove the PBS, add 100 µL/well of the lysis buffer, and incubate the plate on ice for 45 min while shaking.

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4. Transfer the total cell extracts into 1.5-mL Eppendorf tubes and centrifuge them at 18,000 g for 20 min at 4°C (in a prechilled centrifuge). 5. Transfer 20 µL/well of the supernatants into a white polystyrene 96-well plate. 6. Measure the relative light units (RLUs) reading at the plate luminometer. 7. The results are expressed as a percentage of the luciferase activity estimated in the control transfections using only luciferase complementary DNA (cDNA). 8. Use a colorimetric Bradford assay to quantitate the protein content in the samples. Protein concentrations are calculated from a BSA standard curve. 9. To prepare a standard curve, dilute BSA in water to obtain solutions, 160 µL each, with a BSA content ranging from 0 to 4 µg. 10. Transfer each BSA solution (all dilutions are prepared at least in duplicates) in a 96-well culture plate. 11. Transfer 2 µL of lysates in the wells of the same plate (2 wells/ sample), containing 158 µL of water (see Note 14). 12. To all wells, add 40 µL of Bradford reagent and mix well by pipetting. 13. Read the absorbance using an ELISA plate reader at a wavelength of 570 nm. 14. Normalize the RLUs per microgram of total protein. 15. Statistical significance is assessed using a two-tailed Student’s t-test. 3.4.3. MTT Colorimetric Assay

To determine the cell death after the transfections with saporinencoding constructs, a viability MTT colorimetric assay can be performed. 1. Seed B16F1 cells, ~8 × 103 cells/well, in a 96-well plate in 200 μL of complete medium. For every transfection condition, cells are plated in triplicates. Include three control wells of medium alone to provide the blanks for absorbance readings and subtraction. 2. Transfect the cells with increasing amounts of pCI-SAP as described above (see Subheading 3.4.1) 3. Twenty-four hours after transfection, add 10 μL/well 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) and return the plate to a cell culture incubator (37°C and 5% CO2) for 3 h. During this incubation, MTT, a pale yellow substrate, is cleaved by living cells to yield dark blue formazan crystals. Intracellular punctate purple precipitates can be observed under an inverted microscope.


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4. Remove the supernatants and add 200 μL/well of dimethylsulfoxide (DMSO) to dissolve the crystals formed. Swirl gently. 5. Leave the plate in the dark for 2 h at room temperature. 6. Read the absorbance using an ELISA plate reader at a wavelength of 570 nm. The blanks should give values close to zero. 7. The amount of color produced is directly proportional to the number of viable cells. Results are expressed as a percentage of the values obtained with the untreated controls (see Note 15). 3.5. In Vivo Studies

All studies on animals must be first approved and relevant information about any required permit, license, and authorization concerning the use of these animals should be obtained in your country. Our studies were approved by the Ethical Committee of the San Raffaele H Scientific Institute and were performed according to the prescribed guidelines.

3.5.1. Preparation and Inoculation of Melanoma B16F1 Cells Into Mice

B16F1 cells used to establish subcutaneous tumors are handled as described in Subheading 3.2. 1. Cells are kept in culture in 75-cm2 flasks for at least two passages and tested to exclude any mycoplasma or bacterial contamination before they are used for inoculum. 2. When the cells reach 70–80% confluence (see Note 16), the medium is removed by aspiration and the B16 cells are washed with PBS to remove dead and detached cells. 3. Cells are detached by incubation in PBS (10 mL/T75 flask) for 5 min. Flick the flask gently to facilitate cell detachment. 4. Transfer the cell suspension into a 15-mL Falcon tube and pellet the cells by centrifugation (5 min, 300 g). The cell pellet is washed by resuspending the cells in 5 mL of 0.9% NaCl solution and centrifugation at 1,500 rpm for 5 min. 5. The pellet obtained is resuspended in 2 mL of 0.9% NaCl and the cells are counted in a hemocytometer using Trypan blue staining to exclude dead cells (see Note 17). 6. The final cell suspension is prepared such that a final injection volume of 150 μL contains 5 × 104 cells, which are required for tumor implantation. The total amount of cells to prepare depends on the number of animals to be treated. Cells have to be prepared in excess for at least one or two extra animals. The cell suspension is kept in ice, providing the starting material for immediate subcutaneous (s.c.) injection into mice. 7. Prepare 8-week-old C57BL/6 syngenic female mice (see Note 18 and 19). 8. Clean and sterilize the inoculation area of the mice with ethanol. 9. Mix cells and draw the cells into a syringe without a needle (see Note 20). A 1-cc syringe and a 27- or 30-gauge needle are used.

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10. Inject cells (150 μL/5 × 104 cells) s.c. into the lower left flank of the mice (see Note 21). 11. Monitor mice every 2–3 days for tumor growth. Treatments can be started about 10 days after cell inoculation, when most of the tumors have reached an average diameter of 5 mm. 3.5.2. Intratumoral Injection of DNA Complexes in B16 Melanoma-Bearing Mice

Direct intratumoral DNA administration is a reliable and reproducible gene delivery system that combines simplicity and efficient interstitial penetration of the injected agents with local delivery safety benefits (see Note 22). A higher intracellular uptake of DNA complexed to cationic liposomes is achieved, thus, improving antitumor efficacy of therapeutic gene transfer (18).

Optimization of In Vivo Gene Transfer Using the Luciferase Gene

Screening of several cationic polymers and lipids using luciferase (pCILuc) as a reporter gene is performed to determine the best formulation for intratumoral gene delivery (see Fig. 4). 1. When s.c. tumors reach about 5 mm in diameter, mice are randomly divided into groups of five per cage. 2. DNA complexes are prepared immediately before injection. The amount of plasmid DNA injected per animal is 30 μg in an injection volume of 112 µL (see Note 23 and 24). 3. For each group of five mice to be treated with a given formulation, DNA complex solution is prepared in excess for six animals. 4. To prepare DNA complexes, we test several nonviral cationic carriers such as DOTAP and DOTAP derivates (DOTAP-DOPE, DOTAP-cholesterol), DMRI, lipofectin, Lipofectamine, and PEI (22 and 25 K). Figure 4 shows optimal N/P ratios chosen upon testing a limited N/P ratio range for each of these reagents. For N/P ratio calculations, see Note 8. DNA complexes are typically prepared as follows, separately make two solutions: Solution 1: Dilute 180 μg plasmid DNA in apyrogenic, isotonic solution of 0.9% sodium chloride or in 20 mM HEPES buffer (for DOTAP-containing complexes) (see Note 25). Solution 2: Dilute the proper amount of cationic carrier corresponding to the desired N/P ratio in apyrogenic, isotonic solution of 0.9% sodium chloride or in 20 mM HEPES buffer (for DOTAP-containing complexes) (see Note 25). Combine the two solutions, mix gently, and incubate at room temperature for 15 min to allow the formation of the DNA complexes (see Note 26). 5. Mice that are to be injected must be anesthetized by administering tribromoethanol intraperitoneally (i.p); the i.p. dose used is approximately 0.2 mL/10 g (240 mg/kg) body weight (see Note 27 and 28).


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Fig. 4. In vivo comparison of the efficiency of different nonviral vectors in luciferase gene transfer upon direct intratumoral injection. DNA complexes are prepared at the following N/P equivalents: ratio 4.8 for DOTAP and DOTAP/DOPE, 9 for PEI22 and PEI25, 3.3 for DMRI, 10 for lipofectin, and 6.7 for Lipofectamine. Luciferase expression in tumor homogenates after 24 h is expressed in relative light unit (RLU) per microgram of protein. The results are presented as mean ± SE of five animals per group. Naked DNA or lipoplexes were dissolved in saline (except for DOTAP complexes, which were dissolved in 20 mM HEPES) (Reproduced from (1)).

6. After administering the anesthetic, put the mouse back into the box from which it came (see Note 29). The time required for the mice to fail to respond to digital tail or feet pad pinch, and thus to be fully anesthetized, is about 2–3 min. The anesthesia lasts for about 20 min. 7. DNA complexes are injected manually via insulin syringes into the center of established tumors using one or two injection spots depending on the size and shape of the tumor (see Note 30). 8. The animals undergoing anesthesia and treatment are allowed to recover undisturbed in a warmed (see Note 31) and dry cage with water regularly supplied. The animals typically fully recover after 30–40 min. The cages are put in place only when animals are alert, mobile, and breathing normally. 9. Twenty-four hours after the injection, animals are killed in a CO2 chamber. Tumors are harvested and put into 2-mL Eppendorf tubes. Immediately, 1.5 mL of cold lysis buffer is added and tubes are put on ice. 10. Tumor extracts are prepared using a tissue homogenizer by mechanical homogenization of whole tumors in lysis buffer followed by centrifugation in a bench centrifuge at 18,000 g for 15 min at 4°C to remove insoluble particles. Supernatants are collected and frozen by immersion in liquid nitrogen and kept at −80°C.

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11. Luciferase activity in tumor lysates is measured using the Luciferase Assay System (see Note 32). The results are standardized for protein content, and measured using the Bradford protein assay method (see Subheading 3.4.2, steps 5–15). 12. Statistical significance of observed data is assessed using an unpaired-sample Student’s t-test. All data are considered statistically significant at p < 0.05. Injection of SAP-Encoding Constructs in B16F1 Melanomas

The potential of pCI-SAP to promote tumor cell killing in vivo can be evaluated by direct intratumoral injection of the plasmid complexed with either DOTAP (see Fig. 5) or Lipofectamine selected as the most efficient vectors for delivering the luciferase reporter gene. 1. When s.c. tumors reach about 5 mm in diameter, mice are randomly divided into groups of five per cage. 2. To accurately keep records of tumor growth and the state of the animal, mice are weighed and marked for identification purpose (see Note 33).

Fig. 5. Effect on tumor growth of injection of pCI (pmock), pCI-SAP, or pCI-SAPKQ (inactive site mutant) complexed with DOTAP. Subcutaneous B16 tumors with 5 mm of diameter were injected with 50 μg saporin-encoding constructs. The time of injection is indicated by the arrow. The results are presented as mean ± SE of five animals per group. *p < 0.05, **p < 0.01 relative to none; *p < 0.05 relative to mock (Reproduced from (1)).


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3. DNA complexes are prepared immediately before injection. The amount of injected plasmid DNA per animal is 50 μg in the injection volume of 200 µL. 4. For each group of five mice to be treated with a given formulation, DNA complex solution is prepared in excess for six animals. 5. Prepare DNA complexes (separately make the following solutions): Solution 1: Dilute 300 μg of plasmid DNA (see Note 34) in 20 mM HEPES buffer to a final volume of 600 µL (for DOTAP/ DNA complexes) or in apyrogenic, isotonic solution of 0.9% sodium chloride to obtain a final volume of 200 µL (for Lipofectamine/DNA complexes) (see Note 25). Solution 2: Either 600 µL of 5 mg/mL DOTAP (for DOTAP/ DNA complexes) or 1,000 µL of 2 mg/mL Lipofectamine (for Lipofectamine/DNA complexes). Combine the two solutions, mix gently, and incubate at room temperature for 15 min to allow the formation of the DNA complexes (see Note 26). 6. DNA complexes are injected as described in steps 5–8 of Subheading 3.5.2.1. For selected groups, the injection can be repeated after 3 and 5 days. For repeated administrations, DOTAP-based formulations were used because of comparable efficiencies but lower costs with respect to Lipofectamine. 7. Tumor growth is evaluated by daily measures of the main diameters with an electronic digital caliper and the volume is calculated as described (see Note 35). Animals are also monitored for weight loss or for any other sign of systemic pathology or local toxicity (e.g., alterations in animal behavior, signs of pain, ulceration at or near the injection site). 8. Mice are killed in a CO2 chamber when the tumor volume reaches 1,000 mm3. 9. Statistical significance of the observed data must be assessed using unpaired-sample Student’s t-test or ANOVA one-way posttest statistical analysis (GraphPad Prism software) when three groups (treated, mock, and controls) can be compared. All data are considered statistically significant at p < 0.05.

4. Notes 1. Culture medium is usually prepared in bulk, stored at 4°C, and aliquoted into culture flasks just before cultures are set up. All culture mediums are filter sterilized using 0.2-μm

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filters. Before usage, the medium, as well as the PBS and EDTA–trypsin solution, are allowed to warm in the water bath at 37°C or under the hood for 30 min. 2. Lysis buffer is usually prepared as a 5x stock solution and stored at 4°C. 3. Bradford reagent can also be prepared in the laboratory by dissolving 100 mg Coomassie Brilliant Blue G-250 in 50 mL of 95% ethanol and adding 100 mL of 85% (w/v) phosphoric acid. Dilute to 1 L when the dye has completely dissolved, and filter through Whatman #1 paper just before use. “Homemade” reagents work quite well but are usually not as sensitive as the Bio-Rad commercial product. 4. We tested also pSfiSV19-SAP in which the saporin coding sequence was placed under the simian virus 40 (SV40) promoter, but its activity in the in vitro assay was weaker than that of pCI-SAP and was consequently not used for in vivo experiments (1). 5. Bacterial endotoxin markedly decreases the efficiency of transfection both in vitro and in vivo. The endotoxin content should be less than 0.01 EU/μg of plasmid DNA. 6. The plasmid DNA used in transfection studies is typically >90% in circular form. 7. The MW of an average nucleotide is 330 g/mol: 2 µg of DNA contain 6 × 10−9 mol. The MW of 1 monomer of PEI25K is 43 Da and the stock solution is 100 mM. The MW of DOTAP is 698.5 g/mol and the stock solution is 10 mg/mL, which is 14.3 mM. Appropriate amounts of vector for each N/P ratio are calculated as follows: PEI25(μL)=

DOTAP(μL)=

N/P ratio × 6 × 10-9 mol × 43 × 10 4 μg/mol × 10 4 100 × 10-3 mol

N/P ratio × 6 × 10-9 mol × 698.5 × 10 4 μg/mol × 10 4 14.3 × 10-3 mol

8. For Lipofectamine/DNA complexes, the DNA weight-to-lipid volume w:v ratio proposed by the manufacturer is 1–2 μg to 2–25 μL. We use a w:v ratio of 6.7, testing also 5 and 10. 9. The stability of complexes in the presence of serum is addressed by performing the same assay after incubation of each complex formulation for 6 h in complete culture medium as well as in 50% normal mouse serum. 10. The efficiency of transfections observed in vitro cannot be extrapolated to what is expected to be obtained using the


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same polyplexes/lipoplexes in vivo. Thus, after the optimization in a transient transfection system, the same complex formulation needs to be tested for gene transfer efficiency upon intratumoral injection. The final choice of which type of vector as well as the vector-to-DNA ratio is a result of a screening procedure assaying different DNA complexes and complex formulations, using reporter gene activity as a read out of the transfection efficiencies (see Subheading 3.5.2.1). 11. Binding of DNA to PEI typically results in formation of large electroneutralized complexes. To improve the contact/interaction between complexes and cells, and thus their transfection efficiency, plates containing cells treated with DNA–PEI complexes are centrifuged for 2 min at 300 g after adding the DNA complexes to the wells. 12. The effects of protein synthesis inhibition were found to be dependent on both cell type and promoter driving SAP expression (1). 13. From this step onward, keep cells and total cell/tissue extract on ice. Luciferase is a thermo-sensitive protein. To avoid degradation and fully preserve its enzymatic activity, enabling reliable sample comparisons, if not processed immediately, the extracts can be kept at −20°C, but repeated cycles of thawing/freezing must be avoided. 14. If the sample readings are much higher with respect to those of standards, further dilute the lysates. To calculate total protein concentrations from the BSA standard curve, sample protein content should fall within a linear range of the standard curve. 15. A weaker dose-dependent response, ranging from 42% to 56%, was observed by MTT staining of cells treated with increasing amount of SAP-encoding plasmids 24 h after transfection (see Fig. 3d). These results correlate with efficiency of PEI-mediated transfection that was been previously estimated to be around 50%. 16. The number of flasks required depends on the total number of mice to be used in the experiment, keeping in mind that typically more than 107 B16F1 cells are recovered from one 75-cm2 flask. To give rise to s.c. tumors, it is important that the cells are harvested during exponential cell growth so that when they are injected the tumor growth occurs uniformly in all animals with the expected rate. 17. Dilute Trypan blue at 0.8 mM in PBS. Store at room temperature. The solution is stable for 1 month. Mix cells 1:1 with Trypan blue solution. Viable cells exclude Trypan blue, while dead cells stain blue because of Trypan blue uptake.

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18. Animals experience stress as a result of shipping. Allow a period of 3–5 days acclimatization after mice have arrived. It is also recommended that the animals are used to your handling before experimental manipulation. Handle the animal gently but firmly on a regular basis in a nonthreatening situation, e.g., weighing, changing water and food supply. 19. Typically, at least five animals per group are needed to test each treatment condition to achieve a minimum number to allow subsequent statistical analysis. Control groups must include untreated animals as well as mock treated and/or, in our case, those injected with a KQ mutated saporin gene. 20. Using a needle causes a strong, negative pressure that can cause cell damage and lysis. 21. Tumor cell injection is a rapid procedure that does not cause major distress or pain to an animal and thus it is performed without analgesia/anesthesia. The same syringe and needle can be use to inject multiple animals. 22. For clinically oriented research, direct tumor injection might be a method of choice because it represents a simple means of achieving a clinical response in cancer patients. This delivery approach is, in general, applicable to accessible solid tumors, thus limiting the field of action to a few malignancies that, however, are often resistant to conventional therapies (e.g., melanomas, head and neck tumors, mesotheliomas). 23. Besides carrier type and N/P ratio, other transfection parameters that affect physical characteristics and performance of DNA complexes, such as the medium for complex formation (HEPES vs saline vs glucose solution) and plasmid DNA doses (30–80 μg) were examined. The chosen amount of DNA injected per animal is 30 μg for luciferase gene injections and 50 μg for injections of pCI-SAP. 24. In this way, the DNA concentration in the injection mix is 250 ng/μL. If the amount of DNA used to form lipoplexes, in the conditions described here (solvent, incubation time, lipid type, N/P ratio), exceeds this average concentration, DNA complexes will tend to precipitate and this markedly decreases their transfection efficiency. 25. Volumes are adjusted taken into consideration that the final injection mix volume (solution 1 + solution 2) for one treatment/group corresponds to six animals and is 672 µL for luciferase gene injection and 1,200 µL for pCI-SAP-injected animals. 26. DNA complexes are typically prepared in Eppendorf tubes under the hood in a tissue culture room, sealed with Parafilm, and immediately taken to an animal facility.


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27. Intratumoral injection is not painful but causes significant restraint to an animal and thus requires general anesthesia. Anesthetic is formulated according to (19). Ingredients: 2.5 g 2,2,2-tribromethanol 5 mL 2-methyl-2 butanol (tertiaty amyl alcohol) 200 mL distilled water Add tribromethanol to butanol and dissolve by heating (50°C) and stirring. Add distilled water and continue to stir until butanol is completely dispersed. Store in aliquots in the dark at 4°C for up to 4 months. Warm to 37°C and shake before use. 28. Intraperitoneal injection is a commonly used method for small rodents because they do not have readily accessible veins. Intraperitoneal injections should be given in a lower right or lower left quadrant of the abdomen because vital organs are absent in this area. To prevent injection of the intestine, only the tip of the needle should penetrate the abdomen wall. 29. Mice will be more relaxed when placed in a familiar environment and the anesthetic will act more quickly than it would on a distressed mouse. 30. Resistance of the tumor tissue to the influx of the fluid and/ or cells often results in low quantities of material penetrating and remaining in the tumor tissue. To ensure that more material is introduced into the tumor, injection volumes should be minimized, and the injection performed very slowly (using a pump vs manual injection can be considered) keeping the needle inside for at least 20 extra seconds after the injection is finished to prevent backflow along the needle tract. 31. Use of heat lamps requires close observation because anesthetized animals are unable to escape from it, so prolonged exposure can cause hyperthermia and burns. 32. For luciferase measurements, in vivo, also bioluminescence molecular imaging can be used but it, however, requires expensive equipment, such as a Xenogen IVIS imaging system. 33. When we work with limited number of animals (five in each box) and animals are kept for a relatively short time, such as in this case, it is enough to mark their tails with a waterproof pen. 34. The absolute amount per animal of different plasmids used to treat selected groups of animals, namely pCI-SAP, pCISfiRsrT (pmock), and pCI-SAPKQ (see Subheading 2.3 and 3.1) has to be kept the same as do the experimental conditions for DNA complex formation. 35. Different formulas have been proposed to calculate tumor volume from its main diameters, based on the comparison of the calculated volume with an actual volume estimated from the extirpated tumor and tumor weight. We measure

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the relative growth rate of tumors that are large enough to enable us to measure reliably all of the three diameters— width, length, and height.

Acknowledgments The authors are deeply indebted to Lucia Monaco for the development of the nonviral vector strategy, the pCISfiLuc reporter gene construct, and especially for her advice and continuous support. This work has been partially supported by the Italian Cancer Research Association (AIRC, grant 131/01) and by IBBA-CNR, Milano. References 1. Zarovni, N., Vago, R., Soldà, T., Monaco, L., and Fabbrini, M. S. (2007) Saporin as a novel suicide gene in anticancer gene therapy. Cancer Gene Ther. 14, 165–73. 2. Martin, V., Cortes, M. L., de Felipe, P., Farsetti, A., Calcaterra, N. B., and Izquierdo, M. (2000) Cancer gene therapy by thyroid hormone-mediated expression of toxin genes. Cancer Res 60, 3218–24. 3. Yerushalmi, N., Brinkmann, U., Brinkmann, E., Pai, L., and Pastan, I. (2000) Attenuating the growth of tumors by intratumoral administration of DNA encoding Pseudomonas exotoxin via cationic liposomes. Cancer Gene Ther 7, 91–6. 4. Anderson, D. G., Peng, W., Akinc, A., Hossain, N., Kohn, A., Padera, R., Langer, R., and Sawicki, J. A. (2004) A polymer library approach to suicide gene therapy for cancer Proc Natl Acad Sci USA 101, 16028–33. 5. Hartley, M. and Lord, J. (2004) Cytotoxic ribosome-inactivating lectins from plants. Biochim Biophys Acta 1701, 1–14. 6. Sandvig, K. and van Deurs, B. (2002) Transport of protein toxins into cells: pathways used by ricin, cholera toxin and Shiga toxin. FEBS Lett 529, 49–53. 7. Endo, Y., Mitsui, K., Motizuki, M., and Tsurugi, K. (1987) The mechanism of action of ricin and related toxic lectins on eukaryotic ribosomes. The site and the characteristics of the modification in 28 S ribosomal RNA caused by the toxins. J Biol Chem 262, 5908–12. 8. Bagga, S., Seth, D., and Batra, J. K. (2003) The cytotoxic activity of ribosome-inactivating

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protein saporin-6 is attributed to its rRNA N-glycosidase and internucleosomal DNA fragmentation activities. J Biol Chem 278, 4813–20. Hoganson, D. K., Chandler, L. A., Fleurbaaij, G. A., Ying, W., Black, M. E., Doukas, J., Pierce, G. F., Baird, A., and Sosnowski, B. A. (1998) Targeted delivery of DNA encoding cytotoxic proteins through high-affinity fibroblast growth factor receptors. Hum Gene Ther 9, 2565–75. Goergen, J. L. and Monaco, L. (2004) Generation of high-recombinant-protein-producing Chinese hamster ovary (CHO) cells. Methods Mol Biol 267, 477–83. Monaco, L., Tagliabue, R., Soria, M. R., and Uhlen, M. (1994) An in vitro amplification approach for the expression of recombinant proteins in mammalian cells. Biotechnol Appl Biochem 20, 157–71. Fabbrini, M. S., Rappocciolo, E., Carpani, D., Solinas, M., Valsasina, B., Breme, U., Cavallaro, U., Nykjaer, A., Rovida, E., Legname, G., and Soria, M. R. (1997) Characterization of a saporin isoform with lower ribosome-inhibiting activity. Biochem J 322, 719–27. Majoul, I. V., Bastiaens, P. I., and Soling, H. D. Transport of an External Lys-Asp-Glu-Leu (KDEL) (1996) Protein from the Plasma Membrane to the Endoplasmic ReticulumStudies with Cholera Toxin in Vero Cells. J Cell Biol 133, 777–89. Wales, R., Roberts, L. M., and Lord, J. M. (1993) Addition of an endoplasmic reticulum retrieval sequence to ricin A chain significantly

Saporin Suicide Gene Therapy increases its cytotoxicity to mammalian cells. J Biol Chem 268, 23986–90. 15. Vago, R., Marsden, C. J., Lord, J. M., Ippoliti, R., Flavell, D. J., Flavell, S. U., Ceriotti, A., and Fabbrini, M. S. (2005) Saporin and ricin A chain follow different intracellular routes to enter the cytosol of intoxicated cells FEBS J 272, 4983–95. 16. Zhang, S., Xu, Y., Wang, B., Qiao, W., Liu, D., and Li, Z. (2004) Cationic compounds used in lipoplexes and polyplexes for gene delivery. J Control Rel 100, 165–80.

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17. Gebhart, C. L. and Kabanov, A. V. (2001). Evaluation of polyplexes as gene transfer agents. J Control Rel 73, 401–16. 18. Missol, E., Sochanik, A., and Szala, S. (1995) Introduction of murine IL-4 gene into B16 (F10) melanoma tumors by direct gene transfer with DNA liposome complexes. Cancer Lett 97, 189–93. 19. Papaioannou, V. and Fox, J. (1993) Efficacy of tribromethanol anesthesia in mice laboratory. Anim Sci 43, 189–92.

Chapter 15 Using In Vivo Biopanning for the Development of Radiation-Guided Drug Delivery Systems Jerry J. Jaboin, Zhaozhong Han, and Dennis E. Hallahan Summary This chapter illustrates our protocol for in vivo biopanning using T7 bacteriophage libraries for the purpose of selecting recombinant peptides for the tumor-specific delivery of radiosensitizers to radiationinducible antigens within tumor neovasculature. Our goal is to discover peptides binding within tumor vascular endothelium of irradiated tumors. We have previously demonstrated that tumor irradiation increases the spectrum of antigenic targets for drug delivery. To identify candidate peptides with the ability to bind radiation-induced antigens, we inject the phage peptide library intravenously into mice bearing irradiated GL261 and Lewis lung carcinoma (LLC) hind limb tumors. Phage are recovered from excised tumors, amplified, and readministered to mouse-bearing tumors for six total rounds. At least 50 bacterial colonies are selected from each of the tumor types, and prioritized. This prioritization is based on their relative concentrations in tumor versus normal tissues, and then assessment of dominant phage present in both tumor types. These phage are amplified, and the gene sequences determined to deduce the recombinant peptide product. Further prioritization is performed by fluorescence labeling of the selected phage, and injection into irradiated and mock-irradiated tumor-bearing mice for evaluation of in vivo targeting of the candidate phage/peptides. Key words: Biopanning, drug delivery system, mice, phage display, radiosensitization, radiotherapy, treatment-induced biomarker, T7 bacteriophage, tumor cells.

1. Introduction 1.1. Rationale

Concurrent chemoradiotherapy plays a central role in the treatment and cure of various cancer types. Multiple dose escalation studies have demonstrated that increased radiation dose leads to improved local control and survival rates. However, the greatest limitation to dose intensification efforts has been toxicity to


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healthy tissues. In recent years, there has been a strong push to develop strategies to sensitize cancer cells to radiation while minimizing the effects on surrounding normal tissues. In this chapter, we discuss one of our approaches. We have exploited the ability of irradiation to create neoantigens within tumors and their neovasculature. Targeting these radiation-induced proteins with cytotoxic therapies has been able to markedly (>100-fold) improve bioavailability of our drug delivery vectors in animal models (1–3). We propose that exploitation of this technology in cancer patients will greatly improve the bioavailability of cytotoxic drugs and/or radioisotopes, as well as provide a mechanism for the imaging of tumor response. Furthermore, we can accomplish this while minimizing systemic toxicities. 1.2. Tumor Cell Sensitization Strategies

Multiple strategies are being used for the radiosensitization of cancer cells. Some strategies use the targeting of genes (and their protein products) involved in radiation resistance. This has resulted in the successful clinical development of several radiosensitizing compounds. One of the best studied of this group is the family of epidermal growth factor receptor (EGFR) antagonists, which have been developed to reverse the biologic resistance conferred by EGFR overexpression in various tumor types. One agent of this group, cetuximab (Erbitux), received US Food and Drug Administration (FDA) approval in 2006 for concurrent treatment with radiotherapy in advanced head and neck squamous cell carcinomas. This followed a phase III clinical trial demonstrating its ability to improve locoregional control and survival rates over radiation alone (Bonner et al., NEJM 2006). Clinical studies are ongoing with this agent, and others like erlotinib (Tarceva) in multiple EGFR-overexpressing tumor types in a strategy to improve radiosensitivity (4–6). Histone deacetylase (HDAC) inhibitors have also been demonstrated to have in vivo and in vitro monotherapy and radiosensitizing activity against a variety of tumor histologies (7–12). The mechanisms are actively being studied, and clinical trials are underway. There is one open clinical trial for combination with radiotherapy. This is the NCI06-C-0112 trial of valproic acid (VA) followed by daily VA/ Temodar/radiotherapy (American Cancer Society: http://www. cancer.org). Investigators have also developed therapeutic transgene techniques, in which mutant or deleted copies of genes responsible for radiation-induced cell death are replaced with normal gene copies through infection or transfection techniques. An example of this is gene correction with p53. This gene is known to be mutated in at least 60% of solid tumors. Multiple in vitro and animal model studies have proven the principle that gene transfer into affected tumors can restore the apoptotic machinery, and make cells more radiosensitive (13–15). There have been numerous clinical stud-

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ies (phase II/III) demonstrating efficacy of adenoviral p53 gene therapy as stand-alone therapy, chemosensitizing therapy and radiation-sensitizing therapy (16–19). There was a phase II study of 19 patients with nonmetastatic unresectable non-small cell lung carcinoma (NSCLC) at MD Anderson who were not candidates for concurrent chemoradiotherapy (20). These patients received 60 Gy of irradiation with three concurrent intratumoral injections of INGN 201. The treatment was well tolerated, no viable tumor remained after 3 months, and biopsies demonstrated that gene expression was consistent with successful p53 gene transfer. Some radiosensitizing gene therapy techniques involve the exploitation of radiation-sensitizing enzymes. For example, the radiation-inducible nitric oxide synthase enzyme generates toxic metabolites locally from endogenous precursors. In vitro studies and mouse models have effectively demonstrated a therapeutic advantage (21–23). A variant of this method is “suicide gene” technology, in which nontoxic prodrugs are activated to toxic agents within irradiated target tissues of patients. An example of this is the herpes simplex virus thymidine kinase vector, which is introduced into the tumor or tumor bed by gene transfer (24). Ganciclovir is subsequently injected into the patient with the local production of toxic metabolites. Clinical studies (phase I/II/III) in malignant gliomas using this technique have been promising (25–27). A Phase I/II study in high-risk and recurrent prostate cancer after definitive radiotherapy has been undertaken (28). There were acceptable acute toxicities in this study, and 5-year follow-up has suggested clinical efficacy with a lengthening of the prostate-specific antigen (PSA) doubling time and a projected delay in salvage androgen suppression therapy (28). Virotherapeutic agents for sensitization of radiation are another target. The best studied of these are conditionally replicating adenoviruses, which selectively replicate in cancer cells. The ability of these agents to act as radiosensitizers has been demonstrated in vitro and in mouse models (29, 30). Phase I/II studies have been performed with advanced prostate cancer patients (recurrent and metastatic disease) with various PSAselective oncolytic adenoviruses, with evidence of safety and clinical activity (31, 32). 1.3. Radiation Targeting of Tumor Cell and Microenvironment

Irradiation of tumor tissues is known to induce molecular changes in tumor blood vessels (33). These induced cell surface proteins can be recognized and targeted using peptide conjugates (34). This technique has several practical advantages over other delivery techniques. Intratumoral or resection bed injections (used in virotherapeutics, therapeutic transgene, and suicide gene technologies) have a theoretical advantage in that there is local tumor therapy without significant involvement of other organ sites. However, gene expression and drug delivery is often extremely

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limited, with minimal penetration beyond the site of administration (35). This limited biodistribution has been proposed to be related to increased interstitial pressure (Baxter, Boucher), which can reduce the diffusion into the tumor microenvironment. Intravascular administration of gene constructs and proteins (EGFR antagonists, HDAC inhibitors, vascular endothelial growth factor [VEGF] antagonists) has the advantage of being able to more easily bind vascular targets. However, expression can occur in nontarget organs. In previous studies of irradiated tumor microvasculature, we identified receptors and adhesion molecules for site-specific drug delivery (34–37). Although clinically useful, screening with phage-displayed peptide libraries would have an advantage in detecting posttranslationally modified vascular proteins (38, 39). Using these libraries, we have discovered a number of radiationinducible surface proteins in the microvasculature (3, 40). 1.4. Phage Display Technology

Phage display is a powerful research tool for identifying proteins of interest. A small peptide sequence is displayed on the phage surface (capsid). This sequence is encoded by the recombinant DNA genome of the individual phage. The bacteriophage DNA can be easily amplified by in vitro infection of bacteria. We used the T7 bacteriophage in our experiments. It is an icosahedral phage with a capsid shell composed of 415 copies of the T7 capsid protein, which are arranged as 60 hexamers on the shell faces and 11 pentamers at the shell vertices. This system can display peptides as large as 50 amino acids at high copy number. The capsid protein is encoded by gene 10, and the full gene encodes two proteins (10A and 10B), because of a translational frameshift at aa341 of 10A. Functional capsids can be composed entirely of the 10A or 10B proteins, or combinations of the two. However, the region of the capsid protein unique to the 10B protein is on the surface of the phage, and is used for phage display. Gene 10 encoding the capsid protein is cloned with a series of multiple cloning sites at the C terminus of the 10B protein. The natural translational frameshift site within the capsid gene has been removed so that only a single form of the capsid protein is made. This results in a total of 415 peptides expressed on the surface of the phage. The amplified phage library is injected into the host, and recovered by harvesting of the target tissue (tumor and tumor vasculature). The phage is purified, amplified, and can either be enriched by repeat injection into the host or be sequenced to determine putative amino acid sequences (41, 42).

1.5. Drug Delivery Systems

We are evaluating multiple drug delivery vehicles for conjugation to our selected phage, peptide, and streptavidin-labeled peptides. These include, but are not limited to, adenoviral gene delivery constructs, peptide–liposomal conjugates, nanoparticles, and


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minimal binding proteins (e.g., ScFv). Each construct has been approved for clinical application. Above, we discussed the use of adenoviral gene delivery constructs (p53 transgene therapy). Our laboratory has also demonstrated a use for this system when we constructed adenoviral vectors (Fig. 1) with a modification of the fiber coat proteins that increased the tropism of those vectors to malignant gliomas (43). This system is useful for several reasons. Adenovirus is ubiquitous. Adenovirus has a low incidence of pathogenicity, and humans have a high rate of exposure to the most commonly used serotypes (2, 3). When adenovirus does result in pathogenicity, the symptoms are typically self-limited and not more than those of a “cold.” Adenovirus can infect a broad range of human cell types, and can allow for transduction of relatively large DNA segments. In addition, their viral genome does not have a high rate of gene arrangement, so inserted genes tend to have good stability through successive rounds of viral replication. Liposomal conjugates have also been clinically useful. Doxil, a liposomal form of doxorubicin, has been approved for use in multiple myeloma and ovarian cancer. With the advent of Stealth liposomes, or liposomes with PEGylated membrane surfaces, there has been a renewed interest in the use of liposomes as drug delivery vectors. These Stealth liposomes have the advantage of an improved half-life, and an ability to avoid detection by the reticuloendothelial system. Liposomes also seem to have a natural ability to target cancer. Recently, there have been efforts to also

~50 μm A) Hexon

B) Penton Base C) Fiber D) Knob

Fig. 1. The adenoviral capsid coat is composed of: (a) 240 hexons, (b) 12 pentameric penton bases, which are associated with (c) trimeric fibers with a (d) glandular knob responsible for interaction with the target cellular receptor. In Staba et al. (43), the fiber coat region was modified to enhance the tropism of the adenovirus for malignant glioma tumor xenografts. The adenoviral construct is approximately 50 εm in size (see Color Plates).

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add biological moieties to the surface of these molecules, in an attempt to improve the ability to target to a specific site. These ligands include monoclonal antibodies (immunoliposomes), vitamins, and specific antigens. These properties lend themselves naturally to our system. We can decorate liposomes using our selected phage (or peptide), and deliver radiosensitizing drug therapy to tumor cells. The liposomes can carry hydrophilic drugs (e.g., doxorubicin, daunorubicin) in their core, or hydrophobic drugs can be dissolved into the lipid membrane. Then the delivery of the drug can occur by endocytosis with direct fusion to the target cells, or neutralization of charged aqueous drugs and resultant diffusion of the activated drug. In Geng et al. (1), we proved this principle, by decorating Stealth liposomes with wheat germ agglutinin (WGA) (Fig. 2). WGA is known to bind to inflamed endothelial cells. We demonstrated that the WGA– liposomal conjugate was not only able to target irradiated tumor vasculature, but when the liposomes were loaded with cisplatin, the drug delivery system resulted in tumor growth delay in LLC hind limb mouse grafts. Nanoparticles of varied types have also been in development for drug delivery. Abraxane, a formulation of Paclitaxel albuminbound particles, has been approved for use in metastatic breast cancer. This formulation had nearly double the overall response rate (33% vs 19%) compared with solvent-based paclitaxel in a randomized phase III trial (Gradishar et al., JCO 2005). Albumin nanoparticles have multiple technological advantages, including high stability, high carrier capacity, the ability to incorporate both hydrophilic and hydrophobic compounds, and multiple routes of administration (e.g., oral, intravenous, inhalational). We previously used this system in Hallahan et al. (40), demonstrating

Targeting Particle PEGylated Surface ~100-200 nm

Hydrophilic Drug Lipid Bilayer

Fig. 2. This is an illustration of a Stealth liposome. The liposome contains a single lipid bilayer membrane, and the surface is coated with a PEG polymer layer. The aqueous internal compartment contains the cytotoxic agent for delivery. Addition of a peptide, antibody, or phage may be made to the surface of the molecule, allowing for tumor targeting. Liposomal molecules are typically in the 100- to 200-nm size range.


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that fibrinogen-conjugated albumin nanoparticles could target irradiated tumor vasculature. In addition, radioiodination of the fibrinogen-conjugated nanoparticles resulted in tumor growth delay in B16F0 melanoma xenograft cells. Antibody engineering has rapidly evolved over the past decade. Monoclonal antibodies have proven clinically useful, resulting in the FDA approval of several compounds: rituximab (Rituxan), trastuzumab (Herceptin), and bevacizumab (Avastin). These compounds are large, and pose challenges with pharmacokinetics, manufacturing, and stability. Minimal-binding proteins, like ScFv, have become preferred for targeting in conjugation with known drug delivery systems. They are small in size (~30 kD), which allows for tumor penetration, improved renal clearance, and decreased immunogenicity. We recently screened, selected, and characterized ScFvs against one of our previously described radiation-induced antigens, P-selectin (44).

2. Materials 2.1. Tumor Models and Cell Culture

1. We used the Lewis lung carcinoma (LLC) cells (CRL-1435, ATCC, Manassas, VA) and GL-261 murine glioma cells for the biopanning screening process. The MDA-MB-231 breast cancer cell line (HTB016, ATCC) is used as an example of target binding in this protocol. 2. Cell lines are maintained in the following conditions: 3. Dulbecco’s Modified Eagle’s Medium (DMEM) (Gibco/ BRL, Bethesda, MD) is supplemented with 10% fetal bovine serum (FBS, Hyclone, Ogden, UT) and 1% penicillin–streptomycin as recommended by ATCC. 4. Monolayer cells with 80% confluence are segregated and suspended in phosphate-buffered saline (PBS). 5. Heterotopic models are developed by subcutaneously inoculating cell suspension (5 × 105 cells or adjusted for different cells) in nude mice. 6. The tumors are implanted in both hind limbs of mice and used for experiments when the tumor size reached 0.5 cm in diameter. 7. Orthotopic, metastatic, and syngeneic models have been also used according to previously published methods (40).

2.2. Tris-Buffered Saline

1. Add 8 g NaCl and 0.2 g KCl to 800 mL of distilled H2O. 2. Add 3 g of Tris base to the solution.

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3. Adjust the pH to 7.4 using HCl. 4. Add distilled H2O to bring the solution to a total of 1 L. 5. Sterilize by autoclaving. 2.3. PhosphateBuffered Saline

1. Add 8 g NaCl to 800 mL of distilled H2O. 2. Add 0.2 g KCl, 1.44 g Na2HPO4, and 0.24 g KH2PO4 to the solution. 3. Adjust the pH to 7.4 with HCl. 4. Add H2O to make a 1-L solution, and sterilize by autoclaving.

2.4. STE Buffer Solution

1. Add 1 mL of 1 M Tris (pH 8), 0.2 mL of 0.5 M EDTA (pH 8), and 2 mL of 5 M NaCl. 2. Add H2O to make a 100-mL solution, and autoclave.

2.5. 20% PEG-8000/2.5 NaCl Solution

1. Add 20 g polyethylene glycol (PEG)-8000 and 14 g NaCl.

2.6. T7 Bacteriophage

The T7 Select 415-1 EcoRI/HindIII-digested genome and the S peptide insert with cloning site overhangs (EcoRI and HindIII) are purchased and ligated and in vitro packaged (Novagen, Madison, WI) according to the manufacturer’s instructions.

2.7. Polyethylene Glycol Precipitation

1. Add 25% of th\e 20% PEG-8000/2.5 M NaCl solution to the phage solution.

2. Add H2O to make a 100-mL solution, and filter sterilize.

2. Incubate on ice for at least 30 min. 3. Spin down phage at 11 K for 20 min. 4. Re-spin three times to remove all of the PEG solution. 5. Resuspend phage in ~1 mL STE or PBS buffer. 6. Transfer to an Eppendorf tube and spin at 14 K for 10 min. 7. Transfer supernatant to a new Eppendorf tube. 2.8. Near-Infrared Imaging

1. Polyethylene glycol (PEG)-precipitated phages, selected peptides, or streptavidin (Sigma, St. Louis, MO) are labeled with amine-reactive Cy7 dye (Amersham, Arlington Heights, IL) according to the manufacturer’s protocol. 2. Near-infrared imaging (NIR) images are taken with an IVIS imaging system (Xenogen Corp. Alameda, CA). Using this device, radiance (photons/s/cm2) is measured in the region of interest. 3. The region of interest (ROI) is defined using the software provided by Xenogen.


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3. Methods 3.1. T7 Cloning

1. The selected peptide sequences are cloned as a double-stranded oligonucleotide encoding overhangs at each end for cloning into the T7 Select 415–1 arms. 2. Single strands are hybridized by heating at 95°C for 5 min followed by cooling to room temperature for 30 min in NEB 3 buffer (New England Biolabs, Beverly, MA). 3. The double-stranded oligonucleotide is then phosphorylated with 20 U polynucleotide kinase (Boehringer Mannheim, Mannheim, Germany) at 37°C for 30 min followed by heating to 65°C for 15 min in the PNK buffer containing 10 mM ATP. 4. The insert is ligated into T7 Select 415-1 vector and is packaged with the Novagen (Madison, WI) packaging extract according to manufacturer’s instructions. 5. After appropriate dilution, a plaque assay is performed and individual plaques are selected for polymerase chain reaction (PCR) and sequence verified for in-frame insertions. 6. Sequencing reactions are performed with the T7 up and down primers (Novagen) in a thermal cycling protocol, and are analyzed on an Applied Biosystems DNA Sequencer (Applied Biosystems, Foster City, CA).

3.2. T7 Phage Preparation

1. Sequence-verified T7 phage are amplified via infection of culture of the Escherichia coli strain BL-21. To infect 500 mL of a log phase culture of E. coli in Luria broth (1% tryptone, 0.5% yeast extract, and 0.08 M NaCl, pH 7.5), 1.0 × 109 pfu of T7 display phage or wild-type phage are used. 2. The culture is incubated at 37°C for 90 min or until completely lysed. 3. Phage are harvested by adding 50 mL of 5 M NaCl to the culture and the suspension is centrifuged at 8,000×g for 10 min at 4°C. 4. The supernatant is then precipitated by the addition of 50 mL of a 25% polyethylene glycol (PEG, MW 8000), 2.5 M NaCl stock. 5. After 1 h at 4°C, the PEG solution is centrifuged at 11,000×g for 30 min at 4°C. 6. The supernatant is disposed of and the PEG pellet resuspended in 5 mL Tris-buffered saline (TBS) (50 mM Tris, 150 mM NaCl, pH 7.5) followed by a brief spin to remove insoluble material and filtration (0.45-εm filtration followed by 0.22-εm

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filtration, Durapore PVDF membrane filters, Millipore, Bedford, MA). 1. Tumor-bearing mice (xenografts or syngeneic mouse models of cancer) are used for in vivo biopanning (Fig. 3).

3.3. In Vivo Biopanning Process

2. We used two distinct tumor models for the screening process; the GL261 glioma and the Lewis lung carcinoma (LLC) lines. 3. The cells are implanted into the hind limb of mice. 4. Tumors are irradiated with 3 Gy. 5. The amplified T7 phage is administered by intracardiac injection at 4 h after irradiation. 6. After 10 min of circulation, the mice are killed. In Vivo Biopanning of phage-displayed peptides

Injection into

Harvest / Disruption

irradiated xenograft Phage Library

Repeat Injection For Enrichment

Wash & Elute Amplify

After 6 rounds. Isolate Clones

Upper panel: NIR imaging demonstrating Cy7labeled phage binding selectively to irradiated LLC hind limb tumor.

Demonstrate in vivo tumor and tumor vascular tropism

PCR followed by gene sequencing for identification of encoded recombinant proteins

Lower panel: Immunohistochemical staining of phage localized to tumor vasculature

Fig. 3. This is an illustration of our in vivo biopanning system. We begin by irradiating the hind limb tumors of our xenograft models. Four hours after radiotherapy, we inject the amplified phage library into the mice via intracardiac (or tail vein) injection. After 10 min of circulation, the mice are killed. Tumor and organ tissues are collected, disrupted, washed, and the phage is eluted. The collected phage from tumor samples are amplified, and undergoes injection into another irradiated mouse xenograft. This process is repeated for a total of six rounds. The eluted phage is plated on soft agar, and single plaques are picked and PCR amplified. The genes are sequenced, and the recombinant peptides are deduced. Candidate phage were then prioritized by labeling with Cy-7, and injected into irradiated and sham-irradiate tumor-bearing mice. Phage binding within the tumors was normalized to the uptake within the rest of the body. Various time points were taken to determine the kinetics of binding (see Color Plates).


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7. The mice are perfused with 10 mL of PBS (at a rate of 2 mL/ min) into the left atrium; recovered from the right atrium. 8. The mice are subsequently killed, and the organs and tumors are removed. 9. Organs are weighed for normalization of phage to organ weight. 10. Tissues are disrupted using a hand-held homogenizer on ice. 11. To avoid cross-contamination, the homogenizer is cleaned with bleach and rinsed with PBS between homogenization of different organs. 12. The homogenate is then microcentrifuged at 5,000 rpm, and the supernatant is discarded. 13. The pellets are resuspended and washed five times with PBS. 14. The T7 phage bound to tumors are insoluble in pellets. They are amplified by adding E. coli BL21 into the washed pellets. 15. Phage titers recovered from each tissue are then normalized against the weight of each tissue, to determine the phage output per organ/tissue. 16. The phage recovered from treated tumors are amplified at 37°C with shaking until the culture is lysed (~2–3 h). 17. Cultures are then centrifuged at 8,000 rpm for 15 min. The amplified phage will be fractionated into the supernatant. 18. The amplified phage are partially purified by PEG precipitation and resuspended in PBS for another round of biopanning. 19. The entire process is repeated again, for a total of six rounds of in vivo injections. 20. Single plaques are isolated from soft agar. 21. Then gene fragments encoding peptides are amplified with polymerase chain reaction (PCR), using a T7 upstream primer (5 -AAC CCT CAA GAC CCG TTT A-3 ) and a downstream primer (5 -AAC CCT GAC CCG TTT A-3 ). 22. The primer pair solution is prepared in 0.2 pmol/εL with water. 23. The PCR beads are dissolved in 24 εL of primer pair solution. The T7 plaque is suspended in 10 εL of 1X Tris-buffered saline. 24. The PCR reaction mixture is mixed with 1 εL of phage suspension. 25. Sequencing data is collected using an ABI 377 DNA sequencer. 26. The peptide sequences are then deduced from the decoded from the DNA information.

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3.4. Prioritization of Peptides

1. Sequences are identified that appear multiple times in tumor samples after separate biopanning screenings performed in LLC and GL261 animals. 2. We select phage that have high ratios of recovery in tumor versus organs for further analysis. 3. The selected phage have different ratio of enrichment with each graft type. 4. Phage are selected that are highly enriched in both grafts, and they were labeled with amine-reactive Cy7 dye (Amersham) per the manufacturer’s protocol. 5. Labeled phage are then injected into the circulation by tail vein or jugular vein catheter in tumor-bearing mice treated with irradiation. 6. Near-infrared (NIR) images are taken with an IVIS imaging system at various time points after irradiation. 7. Radiance is measured in the region of interest using the Xenogen software. 8. Phage binding (as determined by radiance) is correlated to tumor response (tumor size), and the radiance from phage within tumors is normalized to that within the whole body. 9. Figure 4 represents the irradiated tumor-specific binding of a selected highly enriched phage (-HVGGSSV) to an irradiated

Fig. 4. Near-infrared (NIR) imaging of HVGGSSV peptide in nude mouse-bearing MDAMB-231 tumors in hind limbs. The biotinylated peptide was complexed with Alexa Fluor 750conjugated streptavidin. The complex was injected intravenously after the tumor implanted in the left hind limb was irradiated. Tumor in the right hind limb was used as control. The representative NIR image was acquired with Xenogen’s IVIS system (see Color Plates).


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hind limb with minimal uptake within the nonirradiated hind limb tumor.

4. Notes 1. Our initial studies were performed using the available linear peptide T7 phage library. We subsequently used the cyclic peptide library to increase the diversity of the selected peptides that could bind treated microvasculature. 2. Selecting the 4-h time point after irradiation for biopanning with phage may limit the number of antigenic targets available for radiation. We think that we may increase the number of targets by selecting a later time point, 24 h. 3. Care must be taken to not choose too late a time point. Based on previous studies in our laboratory (2, 45, 46), we had determined that blood flow is reduced at 5 days after irradiation. However, at 24 h, there was no evidence of decreased blood flow by Doppler studies. This may vary with tumor cell subtype. 4. Biopanning was performed serially in our system. Six rounds were performed in LLC, and separately performed in GL261 cells. These all were performed in hind limb graft models. Later studies have varied the tumor microenvironment by performing the screenings also in an orthotopic model. 5. We could also screen several lines in a sequential fashion; for example, by enriching for binding phage within LLC hind limb graft for six rounds, taking those phage, and performing an additional six rounds of biopanning in the GL261 line. 6. Early in our studies, we determined that radiation dose was also very influential in determining selected phage. For example, the threshold dose for E-selectin expression is 1 Gy, whereas it requires at least 5 Gy for induction of intercellular adhesion molecule (ICAM)-1 (47). 7. Performing these studies in mice is limited. Xenografted human tumor cells influence the mouse tumor microenvironment to provide supportive stromal and neovasculature. Thus, the fibroblasts and endothelial cells involved in this process are mouse species specific. Although similar targets may be conserved in human endothelial cells (as exemplified by our in vitro experiments with human umbilical vein endothelial cells), we need to validate the ability of our selected phage to bind in humans. We have previously demonstrated that it was feasible to guide the RGD peptide (another radiation-induced

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target) to an irradiated tumor within a patient (48). We demonstrated that 99mTc-labeled RGD bound in vivo to a breast cancer metastasis after stereotactic radiosurgery (48). 8. Receptor tyrosine kinase (RTK) inhibitors are known to enhance radiosensitivity and endothelial cytotoxicity (2, 45, 46). Thus, we hypothesized that in the response to RTK inhibition and irradiation, cells may have a more robust antigenic response. Han et al. (3) demonstrated that addition of concurrent treatment with SU11248 (a multikinase inhibitor) resulted in increased binding of the HVGGSSV peptide/ phage to treated tumors over radiation alone. It is possible that this increased binding is a result of increased calveoli at the cellular surface. However, it is also known that Src family kinases are the primary tyrosine kinases responsible for regulating calveoli-mediated endocytosis (49). Understanding this process may be useful in determining which drug delivery system to use.

Acknowledgments We thank Errki Ruoslahti (Burnham Institute for Medical Research, La Jolla, CA) for the peptide phage libraries, and Allie Fu, Ling Geng, Helina Onishko for their technical assistance. This work was supported by the US National Cancer Institutes grant R01-CA125757, the Ingram Charitable Fund, and the Vanderbilt–Ingram Cancer Center.

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Chapter 16 Chemosensitization of Tumor Cells: Inactivation of Nuclear Factor-Kappa B Associated with Chemosensitivity in Melanoma Cells After Combination Treatment with E2F-1 and Doxorubicin Hongying Hao, H. Sam Zhou, and Kelly M. McMasters Summary Combination chemotherapy has been shown to be more effective than single-agent therapy for many types of cancer, but both are known to induce drug resistance in cancer cells. Two major culprits in the development of this drug resistance are nuclear factor-κB (NF-κB) and the multidrug resistance (MDR) gene. For this reason, chemogene therapy is emerging as a viable alternative to conventional chemotherapy combinations. We have shown that transduction of the E2F-1 gene in melanoma cells markedly increases cell sensitivity to doxorubicin, thereby producing a synergistic effect on melanoma cell apoptosis. Our microarray results show that the NF-κB pathway and related genes undergo significant changes after the combined treatment of E2F-1 and doxorubicin. In fact, inactivation of NF-κB is associated with melanoma cell apoptosis induced by E2F-1 and doxorubicin, providing a link between the NF-κB signaling pathway and the chemosensitivity of melanoma cells after this treatment. Key words: Apoptosis, doxorubicin, E2F-1, electrophoretic mobility shift assay (EMSA), nuclear factor-κB (NF-κB).

1. Introduction Approximately 40% of all cancer patients are usually treated by surgery and radiotherapy, although the majority (60%) of patients are treated by chemotherapy alone (1). Combination chemotherapy is shown to be more effective than single-agent chemotherapy for many types of cancer, but both single-agent and combination chemotherapy are known to induce drug resistance in cancer cells,


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resulting in treatment failure (2). Innovative treatment strategies are therefore urgently needed to improve the treatment outcome of cancer patients. One emerging treatment strategy that has great scientific interest and promising clinical application is the combination of gene therapy and chemotherapy, or “chemogene therapy” (3). Chemogene therapy has been shown to be a viable alternative to conventional chemotherapy combinations (4, 5). Two major culprits in the development of this drug resistance are nuclear factor-κB (NF-κB) and the multidrug resistance gene (MDR) (2, 6, 7). Recently, solid genetic and biochemical evidence suggests a causative role of NF-κB in malignant conversion and progression (8). NF-κB is responsible for the resistance to chemotherapy in the treatment of aggressive tumors such as melanoma. NF-κB transcription factors are assembled through the dimerization of five Rel-domain-containing subunits: RelA (p65), c-Rel, RelB, p50/NF-κB1, and p52/NF-κB2 (9). Before cell stimulation, most NF-κB dimers are retained in the cytoplasm by binding to specific inhibitors—the inhibitors of NF-κBs (I-κBs). Cell stimulation activates the IκB kinase (IKK) complex, which is composed of two catalytic subunits (IKK-α and IKK-β) and a regulatory subunit (IKK-γ/NEMO) (10). Activated IKK phosphorylates NF-κB-bound IκB proteins and targets them for polyubiquitination and rapid degradation by creating a binding site for the SCFβTrCP ubiquitin ligase complex (11). The phosphorylation, ubiquitination, and degradation of I-κBα then releases the p50–p65 heterodimer. Freed NF-κB dimmers are then activated and translocated to the nucleus, where they coordinate the transcriptional activation of several hundred target genes (12–15). Our previous complementary DNA (cDNA) microarray analyses showed that NF-κB and related genes were involved in the chemogene therapy of E2F-1 and doxorubicin in melanoma cells (16). In this chapter, we address how we use transient transfection of NF-κB reporter plasmid and electrophoretic mobility shift assay (EMSA) to further investigate the inactivation of NF-κB associated with chemosensitivity in melanoma cells after the combination treatment of E2F-1 and doxorubicin.

2. Materials 2.1. Cell Culture

1. Human melanoma cell lines SK-MEL-2 and A375 (American Type Culture Collection [ATCC], Rockville, MD, USA) are cultured in a 5% CO2 incubator at 37°C and subcultured every 3–4 days (~80% confluent) in α-MEM (for SK-MEL-2 cells) or Dulbecco’s Modified Eagle’s Medium (DMEM) (for A375 cells), supplemented with 10% heat-inactivated fetal

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bovine serum (FBS) and a 100 U/mL penicillin/100 μg/mL streptomycin solution. All of the cell culture reagents were obtained from Gibco/BRL (Bethesda, MD, USA). 2. Opti-MEMI reduced serum medium (Invitrogen, Carlsbad, CA, USA). The chemotherapeutic agent doxorubicin (Sigma Chemical Co., St. Louis, MO, USA) is dissolved to 5 mM in tissue-culture H2O and stored in aliquots at –20°C. It is then diluted in culture medium before adding it to the cell cultures for a final concentration of 0.1 μM. 2.2. Recombinant Adenoviral Construct

The Ad5CMV-E2F-1 vector is deleted in the E1 subunit and contains the transgene E2F-1 under the control of the CMV promoter. It was propagated in the 293 cell line, and titers were determined using standard plaque assays (14).

2.3. Transient Transfection of NF-kB Reporter Plasmid

1. pNF-κB-Luc reporter plasmid (Stratagene, catalog number 219077, La Jolla, CA, USA). 2. phRL-CMV plasmid (Promega, catalog number E6271, Madison, WI, USA). 3. Lipofectamine 2000 transfection reagent (Invitrogen). 4. Dual-luciferase reporter assay system (Promega). 1X Passive Lysis Buffer: add 600 μL 5X Passive Lysis Buffer in 2,400 μL distilled H2O to make 3,000 μL of 1X Passive Lysis Buffer. 1X Luciferase Assay Buffer II: add 10 mL Luciferase Assay Buffer II in 1 vial of Luciferase Assay Substrate (lyophilized). After it is completely dissolved, store in aliquots of 1 mL at –20°C. 1X Stop & Glo reagent: add 980 μL Stop & Glo buffer to 20 μL of 50X Stop & Glo substrate, to make 1,000 μL of 1X Stop & Glo reagent. This buffer should be freshly made before use. 5. Tumor necrosis factor-alpha (TNF-α) (R&D Systems, Inc., Minneapolis, MN, USA) is dissolved to 7.5 μg/mL in tissue-culture H2O and stored in aliquots at –20°C. This is then added to the culture medium for a final concentration of 7.5 ng/mL.

2.4. Electrophoretic Mobility Shift Assay

1. 10 mL of 6% acrylamide gel (for 1 mini-gel of 8 × 8 × 0.1 cm): 30 % Acrylamide/bis solution

2 mL

H2O

5.93 mL

5X TBE

2 mL

10 % Ammonium persulfate

70μL

TEMED

3.5 μL

2. 5X TBE: 1.1 M Tris-HCl, 900 mM boric acid, 25 mM EDTA, pH 8.3. Weigh 53.9 g Tris base, 28 g boric acid, and 3.72 g

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disodium EDTA.2H2O. Dissolve in 800 mL distilled H2O. Add slightly less than 28 g boric acid. Mix until completely dissolved. Check and adjust pH to 8.3 with boric acid, and bring the final volume to 1,000 mL with distilled H2O. 3. The sequence of the biotin-end-labeled NF-κB consensus oligonucleotide is: Sense 5′-AGTTGAGGGGACTTTCCCAGGC/3bio/-3′ Antisense 5′-GCCTGGGAAAGTCCCCTCAACT/3bio/-3′ They are synthesized by Integrated DNA Technologies, Inc. (Coralville, IA, USA). 4. NE-PER Nuclear and Cytoplasmic Extraction Reagent can be purchased from Pierce (Pierce, Rockford, IL, USA). 5. LightShift chemiluminescent EMSA kit (Pierce). It contains two kits: one is the LightShift EMSA Optimization and Control Kit; the other is the Chemiluminescent Nucleic Acid Detection Module. Conjugate/blocking buffer: add 66.7 μL of stabilized streptavidin–horseradish peroxidase conjugate to 20 mL blocking buffer (1:300 dilution). 1X wash solution: add 40 mL of 4X wash buffer to 120 mL of ultrapure water. Substrate working solution: add 6 mL of luminol/enhancer solution to 6 mL of stable peroxide solution. 6. BCA protein assay kit (Pierce). 7. Proliferating cell nuclear antigen (PCNA) antibody (1:800) (Santa Cruz Biotechnology, Inc., Santa Cruz, CA, USA). 8. UNSCAN-IT software (Silk Scientific, Inc., Orem, UT, USA).

3. Methods The transient transfection of NF-κB-driven luciferase reporter plasmid was used to determine the NF-κB signaling in transfected cells. The pNF-κB-Luc reporter plasmid contains tandem repeats of the NF-κB enhancer elements. The enhancer elements, together with a TATA box, control the expression of the downstream luciferase reporter gene. The phRL-CMV plasmid, which expresses Renilla luciferase, was used as an internal control to account for variations in transfection efficiency. Electrophoretic mobility shift assay (EMSA) is another supplementary method for detecting NF-κB signaling that did not depend on cell transfection efficiency. If NF-κB subunits


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are present in the nuclear extracts, their interaction with the probe shifts the band of the labeled probe up, because the complex of probe and NF-κB subunit has a lower electrophoretic mobility than the probe alone. To check whether the observed shifted bands are specific for NF-κB, we run a competition test with excess nonlabeled (“cold”) NF-κB oligonucleotide. If the observed signals are NF-κB specific, those signals should disappear in the presence of the excess nonlabeled (“cold”) NF-κB oligonucleotide. To determine which NF-κB subunits are responsible for the observed shifted bands, the EMSA signals can be “supershifted” by adding different NF-κB subunit-specific antibodies (such as p65 and p50) to the binding reactions. The antibodies will bind to the corresponding NF-κB subunit, which results in a NF-κB/ probe/antibody ternary complex; the electrophoretic mobility of this complex is even lower than that of the NF-κB/probe complex, so that a supershifted band can be observed. 3.1. Transient Transfection of pNFkB-Luc Reporter Plasmid

1. SK-MEL-2 and A375 cells are plated in 12-well plates at a density of 1 × 105 cells per well. The next day, the cells are transiently transfected with 0.8 μg of the pNF-κB-Luc reporter plasmid and 0.016 μg of an internal control plasmid (phRL-CMV) using Lipofectamine 2000 reagent as detailed as following (see Notes 1 and 2). 2. Dilute pNF-κB-Luc plasmid DNA and phRL-CMV plasmid DNA in 100 μL of Opti-MEMI medium. Mix gently, then dilute 2 μL of Lipofectamine 2000 in 100 μL of Opti-MEMI medium. Mix gently. Incubate at room temperature for 5 min (see Note 3). 3. After the 5-min incubation, combine the diluted DNA with diluted Lipofectamine 2000 and mix gently. After the combined mixture is incubated for 20 min, add 200 μL of the mixture to each well containing cells and 800 μL of medium. Mix gently by rocking the plate back and forth. 4. Where indicated, TNF-α is then added to the medium after transfection for 5 h, and cells are incubated in TNF-α for an additional 4 h followed by 16 h of mock infection or the combination treatment of Ad-E2F-1 and doxorubicin. 5. For the combined treatment of Ad-E2F-1 and doxorubicin, the cells are first infected with Ad-E2F-1. The media is removed, and the adenoviral vector Ad-E2F-1 is added in 150 μL of α-MEM (no FBS) per well of 12-well plates at an multiplicity of infection (MOI) of 2 and incubated at 37°C for 1 h. Shake the plates every 10 min. One hour after infection, the medium is removed and replaced with 1 mL of fresh media (with 5% FBS) containing doxorubicin at a concentration of 0.1 μM. Mock infection is performed by treatment of cells with media only.

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6. After 16 h of infection, remove the medium completely. Add 250 μL of 1X Passive Lysis Buffer (from the Dual Luciferase Reporter Assay System kit) in each well of 12-well plates. Place the plate on an orbital shaker with gentle shaking to ensure complete and even coverage of the cell monolayer with the lysis buffer. Shake the plate at room temperature for 15 min. 7. Mix the lysis buffer well by pipetting up and down several times. Pipet out 20 μL of passive lysis buffer from the above in a glass tube and put in a luminometer (Zylux Corp., Maryville, TN, USA). Add 100 μL of Luciferase Assay Reagent II (LAR II) to the tube to make a luciferase reading (luciferase activity). Then, add another 100 μL of 1X Stop & Glo reagent into the tube again to make a Renilla luciferase reading. Relative luciferase activities for each sample are calculated as firefly luciferase activity divided by Renilla luciferase activity and are compared to the mean of the control (mock infection) sample. An example result is shown in Fig. 1a and b 3.2. NF-kB DNABinding Activity Measurement by Electrophoretic Mobility Shift Assay 3.2.1. Cytoplasmic and Nuclear Protein Extract

1. SK-MEL-2 and A375 cells are seeded at a density of 1 × 106 cells in 100-mm dishes. After 24 h, the cells are treated with Ad-E2F-1 or a combination of Ad-E2F-1 and doxorubicin as described above. After 24 h of treatments, nuclear protein and cytosolic protein are extracted using NE-PER Nuclear and Cytoplasmic Extraction Reagent, detailed as follows (see Note 4). 2. After 24-h treatment, trypsinize the cells and wash with PBS once. Completely remove PBS from the cell pellet, leaving the cell pellet as dry as possible. 3. Add 50 μL of ice-cold CER I (with 1X protease inhibitor) to the cell pellet. Vortex the tube vigorously on the highest setting for 15 s to fully resuspend the cell pellet. Incubate the tube on ice for 10 min. 4. Add 2.75 μL of ice-cold CER II to the tube. Vortex the tube for 5 s on the highest setting. Incubate the tube on ice for 1 min. Vortex the tube for 5 s on the highest setting again. Then, centrifuge the tube for 5 min at maximum speed (16,000×g) in a microcentrifuge. 5. Immediately transfer the supernatant (cytoplasmic extract) fraction to a clean prechilled tube. Place this tube on ice until use or storage. 6. Resuspend the pellet produced above (contains nuclei) in 25 μL of ice-cold NER (with 1X protease inhibitor). Vortex the tube on the highest setting for 15 s. Return the sample to ice and continue vortexing for 15 s every 10 min, for a total of 50 min. Centrifuge the tube for 10 min at maximum speed (16,000×g) in a microcentrifuge. Immediately transfer the


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Fig. 1. Combination of E2F-1 and doxorubicin inhibited the NF-κB transcriptional activity by luciferase assay. (a) Melanoma cells were co-transfected with a NF-κB reporter plasmid expressing firefly luciferase and a phRL-CMV plasmid expressing Renilla luciferase (as an internal control). After 5 h of transfection, TNF-α was added to the medium, and cells were incubated for an additional 4 h followed by 16 h of mock infection or the combination treatment of E2F-1 and doxorubicin. Luciferase activity was then measured. Relative luciferase activity was calculated as firefly luciferase activity divided by Renilla luciferase activity and shown relative to the control (mock infection). Statistical significance for the combination treatment of E2F-1 and doxorubicin after TNF-α stimulation compared with no treatment after TNF-α stimulation. *p < 0.01, Mock = mock infection, TNF-α = TNF-α stimulation, TNF-α + ED = TNF-α stimulation followed by E2F-1 + doxorubicin. (b) The same assays were carried out in the absence of TNF-α stimulation to determine the effect of the combination treatment on the NF-κB basal levels. The results represent the means ± SD (bars) of at least three different experiments performed in duplicates. Statistical significance for each treated sample was compared with the control (mock infection) (*p < 0.01, ** p < 0.001). Mock = mock infection, ED = E2F-1 + doxorubicin.

supernatant (nuclear extract) fraction to a clean prechilled tube. Place on ice. Store all extracts in aliquots at –80°C (avoid frequent freeze/thaw cycles). 7. Use a BCA protein concentration assay kit to determine the nuclear protein concentration. 3.2.2. DNA-Binding Reaction and Running the Gel

Use the LightShift EMSA Optimization and Control Kit for binding reactions and the Chemiluminescent Nucleic Acid Detection Module for detecting the NF-κB DNA-binding band. We use a Bio-Rad mini-gel running and transferring unit.

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1. Prepare a 6% acrylamide gel. The gel should polymerize in about 30 min. 2. Fill the electrophoresis unit with 0.5X TBE to just above the bottom of the wells (this will reduce the heat generated during electrophoresis). Flush the wells and pre-electrophorese the gel at 100 V for 60 min. 3. Proceed to the binding reaction in step 4 while the gel is preelectrophoresing. 4. Binding reactions are performed using the system described in Table 1. Always add the components in the order listed in Table 1. First, incubate the unlabeled oligo with the nuclear extract at room temperature for 20 min, then incubate with the biotinlabeled oligo for another 20 min at room temperature. Add 5 μL of 5X loading buffer to each 20-μL binding reaction, pipetting up and down several times to mix before preparing to load on a 6% acrylamide gel. Do not vortex or mix vigorously (see Note 5). 5. Switch off the current to the electrophoresis gel. Then, load 25 μL of each sample onto the polyacrylamide gel. Switch on the current (set to 100 V for an 8 × 8 × 0.1-cm gel) and electrophorese the samples until the bromophenol blue dye has migrated approximately 2/3–3/4 down the length of the gel. The free biotin–EBNA (Epstein-Barr Nuclear Antigen) control DNA duplex migrates just behind the bromophenol blue in a 6% polyacrylamide gel. This running step will take about 60 min.

Table 1 Binding reactions for the EMSA system Negative control

Specific competitor

Sample tube

10X binding buffer

2 μL

2 μL

2 μL

50% Glycerol

1 μL

1 μL

1 μL

100 mM MgCl2

1 μL

1 μL

1 μL

1 μg/μL poly (dI.dC)

2 μL

2 μL

2 μL

NP-40

1 μL

1 μL

1 μL

Unlabeled oligo

0

2 pmol

2 pmol

Nuclear extract

0

4 μg

4 μg

Biotin-labeled oligo

40 fmol

40 fmol

40 fmol

Ultrapure water

(Adjust accordingly)



Total volume

20 μL

20 μL

20 μL


309

6. Electrophoretic transfer of binding reactions to nylon membrane: Soak the nylon membrane in 0.5X TBE for at least 10 min. Sandwich the gel and nylon membrane in a clean electrophoretic transfer unit. Wet transfer (in 0.5X TBE) at 380 mA (~100 V) for 60 min for mini-gels (8 × 8 × 0.1 cm). When the transfer is complete, place the membrane with the bromophenol blue-side up on a dry paper towel (there should be no dye remaining in the gel). Allow the buffer on the membrane surface to absorb into the membrane. This will only take a minute. Do not let the membrane dry. Immediately proceed to step 7 (see Note 6). 7. Cross-link the transferred DNA to membrane. Cross-link at 120 mJ/cm2 using a commercial UV light cross-linking instrument equipped with 254-nm bulbs (60 s exposure using the auto cross-link function). After the membrane is crosslinked, proceed directly to Subheading 3.2.3. Alternatively, the membrane may be stored dry at room temperature for several days. Do not allow the membrane to get wet again until ready to proceed with Subheading 3.2.3. 3.2.3. Detect BiotinLabeled DNA-Binding Band

Detect the biotin-labeled DNA-binding band by using horseradish peroxidase-conjugated streptavidin (LightShift chemiluminescent EMSA kit, Pierce, Rockford, IL, USA) (see Note 7). 1. Gently warm the blocking buffer and the 4X wash buffer to 37–50°C in a water bath until all particulate is dissolved. These buffers may be used between room temperature and 50°C as long as all particulate remains in solution. The substrate equilibration buffer may be used between 4°C and room temperature. 2. To block the membrane, add 20 mL of blocking buffer and incubate for 15 min with gentle shaking. 3. Decant the blocking buffer from the membrane and replace it with the conjugate/blocking solution. Incubate the membrane in the conjugate/blocking buffer solution for 15 min with gentle shaking (see Note 8). 4. Transfer the membrane to a new container and rinse it briefly with 20 mL of 1X wash solution. Wash the membrane four times for 20 min each in 20 mL of 1X wash solution with gentle shaking (see Note 9). 5. Transfer the membrane to a new container and add 30 mL of substrate equilibration buffer. Incubate the membrane for 5 min with gentle shaking. 6. Remove the membrane from the substrate equilibration buffer, carefully blotting an edge of the membrane on a paper towel to remove excess buffer. Place the membrane in a clean container or onto a clean sheet of plastic wrap placed on a flat surface.

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7. Pour the substrate working solution onto the membrane, so that it completely covers the surface. Alternatively, the membrane may be placed DNA-side down onto a puddle of the working solution. Incubate the membrane in the substrate working solution for 5 min without shaking (see Note 10). 8. Remove the membrane from the working solution and blot an edge of the membrane on a paper towel for 2–5 s to remove excess buffer. Do not allow the membrane to become dry. Wrap the moist membrane in plastic wrap, avoiding bubbles and wrinkles. Place the membrane in a film cassette and expose to X-ray film for 1 min. Develop the film according to manufacturer’s instructions. The exposure time may be adjusted to obtain the desired signal (see Note 11). 9. The specific NF-κB DNA-binding band is then scanned and quantified with UNSCAN-IT software (Silk Scientific, Inc.).

Fig. 2. Reduced NF-κB DNA-binding activity after the combination treatment of E2F-1 and doxorubicin. After 24 h of treatment as labeled, nuclear extracts were prepared from these treated cells as detailed in Subheading 3 These extracts were then analyzed by electrophoretic mobility shift assay (EMSA). Binding reactions were carried out using 40 fmol of biotin-end-labeled NF-κB DNA and 4 µg of nuclear protein extract. No protein extract was used as negative control, and 50-fold of unlabeled NF-κB probe was used as specific competitor where indicated. (a) EMSA result of SK-MEL-2 cells treated with E2F-1 and doxorubicin. PCNA was used as loading control. The lower graphs represent the means ± SD (bars) of NF-κB densitometric bands performed in five different experiments. (b) Supershift assay of NF-κB showed that NF-κB band was shifted because of the formation of larger molecular complexes after addition of anti-NFκB p65 antibody. Mock = mock infection, ED = E2F-1 + doxorubicin.


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10. For the loading control, 10 µg of nuclear proteins from each sample is subjected to Western blot analysis for PCNA (1:800 from Santa Cruz Biotechnology Inc.), which shows no alternation after different treatments. 11. For the supershift assay, nuclear extracts prepared from the treated cells are first incubated with p65 antibody (sc-372x) (Santa Cruz Biotechnology, Inc.) for 20 min at room temperature before the complex is incubated with the unlabeled oligo and analyzed by EMSA. 12. An example result is shown in Fig. 2a and b

4. Notes 1. For the experiment of transient transfection of pNF-κB-Luc plasmid, the transfection efficiency of the cell lines to be used needs to be checked first. This can be done by transfecting enhanced green fluorescence protein (EGFP) plasmid and counting the positive (green) cells under a fluorescence microscope. If the transfection efficiency is less than 50%, the cell lines cannot be used for this experiment. You have to choose other cell lines that have higher transfection efficiency. 2. The transfection ratio of the pNF-κB-Luc plasmid and the internal control plasmid (phRL-CMV plasmid) needs to be adjusted to different cell lines. 3. The transfection ratio of total plasmid DNA and Lipofectamine 2000 has been optimized in these two cell lines. We did not follow the ratio described in Lipofectamine 2000 manual. We suggest optimizing the ratio in different cell lines for different purposes. 4. If using lesser or larger amounts of cells, the volume of CER I, CER II, and NER needs to be proportionally changed. However, you need to keep a higher protein concentration in the nuclear extract to be used in the following EMSA binding reactions. 5. The volume for each reagent has been optimized in this system. You can increase the amount of nuclear extract (from 4 μg to 6 or 8 μg) if the NF-κB DNA-binding signal is weak. 6. We use the Bio-Rad mini-gel transfer system. Use very clean forceps and powder-free gloves, and handle the membrane only at the corners. Use clean transfer sponges. Avoid using sponges that have been used in Western blots. 7. The recommended volumes are for an 8 × 8-cm membrane. If larger gels are used, adjust the volumes of each buffer

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accordingly. Perform all blocking and detection incubations in clean trays or in plastic weigh boats on an orbital shaker. 8. This conjugate/blocking buffer solution has been optimized for the Nucleic Acid Detection Module and should not be modified. 9. The washing times can be extended to 30 or 40 min. The more thoroughly washed, the clearer the background will be. 10. Exposure to the sun or any other intense light can harm the working solution. For best results, keep the working solution in an amber bottle and avoid prolonged exposure to any intense light. Short-term exposure to typical laboratory lighting will not harm the working solution. 11. We use ECL Hyperfilm from Amersham (Catalog number 28-9068-37) to expose the signals. This kind of film is more sensitive than other films. This nonradioactive detecting method is also sensitive. We suggest using this method to avoid radioactive contamination.

Acknowledgments We are grateful to Mrs. Margaret Abby for her expert manuscript editing. Supported by NIH Grant R01CA90784 to KMM.

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5. Nemunaitis, J., Swisher, S.G., Timmons, T., Connors, D., Mack, M., Doerksen, L., Weill, D., Wait, J., Lawrence, D.D., Kemp, B.L., Fossella, F., Glisson, B.S., Hong, W.K., Khuri, F.R., Kurie, J.M., Lee, J.J. Lee, J.S., Nguyen, D.M., Nesbitt, J.C., Perez-Soler, R., Pisters, K.M.W., Putnam, J.B., Richli, W.R., Shin, D.M., Walsh, G.L., Merritt, J., and Roth, J. (2000) Adenovirus-mediated p53 gene transfer in sequence with cisplatin to tumors of patients with non-small-cell lung cancer. J. Clin. Oncol. 18, 609–622. 6. Yeh, P.Y., Chuang, S.E., Yeh, K.H., Song, Y.C., Ea, C.K., and Cheng, A.L. (2002) Increase of the resistance of human cervical carcinoma cells to cisplatin by inhibition of the MEK to ERK signaling pathway partly via enhancement of anticancer drug-induced NF-κB activation. Biochem. Pharmacol. 63, 1423–1430.

Chemosensitization of Tumor Cells: Inactivation of Nuclear Factor-Kappa 7. Yeh, P.Y., Chuang, S.E., Yeh, K.H., Song, Y.C., and Cheng, A.L. (2003) Involvement of nuclear transcription factor-κB in lowdose doxorubicininduced drug resistance of cervical carcinoma cells. Biochem. Pharmacol. 66, 25–33. 8. Karin, M. (2006) Nuclear factor-kappaB in cancer development and progression. Nature 441(7092), 431–436. 9. Ghosh, S. and Karin, M. (2002) Missing pieces in the NF-κB puzzle. Cell, 109(Suppl.), S81–S96. 10. Rothwarf, D.M. and Karin, M. (1999) The NF-κB activation pathway: a paradigm in information transfer from membrane to nucleus. Sci. STKE (October 26) 1999(5), RE1. 11. Karin, M. and Ben-Neriah, Y. (2000) Phosphorylation meets ubiquitination: the control of NF-κB activity. Annu. Rev. Immunol. 18, 621–663. 12. Werner, S.L., Barken, D. and Hoffmann, A. (2005) Stimulus specificity of gene expression

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programs determined by temporal control of IKK activity. Science. 309, 1857–1861. Park, J.M., Greten, F.R., Wong, A., Westrick, R.J., Arthur, S.C., Otsu, K., Hoffmann, A., Montminy, M. and Karin, M. (2005) Signaling pathways and genes that inhibit pathogen-induced macrophage apoptosis: CREB and NF-κB as key regulators. Immunity 23, 319–329. Covert, M.W., Leung, T.H., Gaston, J.E. and Baltimore, D. (2005) Achieving stability of lipopolysaccharide-induced NF-κB activation. Science 309, 1854–1857. Aggarwal, B.B. (2004) Nuclear factor-κB: the enemy within. Cancer Cell 6, 203–208. Hao, H., Dong, Y.B., Bowling, M.T., Zhou, H.S. and McMasters, K.M. (2006) Alteration of gene expression in melanoma cells following combined treatment with E2F-1 and adriamycin. Anticancer Res. 26(3A), 1947–1956.

Chapter 17 Induction of Tumor Cell Apoptosis by TRAIL Gene Therapy Thomas S. Griffith Summary Members of the tumor necrosis factor (TNF) superfamily influence a variety of immunological functions, including cellular activation, proliferation, and death, upon interaction with a corresponding superfamily of receptors. Whereas interest in the apoptosis-inducing molecules TNF and Fas ligand has peaked because of their participation in events such as autoimmune disorders, activation-induced cell death, immune privilege, and tumor evasion from the immune system, another death-inducing family member, TNF-related apoptosis-inducing ligand (TRAIL), or Apo-2 ligand, has generated excitement because of its unique ability to induce apoptosis in a wide range of transformed cell lines but not in normal tissues. TRAIL is well tolerated when given to healthy animals, and no observable histological or functional changes have been observed in any of the tissues or organs examined. Moreover, multiple injections of soluble TRAIL into mice beginning the day after tumor implantation can significantly suppress the growth of the tumors, with many animals becoming tumor-free. One potential drawback to these findings is that large amounts of soluble TRAIL may be required to inhibit tumor formation, possibly because of the pharmacokinetic profile of soluble TRAIL that indicates that, after intravenous injection, the majority of the protein is rapidly cleared. Increasing the in vivo half-life of recombinant soluble TRAIL or developing an alternative means of delivery may increase the relative tumoricidal activity of TRAIL such that larger, more established tumors could be eradicated as efficiently as smaller tumors. The information presented here describes the production of an adenoviral vector engineered to carry the complementary DNA (cDNA) for murine TRAIL (hTRAIL). Key words: Adenovirus, apoptosis, gene transfer, TRAIL, tumor.

1. Introduction When first described in 1995, Wiley et al. clearly demonstrated that recombinant, soluble tumor necrosis factor (TNF)-related apoptosis-inducing ligand (TRAIL, or Apo-2 ligand [Apo-2L])


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possessed the unique ability of inducing apoptosis in a range of tumor cell lines, while having little to no cytotoxicity against normal cells and tissues (1–3). Since then, the vast majority of the studies published on TRAIL have focused on its tumoricidal activity, with the intention of developing TRAIL into a cancer therapeutic agent. One of the first reports that evaluated the antitumor activity of TRAIL in vivo used immunocompromised mice that were subcutaneously injected with human tumor cells, followed by intraperitoneal or intravenous injections of soluble TRAIL starting various days after tumor implantation (4, 5). Multiple doses of recombinant, soluble TRAIL beginning the day after tumor implantation suppressed tumor outgrowth, with many animals becoming tumor-free. The antitumor activity of the systemically administered TRAIL in these experiments demonstrated that TRAIL could interact with the primary tumor and, potentially, any metastases that would normally be difficult to detect and/or treat. One major drawback to these findings, however, was that large amounts of TRAIL (up to 500 μg/day) were required to inhibit tumor formation, because the intravenously injected TRAIL had an extended distribution half-life of 1.3 h and an elimination half-life of 4.8 h (4). Furthermore, the systemic administration of TRAIL appeared to be most successful only when administered shortly after tumor implantation (4–7). The administration of equivalent doses of recombinant TRAIL protein into humans may be problematic. Thus, as an alternative approach, we were the first to report the development of a nonreplicative recombinant adenoviral vector to deliver the TRAIL gene directly into tumor cells (8, 9). Localized therapy of solid tumors has been successful in a number of settings. For example, prostate cancer is commonly treated with local (intraprostatic) regimens serving as current treatment options, such as cryotherapy, brachytherapy, and several experimental viral-based studies (10–12). Of the viral-based therapies, published data indicate minimal toxicity for adenovirus injection into the prostate up to doses of 1011 plaque forming units (pfu) (13). Recombinant adenoviral vectors infect a wide range of proliferating and quiescent cell types, making this gene delivery system a suitable tool for studying diseases, vaccine therapy, and potential clinical use (14). Recombinant adenovirus is structurally stable, and can be prepared and purified to high titers. Wild-type adenovirus infections are extremely common in the general population, giving adenovirus a well-documented safety record (15). Moreover, no side effects were reported in US military recruits after vaccination of wild-type virus, demonstrating its safety for human use (16). In contrast to wild-type virus, the adenoviral vectors used for gene therapy and vaccine therapy are genetically modified to render the virus replication defective, commonly through the deletion in the E1 region or the E1 and

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E2 regions, and to allow insertion of foreign genes. Adenoviral gene transfer has been used in a variety of experimental conditions that include transfers to the prostate, liver, lung, central nervous system, and cancer cells (17–22). Thus, the information detailed herein describes the production and use of a recombinant, replication-deficient adenoviral vector encoding murine trail driven by a cytomegalovirus (CMV) promoter (Ad5-mTRAIL).

2. Materials 2.1. Cell Culture

1. RPMI 1640, Dulbecco’s Modified Eagle Medium (DMEM), or Modified Eagle Medium (MEM) (Gibco, Invitrogen, Carlsbad, CA) supplemented with 10% fetal bovine serum (FBS, Hyclone, Ogden, UT); 1% penicillin/streptomycin liquid with 10,000 U each (Gibco); 1% sodium pyruvate solution 100 mM (100X) (Gibco); 1% MEM nonessential amino acids solution 10 mM (100X) (Gibco); HEPES buffer solution (1 M) (Gibco); and 0.1% 2-mercaptoethanol (1,000X) (Gibco). When the medium contains all of the above supplements, it will be referred to as “complete” medium, such as complete RPMI.

2.2. PCR and Cloning

1. Qiagen RNeasy Plus Mini Kit (Qiagen, Valencia, CA). 2. Oligo(dT)12–18 primer (Invitrogen). 3. dNTPs (10 mM of each dNTP). 4. 5X First-strand buffer, 0.1 M DTT. 5. Superscript III Reverse Transcriptase (Invitrogen). 6. PfuUltra HF DNA polymerase. 10X PfuUltra HF reaction buffer (Stratagene) 7. Polymerase chain reaction (PCR) primer: forward 5′-CCCCTCGAGATGCCTTCCTCAGGGGCC-3′ (Integrated DNA Technologies, Coralville, IA). 8. PCR primer: reverse 5′-CCCAAGCTTTTAGTTAATTAAAAAGGC-3′ (Integrated DNA Technologies). 9. TOPO cloning vector and kit (Invitrogen). 10. Restriction enzymes: XhoI and HindIII (New England Biolabs, Beverly, MA). 11. QIAquick gel Extraction Kit (Qiagen).

2.3. Adenovirus Production

1. HEPES-buffered saline (HEBS): 5.0 g HEPES, 8.0 g NaCl, 0.37 g KCl, 0.188 g Na2HPO4 7H2O, and 1.0 g glucose in 1,000 mL, pH to 7.1, autoclave, store at 4°C.

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2. 2.5 M CaCl2. 3. 2% FBS/1% penicillin/streptomycin in DMEM. 4. 10% FBS/1% penicillin/streptomycin in DMEM. 5. Light CsCl, 1.2 g/mL in 10 mM Tris-HCl, pH 8.1. 6. Heavy CsCl, 1.45 g/mL in 10 mM Tris-HCl, pH 8.1. 7. 10 mM Tris-HCl pH 8.1. 8. 3% Sucrose/phosphate-buffered saline (PBS). 9. 50% Glycerol/bovine serum albumin (BSA). 10. Slide-A-Lyzer Cassettes (Pierce, Rockford, IL). 11. Beckman Centrifuge Tubes, 1 inch × 3 inch. 12. Beckman Centrifuge Tubes, 9/16 inch × 3 inch. 2.4. Sodium Dodecyl Sulfate (SDS)-Polyacrylamide Electrophoresis (PAGE) and Immunoblotting

1. Cell lysis buffer: 1% NP-40 and Complete Mini protease inhibitor tablet (Roche, Indianapolis, IN) in PBS. 2. SDS-PAGE running buffer: 25 mM Tris-Base, 192 mM glycine, 0.1% (w/v) SDS, pH 8.3. 3. Tris-glycine transfer buffer: 12 mM Tris-Base, 96 mM glycine, 20% (v/v) methanol. 4. Blotting membrane: Ready Gel Blotting Sandwich 0.45-μm nitrocellulose with filter paper (Bio-Rad, Hercules, CA). 5. Blocking buffer: 5% (w/v) nonfat dry milk in Tris-buffered saline with Tween (TBST). 6. Primary antibody: goat anti-TRAIL antiserum (clone K-18; Santa Cruz Biotechnology, Santa Cruz, CA). 7. Wash buffer: Tris-buffered saline with Tween (TBST): 149.96 mM NaCl, 10 mM Tris-Base, pH 7.4, 0.05% (v/v) Tween-20. 8. Secondary antibody: anti-goat-horseradish peroxidase (HRP) antibody (Amersham, Arlington Heights, IL). 9. Chemiluminescence detection: SuperSignal West Pico Chemiluminescence Substrate (Pierce).

2.5. Flow Cytometry

1. PBS: 137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, 1.8 mM KH2PO4, pH 7.4. 2. FACS buffer: PBS with 2% FBS and 0.02% NaN3. 3. Primary antibody: phycoerythrin (PE)-conjugated rat antimouse monoclonal antibody (mAb) (clone N2B2, eBioscience, San Diego, CA) or PE-conjugated rat IgG2a isotype control mAb (Caltag Laboratories, Inc., Burlingame, CA). 4. Sample acquisition and data analysis: FACScan (BD Biosciences, San Jose, CA) and FlowJo (Tree Star, Inc., San Carlos, CA).


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1. Crystal violet: 0.5% (w/v) crystal violet, 25% (v/v) methanol in water. 2. Deoxycholic acid: 2% (w/v) in water. 3. FITC-conjugated annexin V (R&D Systems, Minneapolis, MN) and propidium iodide (50 μg/mL in PBS; Sigma, St. Louis, MO). 4. Annexin binding buffer: 10 mM HEPES/NaOH pH 7.4, 140 mM NaCl, 2.5 mM CaCl2.

3. Methods 3.1. Growth of Cell Lines

1. The transplantable murine renal adenocarcinoma cell line, Renca (23), was obtained from Dr. Robert Wiltrout (National Cancer Institute, Frederick, MD). The mouse B lymphoma cell line 2PK3 transfected with mouse TRAIL cDNA (mTRAIL-2PK3) (24) was obtained from Dr. Hideo Yagita (Juntendo University). The human melanoma cell line, WM 164, was obtained from Dr. Meenhard Herlyn (Wistar Institute, Philadelphia, PA) (3). WM 164 and human embryonic kidney cells (HEK 293) were grown in DMEM containing 10% FBS and 1% penicillin/streptomycin. Renca and mTRAIL-2PK3 were cultured in complete RPMI.

3.2. Production of Adenovirus Encoding the mTRAIL Gene

1. Total RNA is isolated from mTRAIL-2PK3 cells using the Qiagen RNeasy Mini Kit, and then reverse transcribed using Superscript III Reverse Transcriptase and oligo-dT primers. In the first step, mix total RNA (2 μg), oligo-dT (1 μL of 50 μM), and sterile, distilled water to 12 μL in a thin-walled PCR tube. Heat to 70°C for 10 min, and then cool down to 4°C. In the second step, add 1 μL of 10 mM dNTP Mix (final concentration 0.5 mM each), 4 μL of 5X First-Strand Buffer (final concentration 1X), 2 μL of 0.1 M DTT (final concentration 0.01 M), and 1 μL Superscript III Reverse Transcriptase. Heat to 25°C for 10 min, 42°C for 50 min, 70°C for 15 min, and then cool down to 4°C. 2. The cDNA for murine tnfsf10 (mTRAIL) is then amplified by PCR using primers corresponding to the first six codons (the 5′-primer) and the last six codons (the 3′-primer), according to the published sequence (1). In addition, the 5′ and 3′ primers included sequences encoding XhoI and HindIII restriction enzyme sites, respectively (see Note 1). Below are the amounts of each reagent used in one typical PCR reaction mixture. Amounts must be adjusted accordingly for multiple samples. Add each component in the order listed in Table 1 while

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Table 1 Cloning of murine TRAIL gene by PCR Component

Amount per reaction Final concentration

dH2O

32 μL

10X PfuUltra HF reaction buffer

5 μL

1X, Mg2 + 2 mM

dNTPs (10 mM each dNTP)

1 μL

0.2 mM

cDNA

1 μL

Forward primer (10 mM)

5 μL

1 μM

Reverse primer (10 mM)

5 μL

1 μM

PfuUltra HF DNA polymerase (2.5 U/mL)

1 μL

2.5 U

Total reaction volume

50 μL

gently mixing. Immediately before thermal cycling, aliquot 50 μL of the reaction mixture into the appropriate number of sterile thin-walled PCR tubes (see Note 2). Perform PCR using optimized cycling conditions determined to be best for the cycler being used. 3. The PCR product was purified from 1% agarose gel using the QIAquick Gel Extraction Kit, and cloned into a TOPO cloning vector for sequence verification. Once verified, the mTRAIL cDNA was excised by XhoI and HindIII restriction enzyme digestion, and gel purified as above for the subsequent cloning into the pAd5CMVK-NpA shuttle vector. This vector is devoid of the left-hand inverted terminal repeat (ITR), the packaging signal, and the E1 sequences, which greatly reduce or eliminate the production of wildtype, replication-competent adenovirus.

3.3. HEK 293 Transfection

4. A replication-deficient adenovirus encoding the mTRAIL gene (Ad5-mTRAIL) expressed from the cytomegalovirus (CMV) promotor was generated using the RAPAd.I system (25) by the University of Iowa Gene Transfer Vector Core (Iowa City, IA). 1. Plates (150 mm) of HEK 293 cells (1.5 × 106 cells/plate) are prepared the day before the start of transfection. Prepare the 150-mm plates as follows for the cells to reach 60–70% confluency at the time of infection (Table 2).


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Table 2 Recommended plating of HEK 293 cells for transfection Time to transfection

Cells/plate

Ready next day

12 × 106

2 days

5 × 106

3 days

2 × 106

4 days

1.5 × 106

Table 3 Suggested reagents for transfection Shuttle vector

Backbone vector

DNA

15.0 μg

4.0 μg × (n + 1)

Buffer

5.0 μL

5.0 μL

BSA

0.5 μL

0.5 μL

Enzyme

1.0 μL

2.0 μL

Water

To 50.0 μL

To 50.0 μL

n = number of transfections

2. The pAd5CMVK-NpA shuttle vector containing the mTRAIL cDNA and adenovirus backbone vector containing adenovirus type 5 sequences, which have the E1 (E1A and E1B) genes deleted, are digested with either Pac I or Nhe I (Table 3). 3. Add 50 μL digestion reaction of each virus to its corresponding microcentrifuge tube. 4. Vortex for 2 s. 5. Add 25 μL of 2.5 M CaCl2. 6. Vortex for 2 s. 7. Incubate at room temperature for 25 min. 8. Change media on 60-mm plates to be transfected to 2 mL of 2% FBS/1% penicillin/streptomycin DMEM. 9. After incubation, add all of each sample to its corresponding plate in a slow spiral motion. 10. After 4 h, change media to 4 mL of 10% FBS/1% penicillin/streptomycin DMEM and incubate overnight at 37°C with CO2.

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11. The next day, aspirate the media and add 4 mL of 10% FBS/1% penicillin/streptomycin DMEM. 12. Incubate at 37°C with CO2. 13. On day 7, feed the cells with 1 mL of 2% FBS/1% penicillin/ streptomycin DMEM and continue incubating at 37°C with CO2. 14. On day 10, feed the cells with 1 mL of 2% FBS/1% penicillin/ streptomycin DMEM. 15. Check for the presence of plaques. If the plate is ready for harvest (i.e., >60% of the cells lifted), then progress to harvesting. To harvest, use a 5-mL sterile serological pipette and collect the cells and media. Transfer to a 15-mL conical sterile tube. Freeze the media lysate at −20°C (see Note 3). 3.4. Ad5-mTRAIL Purification from Lysates from Five Plates

1. Freeze/thaw collected cells and media from 150-mm plates transfected earlier. 2. Freeze/thaw two more times using a 37°C water bath and a dry ice/EtOH bath. 3. Centrifuge media lysate at 2,500 rpm (600g) for 10 min at 4°C. 4. Use 15 mL of 2% FBS/1% penicillin/streptomycin DMEM per plate for a total of 75 mL for five 150-mm plates. 5. Add supernatant from the media lysate to the media. 6. Remove media from plates and very gently add 15 mL of the 75 mL media plus virus to each plate. 7. Incubate at 37°C with 5% CO2 until at least 60% of the cells are lifted from the plate. 8. Using a 20-mL sterile serological pipette, collect the cells and media from the plates (five 150–mm plates). Transfer to two 50-mL conical centrifuge tubes. 9. Centrifuge at 1,500 rpm (217g) for 10 min at 4°C. 10. Remove the media without disturbing the cell pellet. 11. Very gently wash the cells with 5 mL of PBS. 12. Centrifuge at 1,500 rpm (217g) for 5 min at 4°C. 13. Remove the PBS without disturbing the pellet. 14. Resuspend the cells in 5 mL of 10 mM Tris-HCl. 15. Freeze cell lysate at −20°C.

3.5. Viral Particle Purification

1. Freeze/thaw the lysate three times using a 37°C water bath and a dry ice/EtOH bath. 2. Spin the lysate at 3,500 rpm (1,178 g) for 10 min.

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1. Add 3 mL of light CsCl to the centrifuge tube. 2. Add 3 mL of heavy CsCl very slowly under the light CsCl (see Note 4). 3. Add the clarified supernatant very gently against the side of the tube on top of the CsCl gradient. Supplement with 10 mM Tris-HCl to about 0.25 inches below the top of the tube. Make sure the tubes are balanced. 4. Spin in the ultracentrifuge at 20,000 rpm (33,735g) at 4°C for at least 3 h. 5. Pull the viral band (see Note 5). 6. Remove the cell debris at the top of the gradient, to about half an inch above the viral bands with a glass Pasteur pipette. 7. Place solution in waste bottle. 8. Use a 5-mL syringe with an 18-gauge 1-inch needle. Insert the needle through the side of the tube about four needle widths below the virus band. 9. Extract the virus in as small a volume as possible, leaving behind incomplete bands higher up the tube. 10. Dilute the band by raising the volume to 3 mL of 10 mM Tris-HCl in a snap-cap tube. 11. Place the syringe and needle inside the snap-cap tube and pull up the 10 mM Tris-HCl. The band must be diluted by 50% with 10 mM Tris-HCl (see Note 6).

3.5.2. Second CsCl Gradient

1. Use a Beckman SW60 or equivalent rotor, buckets, and tubes. 2. For 1-mL gradients, follow the above instructions to form the gradient and add the virus. 3. Spin in the ultracentrifuge at 20,000 rpm (30,195g) at 4°C overnight. 4. Pull the viral band as above using a 5-mL syringe. If not dialyzing, put the virus in a small, labeled snap-cap tube, or Eppendorf microcentrifuge tube. 5. Store at −80°C.

3.5.3. Dialysis

1. In the syringe, add 1 mL of 3% sucrose/PBS, about half the volume of the band (see Note 7). 2. Inject the contents of the syringe into a “Slide-A-Lyzer” dialysis cassette (Pierce). Using the 5-mL syringe, pull out the remaining air in the slide. 3. Place the dialyzer slide with a buoy in a bucket of 3% sucrose/ PBS. Use 1 L sucrose/PBS per slide.

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4. Change the sucrose/PBS once an hour, two times. A total of 3 L of 3% sucrose/PBS will be needed for each virus preparation. 5. To remove the virus from the dialyzer slide, use a 5-mL syringe and add 1–2 mL of air to the slide (see Note 8). 6. With syringe still in the Slide-A-Lyzer, extract the virus (see Note 9). 7. Place virus in a snap-cap tube. 3.5.4. For Glycerol Stock

1. Transfer 250 mL of virus into a 2-mL screw cap tube. 2. Label the tube with virus name. 3. Add approximately 300 mL of 50% glycerol/BSA to the virus (see Note 10). 4. Turn the spectrophotometer on and allow it to warm up. 5. Prepare a blank of 13 μL of sucrose/PBS in 987 μL of 10 mM Tris. 6. Dilute the virus to 1:75 dilution (13 μL of virus in 987 μL of 10 mM Tris). 7. Measure the absorbance at 260 nm wavelength (see Note 11). 8. Dilute in 3% sucrose/PBS or glycerol/PBS to the virus to 1.0 × 1012 particles/mL.

3.5.5. Target Values

1. 3% Sucrose/PBS preps: 1012 particles/mL for the working stock (between 1 × 1012 and 1.5 × 1012 particles/mL is acceptable). 2. Glycerol/BSA stocks: 3 × 1012 particles/mL (between 3 × 1012 and 4 × 1012 particles/mL is acceptable). 3. Aliquot and freeze at −80°C (sucrose/BSA) or −20°C (glycerol/BSA). For 3% sucrose/PBS preps: 1 mL, 500 μL, and 200 μL aliquots.

3.6. Quantitation of Viral Titer

1. Viral titer is quantitated using the Adeno-X rapid titer kit (Clontech), which is a hexon-specific antibody-based assay. The hexon protein is encoded by the adenoviral genome and is an essential component of the adenoviral capsid required for adenoviral replication; however, hexon protein expression depends on the E1 gene product. Therefore, infectious activity can only be measured in E1 transcomplementing cell types, such as HEK 293, because only the infected cells will produce the hexon protein. 2. Recombinant adenovirus is screened for replication competent virus by A549 plaque assay. A typical titering schedule is listed in Table 4.


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Table 4 Typical adenovirus titering schedule Monday

Tuesday

Wednesday

Thursday

Week #1

Prepare 24-well plates

Make and plate A549 cells: 1st dilutions overlay (day 1)

Week #2

A549 Cells: 2nd overlay (day 6)

Week #3 A549 cells: 3rd overlay (day 11)

Friday

Read titer (day 12)

Table 5 Sample dilution series Dilution Type of media Amount of media

Amount to transfer

Sample

—

—

15 μL

10−2

2% MEM

1485 μL (742.5 μL × 2)

15 μL

10−4

2% MEM

1485 μL (742.5 μL × 2)

15 μL

−6

10

2% DMEM

1485 μL (742.5 μL × 2)

15 μL

10−8

2% DMEM

1485 μL (742.5 μL × 2)

150 μL

−9

10

2% DMEM

1350 μL (675 μL × 2)

150 μL

10−10

2% DMEM

1350 μL (675 μL × 2)

N/A

3. The human carcinoma airway epithelial cell line, A549, is used to determine wild-type adenovirus pfu/mL. Prepare 24-well plates the day before the start of the titers by adding A549 cells at 5 × 104 cells/well (see Note 12). 3.6.1. Viral Dilution Preparation

1. For each virus to be titered, label a set of small snap-cap tubes. The dilutions vary from virus to virus depending on which dilution needs to be plated. The starting concentration of virus should be 1.0–1.5 × 1012 particles/mL, where the dilutions are usually 10−2, 10−4, 10−6, 10−8, 10−9, and 10−10. The 10−4 dilution will be plated on A549 cells. 2. Add media (MEM with 2% FBS and 1% penicillin/streptomycin) into tubes according to Table 5. 3. To plate the dilutions onto the A549 cells, transfer 500 μL from the 10−4 dilution tube to the tube containing 2 mL of MEM with 2% FBS and 1% penicillin/streptomycin. Aspirate

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the media from six wells of cells, and add 200 μL to each well. Repeat on another six wells. 4. Incubate cells overnight at 37°C with 5% CO2. 5. Overlay cells at 24 h. 3.6.2. Overlay Schedule

1. The plates of A549 cells are to be overlaid three times according to the schedule below. The solutions for the first and second overlays are the same, whereas the third overlay contains Neutral Red dye to stain living cells (Table 6). 2. First and second overlay media contains 1.6% Agar Noble and MEM with 1% penicillin/streptomycin and 8% FBS. 3. Microwave the 1.6% Agar Noble (it is a gel at room temperature) for approximately 4 min until it liquefies. Place the Agar Noble in a 45°C water bath to cool for ~1 h before using. 4. Warm up the MEM with FBS and penicillin/streptomycin at 37°C. 5. Calculate the amount of media needed for the number of plates to be overlaid (see chart above). Aliquot media in 50-mL conical tubes. For the first and second overlays, the composition is 50% MEM/FBS/penicillin/streptomycin media and 50% Agar Noble. 6. Get the plates to be overlaid from the incubator, because the Agar Noble will not stay liquid for more than approximately 10–15 min after it is out of the water bath. 7. Calculate the amount of Agar Noble needed per 50-mL conical tube. Add Agar Noble to conical tube containing MEM/ FBS/penicillin/streptomycin media (one conical tube at a time), mix contents. Remove media from the 24-well plates (see Note 13). 8. Allow the plates to sit until the Agar Noble has solidified. 9. Place plates back into incubator.

Table 6 Typical schedule for A549 overlays to detect replicationcompetent adenovirus (RCA) A549 overlay

Day

Amount/well

Amount/24-well plate

1st

1

1,000 μL

24 mL

2nd

6

500 μL

12 mL

3rd w/neutral red

11

300 μL

7.2 mL


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10. The third overlay media contains 1.6% Agar Noble, MEM with 1% penicillin/streptomycin and 6% FBS, and 1.4% Neutral Red. 11. Prepare Agar Noble and media the same way as described above, except the third overlay media consists of 47% MEM/ FBS media, 3% Neutral Red, and 50% Agar Noble. Calculate amount of Neutral Red needed per 50-mL tube. Filter the dye using a 5-mL sterile syringe and the 0.45-μm filter. Add dye immediately before adding the Agar Noble one tube at a time. Neutral Red will precipitate in the MEM/FBS media. 3.6.3. Analyze Results

1. Look at the plates under the inverted microscope at 12 days. You should see an even cell monolayer. 2. Estimate the extent of cell death. If the percentage of death cells is greater than 40%, the wild-type titer determination should be repeated at a higher dilution (10−5). 3. Plaques in A549 cells appear as a clear, almost perfectly round circle of dead cells. They can often be seen as clear circles on the backside of the plate. 4. If any plaques are seen, record the total number of plaques. For example, if there were 3 plaques in 12 wells at the 10−3 dilution, multiply the number of plaques times two (500 μL plated on 12 wells), making the titer 6 × 103 pfu/mL (see Note 14). 5. If no foci are seen, the titer is reported as 95% as assessed by propidium iodide exclusion.

3.8. In Vitro Killing of Tumor Cell Lines with Adenoviral Vectors

1. Tumor cell sensitivity to Ad5-mTRAIL is assayed using the following procedure. Renca cells are added to 96-well plates (2 × 104 cells/well) in complete medium, and then allowed to adhere for at least 6 h before infection. Ad5-mTRAIL is added at the appropriate MOI (pfu/cell) in 0.1 mL RPMI containing 2% FBS for 4 h. Cells are washed with PBS, and then 0.2 mL complete RPMI is added to each well for the duration of the experiment. Cell death is determined after 24 h by crystal violet staining as described (26). Briefly, plates are gently inverted to remove excess media, washed with PBS to remove any nonadherent cells, and then gently blotted on a paper towel. Crystal violet (0.1 mL/well) is then added for 5 min at room temperature. The plates are gently washed with water until the water is clear. The plates are again gently blotted on paper towels to remove any remaining water in the wells. Finally, deoxycholic acid (0.1 mL/ well) is added to each well, and the plate is gently rocked/ shaken for at least 15 min at room temperature to solubilize the stained cells remaining in each well. The plate is read on a microtiter plate reader at 590 nm, and the results are presented as percent cell death: [1 – (OD cells treated/OD cells not treated)] × 100. An example of the results produced is shown in Fig. 3.

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Griffith 100

% Cell death

80

60

40

20

0 30

100

300

1000

3000

M.O.I. (pfu/cell)

Fig. 3. Renca cells (2 × 104 cells/well) were added to a 96-well flat-bottom plate, and the cells were infected at the indicated Ad5-mTRAIL MOI (pfu/cell). Cells were then stained with crystal violet after 24 h to measure cell death, and each symbol is an average of three wells.

Fig. 4. Measurement of phosphatidylserine (PS) externalization and propidium iodide (PI) exclusion on Renca tumor cells infected with Ad5-mTRAIL. Renca tumor cells were infected with 1,000 pfu/cell Ad5-mTRAIL, and then cultured for 18 h before staining with FITC–annexin V and PI and analyzed by flow cytometry. The percentage of cells in each quadrant is indicated in the upper right corner.

To specifically assess for apoptotic cell death, Renca tumor cell targets are seeded in 24-well plates and infected with Ad5mTRAIL as described in Subheading 3.3.1 totic cell death is measured by flow cytometry using FITC-conjugated annexin


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V (R&D Systems, Minneapolis, MN) and propidium iodide (Sigma) as described (27). Briefly, cells are removed from the plate, washed twice with cold PBS, and then resuspended in 100 μL annexin V binding buffer. FITC-conjugated annexin V (5 μL) and propidium iodide (5 μL) is added to the cells, which are incubated in the dark for 15 min at room temperature. After the incubation and without washing the cells, an additional 400 mL of annexin V binding buffer is added to the cells. The cells are analyzed on a flow cytometer within 1 h of completing the cell staining. An example of the results produced is shown in Fig. 4.

4. Notes 1. This describes a recombinant adenovirus encoding the murine tnfsf10 (mTRAIL) cDNA (accession #U37522). We have also produced a similar virus encoding the human TNFSF10 (hTRAIL) cDNA (accession #U37518), which was produced via the same procedure (8, 9, 25). 2. If the thermal cycler is not equipped with a heated cover, overlay each reaction with 50 mL of DNase-, RNase-, and protease-free mineral oil. 3. If the cells are not ready, keep checking the plate daily for the presence of plaques. Do not keep plates longer than 15 days. If there is no growth at day 15, harvest and transfer lysate and media to a single 150-mm plate. 4. Always take 4–6 mL of heavy CsCl in the pipette so no air bubbles disrupt the gradient. 5. The viral band is the bottom white colored band that is present where the heavy and light CsCl meet. 6. If you started with 5- or 10-plate lysates, dilute your pulled band to only 2 mL. 7. If the virus is too concentrated, it will precipitate out. 8. Use a different port from step 7.5.2. 9. Note the total volume of virus in the syringe. Subtract 0.4 mL from the total volume. 10. For very thick bands, add 500–1,000 mL. 11. OD260 = 1.1 × 1012 particles/mL. Calculate the particle per milliliter concentration taking into consideration the 1:75 dilution factor. 12. Cells should be 50–70% confluent at time of use. 13. Do not remove media from more than six wells at a time. SLOWLY add MEM/Agar Noble to the wells. You do not

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want cells to be without media for too long. Add the Agar Noble to the next conical tube and continue overlaying as needed. The Agar Noble can be placed back into the water bath between uses, so it will stay liquid. 14. If a viral prep was diluted to a concentration of 1012 before titering, the initial dilution must be taken in consideration when calculating the final titer. 15. Because the principal viral vector for cancer gene therapy is adenovirus, the success of adenoviral-based therapies is primarily dictated by CAR recognition (15). CAR is ubiquitously expressed in most benign epithelial tissues. Yet, marked variations in CAR levels have been demonstrated using different cancer cell lines of the same tissue origin (28). For example, Okegawa et al. examined three human prostate cancer cell lines and found that CAR is downregulated in the tumorigenic cell line PC-3 compared with DU-145 and LNCaP (29). Despite these variations, adenoviral infection of human prostate tumor cells appears to be very efficient, and numerous reports have evaluated the use of recombinant adenoviral vectors in the treatment of prostate cancer (30–37). 16. It is essential to use nonreducing sample buffer to ensure the maintenance and detection of TRAIL multimers in the SDSPAGE/immunoblot. The use of a reducing sample buffer will result in most/all of the TRAIL produced showing up as a monomer on the blot. 17. The listed primary antibody incubation time is what we have found to be a minimum time for detection in Western blot. Longer periods of time can easily be used, for example, overnight at 4°C. It is critical, however, to keep the membrane wet during the entire incubation time. 18. Be sure to thoroughly wash the membrane after the secondary antibody step. Any nonspecifically bound secondary antibody can react with the chemiluminescent reagent and increase the background noise on the blot. 19. Whereas surface expression of TRAIL is essential for the tumoricidal activity of Ad5-TRAIL, the sensitivity of the infected cell to TRAIL-induced apoptosis is also an important component of this phenomenon. In general, TRAILsensitive tumor cells will also be killed by Ad5-TRAIL, even though recent reports have shown that cells resistant to soluble TRAIL can be killed after adenoviral delivery of fulllength TRAIL (38). Thus, the results presented in Fig. 2 were generated using the TRAIL-resistant human melanoma cell line WM 164 (3). Alternatively, TRAIL-sensitive cells can be treated with the pan-caspase inhibitor, zVAD-fmk, to block apoptosis and allow cell surface analysis of TRAIL.


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Acknowledgments I thank Dr. Troy K. Kemp, Tamara Kucaba, and Dr. Rebecca L. VanOosten for their technical assistance; and Maria Scheel, Dalyz Ochoa, and the University of Iowa Gene Transfer Vector Core for virus production. This work was supported by the National Cancer Institute (CA109446).

References 1. Wiley S.R., Schooley K., Smolak P.J., et al. (1995) Identification and characterization of a new member of the TNF family that induces apoptosis. Immunity 3:673–682. 2. Pitti R.M., Marsters S.A., Ruppert S., Donahue C.J., Moore A., Ashkenazi A. (1996) Induction of apoptosis by Apo-2 ligand, a new member of the tumor necrosis factor cytokine family. J Biol Chem 271:12687–12690. 3. Griffith T.S., Chin W.A., Jackson G.C., Lynch D.H., Kubin M.Z. (1998) Intracellular regulation of TRAIL-induced apoptosis in human melanoma cells. J Immunol 161:2833–2840. 4. Walczak H., Miller R.E., Ariail K., et al. (1999) Tumoricidal activity of tumor necrosis factorrelated apoptosis-inducing ligand in vivo. Nat Med 5:157–163. 5. Ashkenazi A., Pai R.C., Fong S., et al. (1999) Safety and antitumor activity of recombinant soluble Apo2 ligand. J Clin Invest 104:155–162. 6. Gliniak B., Le T. (1999) Tumor necrosis factor-related apoptosis-inducing ligand’s antitumor activity in vivo is enhanced by the chemotherapeutic agent CPT-11. Cancer Res 59:6153–6158. 7. Chinnaiyan A.M., Prasad U., Shankar S., et al. (2000) Combined effect of tumor necrosis factor-related apoptosis-inducing ligand and ionizing radiation in breast cancer therapy. Proc Natl Acad Sci USA 97:1754–1759. 8. Griffith T.S., Anderson R.D., Davidson B.L., Williams R.D., Ratliff T.L. (2000) Adenoviralmediated transfer of the TNF-related apoptosisinducing ligand/Apo-2 ligand gene induces tumor cell apoptosis. J Immunol 165:2886–2894. 9. Griffith T.S., Broghammer E.L. (2001) Suppression of tumor growth following intralesional therapy with TRAIL recombinant adenovirus. Mol Ther 4:257–266. 10. Denis L.J. (2000) The role of active treatment in early prostate cancer. Radiother Oncol 57:251–258.

11. Stone N.N., Stock R.G. (2000) Prostate brachytherapy in patients with prostate volumes >/= 50 cm(3): dosimetic analysis of implant quality. Int J Radiat Oncol Biol Phys 46:1199–1204. 12. Steiner M.S., Gingrich J.R. (2000) Gene therapy for prostate cancer: where are we now? J Urol 164:1121–1136. 13. Herman J.R., Adler H.L., Aguilar-Cordova E., et al. (1999) In situ gene therapy for adenocarcinoma of the prostate: a phase I clinical trial. Hum Gene Ther 10:1239–1249. 14. Tripathy S.K., Black H.B., Goldwasser E., Leiden J.M. (1996) Immune responses to transgene-encoded proteins limit the stability of gene expression after injection of replicationdefective adenovirus vectors. Nat Med 2:545–550. 15. Bergelson J.M., Cunningham J.A., Droguett G., et al. (1997) Isolation of a common receptor for Coxsackie B viruses and adenoviruses 2 and 5. Science 275:1320–1323. 16. Rubin S.A., Rorke L.B. Adenoviral vaccines. In: Plotkin M, ed. Vaccines. Philadelphia: W.B. Saunders; 1990:492–512. 17. Akli S., Caillaud C., Vigne E., et al. (1993) Transfer of a foreign gene into the brain using adenovirus vectors. Nat Genet 3:224–228. 18. Baratin M., Ziol M., Romieu R., et al. (2001) Regression of primary hepatocarcinoma in cancer-prone transgenic mice by local interferon-gamma delivery is associated with macrophages recruitment and nitric oxide production. Cancer Gene Ther 8:193–202. 19. Cordier L., Duffour M.T., Sabourin J.C., et al. (1995) Complete recovery of mice from a pre-established tumor by direct intratumoral delivery of an adenovirus vector harboring the murine IL-2 gene. Gene Ther 2:16–21. 20. Le Gal La Salle G., Robert J.J., Berrard S., et al. (1993) An adenovirus vector for gene transfer into neurons and glia in the brain. Science 259:988–990. 21. Rosenfeld M.A., Yoshimura K., Trapnell B.C., et al. (1992) In vivo transfer of the human

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Griffith cystic fibrosis transmembrane conductance regulator gene to the airway epithelium. Cell 68:143–155. Timme T.L., Hall S.J., Barrios R., Woo S.L., Aguilar-Cordova E., Thompson T.C. (1998) Local inflammatory response and vector spread after direct intraprostatic injection of a recombinant adenovirus containing the herpes simplex virus thymidine kinase gene and ganciclovir therapy in mice. Cancer Gene Ther 5:74–82. Murphy G.P., Hrushesky W.J. (1973) A murine renal cell carcinoma. J Natl Cancer Inst 50:1013–1025. Kayagaki N., Yamaguchi N., Nakayama M., et al. (1999) Expression and function of TNFrelated apoptosis-inducing ligand on murine activated NK cells. J Immunol 163:1906–1913. Anderson R.D., Haskell R.E., Xia H., Roessler B.J., Davidson B.L. (2000) A simple method for the rapid generation of recombinant adenovirus vectors. Gene Ther 7:1034–1038. Flick D.A., Gifford G.E. (1984) Comparison of in vitro cell cytotoxic assays for tumor necrosis factor. J Immunol Methods 68:167–175. Martin S.J., Reutelingsperger C.P., McGahon A.J., et al. (1995) Early redistribution of plasma membrane phosphatidylserine is a general feature of apoptosis regardless of the initiating stimulus: inhibition by overexpression of Bcl-2 and Abl. J Exp Med 182:1545–1556. Hemminki A., Alvarez R.D. (2002) Adenoviruses in oncology: a viable option? BioDrugs 16:77–87. Okegawa T., Li Y., Pong R.C., Bergelson J.M., Zhou J., Hsieh J.T. (2000) The dual impact of coxsackie and adenovirus receptor expression on human prostate cancer gene therapy. Cancer Res 60:5031–5036. Eastham J.A., Hall S.J., Sehgal I., et al. (1995) In vivo gene therapy with p53 or p21 adenovirus for prostate cancer. Cancer Res 55:5151–5155.

31. Hall S.J., Mutchnik S.E., Yang G., et al. (1999) Cooperative therapeutic effects of androgen ablation and adenovirus-mediated herpes simplex virus thymidine kinase gene and ganciclovir therapy in experimental prostate cancer. Cancer Gene Ther 6:54–63. 32. Nasu Y., Bangma C.H., Hull G.W., et al. (1999) Adenovirus-mediated interleukin-12 gene therapy for prostate cancer: suppression of orthotopic tumor growth and preestablished lung metastases in an orthotopic model. Gene Ther 6:338–349. 33. Anello R., Cohen S., Atkinson G., Hall S.J. (2000) Adenovirus mediated cytosine deaminase gene transduction and 5-fluorocytosine therapy sensitizes mouse prostate cancer cells to irradiation. J Urol 164:2173–2177. 34. Shariat S.F., Desai S., Song W., et al. (2001) Adenovirus-mediated transfer of inducible caspases: a novel “death switch” gene therapeutic approach to prostate cancer. Cancer Res 61:2562–2571. 35. Cao G., Su J., Lu W., et al. (2001) Adenovirusmediated interferon-beta gene therapy suppresses growth and metastasis of human prostate cancer in nude mice. Cancer Gene Ther 8:497–505. 36. Katner A.L., Hoang Q.B., Gootam P., et al. (2002) Induction of cell cycle arrest and apoptosis in human prostate carcinoma cells by a recombinant adenovirus expressing p27(Kip1). Prostate 53:77–87. 37. Flynn V., Jr., Ramanitharan A., Moparty K., et al. (2003) Adenovirus-mediated inhibition of NF-kappaB confers chemo-sensitization and apoptosis in prostate cancer cells. Int J Oncol 23:317–323. 38. Voelkel-Johnson C., King D.L., Norris J.S. (2002) Resistance of prostate cancer cells to soluble TNF-related apoptosis-inducing ligand (TRAIL/Apo2L) can be overcome by doxorubicin or adenoviral delivery of fulllength TRAIL. Cancer Gene Ther 9:164–172.

Chapter 18 Silencing Epidermal Growth Factor Receptor by RNA Interference in Glioma Chunsheng Kang, Peiyu Pu, and Hao Jiang Summary Glioblastoma multiforme (GBM) can arise de novo or progress from a lower to higher grade and can possess a series of genetic alterations and dynamic progressions, which have been correlated with the molecular pathology of GBM. Epidermal growth factor receptor (EGFR) has been shown to be overexpressed in a variety of tumors and is one of the important mediators responsible for the development of high-grade gliomas, especially in primary glioblastomas. Most recently, RNA interference (RNAi), in which double-stranded RNA (dsRNA) induces sequence-specific degradation of the targeting messenger RNA (mRNA), has been extensively developed and studied. RNAi is able to silence the targeted gene expression more efficiently and specifically. In the present study, we silence the EGFR expression using two separate short interfering RNAs (siRNAs) targeting the extracellular ligand-binding domain and intracellular tyrosine kinase domain, respectively. We demonstrate that suppression of EGFR expression, by using either antisense or siRNA approaches, inhibits U251 glioblastoma cell growth in vitro and in vivo, and siRNA seems to be more effective than the antisense approach. Key words: EGFR, glioma, invasion, proliferation, siRNA.

1. Introduction Gliomas are the most common primary tumors in the brain. Current therapeutic modalities include surgical resection, radio surgical/ external irradiation, chemotherapy, and biological therapy. Most patients with high-grade gliomas (anaplastic gliomas and glioblastoma multiformes [GBMs]) receive radiation and chemotherapy regardless of the extent to which the visible tumor has been removed. However, in a meta-analysis of 12 randomized clinical trials, the overall survival rate of high-grade glioma patients after Wolfgang Walther and Ulrike S. Stein (eds.), Methods in Molecular Biology, Gene Therapy of Cancer, vol. 542 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-561-9_18

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surgical resection is 40% at 1 year after surgical resection and is only slightly higher (46%) after combined treatment with radiotherapy and chemotherapy (1). Epidermal growth factor receptor (EGFR) is overexpressed in a variety of tumors and has been demonstrated to be one of the important mediators responsible for the development of malignant gliomas, especially in primary glioblastomas (2). EGFR amplification, overexpression, or mutation, is the early and major molecular event in glioblastomas. Previous studies demonstrate that EGFR is overexpressed in 90% of glioblastomas (3). Antisense approaches targeting EGFR significantly inhibit not only the EGFR expression but also the proliferation of malignant glioma cells in vitro and in vivo (4). Therefore, EGFR is an important therapeutic target for molecular therapy of malignant gliomas. RNA interference (RNAi), in which double-stranded RNA (dsRNA) induces sequence-specific degradation of the targeting messenger RNA (mRNA), has been widely studied and used (5). RNAi technology can silence the targeted gene expression more efficiently and specifically (6, 7). Short interference RNA (siRNA) by plasmidbased methodology will trigger RNAi by enzymatic dicing. To date, a few reports have been published on the treatment of malignant gliomas using an RNAi-based approach. In the present study, we silence the EGFR expression using two separate siRNAs targeting the extracellular ligand-binding domain and intracellular tyrosine kinase domain, respectively. Furthermore, suppression of EGFR expression, by using either an antisense or an siRNA approach, inhibits the U251 glioblastoma cell growth in vitro and in vivo, and siRNA seems to be more effective than the antisense approach.

2. Materials 2.1. Cell Lines and Transfection

1. TJ905 glioblastoma cell line has been established and characterized in the laboratory of Neuro-Oncology, Tianjin Neurological Institute, Tianjin, China. Human glioblastoma (see Note 1). U251-MG cells are obtained from Institute of Biochemistry and Cell Biology, Chinese Academy of Sciences. 2. Dulbecco’s Modified Eagle’s Medium (DMEM) (Gibco/BRL, Bethesda, MD, USA). 3. Fetal bovine serum (FBS, HyClone, Ogden, UT, USA). 4. Trypsin (0.25%). 5. Ethylenediamine tetraacetic acid (EDTA) (Gibco/BRL). 6. The short hairpin RNA (shRNA) expression plasmid psiRNANeoG2 plasmid and Escherichia coli GT116 competent cells are from InvivoGen (San Diego, CA, USA).

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7. Lipofectamine (Invitrogen, Carlsbad, CA, USA). 8. Teflon cell scrapers (Fisher, Springfield, NJ, USA). 2.2. Sodium Dodecyl Sulfate (SDS)-Polyacrylamide Gel Electrophoresis (PAGE)

1. Extraction buffer: 1% Nonidet P-40 lysis buffer (20 mM Tris, pH 8.0, 137 mM NaCl, 1% Nonidet P-40, 10% glycerol, 1 mM CaCl2, 1 mM MgCl2, 1 mM phenylmethylsulfonyl fluoride, 1 mM sodium fluoride, 1 mM sodium orthovanadate), add a protease inhibitor mixture to the buffer. 2. Running buffer: 25 mM Tris (1.51 g Tris), 250 mM glycine (9.4 g glycine), 0.1% SDS (0.5 g SDS), adjust volume to 1 L with dH2O. 3. Transfer buffer: 25 mM Tris (3.03 g Tris), 192 mM glycine (14.4 g glycine), 20% methanol (200 mL methanol), 0.02% SDS (0.2 g SDS), adjust volume to 1 L with dH2O. 4. TBST: NaCl (29.22 g NaCl), Tris (2.42 g Tris), add dH2O to 800 mL, adjust pH to 7.5, and adjust the volume to 1 L with dH2O, add 0.1 mL Tween-20 to 100 mL TBS. 5. Blocking solution: 5% nonfat dry milk in TBST. 6. Antibody buffer: 4% nonfat dry milk in TBST. 7. Millipore PVDF membrane (Millipore, Bedford, MA) and 3M Whatman chromatography paper (Whatman, Maidstone, UK). 8. Antibody for EGFR and secondary antimouse IgG conjugated to horseradish peroxidase (Santa Cruz Biotechnology, Santa Cruz, USA). 9. Enhanced chemiluminescent (ECL) reagents from SuperSignal protein detection kit (Pierce, Rockford, IL, USA).

2.3. Stripping and Reprobing Blots for EGFR

1. Stripping buffer: 62.5 mM Tris-HCl, pH 6.8, 2% (w/v) SDS. Store at room temperature. Warm to working temperature of 70° and add 100 mM β-mercaptoethanol. 2. Wash buffer: 0.1% (w/v) BSA in TBS-T. 3. Primary antibody: Anti-EGFR and β-actin (Santa Cruz Biotechnology).

2.4. Flow Cytometry for Cell Cycle

1. FACS flow cytometer (Becton–Dickinson, San Jose, CA, USA).

2.5. Immunohistochemistry and Immunofluorescence Staining

1. Microscope cover slips (40 × 40 × 0.15 mm) from Zhongshan Corp., Beijing, China.

2. Propidium iodide (PI) staining solution: PI (100 mg/mL, Sigma, St. Louis, MO, USA), Triton X-100 (1.0%), NaCl (0.9%).

2. Ca2+-, Mg2+-free phosphate-buffered saline (CMF-PBS): Prepare 10X stock with 1.37 M NaCl, 27 mM, KCl, 100 mM Na2HPO4, 18 mM KH2PO4 (adjust to pH 7.4 with HCl if necessary) and autoclave before storage at room temperature. Prepare working solution by dilution of one part with nine parts water.

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3. Paraformaldehyde (Sigma): Freshly prepare a 4% (w/v) solution in PBS for each experiment. The solution needs to be carefully heated (use a stirring hot plate in a fume hood) to dissolve, and then cool down to room temperature before use. 4. Permeabilization solution: 0.5% (v/v) Triton X-100 in PBS. 5. Antibody dilution buffer: 3% (w/v) BSA in PBS. 6. Secondary antibody: Antimouse IgG conjugated to Cy3 (Zhongshan Corp., Beijing, China). 7. Nuclear stain: 300 nM 4,6-diamidino-2-phenylindole (DAPI) in water. 2.6. Transwell Invasion Assay

1. The transwell filters (Costar, Corning, NY, USA) were coated on the upper surface with polycarbonic membrane (diameter 6.5 mm, pore size 8 μm). 2. Matrigel basement membrane matrix (Becton Dickinson, Lincoln Park, NJ, USA).

3. Methods 3.1. Cell Culture

Cell are cultured in DMEM supplemented with 10% FBS at 37° in 5% CO2. Cells are subcultured every 2–3 days using trypsin (0.25%) and 1 mM EDTA.

3.2. The siRNA Expression Plasmid Construction

1. Two siRNA targeting sequences are selected, corresponding to the open-reading frame of the human EGFR gene (NM005228, 247–3879) (Fig. 1) (see Note 2). One is targeting the extracellular ligand-binding domain (clone 516), the sequence for 516 clone is 5¢-TGCCTTAGCAGTCTTATCTA-3¢, with GC content 38.10%. The other sequence is targeting the intracellular tyrosine kinase domain (clone 2400). The sequence for the 2400 clone is 5¢-AGGAATTAAGAGAAGCAACAT-3¢, with GC content 33.33%. Both targeting sequences are Blasted to avoid silencing other unrelated genes. 2. The siRNA-Neo2 plasmid (InvivoGen) is used to express short hairpin RNA (siRNA) in cells. The Acc651/HindIII double-digesting protocol is used for constructing the siRNA plasmid in psiRNA-neo2. Two oligonucleotides are synthesized. The 5¢ terminal contains the Acc651 restriction enzyme site and the 3¢ terminal contains the Hind III restriction enzyme site. AN refers to the selected targeting


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Fig. 1. Schematic example of plasmid construction for siRNA constructs. Plasmid construction: downstream Acc651, upstream Hind III.

sequences, N refers to its antisense sequences, and TN refers to its sense sequences: 5 ¢ -GTACCTC-AN(21)-TCAAGAG-N(21)T-TTTTTGGAAAGCT-3¢ 3 ¢ -CATGGAG-TN(21)-AGTTCTC-N(21)A-AAAAACCTTTTCGA-5¢. 3. Equal amounts of the forward and reverse oligonucleotides for each designed siRNA are mixed, denatured, and annealed to form the double-stranded siRNA insert. Each insert is ligated into plasmid psiRNA-NeoG2 by T4 DNA ligase at Acc651/HindIII site of psiRNA-NeoG2 (Fig. 1), and named as psiRNA-NeoG2-516 and psiRNA-NeoG2-2400, respectively (Fig. 2). Ligated psiRNA-NeoG2 constructs are transformed into E. coli GT116 competent cells (InvivoGen). Recombinant plasmid siRNA (psiRNA) plasmids are selected, extracted, and purified. The sequences of the siRNA inserts are verified by DNA sequencing. The control siRNA plasmid is also provided by InvivoGen, and contained the mismatched siRNA sequences. 3.3. Antisense EGFR Constructs

The antisense orientation of the EGFR complementary DNA (cDNA) fragment (552 bp) carryied in the pactQsR3 plasmid (kindly provided by Dr Laura Beguinot, Italy) (8) is complementary to the 3¢ coding region of EGFR mRNA. The pcDNA3 plasmid

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Fig. 2. Electrophoresis of p-anti-hEGFR, psiRNA-NeoG2-516, and psiRNA-NeoG2-2400. 1,12: low molecular weight DNA marker DL2000, XbaI/XhoI double enzyme-digested donor plasmid, antisense EGFR donor plasmid, XbaI/XhoI double enzyme-digested p-anti-hEGFR, p-anti-hEGFR; 6: pcDNA3; 7,8: high molecular weight DNA marker λ/Hind III; 9: psiRNANeoG2-516; 10: psiRNA-NeoG2-2400; 11: psiRNA-NeoG2 empty vector.

is double-digested with XbaI/XhoI, electrophoresed in 2% low melting point (LMP) agarose in 1X TAE solution for 20 min. The fragment of interest is recovered using Gel Recovery Kit (Takara, Dalian, China), and the antisense fragment of EGFR is then subcloned into pcDNA3 using a T4 Ligation Kit (Takara) at 16° overnight and the sequencing is verified, and named p-antihEGFR (Fig. 2). 3.4. Transfection of siRNA and Antisense Constructs (see Note 3)

1. In a 6-well plate or 35-mm tissue culture dishes, 2 × 105 cells/ well are plated and grown until they are 50–80% confluent. The medium is changed to serum-free DMEM before transfection. 2. Prepare the following solutions in 2-mL sterile tubes: Solution A: For each transfection, dilute 2 mg of plasmid DNA in 100 mL of serum-free DMEM. Solution B: For each transfection, dilute 10 mL of Lipofectamine Reagent (Invitrogen) in 100 mL serum-free DMEM. Combine the two solutions, mix gently, and incubate at room temperature for 15–45 min. Wash the cells twice with 2 mL of serum-free DMEM. 3. For each transfection, add 800 mL serum-free DMEM to each tube containing the lipid–DNA complexes. Mix gently and overlay the 1 mL lipid–DNA complex solution onto the washed cells. Incubate the cells for 4–8 h at 37° in a CO2 incubator. 4. Remove the transfection mixture and replace with normal growth medium for another 24 h. Approximately 24 h after transfection, exchange the medium to regular growth medium supplemented with 300 µg/mL of neomycin (G418, dissolved in Hanks solution at 50–100X stock solution and filtered through 0.22-mm filters for sterilization). Cells are subcultured at a 1:5 dilution in 300 µg/mL G418containing medium.


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5. Keep exchanging the medium every day for 7–10 days, until all cells have died in the dish of the negative control (nontransfected cells). Individual cell clones can be found in the transfected dishes. Carefully pick and mark large, healthy, and well-separated clones using a cloning separator, expand up to more than 1 × 108 cell pools in G418-containing medium for further analysis. 3.5. Detection of EGFR Downregulation After Transfection with siRNA and Antisense EGFR Constructs 3.5.1. Western Blotting Analysis

1. Parental and transfected cells are washed with ice-cold PBS three times (Fig. 3). Cells are then solubilized in extraction buffer solution. Place the above solution on ice for 30 min and suck–blow with a fine syringe (No. 10) for at least 20 times. Homogenates are centrifuged at 20,000×g for 15 min at 4°. Carefully transfer the supernatant into a fresh tube. Centrifuge the solution at 10,000×g for 2 min to remove the cell debris. Protein concentrations are determined by a bicinchoninic acid protein assay kit (Pierce). 2. Separate the proteins on the basis of molecular weight by SDSPAGE (in a standard stacking gel and an 8–10% separating gel). 3. Transfer the proteins from the gel to a PVDF membrane (Millipore) by tank electroblotting. Incubate with primary antibody against target protein (Santa Cruz Biotechnology), followed by incubation with HRP-conjugated secondary antibody (Zymed, South San Francisco, CA, USA). The specific protein is detected by using a SuperSignal protein detection kit (Pierce). The membrane is stripped and reprobed with an antibody against b-actin (Santa Cruz Biotechnology).

3.5.2. Immunofluorescence Staining

Cells are grown on glass coverslips in culture dishes overnight, washed with Ca2+-, Mg2+-free phosphate-buffered saline (CMFPBS) and fixed with freshly prepared 4% paraformaldehyde for 30 min at room temperature. Fixed cells are incubated with 0.3%

A

B

C

D

E EGFR(175kd)

Actin(42kd)

Fig. 3. Western blot analysis of EGFR expression in TJ905 glioma cells after transfection with p-anti-hEGFR, psiRNANeoG2-516, and psiRNA-NeoG2-2400 A. EGFR expression of parental TJ905 cells B. EGFR expression of TJ905 cells transfected with empty vector C. EGFR expression of TJ905 cells transfected with antisense EGFR p-anti-hEGFR D. EGFR expression of TJ905 cells transfected with siRNA expression plasmid psiRNA-NeoG2-516 E. EGFR expression of TJ905 cells transfected with siRNA expression plasmid psiRNA-NeoG2-2400.

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H2O2 and diluted goat serum for 30 min each at room temperature, and then with a 1:100 dilution of rabbit anti-EGFR polyclonal antibody (Santa Cruz Biotechnology) in the presence of 0.1% Triton X-100 and 0.5% BSA. After incubation at 4° overnight, a fluorescein isothiocyanate (FITC)-conjugated secondary antibody (1:50 dilution, Zymed) is added to the slides for 60 min at 37°. EGFR expression is examined by laser scanning confocal microscope (BioRad Radiance 2000 MP, Hercules, CA, USA) with excitation and emission wavelengths of 488 nm and 530 nm, respectively. 3.6. Evaluation of the Effect of siRNA and Antisense Constructs on Cell Proliferation and Invasion 3.6.1. Cell Cycle Analysis

Parental and transfected cells in the log phase of growth are harvested by trypsinization, washed with PBS, and fixed with absolute ethanol overnight (Fig. 4). Ethanol is removed by centrifugation at 800 rpm for 5 min at 4°. Cells are washed with PBS (0.01 M, pH 7.2) twice and incubated with 200 mL of 1 mg/mL RNase for 30 min at 37°. Nuclei of cells are stained with 800 mL of propidium iodide for an additional 30 min at 4°, avoiding light. A total of 10,000 nuclei are examined in a FACS flow cytometer (Becton–Dickinson) and DNA histograms are analyzed by Modifit software.

3.6.2. In Vitro Growth on Matrigel Matrix

Parental and transfected cells in the log phase of growth are washed in medium and seeded in duplicate in 24-well plates (1 × 105 cells/ well) precoated with 250 mL of Matrigel basement membrane matrix (Becton–Dickinson) (Fig. 5). After incubation for 36 h, photographs are taken at a 10× magnification with an Olympus camera attached to an IX70 inverted phase-contrast microscope.

3.6.3. Transwell Invasion Assay

Transwell filters (Costar) are coated on the upper surface of a polycarbonic membrane (diameter 6.5 mm, pore size 8 mm) with Matrigel (3.9 mg/mL, 60–80 mL) (Fig. 6). After incubation for

Fig. 4. Suppression of EGFR expression by antisense and psiRNA-NeoG2-2400 plasmids in U251 cells induces G2/M arrest and inhibits cell growth. The G2/M phase fraction of parental U251 cells and cells transfected with scrambled vector was 9.6% and 10.8%, respectively, whereas the percentage of antisense and siRNA plasmids transfected cells in G2/M phase was increased to 22.7% and 21.3%, respectively. The S phase fraction in parental U251 cells and cells transfected with scrambled vector was 32.3% and 31.6%, respectively, but decreased significantly in antisense and siRNA plasmid-transfected cells to 19.1% and 17.3%, respectively.


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Fig. 5. U251 glioma cell growth on extracellular matrix (ECM) after transfection with psiRNA-NeoG2-2400 and antisense plasmids. The attached growth pattern indicates normal cell growth, the spotted cell clustering signifies poor cell growth. The experiment was repeated in triplicate; representative pictures are shown after 36 h of incubation.

a

Invasion Capability

b

20 18 16 14 12

Control psiRNA-scr psiRNA-NeoG2-2400 p-anti-EGFR

10 8 6 4 2 0

Fig. 6. The invasive capability of U251 cells and cells transfected with psiRNA-scr, panti-EGFR, and psiRNA-NeoG2-2400 constructs was examined by a transwell invasion assay. Cell invasion was decreased in cells of panti-EGFR and psiRNANeoG2-2400 transfected groups as assessed by the number of cells invading into the lower surface of the polycarbonic membrane via Matrigel (one-way ANOVA, F = 12.067, p < 0.001).

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30 min at 37°, Matrigel becomes solidified and acts as the extracellular matrix (ECM) for tumor cell invasion analysis. Harvested cells (1 × 105 cells) suspended in 100 mL of serum-free DMEM are added into the upper compartment of the chamber. A total of 200 mL of conditioned medium from NIH3T3 cells is used as a source of chemoattractant and placed in lower compartment of the chamber. After 24 h of incubation at 37° in 5% CO2, the medium is removed from upper chamber. The noninvaded cells on the upper side of the chamber are scraped off with a cotton swab. The cells that had migrated from the Matrigel into the pores of the inserted filter are fixed with 100% methanol, stained with hematoxylin, mounted, and dried for 30 min at 80°. The number of cells invading through the Matrigel is counted in three randomly selected visual fields, each from the central and peripheral portion of the filter, using an inverted microscope at 200× magnification (Fig. 6a and b). Each test is repeated three times. 3.7. Identification of the Effect of EGFR siRNA In Vivo 3.7.1. Establishment of Subcutaneous Glioma Model

3.7.2. Gene Therapy with EGFR siRNA and Antisense Constructs

Six-week-old female immunodeficient nude mice (BALB/c-nu) are purchased from the animal center of the Cancer Institute of Chinese Academy of Medical Sciences, bred at the facility of laboratory animals, Tianjin University, and housed in microisolator individually ventilated cages with water and food. All experimental procedures are carried out according to the regulations and internal biosafety and bioethics guidelines of Tianjin Medical University and the Tianjin Municipal Science and Technology Commission. To ensure the homogeneity of the experimental results, we use the tumor-implantation method to create a subcutaneous glioma model. A direct plasmid–Lipofectamine mixture administration is used to simulate clinical application. Four mice are subcutaneously injected with 1 × 108 U251 cells, in a volume of 50 mL of PBS premixed with equal volume of Matrigel matrix. Mice are monitored daily, and three out of four mice form subcutaneous tumors. When the tumor size reaches approximately 5 mm in length, the tumors are surgically removed, cut into pieces of 1–2 mm3, and reseed individually into the other 40 mice. When the tumor size reaches approximately 5 mm in length in these mice, the following procedures are performed. The mice with subcutaneous tumors are randomly divided into U251 (control), psiRNA-scr (scrambled siRNA), p-anti-EGFR (antisense EGFR), and psiRNA-EGFR (EGFR siRNA) groups. In the p-anti-EGFR, psiRNA-scr, and psiRNA-EGFR groups, 10 mg of plasmid DNA in 10 mL of Lipofectamine are injected into subcutaneous tumors using a multisite injection manner. Mice in the U251 group receive 10 mL of PBS only. Another administration is conducted on the third day. The tumor volume is measured with a caliper, using the formula volume = length × width2/2 (Fig. 7a).


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Fig. 7. Compared with the group of PBS and psiRNA-scr, tumor growth in nude mice in the psiRNA-NeoG2-2400 and panti-EGFR groups was inhibited (p < 0.001) (a), EGFR expression of tumors was downregulated (b), apoptosis cells become obvious with TUNEL staining (c) (see Color Plates).

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3.8. Immunohistochemistry of Tumor Samples

After 28 days, mice are killed and paraffin-embedded tumor tissue sections are prepared for examination of EGFR, proliferating cell nuclear antigen (PCNA), and glial fibrillary acidic protein (GFAP) expression. Sections are dewaxed, treated with 3% H2O2 for 10 min, and incubated with appropriate antibodies (1:200 dilution) overnight at 4°. A biotinylated secondary antibody (1:200 dilution) is added at room temperature for 1 h, followed by the incubation with ABC–peroxidase for an additional 1 h. After washing with Tris buffer, the sections are incubated with 3,3¢ diaminobenzidine (DAB, 30 mg dissolved in 100 mL Tris buffer containing 0.03% H2O2) for 5 min, rinsed in water, and counterstained with hematoxylin (Fig. 7b). Apoptosis in tumor samples is detected by the terminal deoxynucleotide transferase-mediated dUTP nick-end labeling (TUNEL) method. Briefly, sections are dewaxed, incubated with blocking solution (0.3% H2O2 in double distilled water) for 30 min, and permeabilized with 0.1% Triton X-100 in PBS for another 2 min on ice. Apoptosis is detected using an in situ cell death kit (Roche, Indianapolis, IN, USA). Positive cells are visualized by fluorescent microscopy (Fig. 7c). The reaction mixture is incubated without enzyme in a control coverslide to detect nonspecific staining.

4. Notes 1. The tumor sample we used was from a male patient with a parietal glioblastoma, the tumor tissue obtained half an hour after surgical resection was minced mechanically by surgical scissors into 1-mm3 pieces and incubated in Way Mouth MAB 87/3 supplemented with 15% fetal calf serum at 37°, in 5% CO2. After 1 week of incubation, the tumor cells reached confluency and the cells were passaged at a split ratio of 1:2 using 0.125% trypsin in 0.02% EDTA. The doubling time of TJ905 cells is 41.03 h, and the colony formation rate is 2.5%. Histopathological examination demonstrates that Ki67 labeling index is 72.8%, and the cells are immunopositive for S-100, GFAP, and vimentin by immunohistochemical staining. We use the TJ905 glioblastoma cell line for screening the silencing effect of the above-mentioned siRNA constructs by Western blot, and U251 glioblastoma cells were used for further in vitro and in vivo study. 2. Target sequence selection: Targeted regions on the cDNA sequence of a targeted gene should be located 50–100 nucleotides (nt) downstream of the start codon (ATG). Search for sequence motif AA(N19)TT or NA(N21), or NAR(N17)YNN,


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where N is any nucleotide, R is purine (A, G), and Y is pyrimidine (C, U), A/U at the 5¢ end of the antisense strand, G/C at the 5¢ end of the sense strand; at least five A/U residues in the 5¢ terminal one third of the antisense strand, and the absence of any GC stretch of more than 9 nt in length, 30–50% GC content are more active than those with a higher G/C content. Targeting introns should be avoided, because RNAi only works in the cytoplasm and not within the nucleus (9, 10). To minimize off-target effects, we suggest using BLAST to eliminate homology to other coding sequences. Programs can be found at: www.ncbi.nlm.nih.gov/BLAST (NCBI), http://genome.ucsc.edu/cgi-bin/hgBlat (UCSC), and http://www.ensembl.org/Multi/blastview (Ensembl). 3. There are several methods for preparing siRNA, such as chemical synthesis, in vitro transcription, siRNA expression vectors, and PCR expression cassettes. Nowadays, chemical synthesis siRNA is widely used because it is ready-to-use, suitable for gene function study, and commercial available from many companies. Suppliers of RNA synthesis reagents are Proligo (Hamburg, Germany), Dharmacon Research (Lafayette, CO, USA), Ambion (part of ABI, Austin, TX, USA), and many more. Even siRNA libraries in oligo, plasmid, or virus form can be purchased from companies if provided the RefSeq mRNA sequences from NCBI website. To transfect tumor cells in a 24-well format, use the following conditions as a starting point. In RNAi studies using these conditions, >80% knockdown of an endogenous gene is observed within 48 h of transfection. (a) Cell density: 3 × 104 cells/well (cells will be about 50% confluent at the time of transfection). (b) Amount of Oligofectamine: 3 mL; amount of siRNA of interest: 60 pmol (20 pmol/mL). (c) Dilute 60 pmol of siRNA in 50 mL of serum-free DMEM and mix gently. Mix Oligofectamine gently before use, and then dilute 3 mL in 12 mL of serum-free DMEM. Mix gently and incubate for 5 min at room temperature. After incubation, combine the diluted siRNA with the diluted Oligofectamine (total volume is 68 mL). Mix gently and incubate for 20 min at room temperature to allow the formation of the siRNA–Oligofectamine complex. Add the 68 mL of siRNA–Oligofectamine complex to each well. Mix gently by rocking the plate back and forth. (d) Incubate the transfected cells at 37° in a CO2 incubator for 24–72 h until they are ready to assay for gene knockdown. It is generally not necessary to remove the siRNA–Oligofectamine complex or change the medium.

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Fig. 8. Transfection efficiency detection (200×).

However, growth medium may be replaced after 4–6 h without the loss of transfection activity. (e) FITC-labeled siRNA corresponding to the human EGFR open reading frame 5¢-AGGAATTAAGAGAAGCAACAT-3¢ is purchased from GenePharma (Shanghai, China). After transfection with Oligofectamine for 48 h, U251 cells are visualized by a DP70 cooled-CCD attached to an IX71 (Olyumpus, Tokyo, Japan). More then 70% cells are FITC positive, indicating that the transfection efficiency is approximately 70% (Fig. 8).

Acknowledgments This work is supported by Tianjin Science and Technology Committee, grant numbers 05YFJZJC1002 and 06YFSZSF01100; the China National Natural Scientific Found (30772231); and the Program for New Century Excellent Talents in University, The Ministry of Education of the People`s Republic of China (grant number NCET-07-0615).

References 1. Stewart LA . (2002) Chemotherapy in adult high-grade glioma: a systematic review and meta-analysis of individual patient data from 12 randomised trials. Lancet . 359, 1011– 1018.

2. Mendelsohn J, Baselga J. (2000)The EGF receptor family as targets for cancer therapy. Oncogene. 19, 6550–6665. 3. Wells A. (1999) EGF receptor. Review. Int J Biochem Cell. 31, 637–643.

Silencing Epidermal Growth Factor Receptor by RNA Interference in Glioma 4. Pu P, Liu X, Liu A, Cui J, Zhang Y. (2000) Inhibitory effect of antisense epidermal growth factor receptor RNA on the proliferation of rat C6 glioma cells in vitro and in vivo. J Neurosurg. 92, 132–139. 5. Fire A, Xu S, Montgomery MK, Kostas SA, Driver SE, Mello CC. (1998) Potent and specific genetic interference by double-stranded RNA in Caenorhabditis elegans. Nature. 391, 806–811. 6. Zamore PD. (2001)RNA interference: listening to the sound of silence. Nat Struct Biol. 8, 746–751. 7. Sijen T, Fleenor J, Simmer F, Thijssen KL, Parrish S, Timmons L, Plasterk RH, Fire A. (2001) On the role of RNA amplification in dsRNA-triggered gene silencing. Cell. 107, 465–476.

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8. Moroni MC, Willingham MC, Beguinot L. (1992) EGFR antisense RNA blocks expression of epidermal growth factor receptor and suppresses the transforming phenotype of a human carcinoma cell line. J Biol Chem. 267, 2714–2722. 9. Ui-Tei K, Naito Y, Takahashi F, Haraguchi T, Ohki-Hamazaki H, Juni A, Ueda R, Saigo K. (2004) Guidelines for the selection of highly effective siRNA sequences for mammalian and chick RNA interference. Nucleic Acids Res. 32, 936–948. 10. Elbashir SM, Harborth J, Lendeckel W, Yalcin A, Webe K, Tuschl T. (2001) Duplexes of 21-nucleotide RNAs mediate RNA interference in mammalian cell culture. Nature. 411, 494–498.

Chapter 19 Delivery of Phosphorodiamidate Morpholino Antisense Oligomers in Cancer Cells Gayathri R. Devi Summary Phosphorodiamidate morpholino oligomers (PMO), which have a neutral chemistry, are extensively being used as tools for selective inhibition of gene expression in cell culture models and are currently in human clinical trials. PMO oligomers possess a unique structure, in which the deoxyribose moiety of DNA is replaced with a six-membered morpholine ring and the charged phosphodiester internucleoside linkages are replaced with neutral phosphorodiamidate linkages. PMO internalization in uptake-permissive cells has been observed to be specific, saturable, and energy-dependent, suggesting a receptor-mediated uptake mechanism. Understanding PMO transport should facilitate the design of more effective synthetic antisense oligomers as therapeutic agents. Key words: Apoptosis, delivery, fluorescence, immunoblots, morpholino, oligonucleotides, phosphorodiamidate, PMO.

1. Introduction Antisense technology is now widely used for not only gene functional analysis (1) and target validation in drug discovery approaches, but is also being developed for various therapeutic applications (2, 3). Antisense agents can be broadly divided into two categories based on their mechanism of action. Those that bind to their target sequences and inhibit gene expression by RNAse H-dependent cleavage mechanisms (4) such as phosphorothioates, phosphorodiesters, and others, such as methyl phosphonates, peptide nucleic acids, 2′-O-alkyl RNA derivatives and the phosphorodiamidate morpholino oligomers (PMO) that


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inhibit gene expression by steric interference with translation of messenger RNA (mRNA) or progression of splicing (5–10). Because antisense oligonucleotides act by binding to their target mRNA, which is localized within cytosol or nucleus, the antisense oligonucleotides need to cross the cell membrane. Previous studies, exclusively with charged oligonucleotides (phosphodiester and phosphorothioates) have shown evidence of various DNA entry mechanisms in specific cell lines, which include fluid phase endocytosis in HL60 cells, both receptor-mediated endocytosis and receptor-mediated but endocytosis-independent uptake in erythroleukemia cells, cell membrane nucleic acid channel in the renal brush border, and scavenger receptors in the liver (11–13). In general, the charged oligodeoxynucleotides are taken up actively by cells in a calcium-dependent manner, inhibited by metabolic inhibitors and are temperature dependent. Attempts to identify the still-elusive receptor protein have revealed potential candidates, again specific for the particular cell lines studied in each case (14–18). In the case of other uncharged nucleic acid analogs, Peptide nucleic acid (PNA) show poor uptake compared with methylphosphonates (19). Cellular uptake of methylphosphonate oligonucleosides has been reported to be cell type dependent, predominantly non-receptor-mediated, and by fluid phase pinocytic mechanism, and methylphosphonate oligonucleosides have shown inhibition of target protein by unassisted delivery in various cellular models (20, 21). This chapter attempts to present methods that relate to delivery and analysis of PMO antisense agents in cancer cells. The PMO backbone is neutral in nature because the deoxyribose moiety of DNA is replaced with a six-membered morpholine ring and the charged phosphodiester internucleoside linkages are replaced with neutral phosphorodiamidate linkages. Different PMO strategies for delivery into cancer cells are presented here, which include physical and charge-based methods to assist PMO uptake in culture (22–24), along with methods for analyzing target inhibition (25–35) after PMO treatment, which will allow for use of these agents in rapid screening for therapeutics and as a research tool for a variety of cancer cell models.

2. Materials 2.1. Cell Culture and Lysis

1. Medium for primary prostate cells, Clonetics PrEGM Bullet kit media (Clonetics, San Diego, CA). 2. Media for primary hepatocytes from In Vitro Technology (Baltimore, MD). 3. RPMI 1640 (Gibco/BRL, Bethesda, MD).

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4. DMEM F12 and DMEM (Hyclone Laboratories, Logan, UT). 5. 10% FBS (Life Technologies, Gaithersburg, MD). 6. 100 U/mL penicillin, 75 U/mL streptomycin (Life Technologies, Gaithersburg, MD). 7. Cell solubilization/lysis buffer: 20 mM Hepes, pH 7.4, 150 mM NaCl, 1% sodium deoxycholic acid, 1% Triton X-100, 0.2% sodium dodecyl sulfate [SDS]) (see Note1). 8. Protease inhibitor cocktail (Sigma, St. Louis, MO). 9. 1 mM PMSF (Sigma). 10. Pierce BCA Protein Assay Kit (Pierce, Rockford, IL). 11. Teflon cell scrapers (Fisher, Pittsburgh, PA). 12. Cell scrapers for PMO loading (Sarstedt, Newton, NC). 13. Cell culture plates (Falcon, Becton Dickinson, Lincoln Park, NJ). 14. Dimethyl sulfoxide (DMSO) (Sigma). 15. Lipofectamine (Gibco/BRL). 16. PMO Special Delivery Reagent, ethoxylated polyethylenimine (Gene Tools, Corvallis, OR). 17. 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) reagent (Sigma). 2.2. Phosphorodiamidate Morpholino Oligomers

1. PMO can be custom ordered from Gene Tools (http:// www.gene-tools.com).

2.3. Plasmids

1. pCiNeo expression vector (Promega, Madison, WI).

2. In addition, research support and related publications can be accessed from AVI BioPharma, Inc., Corvallis, OR, which is a biopharmaceutical company developing drugs to treat lifethreatening diseases (http://www.avibio.com).

2. T7 Mega script for in vitro transcription (Ambion, Austin, TX). 2.4. SDSPolyacrylamide Gel Electrophoresis (PAGE)

Preparation of buffers (store all buffers at room temperature, prepare solutions with Mill-Q or equivalently purified water): 1. Running buffer: 25 mM Tris, 250 mM glycine, 0.1% SDS. 2. Transfer buffer: 25 mM Tris, 0.2 M glycine, 20% methanol (pH 8.5) (with cooling during use, see Note 2). 3. 10X Tris-buffered saline (TBS): To prepare 1 L of 10X TBS: 24.2 g Tris base, 80 g NaCl, adjust pH to 7.6 with HCl (use at 1X). 4. Wash buffer: (TBS/T): 1X TBS, 0.1% Tween-20. 6. 1X Phosphate-buffered saline (PBS).

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7. NP40 lysis buffer (Biosource) supplemented with fresh protease inhibitor cocktail (Sigma) and 1 mM PMSF (Sigma). 7. Nonfat dry milk (weight to volume [w/v]). 8. Blocking buffer: 1X TBS, 0.1% Tween-20 with 5% w/v nonfat dry milk; for 150 mL, add 15 mL of 10X TBS to 135 mL water, mix. Add 7.5 g nonfat dry milk and mix well. While stirring, add 0.15 mL Tween-20 (100%). 9. Wash buffer: 1X TBS, 0.1% Tween-20 (TBS/T). 10. Bovine serum albumin (BSA). 11. Primary antibody dilution buffer: 1X TBS, 0.1% Tween-20 with 5% nonfat dry milk; for 20 mL, add 2 mL of 10X TBS to 18 mL water, mix. Add 1.0 g nonfat dry milk and mix well. While stirring, add 20 μL Tween-20 (100%). 12. Kaleidocope prestained protein marker, broad range (premixed format) (Biorad). 13. Biotinylated protein ladder (Invitrogen, Carlsbad, CA). 14. PVDF membrane (Millipore, Billerica, PA). 15. SuperSignal West Pico Chemiluminescent Substrate (Pierce) or Enhanced Chemiluminescence Kit (Amersham, Arlington Heights, IL) and Bio-Max Film (Kodak, Rochester, NY) (see Note 3). 2.5. Stripping and Reprobing Blots

1. Stripping buffer: 62.5 mM Tris-HCl, pH 6.8, 2% (w/v) SDS. Store at room temperature. Warm to working temperature of 70°C and add 100 mM β-mercaptoethanol (see Note 4). 2. Wash buffer: 0.1% (w/v) BSA in TBS-T.

3. Methods 3.1. Delivery of PMO into Cancer Cells

1. Plate cells at 5 × 105 cells/well in a 6-well plate 24 h before the start of the experiment.

3.1.1. Scrape Loading

2. Add the PMO to the medium to give the desired concentration in a total volume of 2 mL. 3. Scrape cells with a cell scraper (Sarstedt) (see Note 5). 4. The cells are then gently transferred with the medium into a new well and left for a desired time before processing for further analysis.

3.1.2. Syringe Loading

1. Incubate 1 × 106 cancer cells/mL of growth medium for 20 min at 37°C. 2. Add the desired amounts of PMO and PF-127 (2% w/v) and mix.


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3. Draw up the cell suspension in a sterile 1-mL syringe through a 25-5/8-gauge needle and then expel by steady pressure on the plunger. Repeat this procedure four times. 4. Add the growth medium recommended for the type of cancer cells (2 mL) to each sample. 5. Pellet the cells by centrifugation. 6. Resuspend the cell pellet in 2 mL culture medium and plate in a 6-well plate. 7. The cells can then be grown for 24–96 h time periods and processed subsequently for desired functional assays. 3.1.3. Charged-Based PMO Delivery

1. Partially complementary DNA molecules with a 10-base adenine 5′ overhang need to be synthesized to serve as carriers for each PMO sequence (24). 2. The PMOs are delivered into cancer cells at a concentration of 1.4 or 2.8 µM. 3. A PMO/DNA duplex is formed by incubating 1 mM stocks of the PMO and the respective partially complementary DNA in a 3:2 ratio for 10 min at room temperature. 4. For preparing 1.4 µM treatment mixture, 9.3 µL of the duplex stock should be diluted in 200 µL deionized water. 5. Add 10 µL of the weakly basic special delivery reagent ethoxylated polyethylenimine. 6. Vortex the tube for 30 s and incubate for 20 min at room temperature. 7. Add serum-free media (3.6 mL) and deionized water (180.7 µL) to the tube to bring the final volume to 4.0 mL. 8. Expose subconfluent-cultured cells to the above mixture for 3 h at 37°C (1.5 mL/well for a 6-well plate and 150 µL/well for a 96-well plate). 9. Aspirate the mixture and replace with the normally recommended serum-containing media for the cells. Harvest the cells at different time points for subsequent experimental analysis as needed.

3.1.4. Unassisted PMO Delivery for Primary Cells

The PMO delivery strategies may be toxic to certain primary cultures. In addition, primary cultures seem to take up PMO agents without the need for any assisted delivery strategies (29, 36, 37). 1. Seed primary cells in a 24-well (20,000 cells) or 6-well (2.0–2.5 × 10 5 cells) plates in their respective serumcontaining media for 24 h. 2. Add the PMO agent to the cells in serum-containing media at various concentrations and incubate for different time points. 3. The cells can then be processed as needed for further analysis.

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3.2. Analysis of PMO Effect on Target Protein in Cells

3.2.1. MTT Cell Proliferation Assay

After delivery of PMO agents in cells in culture, there are various ways to analyze whether the PMO agent was effective in inducing a biological effect. These include measuring the viability of cells after PMO treatment to ensure that any change in viability is caused by inhibition of the target protein and not by toxic effects caused by delivery of the PMO agent itself; measurement of the level of inhibition of target protein in the presence of the PMO agents in cells by Western immunoblots; use of a plasmid-based screening system to ensure antisense specificity; if fluorescencelabeled PMO agents are used, photomicrographs can be taken of cells to detect PMO localization. 1. At different time intervals after treatment of cells with PMO or appropriate vehicle in serum-free media, add 200μl of 5 mg/mL MTT to each well. 2. Incubate at 37°C until blue coloration starts to appear in the cells (see Note 6). 3. If working with adherent cell lines, the media can be aspirated gently without touching the cells or the plates can be inverted over a sink to remove media. Any small remaining media in the plate will not affect the next step. In the case of suspension cultures, skip this step and proceed to next step. 4. Replace with 100 μL dimethyl sulfoxide (DMSO)/well. The converted dye is solubilized with DMSO. Put the plate in a 37°C incubator for 5 min to get rid of any air bubbles. 5. Transfer the plate to a plate reader (we have used a Molecular Devices reader). The absorbance should be read around 540 nm and can be analyzed by SOFTmax Program.

3.2.2. Propidium Iodide Staining for Cell Viability

Trypsin 1. Collect detached and adherent cells using trypsin. 2. Centrifuge cells at 18 g for 3 min, wash with 1% BSA in PBS (repeat twice). 3. Add 4 μL of 0.5 μg/mL PI stock solution and 36 μL of 1% BSA/PBS to each tube. 4. Incubate tubes in the dark for 20 min at room temperature. 5. Centrifuge cells at 1,000 rpm for 3 min, wash with 1% BSA in PBS (repeat twice). 6. Resuspend pellet in 200 μL of 1% BSA in PBS and transfer to a 1.2-mL microfuge tube. 7. Analyze by flow cytometry.

3.2.3. Western Immunoblotting for Target Proteins

To test whether the PMO agent inhibited the target protein expression, Western immunoblotting (SDS-PAGE) can be carried out in the cell lysates prepared from cells treated with PMOs.

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Blotting membrane: This protocol has been optimized for PVDF membranes. A general protocol for sample preparation is described below. 1. Treat cells by adding fresh media containing the regulator for the desired time. Aspirate media from cultures, wash cells with 1X PBS, aspirate. 2. Lyse cells by adding NP40 lysis buffer (40 μL per well of a 6-well plate). Immediately scrape the cells off the plate and transfer the extract to a microcentrifuge tube. Keep on ice. Sonicate for 10–15 s to shear DNA and reduce sample viscosity. Microcentrifuge for 10 min. Transfer supernatant to a new microcentrifuge tube. 3. Perform protein estimation (using BCA Protein Estimation Kit; Pierce Cat # 23227). 4. Dilute appropriate amount of protein 1:1 in Laemmli loading buffer (BioRad cat # 161-0737). 5. Heat sample 95–100°C for 5 min; cool on ice. 6. Microcentrifuge for 20 s. 7. Load onto SDS-PAGE gel (10 × 10 cm). 8. Use prestained molecular weight marker (10 μL/lane) to verify electrotransfer and biotinylated protein ladder (2 μL/ lane) to determine molecular weights. 9. Electrotransfer to PVDF membrane.

Membrane Blocking and Antibody Incubations

Volumes indicated here are for 10 × 10 cm (100 cm2) of membrane; for different-sized membranes, adjust volumes accordingly. 1. (Optional) After transfer, wash nitrocellulose membrane with 25 mL TBS for 5 min at room temperature. 2. Incubate membrane in 25 mL of blocking buffer for 1 h at room temperature. 3. Incubate membrane and primary antibody (at the appropriate dilution) in 5 mL primary antibody dilution buffer with gentle agitation overnight at 4°C or 1 h at room temperature. 4. Wash three times for 10 min each with 15 mL of TBS/T. 5. Incubate membrane with HRP-conjugated secondary antibody (1:2,000) in 5 mL of blocking buffer with gentle agitation for 1 h at room temperature. 6. Wash three times for 10 min each with 15 mL of TBS/T.

Detection of Proteins

1. Mix 2 mL enhancer and 2 mL peroxide solution (Pierce kit) together. 2. Incubate membrane (in the dark) for 5 min at room temperature with gentle agitation. 3. Drain membrane of excess developing solution (do not let dry), wrap in plastic wrap and expose to x-ray film. An initial 1-min exposure should indicate the proper exposure time.

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3.2.4. Photomicrography for Localization of Fluorescent PMO in Cells (30, 33)

1. Wash PMO-treated cells three times with phosphate-buffered saline. 2. Look at various fields under an inverted microscope with appropriate filter for FITC label. We have used the Nikon Diaphot 300 microscope (Nikon Instruments, Melville, NY) connected to an Olympus Magnafire SP-brand digital camera (Olympus America Inc., Melville, NY) with a high success rate, although any digital or manual camera attached to a microscope can be used. 3. The exposure times should be kept constant for all fluorescent pictures, at 8 s.

3.2.5. Plasmid-Based Test System for Screening PMO Antisense Activity (25, 32, 38)

To ascertain the sequence specificity of any PMO, a plasmid-based test system can be used for both cell-free and cellular screening to see whether the PMO is able to specifically inhibit target protein expression in a sequence-dependent manner. A fusion construct needs to be generated for the target gene. In general, this is carried out by subcloning 29–40 bases of the 5′ untranslated region, AUG translational start site, and the first 16–20 bases of the protein coding sequence of the target gene followed by luciferase into the pCiNeo expression vector. This plasmid features a T7 promoter capable of generating in vitro transcribed RNA from a cloned insert for use in cell-free rabbit reticulocyte in vitro translation reactions and a cytomegalovirus (CMV) promoter for constitutive expression in mammalian cells. Luciferase Assay in Cell culture: HeLa cells (70–80% confluence) work well for this protocol. 1. Transfect the pCiNeo-LucΔA plasmid with the gene of interest in HeLa cells using Lipofectamine according to the manufacturer’s directions. 2. Incubate for 24 h and trypsinize the cells, count, and plate 6 × 105 cells/well in 6-well plates. 3. Allow the cells to adhere overnight. 4. After incubation, use the scrape-loading technique as elaborated in the previous section with vehicle or PMOs at different concentrations. 5. Prepare cell lysates 24 h later, normalize for protein content, and determine luciferase activity using a luminometer.

4. Notes 1. Alternately, NP40 cell lysis buffer from BioSource, Camarillo, CA can also be used.


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2. Transfer buffer can be reused for up to five transfers within 7–10 days if the voltage is maintained constant in each successive run. However, adequate cooling by using a refrigerated bath or transferring in the cold room to keep the buffer at no more than room temperature is required for optimal results and protection of the apparatus from heat damage. 3. In case quantification of data is desired, scanning densitometry of the films can be carried out. Care must be taken that the signals in the film are not saturated to get accurate reading. 4. Open and use β-mercaptoethanol in a fume hood. The reagent generates a significant, unpleasant smell in the laboratory and is also toxic to eyes and skin on contact. 5. Use of this particular cell scraper is extremely important for maintaining good cell viability after cell scraping. 6. This step needs to be optimized initially for each cell type to determine the length of incubation.

Acknowledgments I thank Katherine Aird and Rami Ghanayem for editorial and technical assistance and Dr. Patrick Iversen (AVi BioPharma Inc.). Some of the work presented here was supported by Department of Defense (DOD) grants DAMD17-01-1-0017 (GRD) and W81XWH-07-1-0392 (GRD). References 1. Nasevicius A, Ekker SC. (2000) Effective targeted gene “knockdown” in zebrafish. Nat Genet 26:216–220. 2. Sahu NK, Shilakari G, Nayak A, Kohli DV. (2007) Antisense technology: a selective tool for gene expression regulation and gene targeting. Curr Pharm Biotechnol 8:291–304. 3. Tillman LG, Geary RS, Hardee GE. (2008) Oral delivery of antisense oligonucleotides in man. J Pharm Sci 97:225–236. 4. Stein CA, Benimetskaya L, Mani S. (2005) Antisense strategies for oncogene inactivation. Semin Oncol 32:563–572. 5. Ghosh C, Stein D, Weller D, Iversen P. (2000) Evaluation of antisense mechanisms of action. Methods Enzymol 313:135–143. 6. Arora V, Devi GR, Iversen PL. (2004) Neutrally charged phosphorodiamidate morpholino antisense oligomers: uptake, efficacy

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8.

9.

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and pharmacokinetics. Curr Pharm Biotechnol 5:431–439. Summerton J. (1999) Morpholino antisense oligomers: the case for an RNase H-independent structural type. Biochim Biophys Acta 1489:141–158. Summerton J, Weller D. (1997) Morpholino antisense oligomers: design, preparation, and properties. Antisense Nucleic Acid Drug Dev 7:187–195. Giles RV, Spiller DG, Clark RE, Tidd DM. (1999) Antisense morpholino oligonucleotide analog induces missplicing of C-myc mRNA. Antisense Nucleic Acid Drug Dev 9:213–220. Aartsma-Rus A, van Ommen GJ. (2007) Antisense-mediated exon skipping: a versatile tool with therapeutic and research applications. RNA 13:1609–1624.

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11. Stein CA. (1997) Controversies in the cellular pharmacology of oligodeoxynucleotides. Antisense Nucleic Acid Drug Dev 7:207–209. 12. Hans SS, Hans BA, Dhillon R, Dmuchowski C, Glover J. (1998) Effect of dopamine on renal function after arteriography in patients with pre-existing renal insufficiency. Am Surg 64:432–436. 13. Biessen EA, Vietsch H, Kuiper J, Bijsterbosch MK, Berkel TJ. (1998) Liver uptake of phosphodiester oligodeoxynucleotides is mediated by scavenger receptors. Mol Pharmacol 53:262–269. 14. Yakubov LA, Deeva EA, Zarytova VF, Ivanova EM, Ryte AS, Yurchenko LV, et al. (1989) Mechanism of oligonucleotide uptake by cells: involvement of specific receptors? Proc Natl Acad Sci USA 86:6454–6458. 15. Loke SL, Stein CA, Zhang XH, Mori K, Nakanishi M, Subasinghe C, et al. (1989) Characterization of oligonucleotide transport into living cells. Proc Natl Acad Sci USA 86:3474–3478. 16. Benimetskaya L, Loike JD, Khaled Z, Loike G, Silverstein SC, Cao L, et al. (1997) Mac-1 (CD11b/CD18) is an oligodeoxynucleotidebinding protein. Nat Med 4:414–420. 17. Laktionov PP, Dazard JE, Vives E, Rykova EY, Piette J, Vlassov VV, et al. (1999) Characterization of membrane oligonucleotide-binding proteins and oligonucleotide uptake in keratinocytes. Nucleic Acids Res 27:2315–2324. 18. de Diesbach P, N’Kuli F, Delmee M, Courtoy PJ. (2003) Infection by Mycoplasma hyorhinis strongly enhances uptake of antisense oligonucleotides: a reassessment of receptor-mediated endocytosis in the HepG2 cell line. Nucleic Acids Res 31:886–892. 19. Gray GD, Basu S, Wickstrom E. (1997) Transformed and immortalized cellular uptake of oligodeoxynucleoside phosphorothioates, 3′-alkylamino oligodeoxynucleotides, 2′-O-methyl oligoribonucleotides, oligodeoxynucleoside methylphosphonates, and peptide nucleic acids. Biochem Pharmacol 53:1465–1476. 20. Levis JT, Butler WO, Tseng BY, Ts’o PO. (1995) Cellular uptake of oligodeoxyribonucleoside methylphosphonates. Antisense Res Dev 5:251–259. 21. Delong RK, Miller PS. (1996) Inhibition of human collagenase activity by antisense oligonucleoside methylphosphonates. Antisense Nucleic Acid Drug Dev 6:273–280. 22. Partridge M, Vincent A, Matthews P, Puma J, Stein D, Summerton J. (1996) A simple method for delivering morpholino antisense oligos into the cytoplasm of cells. Antisense Nucleic Acid Drug Dev 6:169–175.

23. Ghosh C, Iversen PL. (2000) Intracellular delivery strategies for antisense phosphorodiamidate morpholino oligomers. Antisense Nucleic Acid Drug Dev 10:263–274. 24. Morcos PA. (2001) Achieving efficient delivery of morpholino oligos in cultured cells. Genesis 30:94–102. 25. Hudziak RM, Summerton J, Weller DD, Iversen PL. (2000) Antiproliferative effects of steric blocking phosphorodiamidate morpholino antisense agents directed against c-myc. Antisense Nucleic Acid Drug Dev 10:163–176. 26. Arora V, Iversen PL. (2000) Antisense oligonucleotides targeted to the p53 gene modulate liver regeneration in vivo. Drug Metab Dispos 28:131–138. 27. Arora V, Iversen PL. (2001) Redirection of drug metabolism using antisense technology. Curr Opin Mol Ther 3:249–257. 28. Arora V, Knapp DC, Reddy MT, Weller DD, Iversen PL. (2002) Bioavailability and efficacy of antisense morpholino oligomers targeted to c-myc and cytochrome P-450 3A2 following oral administration in rats. J Pharm Sci 91:1009–1018. 29. Arora V, Cate ML, Ghosh C, Iversen PL. (2002) Phosphorodiamidate morpholino antisense oligomers inhibit expression of human cytochrome P450 3A4 and alter selected drug metabolism. Drug Metab Dispos 7:757–762. 30. Arora V, Hannah TL, Iversen PL, Brand RM. (2002) Transdermal use of phosphorodiamidate morpholino oligomer AVI-4472 inhibits cytochrome P450 3A2 activity in male rats. Pharm Res 10:465–1470. 31. Devi GR, Oldenkamp JR, London CA, Iversen PL. (2002) Inhibition of human chorionic gonadotropin beta-subunit modulates the mitogenic effect of c-myc in human prostate cancer cells. Prostate 53:200–210. 32. London CA, Sekhon HS, Arora V, Stein DA, Iversen PL, Devi GR. (2003) A novel antisense inhibitor of MMP-9 attenuates angiogenesis, human prostate cancer cell invasion and tumorigenicity. Cancer Gene Ther 10:823–832. 33. Iversen PL, Arora V, Acker AJ, Mason DH, Devi GR. (2003) Efficacy of antisense morpholino oligomer targeted to c-myc in prostate cancer xenograft murine model and a Phase I safety study in humans. Clin Cancer Res 9:2510–2519. 34. Amantana A, London CA, Iversen PL, Devi GR. (2004) X-linked inhibitor of apoptosis protein inhibition induces apoptosis and enhances chemotherapy sensitivity in human prostate cancer cells . Mol Cancer Ther 3 : 699 – 707 . 35. Sekhon HS, London CA, Sekhon M, Iversen PL, Devi GR. (2007) c-MYC antisense phosphosphorodiamidate morpholino oligomer

Delivery of Phosphorodiamidate Morpholino Antisense inhibits lung metastasis in a murine tumor model. Lung Cancer Dec 18 (Epub ahead of publication). 36. Suwanmanee T, Sierakowska H, Lacerra G, Svasti S, Kirby S, Walsh CE, et al. (2002) Restoration of human beta-globin gene expression in murine and human IVS2–654 thalassemic erythroid cells by free uptake of antisense oligonucleotides. Mol Pharmacol 62:545–553.

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Chapter 20 Use of RNA Aptamers for the Modulation of Cancer Cell Signaling Sunjoo Jeong, Hee Kyu Lee, and Mee Young Kim Summary Aptamers are in vitro evolved molecules that bind to target proteins with high affinity and specificity by adapting three-dimensional structures upon binding. Because cancer cells exhibit the activation of signaling pathways that are not usually activated in normal cells, RNA aptamers against such a cancer cell-specific signal can be useful lead molecules for cancer gene therapy. The Wnt/b-catenin signaling pathway plays important roles in a critical initiating event in the formation of various human cancers. Because mutations in b-catenin have been found to be responsible for human tumorigenesis, b-catenin is the molecular target for effective anticancer therapies. Here, we describe the selection of RNA aptamers against b-catenin/T-Cell Factor (TCF) proteins and their intracellular expression as intramers. The RNA aptamers acted as central inhibitory players for multiple oncogenic functions of b-catenin in colon cancer cells. These data provide the proof-of-principle for the use of RNA aptamers for an effective anticancer gene therapy. Key words: Anticancer therapy, catenin, colon cancer, RNA aptamer, RNA intramer, TCF, tumorigenesis.

1. Introduction Aptamers are relatively short, single-stranded DNA or RNA oligonucleotides that were derived by means of the in vitro selection method referred to as systemic evolution of ligands by exponential enrichment (SELEX, (1–3)). Because aptamers bind to molecular target proteins with high affinity and specificity, the functions of the target proteins can be mediated after binding as a result of the stable tertiary structure of the aptamers. In addition to such tight binding to the pathogenic target proteins, low toxicity and lack Wolfgang Walther and Ulrike S. Stein (eds.), Methods in Molecular Biology, Gene Therapy of Cancer, vol. 542 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-561-9_20

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of immunogenicity renders aptamers excellent gene therapeutic agents, making them an attractive alternative to antibody or small molecule-based therapy (4–6). Their chemical nature also offers them a unique capacity for application in the fields of diagnostics and sensors, as well as molecular imaging (7–9). The first US Food and Drug Administration (FDA)-approved aptamer drug, Macugen, anti-vascular endothelial growth factor (VEGF) RNA aptamer, is now on the market, demonstrating the potential utility of an RNA aptamer as a novel therapeutic modality. The Wnt/b-catenin signaling pathway has attracted significant attention from cancer researchers and pharmacologists in recent years (10–12). Activation of the Wnt signaling pathway is considered the initiating event in the transformation of intestinal epithelial cells, and b-catenin is a key regulator protein in this process. According to the most widely accepted current model, glycogen synthase kinase (GSK)-3b phosphorylates b-catenin in the cells Adenomatous Polyposis Coli (APC)/Axin complex. Phosphorylated b-catenin is ubiquitinated and degraded by the proteasome; therefore, the level of cytoplasmic b-catenin protein is quite low in inactivated normal cells. When Wnt acts on the frizzled receptor, dishevelled antagonizes the action of GSK-3b, leading to the accumulation of cytoplasmic b-catenin through the dissociation of b-catenin from the APC/Axin complex and translocation to the nucleus. In the nucleus, b-catenin binds to the DNA binding transcription factor TCF, and thereby stimulates the expression of various target genes. It is conceivable that b-catenin functions as an oncogene product by activating the transcription of many oncogenic target genes, such as cyclin D1 and c-myc (13, 14). In addition to its role in transcription, b-catenin was found to be involved in alternative splicing in colon cancer cells (15, 16) and stabilization of some messenger RNAs (mRNAs), such as cyclooxygenase-2 (COX-2) and cyclin D1 (17, 18). During the last decade, various anticancer agents have been developed to block the unlimited activation of Wnt/b-catenin signaling. The most prominent candidates are the nonsteroidal anti-inflammatory drugs (NSAIDs), including selective COX-2 inhibitors. Despite the discovery of powerful NSAID-based cancer therapies, considerable disadvantages remain. Therefore, specific mechanism- and structure-based inhibitors must be developed to selectively modulate Wnt/b-catenin activation in tumorigenesis. Notably, one RNA aptamer, known as AS1411, is the first aptamer to be tested as a cancer therapeutic in clinical trials (19). Here, we describe the methods used for the selection of RNA aptamers for b-catenin and TCF proteins. RNA aptamers are also expressed in the cells; in this case, they are referred to as RNA intramers (20, 21). RNA aptamers have been shown to selectively modulate multiple functions of b-catenin and TCF proteins in colon cancer cells. Thus, we propose that RNA aptamers reduce

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the tumorigenic potential of oncoproteins, and could be further developed as cancer gene therapeutic agents.

2. Materials 2.1. Preparation of Glutathione S-Transferase (GST)–TCF and GST–b-Catenin Proteins

1. 5X extraction buffer: 250mM Tris-HCl (pH 8.5), 500 mM NaCl, 5mM EDTA. Add DTT fresh to a final concentration of 1mM when making a 1X solution. 2. 100 mM phenylmethylsulfinyl fluoride (PMSF, Sigma, St. Louis, MO, USA): Saturated solution in isopropanol. 3. 50 mg/mL lysozyme. 4. 10% (w/v) sarcosyl: Dissolve 10g of sarcosyl in 100 mL of 1X extraction buffer (without DTT). 5. 20% (v/v) Triton X-100. 6. Glutathione-Sepharose 4B (Amersham Pharmacia, Arlington Heights, IL, USA): Swell beads in dH2O at 4°C for 1 h. Equilibrate in lysis buffer before use. 7. Reduced glutathione: Dissolve fresh in 1X lysis buffer.

2.2. Amplification of DNA Oligonucleotide Library

1. DNA oligonucleotide library: A DNA library was designed to have the embedded T7 promoter sequences (5¢-TAATACGACTCACTATAGGG-3¢), the 5¢-terminal restriction enzyme cloning site, random 50–70 nucleotide sequences, and the 3¢-terminal restriction enzyme cloning site. DNA oligonucleotides can be ordered from a commercial company and diluted in dH2O to a 100µM as a stock solution. 2. Polymerase chain reaction (PCR) primers for DNA library: The forward primer should contain T7 promoter sequences, and each primer is recommended to include restriction enzyme sites for the cloning of RNA aptamers to the common vector. 3. 5U/µL Taq polymerase and commercial 10X buffer. 4. dNTP mixture: 10 mM each of dATP, dCTP, dGTP, and dTTP in dH2O.

2.3. In Vitro Transcription

1. 10X T7 RNA polymerase buffer: 400 mM Tris-HCl (pH 8.0), 60 mM MgCl2, 50 mM DTT, 10 mM spermidine, 0.1% (v/v) Triton X-100. 2. 10mM stock mixture of ATP, CTP, GTP, and UTP. 3. 1U/µL RQ1 DNase. 4. RNA gel-loading buffer: 95% (v/v) deionized formamide, 0.025% (w/v) Bromophenol blue, 0.025% (w/v) Xylen cyanol

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FF, 5 mM EDTA (pH 8.0), 0.025% (w/v) sodium dodecyl sulfate (SDS). 5. RNA elution buffer: 0.5M Ammonium acetate, 1mM EDTA (pH 8.0), 0.2% (w/v) SDS. 2.4. In Vitro Selection of RNA Aptamers

1. RNA binding buffer: For b-catenin, 25 mM HEPES (pH 7.5), 150 mM NaCl, 1 mM MgCl2, 2 mM DTT, and 200 U/mL RNase inhibitor (Takara); for TCF-1, 30 mM Tris-HCl (pH 7.5), 150 mM NaCl, 1.5 mM MgCl2, 2 mM DTT, 0.1% bovine serum albumin (BSA), 100 mg/mL transfer RNA (tRNA), and 200 U/mL RNase inhibitor. 2. Purified GST recombinant proteins. 3. Glutathione-Sepharose 4B (Amersham Pharmacia): Swell beads in dH2O at 4°C for 1 h. Equilibrate in binding buffer before use. 4. Phenol:chloroform:isoamyl alcohol (50:48:2). 5. 100% and 70% ethanol. 6. 200 U/µL M-MuLV reverse transcriptase and the appropriate 10X reaction buffer. 7. Cloning vector: After confirming the enrichment of RNA binding to the target protein, RNA aptamers are cloned to the vector; for example, pU6 vectors generating U6-Aptamer clones.

2.5. GST Pull-Down RNA Binding Assay

1. 10 U/mL calf intestinal alkaline phosphatase. 2. 10 U/mL T4 polynucleotide kinase. 3. 6,000 Ci/mmol [g-32P]ATP (Amersham Biosciences). 4. RNA binding buffer: For b-catenin, 25 mM HEPES (pH 7.5), 150 mM NaCl, 1 mM MgCl2, 2 mM DTT, and 200 U/mL RNase inhibitor (Takara, Tokyo, Japan); for TCF-1, 30 mM TrisHCl (pH 7.5), 150 mM NaCl, 1.5 mM MgCl2, 2 mM DTT, 0.1% BSA, 100 mg/mL tRNA, and 200 U/mL RNase inhibitor. 5. Purified GST recombinant proteins. 6. Glutathione-Sepharose 4B (Amersham Pharmacia): Swell beads in dH2O at 4°C for 1 h. Equilibrate in binding buffer before use.

2.6. RNA-Electrophoretic Mobility Shift Assay (EMSA)

1. Forty percent acrylamide/bis solution (19:1) and N,N,N,N¢tetramethyl-ethylenediamine (TEMED). 2. Ammonium persulfate: prepare 10% solution in dH2O. 3. Running buffer (TBE buffer): Prepare 5X stock with 450 mM Tris-borate, 10 mM EDTA, and autoclave before storing at room temperature. 4. MgCl2: Prepare 1M stock solution in dH2O and autoclave before storing at room temperature.


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5. Glycerol: Prepare 50% stock solution in dH2O and autoclave before storing at room temperature. 6. Native acrylamide gel: 5% acrylamide mix (19:1), 0.5X TBE, 10mM MgCl2, 2% glycerol. 2.7. Construction of RNA Intramers

1. pU6 vector: RNA expression vector containing U6 promoter and the stem-loop structure of U6 RNA. 2. pU6 vector cloning primers: To construct the U6-based RNA intramers (U6-Aptamer), amplify DNA from the pUC19-Aptamer with primers, U6-F1 (5¢-TGATGTCGACTAGGGACGCGTGGT-3¢) and U6-R1 (5¢-GACTCTAGAGGATCCCCG-3¢). 3. pDHFR vector: RNA expression vector containing the cytomegalovirus (CMV) promoter and the aberrantly spliced DHFR complementary DNA (cDNA). 4. pDHFR vector cloning primers: To construct the dihydrofolate reductase pDHFR vectors contain a splice variant of dihydrofolate reductase (DHFR) cDNA (DHFR)-based RNA intramer (DHFRAptamer), the DNA was amplified from the pUC19-Aptamer with primers DHFR F1 (5¢-TCACCGCGGGAGCTCGGTACC-3¢) and DHFR R1 (5¢-CCCCGCGGATCCTCCTCTGCAAAGCTT-3¢).

2.8. Cell Culture

1. Media: DMEM or RMPI-1640. 2. Fetal bovine serum (FBS): 10% in DMEM or RPMI-1640. 3. Human HCT116 colorectal carcinoma, HT-29 colorectal adenocarcinoma, embryonic kidney 293T, and mouse NIH3T3 cells (American Type Culture Collection [ATCC]) were cultured in DMEM with 10% FBS. Human LoVo colorectal adenocarcinoma cells were cultured in RPMI-1640 with 10% FBS.

2.9. RNA Immunoprecipitation

1. Formaldehyde: 0.3–1% (v/v). 2. Glycine: 0.25M (pH 7.0). 3. Cell Lysis Buffer: 50mM HEPES (pH 7.5), 1% NP-40, 0.5% sodium deoxycholate, 0.05% SDS, 1mM EDTA, 50 mM NaCl, protease inhibitor cocktail (Sigma), RNase inhibitor (Takara). 4. Antibodies: Normal IgG or anti-b-catenin antibody. 5. Washing Buffer: 50 mM HEPES (pH 7.5), 1% NP-40, 0.5% sodium deoxycholate, 0.05% SDS, 1mM EDTA, 1M NaCl. 6. Protein G-Sepharose: Previously washed in washing buffer (40 µL of bed volume). 7. RIP Resuspension Buffer: 50mM HEPES (pH 7.5), 5 mM EDTA, 10mM DTT, 1% SDS. 8. TRIzol (Invitrogen, Carlsbad, CA, USA) for RNA purification. 9. RNase-free glycogen (10 mg/mL, Ambion, Austin, TX, USA).

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3. Methods 3.1. Preparation of GST Recombinant Proteins

1. Escherichia coli BL21 are cultured, and 0.1mM IPTG is added for 3 h at 28–37°C to OD600 ~0.5–0.6 to induce the expression of the GST recombinant proteins (see Note 1). 2. Cells are pelleted by spinning down at 5,000×g at 4°C and resuspended in 1X extraction buffer (3–5 mL per gram of cell pellet). All steps should be performed on ice. 3. Lysozyme (final concentration, 1mg/mL) and PMSF (final concentration, 1mM) are added to the resuspended cell pellets. 4. Extracts are incubated on ice for 1 h and mixed by occasional inversion. 5. Sarcosyl is added to 1% from a 10% stock and mixed by inversion. 6. Triton X-100 is added to 1% from a 20% stock, and the mixture is then incubated for 30 min at 4°C. 7. Lysates are spun for 15 min at 15,000×g. 8. Supernatants are loaded onto a glutathione-Sepharose column or slurry that has previously been equilibrated with extraction buffer. One milliliter of swelled glutathione-Sepharose 4B per liter of bacterial culture should generally be used. 9. Columns are washed with 10 bed volumes of extraction buffer with 0.5% Triton X-100 and PMSF. Washing is repeated twice. 10. GST proteins are eluted with 10 bed volumes of extraction buffer containing 10mM glutathione. 11. Recombinant proteins are concentrated using a Centricon-30 (Chemicon, Billerica, MA, USA), and the concentrations of proteins are assessed by SDS-polyacrylamide gel electrophoresis (PAGE) and Bradford assay.

3.2. Amplification of the DNA Oligonucleotide Library

1. A DNA library (100 µM) consisting of random sequences of 50–70 nucleotides is amplified by PCR (95°C for 30s, 55°C for 30s, and 72°C for 30s, limit to run 15 cycles). The cycle program, the amount of template, and the concentrations of primers should be adjusted for optimal PCR performance. 2. PCR products are analyzed on a 2% agarose gel to confirm the amplification of the single band of the expected size. 3. The amplified DNA library is recovered by phenol extraction and ethanol precipitation. It is recommended that the starting DNA library contains at least 1 × 1014 different DNA molecules.


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1. The transcription reaction is set up with 0.5–1µg of the DNA template, 10X RNA polymerase buffer, NTP mix (10mM each), 20 U/µL T7 RNA polymerase, and 40 U/µL RNase inhibitor. The final volume of the reaction is brought up to 100µL with DEPC-treated dH2O, and the reaction is then incubated for 3 h at 37°C. 2. 5 µL RQ1 DNase is added to the mixture and incubated for 30 min at 37°C. 3. Equal volumes of phenol:chloroform are added and the mixture is vortexed thoroughly. 4. Aqueous phases are transferred to fresh tubes, and RNA is precipitated with two volumes of 100% ethanol. Samples are stored for 20min at −70°C and centrifuged at 12,000 × g for 25 min at 4°C. Pellets are washed with 0.5mL of 70% ethanol by centrifugation. 5. RNA pellets are dissolved in 15 µL of RNA gel-loading buffer. 6. RNA samples are loaded on a denaturing 7M urea–6% polyacrylamide gel. 7. RNA bands are eluted with RNA elution buffer and incubated at 37°C for 2 h. 8. RNA samples are precipitated as described in step 4. 9. Purified RNA is dissolved in 30 µL of DEPC-treated dH2O, and the concentration of RNA is determined using a UV spectrophotometer.

3.4. In Vitro Selection (SELEX)

1. Purified RNA (3 µg) is denatured for 5 min at 65°C, and is subsequently renatured at room temperature for 20 min. 2. RNA is preincubated with an excess amount of GST protein (5 mg) in 200 mL of binding buffer, and is then shaken for 30 min at room temperature (see Note 2). 3. Previously swelled glutathione-Sepharose 4B (40 mL) is added to the reaction, and the reaction is then incubated for another 30 min. The beads are centrifuged at 5,000×g for 1min, and the supernatants are separated to remove RNA that is bound to GST or to the beads. 4. Purified GST-tagged target protein (0.3 µM in cycles 1–5 or 0.03µM in the following cycles) is added to the unbound RNA sample and incubated for 30 min at room temperature with slow agitation. 5. Glutathione-Sepharose 4B beads (30 mL) are incubated for 30min at room temperature. 6. Beads are centrifuged at 5,000×g for 2 min to pull down protein– RNA complexes.

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7. Pellets are washed three times with 500 mL of the binding buffer, and the bound RNA is eluted by incubation with 5mM EDTA for 5 min at room temperature. 8. RNA samples are extracted by the addition of equal volumes of phenol:chloroform, and are then filtered by Sephadex G-50 quick spin column chromatography (see Note 3). 9. Two volumes of 100% ethanol and 1µL of 10 mg/mL RNase-free glycogen are added. The mixture is stored for 20 min at −70°C and centrifuged at 13,000 rpm for 25 min at 4°C. 10. The pellets are washed with 0.5 mL of 70% ethanol and air-dried. 11. Pellets are resuspended in 33 µL of DEPC-treated dH2O. 12. Reverse transcription reactions are set up with 33µL of the selected RNA, 10 µL of 5X reverse transcriptase buffer, 5 µL of dNTP mix (10mM each), and 1 µL of reverse primer (25 µM). 13. The mixture is incubated for 5 min at 65°C, and is then incubated on ice for 10 min. 14. 1µL M-MuLV reverse transcriptase is added and incubated for 1 h at 42°C. 15. The selected cDNA (10µL) is amplified as the template. The PCR program is comprised of: 25 cycles of 30s at 95°C, 30s at 55°C, and 30s at 72°C, followed by 1 cycle of 7 min at 72°C. 16. Amplified DNA is precipitated as described in step 9. 17. In vitro transcription is performed using 15 µL of the amplified DNA, as described in Subheading 3.3. 18. The in vitro selection procedure is repeated as described in steps 1–16 (see Note 4). 19. Enrichment of RNA binding to the target protein is monitored at each of the four rounds of selection using RNA-EMSA or GST pull-down assay. 20. The selected RNA molecules are cloned after confirmation of the enrichment of the RNA pool to the target protein. 21. Amplified DNA is digested with restriction enzymes present in the primer sites, and is then ligated into appropriate vectors (such as pUC19 vector for sequencing and in vitro analysis). 3.5. RNA Binding Assay; GST Pull-Down

1. In vitro transcribed RNA (5 mg) is labeled with [g-32P] ATP and T4 polynucleotide kinase. 2. Radiolabeled RNA is separated from unincorporated ATP by spin-chromatography through Sephadex G-50. 3. RNA is heated at 65°C for 5 min, and is then slowly cooled down at room temperature to allow the folding of the RNA into its tertiary structure.


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4. RNA is incubated with an appropriate amount of GSTtagged target protein or GST in binding buffer for 30 min at room temperature. 5. Glutathione-Sepharose 4B is added to each reaction and incubated for 30 min at room temperature with shaking. 6. The beads are centrifuged at 5,000×g for 5 min, and the supernatants containing unbound RNA are discarded. 7. The beads are washed three times with 400 µL of binding buffer. 8. Bound RNAs are eluted with 5 mM EDTA for 5 min at room temperature. 9. The elutes are extracted with phenol:chloroform. 10. RNAs are precipitated with two volumes of ethanol in the presence of 0.3M sodium acetate (pH 5.2), incubated for 30min at −80°C, and centrifuged at 13,000 rpm for 30 min. 11. The pellets are washed with 70% ethanol and centrifuged again. 12. The pellets are resuspended in 10 µL of DEPC-treated dH2O, and RNA loading dyes are added. RNA samples are heated at 65°C for 5 min and incubated on ice for 5 min. 13. RNA molecules are analyzed by 6% polyacrylamide–7M urea gel electrophoresis and exposed to autoradiography. 3.6. RNA Binding Assay; RNA-EMSA

1. Radiolabeled RNA (20pM) is incubated with various concentrations of the protein in the binding buffer for 30 min at room temperature. 2. The gel is pre-electrophoresed to remove any charged molecules left from the polymerization reaction. 3. Samples are loaded directly on the running gel. The RNA– protein complexes are separated on 5% native polyacrylamide gels in 0.5X TBE at 250V for 3 h (see Note 5). 4. One of the plates is removed, and the gel is then transferred to 3M paper. The gel is dried and exposed to autoradiography. 5. To determine the binding affinity of the RNA, different concentrations of proteins are incubated with the radiolabeled RNA, and the fractions of protein bound are plotted. 6. Competitive binding assays are also performed as above, with various concentrations of unlabeled competitor RNA (Fig. 1a).

3.7. Construction and Expression of the RNA Intramer

1. The expression vectors for the RNA aptamer are the pDHFR or pU6 vectors. pDHFR vectors contain a splice variant of dihydrofolate reductase (DHFR) cDNA that can be highly expressed in the cells. The pU6 vector contains a U6 promoter and the stem-loop structure of U6 RNA, which can express nuclear localized RNA and can protect RNA against exonucleases, respectively.

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Fig. 1. In vitro selection of RNA aptamer and expression of RNA intramer. (a) RNA-EMSAs of the selected RNA aptamer to GST–b-catenin protein. [32P]ATP-labeled SE-0 and SE-8 RNA (aptamer, 80 pM) were incubated with increasing concentrations of GST–b-catenin protein. After incubation, RNA–protein complexes were resolved on 5% native gels. Lane 1, labeled SE-0 RNA only; lanes 2–4, b-catenin (10, 25, 50 nM); lane 5, labeled SE-8 RNA only; lane 6–11, b-catenin (1, 2.5, 5, 10, 25, 50 nM); lane 12, Arm 1-12 (200 nM); lane 13, GST (200 nM). (b) Stability of the U6-RNA intramer. 293T cells were transfected with the RNA aptamer expression vector (pU6-Aptamer, 0.5 µg), and the expression level of the RNA intramer was detected by RT-PCR until 5 days after transfection. (c) Northern blot analysis of the RNA intramer. 293T cells were transfected with either pU6-vector or pU6-Aptamer, and total RNA was isolated. Lanes 1–3, total RNA from the transfected cells as indicated at top. Lane 4, in vitro transcribed (IVT) RNA aptamer (10 ng). The size of the pU6 -Aptamer is 164 nucleotides (nt), and that of the IVT aptamer is 80 nt.

2. To insert the aptamer into the pDHFR or pU6 vectors, the DNA sequence of the aptamer is amplified from the pUC19aptamer with pDHFR or pU6 primers, respectively. 3. Each amplified fragment is digested with SacII (in the case of cloning to the pDHFR vector) or SalI/XbaI (in the case of cloning to the pU6 vector). 4. PCR products are ligated to vectors, resulting in pDHFRAptamer or pU6-Aptamer. 5. The expression level of the RNA aptamer is estimated by reverse transcriptase PCR (RT-PCR) and real-time PCR using the aptamer-specific primer sets as described below (Fig. 1b and c). 3.8. RT-PCR and Real-Time PCR

1. To confirm the expression of RNA intramers, total RNA is extracted from cells after transfection with pU6-Aptamer or pDHFR-Aptamer using TRIzol reagent (Invitrogen). 2. After reverse transcription, cDNA is amplified by using primer pairs U6-F1 (5¢-TGATGTCGACTAGGGACGCGTGGT-3¢) and U6-R1 (5¢-GACTCTAGAGGATCCCCG-3¢) for pU6-Aptamer,


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or DHFR-FOR (5¢-TCACCGCGGGAGCTCGGTACC-3¢) and DHFR-REV (5¢-TTGGATCCCCGCGGAAGCTT-3¢) for pDHFR-Aptamer. 3. Real-time PCR is performed with a Rotor-Gene RG-3000A system (Corbett Research, Sydney, Australia). Reactions are amplified with the selective primers described above and an LC FastStart reaction mix SYBR Green I kit (Roche, Indianapolis, IN, USA), used according to the instructions of the manufacturer. 4. The PCR amplification cycle includes denaturation at 95°C for 15s, annealing at 47°C for 10s (pU6-Aptamer) or at 55°C for 10s (pDHFR-Aptamer), and extension at 72°C for 15s. 5. Quantification is carried out with Rotor-Gene 6 software (Corbett Research). Relative levels of RNA intramers are expressed as the ratio of the comparative threshold cycle (CT) to the internal control GAPDH mRNA. 3.9. Immunoprecipitation and Western Blotting

1. Cytoplasmic (350 µg) or nuclear (350 µg) extracts are incubated (2 h, 4°C) with a 50% (v/v) suspension of protein G-Sepharose beads (Amersham) that have been precoated with 3 µg of antib-catenin monoclonal antibody. 2. After beads are washed, the immunoprecipitated proteins are resolved by 10% SDS-PAGE and transferred to a PVDF membrane. 3. The blots are probed with specific antibodies and revealed by enhanced chemiluminescence (ECL).

3.10. RT-PCR Analysis

1. Total cellular RNA is isolated with TRIzol (Invitrogen), reverse transcribed with M-MuLV Reverse Transcriptase, and used in the PCR reactions. 2. The following PCR primers are used; cyclin D1, 5¢-CTGGCCATGAACTACCTGGA-3¢ (forward) and 5¢-GTCACACTTGATCACTCTGG-3¢ (reverse); b-catenin, 5¢-CGGGATCCACAAGAAACGGCTTTCA-3¢ (forward) and 5¢-GAGAATTCCAGGTCAGTATCAAACCA-3¢ (reverse); c-myc, 5¢-CTTCTGCTGGAGGCCACAGCAAACCTCCTC-3¢ (forward) and 5¢-CCAACTCCGGGATCTGGTCACGCAGGG-3¢ (reverse). 3. PCR products are analyzed on a 1.5% agarose gel, followed by ethidium bromide staining (Fig. 2).

3.11. RNA Immunoprecipitation

1. HCT116 cells are transiently transfected with the various pU6 RNA expression vectors. Cells are scraped, spun down (4,000 rpm, 3 min, 4°C), and resuspended with 1 mL of cold 1X PBS. 2. Formaldehyde (0.3–1%) is added to the mixture and incubated at room temperature for 5 min with slow mixing (see Note 6). 3. Reactions are quenched with glycine (pH 7.0) and incubated at room temperature for 5 min.

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Fig. 2. Regulation of Wnt/b-catenin target genes by the RNA intramers. (a) HEK 293T cells were transfected with pU6-Vector or the pU6-Aptamer (each 0.5 mg). After incubation with 20 mM LiCl for 16 h, total RNA was purified, and the expression levels were measured by RT-PCR. (b) RT-PCR analysis of HCT116 cell lines stably expressing the RNA aptamer. The levels of c-myc, cyclin D1, and RNA aptamer were determined. GAPDH was used as a loading control.

4. Cells are spun down (4,000 rpm, 3 min, 4°C) and washed with 1X PBS. This process is repeated once. 5. Supernatants are discarded, and Cell Lysis Buffer (~600 µL for a 60-mm culture) is subsequently added. This is followed by incubation at 4°C for 30 min (see Note 7). 6. Cell extracts are sonicated to solubilize the Ribonucleic acidprotein (RNP) complex (output: 8–9 W, each 15s). 7. Extracts are spun down (13,000 rpm, 15 min, 4°C), and the supernatants are transferred to new tubes. 8. For preclearing, extracts are incubated with protein G-Sepharose beads (40 µL of bed volume) and rotated at 4°C for 2–3 h. 9. Extracts are spun down (4,000 rpm, 3 min, 4°C), and the supernatants are removed. 10. Normal IgG or anti-b-catenin antibody is added to the precleared cell lysates, and the mixture is rotated at 4°C for overnight (see Note 8). 11. Protein G-Sepharose beads (40 mL of bed volume) are equilibrated in Cell Lysis Buffer by rotation at 4°C for 2–3 h, and are spun down at 4,000 rpm for 3 min in 4°C. 12. Beads are washed with the 500 mL of washing buffer, rotated at 4°C for 15 min, and spun down (4,000 rpm, 3 min, 4°C). The supernatants are then removed. This process is repeated three times.


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13. Washed beads are resuspended with 100 mL of RIP Resuspension Buffer. 14. To remove cross-linkers, extracts are incubated at 70°C for 45 min. 15. To isolate RNA, 300 mL of TRIzol solution is added, followed by the addition of 80 mL of chloroform. The mixture is vortexed vigorously and incubated at room temperature for 10 min. 16. After centrifugation at 13,000 rpm for 15min at 4°C, the aqueous phase is transferred to a new tube. 17. RNase-free glycogen (1 mL) is added as a carrier. An equal volume of isopropanol is added and spun down (13,000 rpm) for 30 min at 4°C to precipitate RNA. 18. Pellets are washed with 400 mL of 70% ethanol and spun down (13,000 rpm) for 10 min at 4°C. The pellets are then dried. 19. Pellets are dissolved with 30 mL of DEPC-treated water (see Note 9). 20. One-third of the total RNA sample is used as a template for reverse transcription. 21. RT-PCR is performed with a specific primer set and analyzed to detect the bound RNA. 3.12. Selection of Stable Aptamer Transfectants

1. HCT116 cells are co-transfected with pU6-Aptamer, pU6-NC, or pU6 vector in the presence of pTK-Hyg (Clontech, Palo Alto, CA, USA). 2. Stably transfected clones are selected with hygromycin B (Invitrogen). 3. After 2 weeks, hygromycin-resistant clones are tested for expression of the RNA aptamer by RT-PCR.

4. Notes 1. High-level expression of foreign proteins in E. coli often results in the formation of an insoluble product, termed an inclusion body. An undesired inclusion body may be reduced by: (1) lowering the growth temperature between 20°C and 30°C, (2) shortening the induction time, and (3) increasing aeration. 2. The composition of the selection buffer should be optimized for each target protein. Divalent metal ions, such as Mg2+, are critical for optimal binding of the RNA molecule to the target protein. Therefore, it is recommended that MgCl2 be included in the binding buffer.

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3. Selected RNA should be isolated by gel purification at each round of selection to remove undesired RNA species. Failure to purify the selected RNA band often results in the enrichment of RNA molecules with aberrant size. 4. It is critical that the affinity and specificity of the RNA binding to the target protein be monitored until sufficient enrichment is achieved. We generally perform a GST pull-down assay or RNA-EMSA in every three to four rounds of the selection. 5. Concentrations of acrylamide and bis-acrylamide can be modified to improve the resolution of RNP complexes. We recommend using a gel plate with a sufficient length to fractionate various RNP complexes, as well as unbound RNA. The voltage gradient described here (13V/cm) can be increased substantially without problems such as heat dissipation and dissociation of the RNP complex. It is recommended that the gel be run at a low temperature, which tends to form a tighter band and a reduced rate of diffusion. 6. The concentration of formaldehyde and the time for crosslinking can be adjusted to achieve optimal cross-linking. Excessive cross-linking may result in the loss of material due to poor solubilization, or may mask the epitopes that should be recognized during the immunoprecipitation procedure. 7. We recommend proceeding immediately to the immunoprecipitation procedure once the cell extracts are prepared. Although the extracts can be stored at −80°C until future use, there is an increasing risk of the degradation of RNA and proteins during long storage. 8. It is important to demonstrate the specificity of the target protein binding by the intracellularly expressed RNA aptamer. Therefore, we always include control immunoprecipitation reactions using preimmune serum or unrelated antibody. 9. Transfected RNA expression vector DNA may be co-purified during total RNA isolation, which could obscure the binding of the expressed RNA aptamer to the target protein in the cells. To avoid this problem, isolated RNA samples are digested with DNaseI, followed by the removal of DNaseI using DNaseinactivating reagent (Ambion).

Acknowledgments This study was supported by grants from the Korea Science and Engineering Foundation from Ministry of Education, Science and Technology (R0A-2008-000-20053-0, R01-2006-000-10194-0) and the National Cancer Center from Ministry for Health, Welfare and Family Affairs (00000316) of Korean Government.


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References 1. Tuerk, C. and Gold, L. (1990) Systemic evolution of ligands by exponential enrichment: RNA lignads to bacteriophage T4 DNA polymerase. Science 249, 505–510. 2. Ellington, A. D. and Szostak, J. W. (1990) In vitro selection of RNA molecules that bind specific ligands. Nature 346, 818–822. 3. Robertson, D. L. and Joyce, G. F. (1990) Selection in vitro of an RNA enzyme that specifically cleaves single-stranded DNA. Nat. Struct. Biol. 344, 467–468. 4. Bunka, D. H. and Stockley, P. G. (2006) Aptamers come of age - at last. Nat. Rev. Microbiol. 4, 588–596. 5. Lee, J. F., Stovall, G. M. and Ellington, A. D. (2006) Aptamer therapeutics advance. Curr. Opin. Chem. Biol. 10, 282–289. 6. Que-Gewirth, N. S. and Sullenger, B. A. (2007) Gene therapy progress and prospects: RNA aptamers. Gene Therapy 14, 283–291. 7. Farokhzad, O. C., Cheng, J., Teply, B. A., Sherifi, I., Jon, S., Kantoff, P. W., Richie, J. P., and Langer, R. (2006) Targeted nanoparticle-aptamer bioconjugates for cancer chemotherapy in vivo. Proc. Natl. Acad. Sci. U.S.A. 103, 6315–6320. 8. Proske, D., Blank, M., Buhmann, R., and Resch, A. (2005) Aptamers-basic research, drug development, and clinical applications. Appl. Microbiol. Biotechnol. 69, 367–374. 9. Chu, T. C., Marks, J. W. III, lavery, L, A., Faulkner, S., Rosenblum, M. G., Ellington, A. D., and Levy, M. (2006) Aptamer:Toxin conjugates that specifically target prostate tumor cells. Cancer Res. 66, 5989–5992. 10. Polakis, P. (2000) Wnt signaling and cancer. Genes Dev. 14, 1837–1851. 11. Huelsken, J. and Birchmeier, W. (2001) New aspects of Wnt signaling pathways in higher vertebrates. Curr. Opin. Genet. Dev. 11, 547–553. 12. Gregorieff, A. and Clevers, H. (2005) Wnt signaling in the intestinal epithelium: from endoderm to cancer. Genes Dev. 19, 877–890.

13. Tetsu, O. and McCormick, F. (1999) bcatenin regulates expression of cyclin D1 in colon carcinoma cells. Nature 398, 422– 426. 14. He, T. C., Sparks, A. B., Rago, C., Hermeking, H., Zawel, L., da Costa, L. T., Morin, P. J., Vogelstein, B., and Kinzler, K. W. (1998) Identification of c-MYC as a target of the APC pathway. Science 281, 1509–1512. 15. Sato, S., Idogawa, M., Honda, K., Fujii, G., Kawashima, H., Takekuma, K., Hoshika, A., Hirohashi, S., and Yamada, T. (2005) b-Catenin interacts with the FUS proto-oncogene product and regulates pre mRNA splicing. Gastroenterology 129, 1225–1236. 16. Lee, H. K., Choi, Y. S., Park, Y. A., and Jeong, S. (2006) Modulation of oncogenic transcription and alternative splicing by b-catenin and an RNA aptamer in colon cancer cells. Cancer Res. 66, 10560–10566. 17. Lee, H. K. and Jeong, S. (2006) b-Catenin stabilizes Cyclooxygenase-2 mRNA by interacting with AU-rich elements of 3’-UTR. Nucleic Acids Res. 34, 5705–5714. 18. Lee, H. K., Kwak, H. Y., Hur, J., Kim, I. A., Yang, J. S., Park, M. W., Yu, J., and Jeong, S. (2007) b-Catenin regulates multiple steps of RNA metabolism as revealed by the RNA aptamer in colon cancer cells. Cancer Res. 67, 9315–9321. 19. Miller, D., Laber, D., Bates, P., Trent, J., Taft, B., and Kloecker, G.H. (2006) Extended phase I study of AS1411 in renal and non-small cell lung cancers. Ann. Oncol. 17, ix147–148. 20. Famulok, M. and Mayer, G. (2005) Intramers and aptamers: applications in protein-function analyses and potential for drug screening. Chem. Bio. Chem. 6, 19–26. 21. Choi, K. H., Park, M. W., Lee, S. Y., Jeon, M. Y., Kim, M. Y., Lee, H. K., Yu, J., J. W. (2006) In vitro selection of RNA molecules that bind specific ligands. Mol Cancer Ther 5, 2418–2434.

Chapter 21 G-Rich Oligonucleotides for Cancer Treatment Paula J. Bates, Enid W. Choi, and Lalitha V. Nayak Summary Oligonucleotides with guanosine-rich (G-rich) sequences often have unusual physical and biological properties, including resistance to nucleases, enhanced cellular uptake, and high affinity for particular proteins. Furthermore, we have found that certain G-rich oligonucleotides (GROs) have antiproliferative activity against a range of cancer cells, while having minimal toxic effects on normal cells. We have investigated the mechanism of this activity and studied the relationship between oligonucleotide structural features and biological activity. Our results indicate that the antiproliferative effects of GROs depend on two properties: the ability to form quadruplex structures stabilized by G-quartets and binding affinity for nucleolin protein. Thus, it appears that the antiproliferative GROs are acting as nucleolin aptamers. Because nucleolin is expressed at high levels on the surface of cancer cells, where it mediates the endocytosis of various ligands, it seems likely that nucleolin-dependent uptake of GROs plays a role in their activity. One of the GROs that we have developed, a 26-nucleotide phosphodiester oligodeoxynucleotide now named AS1411 (formerly AGRO100 or GRO26B-OH), is currently being tested as an anticancer agent in Phase II clinical trials. Key words: Aptamer, cancer, G-rich oligonucleotides, G-quartets, quadruplex, nucleolin.

1. Introduction Since the 1980s, when automated DNA synthesis became widely accessible, researchers have been trying to develop oligonucleotidebased therapies for cancer. Initially, these efforts focused on developing single-stranded oligomers that could inhibit the transcription or translation of specific cancer-associated genes by antigene (triple helix) or antisense approaches (1–4). More recently, there has been interest in synthetic double-stranded RNA oligonucleotides for RNA interference and in protein-binding oligonucleotides for


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use as transcription factor decoys, immunostimulatory agents (often containing CpG motifs), or as aptamers for particular protein targets (4–10). As with other gene therapy approaches, a major limitation of oligonucleotide-based strategies has been the generally poor cellular uptake of “naked” oligomers, and transfection reagents are usually required for biological activity in cell culture models. Another obstacle is the limited stability of most phosphodiester oligonucleotides in the presence of serum nucleases, which has led to the use of modified nucleic acid analogs, such as phosphorothioates. We have been studying a subset of potentially therapeutic oligonucleotides, which we have termed G-rich oligonucleotides (GROs) (11–17). These are quite distinct from other classes of therapeutic oligonucleotides in that growth inhibitory effects on cancer cells can be observed using unmodified (phosphodiester) sequences, without the need for transfection. After observing that G-rich sequences frequently had potent and selective antiproliferative effects on cultured cancer cells, we set out to determine the mechanism of this phenomenon and evaluate the possible utility of GROs as novel treatments for cancer. During the course of our studies, we have examined a large number of G-rich oligonucleotides to determine correlations between biological activity and structural or physical properties (11, 13, 14). It emerged that the formation of a stable G-quadruplex structure was necessary for antiproliferative activity. However, this type of structure was not sufficient for activity and there was a strong positive correlation between anticancer activity and binding to nucleolin protein. For example, an oligonucleotide containing 2¢-O-methyl-guanosines in place of 2¢-deoxyguanosines was able to form a stable quadruplex structure (as assessed by circular dichroism [CD] and UV melting techniques), but did not bind well to nucleolin and had no antiproliferative activity against cancer cells (13). The culmination of our research on G-rich oligonucleotides has been the development of AS1411 (previously called AGRO100 or GRO26B-OH), a 26-nucleotide phosphodiester oligodeoxynucleotide that is currently being developed by Antisoma PLC (London, UK). In a Phase I clinical trial of this agent, 30 patients who had advanced cancers were treated by continuous infusion of AS1411 at escalating doses up to 40 mg/kg/day for 7 days. No serious side effects related to AS1411 were observed in any of the patients. Furthermore, there was evidence of clinical activity and several patients with metastatic cancer had objective clinical responses or prolonged disease stabilization after treatment with AS1411 (18, 19). This GRO has now advanced to Phase II trials. This chapter describes methods to assess the antiproliferative effects of G-rich oligonucleotides, as well as strategies to determine whether these effects are related to specific properties, such as protein binding, enhanced nuclease resistance, and quadruplex structure.

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2. Materials 2.1. Oligodeoxyribonucleotides

1. Typically, we purchase G-rich oligonucleotides in the “desalted” form on a 1 mmol scale from Integrated DNA Technologies (IDT, Coralville, IA) or Oligos, etc. (Wilsonville, OR). Many other companies also offer comparable custom oligonucleotide synthesis. 2. A UV-visible spectrophotometer capable of recording absorbance at 260 nm is required to measure concentration.

2.2. Cell Culture and MTT Assay

1. Dulbecco’s modified Eagle’s medium (DMEM) (GIBCO, Grand Island, NY) supplemented with 10% heat-inactivated fetal bovine serum (FBS, 15 min at 65°C, GIBCO) and 1% penicillin/streptomycin (10,000 U each per 1 mL; GIBCO). 2. 10X (0.5%) trypsin and ethylenediamine tetraacetic acid (EDTA •4 Na). 3. 1X Dulbecco’s phosphate buffered saline (DPBS), without salts (GIBCO). 4. 5 mg/mL of 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl tetrazolium bromide (MTT) (Sigma, St. Louis, MO) in water passed through 0.2-mm filter. Protect from light. Long-term storage should be at −20°C. Thaw vials as needed and store at 4°C. 5. MTT Lysis Buffer: 10% sodium lauryl sulfate (SDS, Sigma) and 0.01 N HCl. To make 500 mL buffer, weigh out 50 g SDS, add 5 mL of 1 N HCl, and fill with water to 500 mL. Filter through a 0.2-mm filter.

2.3. Radiolabeling and Polyacrylamide Gel Electrophoresis for Electromobility Shift Assay and Stability Assay

1. Resuspended G-rich oligonucleotides. 2. Standard quadruplex-forming oligodeoxynucleotide (human telomere sequence), “TEL”: 5¢-TTA GGG TTA GGG TTA GGG TTA GGG-3¢. 3. Standard protective equipment for use of 32P. 4. [g-32P] ATP (PerkinElmer, Waltham, MA). 5. T4 polynucleotide kinase, supplied with 5X forward buffer (Invitrogen, Carlsbad, CA). 6. TE Midi SELECT-D, G-25 Microcentrifuge Spin Columns (Shelton, Peosta, IA). 7. Scintillation fluid, vials, and counter. 8. Reagents for PAGE: 40% acrylamide solution (wearing a face mask, weigh out 380 g of DNA-sequencing grade acrylamide and 20 g of N,N¢-methylene bisacrylamide, add 600 mL H2O, heat solution at 37°C to dissolve, adjust volume to 1,000 mL with H2O); 10X TBE buffer (108 g Tris base, 55 g boric acid, 40 mL 0.5 M EDTA [pH 8.0] per liter); 10% (w/v) ammonium

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persulfate in water (APS, store at 4°C for up to 1 month); N,N,N,N,-tetramethylethylenediamine (TEMED, Invitrogen). 9. Urea (Sigma) for denaturing PAGE. 10. Nuclear or S-100 extracts for electromobility shift assay (EMSA). These can be purchased from many companies (e.g., Promega Corp.) or prepared by standard methods (20). 11. Dry ice and ethanol for stability studies. 12. 6X DNA loading buffer, 0.25% bromophenol blue, 0.25% xylene cyanol FF, 30% glycerol in water for native PAGE. For denaturing PAGE, a buffer consisting 98% deionized formamide, 10 mM EDTA, pH 8.0, 0.025% bromophenol blue, and 0.025% xylene cyanol FF is used. 13. Gel fixing solution: 10% (v/v) methanol + 10% (v/v) glacial acetic acid in water, and plastic container with lid. This may be stored and reused several times. It should be ultimately disposed of in an approved sink, because it may contain traces of radioactivity after use. 14. Equipment for vertical gel electrophoresis: glass plates, spacers, comb, stand, and power supply. 15. Autoradiographic film (Biomax MR or X-Omat, Kodak), film cassettes with intensifying screen (Kodak) and developer; or phosphorimager and exposure cassette. 2.4. Circular Dichroism Spectroscopy and Thermal Denaturation–Renaturation

1. Resuspended oligomers. 2. Buffers of interest. 3. CD spectropolarimeter. 4. UV-visible spectrophotometer (or CD spectrophotometer) equipped with Peltier effect heating block and temperature controller. 5. Suitable quartz cuvettes (with stopper, for denaturation– renaturation studies).

3. Methods

3.1. Resuspension of Oligodeoxynucleotides, Measurement of Concentration, and Sterilization

1. Resuspend oligonucleotides in H2O (see Notes 1 and 2) estimated to give the concentration for the desired stock solution (based on the amount of oligomer received), typically 500–1,000 µM. Incubate at 65°C for 15 min or until fully dissolved. 2. To sterilize the stock solution, pass through a small 0.2-mm syringe filter into a sterile microfuge tube.


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3. To determine the concentration of the stock solution: Prepare a sample (typically 1–2 mL) from a stock solution that is 500- to 1,000-fold diluted in H2O. Using water as reference, determine the absorbance at 260 nm (A260) using a quartz cuvette. The concentration of the stock solution, C, can be derived using the formula: C=

A 260 Xdilutionfactor eL

Dilution factor = dilution of sample from stock e = molar extinction coefficient of oligonucleotide (supplied by the manufacturer or calculated as described (14)) L = pathlength of cuvette in centimeter (usually 1) 4. A more accurate method of calculating the concentration is as follows: Make several independent dilutions from stock solution, e.g., 1:250, 1:333, 1:500, and 1:1,000, and check the absorbance at 260 nm. These values can then be graphed as A260 (on the y-axis) against 1/dilution (x-axis). According to the equation above, the slope (a) of the line, y = ax + b, will then be equal to e × C × L. Thus, the concentration of the stock can be calculated. An example is shown in Fig. 1 3.2. Treating Cells with Oligonucleotides

The stock solution is added directly into the medium containing the cells so as to make the desired final concentration needed to treat the cells. Typically, we add 5 µL of a concentrated stock solution (31X final concentration) to wells containing 150 µL of medium.

3.3. MTT Assay

1. Cells are plated in 150 µL of medium in 96-well plates. The number of cells per well is based on the doubling time of the cells and ability of the cells to reach confluence by the end of the assay. Most cancer cells, like HeLa and DU145, are plated

A260

Dilution Factor (DF)

0.8 1/DF

0.7 0.6 0.5 0.4

0.749

250

0.004

0.568

333

0.003

0.433

500

0.002

0.209

1000

0.001

0.3 0.2

y = 175.5x + 0.051 R2 = 0.9912

0.1 0 0

0.001

0.002

0.003

0.004

0.005

Fig. 1. Accurate determination of oligonucleotide concentration. Four independent dilutions of the stock solution were made (2, 4, 6, and 8 µL in 2 mL water) and the absorbance at 260 nm (A260) was measured. Plotting the A260 on the y-axis against 1/(dilution factor) on the x-axis should give a linear slope, the gradient of which is related to the stock concentration. In this example, the extinction coefficient is 250,800 M/cm and the pathlength is 1 cm, so the stock concentration = 175.5/(1 × 250,800) = 0.0006998 M = 699.8 µM.

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at 1 × 103 cells per well, whereas cells that proliferate at a slower pace, e.g., Hs27, need to be plated at 3 × 103 cells per well. Usually, triplicate wells per sample are used and typical concentration ranges are 0.2–20 µM. 2. Plating is followed by overnight incubation (approximately 16 h) and then oligonucleotide is added directly to the culture medium to give the final desired concentration. Culture medium is not changed throughout the duration of the experiment. 3. After 120 h (5 days after addition of oligonucleotide) or when the untreated cells reach confluence, add 15 µL (10% of culture medium volume) of MTT solution (see Note 3) to each well. Incubate at 37°C for 4 h. 4. Add 75 µL (50% of culture medium volume) of MTT lysis buffer to each well and incubate for 16 h (overnight) at 37°C. 5. Determine absorbance at 570 nm (A570) using a microplate reader. Use 150 µL of media + 15 µL of MTT solution + 75 µL of MTT lysis buffer as the blank. 6. Analysis of results: The A570 values are proportional to the number of viable cells remaining in the well. The relative activity of various oligonucleotides at a fixed concentration can be compared. Alternatively, the GI50 values (the concentration for 50% inhibition of cell growth) can be calculated by plotting A570 vs concentration. In this case, the A570 at the initial cell density, immediately before addition of oligonucleotide, should be determined (on a separate plate) and subtracted as background. Cytostatic activity against cancer cells is seen for active GROs; typical GI50 values are in the range of 1–10 µM. 3.4. Radiolabeling of Oligomers

1. All manipulations of radioactive materials should be carried out using standard safety precautions for 32P, including use of Plexiglas shields and test tube holders; routine monitoring of personnel, equipment, and benches for radiation exposure or contamination; and disposal of radioactive materials in approved containers. 2. To 5¢-radiolabel an oligonucleotide: In a 1.5-mL tube (see Note 4), combine 1 mL of 1 mM oligonucleotide, 5 mL water, 2 mL of 5X forward buffer, and 1 mL T4 kinase (keep kinase on ice, and minimize exposure to warmth). Using appropriate precautions, add 1 mL [g-32P] ATP. Incubate at 37°C for 30 min, then at 65°C for 10 min. 3. Meanwhile, prepare the TE-Midi column: Remove the bottom cap first then the top cap from the spin column. Discard the caps. Place the column in a collection tube. Spin for 2 min at 16,000 g in a tabletop microcentrifuge. Discard the collection tube with buffer and place the prepared column in a fresh collection tube.


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4. Add 40 mL of water to the reaction tube, and transfer the entire contents of tube (50 mL) to the prepared column. Incubate for 5 min at room temperature. 5. Spin the column again, as before, and discard the column in a radioactive waste container. Keep the collection tube containing end-labeled TEL on ice. Take a 1-µL aliquot and place it in a scintillation vial to determine activity (counts per min [cpm]/µL) in a scintillation counter. Store labeled oligonucleotides at −20°C with appropriate shielding for up to 2 weeks. 6. Expected specific activities are in the range of 200,000– 2,000,000 cpm/µL. 3.5. Electrophoretic Mobility Shift Assay

1. Prepare a 5% polyacrylamide, nondenaturing (native) gel: Assemble gel plates and spacers using a preferred method (e.g., by sealing with tape or melted 1% agarose gel). In a glass flask, combine 6.25 mL of 40% acrylamide, 5 mL of 10X TBE, and 38.7 mL water. Swirl to mix. Add 350 mL of 10% APS and 35 mL TEMED. Working quickly, swirl flask to mix and pour to assembled plates. Avoid air bubbles. Place the comb and allow the gel to polymerize for 30 min or more. After polymerization, the assembled gel may be stored at room temperature in an airtight bag or covered in plastic wrap for up to 2 days. 2. Prepare the samples: Samples typically contain 50,000 cpm of labeled oligo, 2.5 µg of protein extracts, unlabeled competitor oligonucleotide (optional) at 40 nM final concentration, 4 µL of 5X protein binding buffer (1X is 20 mM Tris-HCl, pH 7.4, 140 mM Kl, 2.5 mM MgCl2, 1 mM dithiothreitol, 0.2 mM phenylmethylsulfonyl fluoride [PMSF]) in a total volume of 20 µL. Poly(dI-dC) may also be added in some cases (see Note 5). 3. For a regular EMSA (to compare protein binding of multiple labeled oligonucleotides), dilute each labeled oligonucleotide to 10,000 cpm/µL and add 5 µL of each individual sequence to separate tubes. Make a protein solution mix containing protein extracts (2.5 µg/reaction), 5X buffer (4 µL/reaction), and water (sufficient that the final volume of the protein solution mix will be 15 µL per reaction). Add 15 µL of the protein solution mix to each tube and incubate at 37°C for 30 min. Briefly centrifuge the tube before opening. 4. For a competition EMSA (to evaluate how well unlabeled oligonucleotides compete for protein binding using singlelabeled oligonucleotides), the procedure is very similar. In this case, the labeled oligonucleotide is TEL (a standard quadruplex sequence, see Subheading 2.3.2). Components are prepared as above, except that the water in the protein solution mix is sufficient to give 13 µL per reaction. Add 5 µL of labeled TEL to each tube, then 2 µL of 400 nM unlabeled

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competitor oligonucleotide, then 13 µL of protein mix solution, and incubate as above. The final concentration of unlabeled oligonucleotide may need to be optimized and is typically in the range of 10–100 nM. 5. Control samples should include a sample with no protein added and a sample with no competitor (for competitive EMSA). 6. While the samples are incubating: Assemble gel and tank, fill the top and bottom tanks with 1X TBE buffer, and ensure there are no leaks. Flush the wells with buffer using a syringe and needle. If necessary (when a bottom spacer is used), remove trapped air from the bottom of the gel by injecting buffer using a syringe and needle (bend the needle slightly, if needed). 7. When the sample incubation is complete, add an appropriate volume of 6X native PAGE loading buffer to samples (4 µL per 20 µL sample), mix and load samples into wells. (Usually, we load half of the sample, i.e., 12 µL, and retain half in case the gel needs to be rerun). Run the gel at 180 V (~10 mA) for 2 h, or until the dye front has traveled two thirds the length of the gel. 8. Pour fixing solution to a depth of 2 cm in a container that is large enough to hold the gel (e.g., a shallow glass dish or plastic container with lid). Disassemble the gel apparatus, bearing in mind that the gel (and possibly the buffer) is radioactive. Take care not to rip the gel, which is extremely fragile. Carefully remove one gel plate (gently lever off with a spatula or spacer) and cut off a small corner of the gel (e.g., bottom right corner) to mark the orientation. Invert the gel and plate into container. With care, loosen one corner of the gel with a thin spatula. The gel should release from the glass plate and drop into the solution. Ensure the gel is completely submerged with no trapped air bubbles. Place on a slow circular agitator (5 rpm) and incubate for 30 min at room temperature. 9. Lay out plastic film large enough to wrap the gel. Using both hands, carefully gather the gel and transfer it to plastic film. Very gently, spread the gel flat. Use Kimwipes to remove excess solution (which may be radioactive). Fold the film around the gel, gently remove air bubbles, and wrap the gel completely. 10. Place the gel in an X-ray film cassette and, in the dark room, place the film against the gel and close the cassette. Expose for 4–48 h at −80°C and develop using a suitable processor. Alternatively, place the gel in a phosphorimager cassette and expose for 1–12 h at room temperature, then analyze. 11. Analysis of results: The purpose of the EMSAs is to try to identify shifted bands whose presence correlates with antiproliferative activity to identify candidate target proteins for GROs. In the regular EMSA, binding is indicated by


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the presence of a shifted band, whereas in the competitive EMSA, binding is indicated by disappearance of shifted band(s) seen when TEL is incubated with nuclear extracts. An example is shown in Fig. 2 3.6. Stability Assay (Nuclease Resistance)

1. End-label oligomers, pass them through columns, and count the activity as described (see Subheading 3.4). 2. Prepare a flash-freeze bath: in a Styrofoam box with a lid or another appropriate container, place dry ice. Pour 100% ethanol over the dry ice. Replenish as needed. 3. Prepare stock solutions containing 1 × 106 cpm of each oligonucleotide in 100 µL of the medium of interest (e.g., serumsupplemented cell culture medium or buffer containing S-100 extracts). Immediately remove a 5-µL aliquot and flash freeze. Label the tube with the name of the oligonucleotide and the time point (zero, in this case), then store at −80°C until all samples have been harvested.

Relative Cell Number

4. Remove additional 5-µL aliquots at the designated time points (see Subheading 3.6.6), flash freeze, and store as above. When all samples have been frozen, prepare a denaturing 12% polyacrylamide gel (see Subheading 3.6.5). Thaw samples on ice, centrifuge briefly, add 5 mL of denaturing PAGE loading buffer to each tube. Heat samples at 65°C for 15 min, then place on ice for 5 min. Load samples on the denaturing gel and run as described below.

1.2

MTT Assay in HeLa Cells

1 0.8 0.6 0.4 0.2 0

Oligo Name

29A 15B

-

A

B

C

D

E

F

G

H

I

T30 20A 23A EMSA

TEL-BP (nucleolin)

Fig. 2. Example of MTT assay and competition EMSA. The top panel shows an MTT assay to determine the relative cell number remaining after a 5-day incubation with 10 µM of various G-rich oligonucleotides. The bottom panel shows part of a gel from a competition EMSA using the same oligonucleotides as unlabeled competitors (40 nM). These data indicate that there is a good correlation between antiproliferative activity and ability to compete with the TEL sequence for binding to a specific protein complex (TEL-BP, which contains nucleolin). See ref.14 for further details.

388


5. For denaturing PAGE: In a glass flask or beaker, combine 31.5 g urea, 23.5 mL of 40% acrylamide solution (see Subheading 2.3.8), 7.5 mL of 10X TBE, and 14 mL H2O. Cover the top with foil and place on a hotplate-stirrer at low heat with stirring until urea is dissolved. Filter using a bottle-top nitrocellulose filter and vacuum flask, then allow the solution to remain under partial vacuum for 5 min to degas. Cool the solution to room temperature (if the solution is still warm, polymerization will occur too quickly). Add 35 µL TEMED, mix well, then add 350 µL of 10% APS. Pour into plates, insert comb, and wait for polymerization (about 30 min). Assemble the apparatus, add 1X TBE buffer to the reservoirs, remove the comb, and rinse out the wells. Run the gel for 30 min at 200 V before loading the samples. Turn off the current and flush out the urea from the wells with 1X TBE immediately before loading. Run the gel at 400 V for 1–2 h or until the bromophenol blue has run two thirds of the gel. The gel will become hot, so cooling may be needed to prevent the plates from cracking. This can be achieved by running the gel in a cold room, using an oscillatory fan, or clamping a metal plate to the front glass plate to disperse heat. Once the gel has run, the apparatus is disassembled, and the gel is fixed and analyzed exactly as described for the EMSA (see Subheadings 3.5.8–3.5.10). 6. Analysis of results: Unstructured phosphodiester oligonucleotides are usually digested by exonucleases in serum-containing medium within a few hours, whereas quadruplex oligonucleotides are much more stable and are intact even after several days (see (13), for example). An alternative source of nucleases is S-100 extracts (we typically use 2 µg/100 µL in 1X protein binding buffer [see Subheading 3.5.2]). In this case, samples are incubated for up to 8 h. General examples of these assays are shown in Fig. 3 3.7. Analysis of G-Quadruplex Formation by Circular Dichroism Spectroscopy and Thermal Denaturation–Renaturation

1. Oligomer solutions are made at a fixed concentration (typically, we make 2 mL with 2–5 µM oligonucleotide for a 1-cm cuvette) in the buffer of interest. Buffers usually contain either 10 mM Tris-HCl or 10 mM phosphate buffer at pH 7.4, supplemented with the desired salt (e.g., 0.1 M KCl or 0.1 M NaCl). For annealing, a solution of oligonucleotide in water (at 2X desired final concentration) is heated at 95°C for 5 min to denature it, then placed on ice. An equal volume of 2X buffer is added and the samples are incubated at 37°C for ³15 min. A longer annealing may be required for some sequences (see Note 6). 2. CD spectra are collected (e.g., on a Jasco J-715 spectropolarimeter) between 340 and 200 nm, using 200 nm/min scanning speed, 2 s response time and 5 nm bandwidth. The reference sample is the same buffer used to prepare oligomer samples.


a

PS-CT

PO-CT 0 15 60 120

389

480

0 15

Incubation Time (min)

60 120 480

Intact Oligo

Degraded Oligo

PS-CT

PO-CT

b 0

15

30

60

90 120

0

15

30

60

90 120

Incubation Time (min)

Intact Oligo

Fig. 3. Stability assay to determine nuclease resistance. This is a general example comparing the stability of a C/T-rich DNA sequence (5¢-CCCTTTTTCTTCCCTCCCCTCCCT-3¢) with either a phosphodiester backbone (PO-CT) or a nucleaseresistant phosphorothioate backbone (PS-CT). Note that G-rich phosphodiester oligonucleotides display a much increased nuclease resistance (e.g., intact for 5 days in serum-containing medium) compared with nonquadruplex sequences (not shown, see (13)). In this figure, panel (a) shows radiolabeled oligonucleotides incubated in DMEM with 10% fetal calf serum (FCS; not heat-inactivated) and panel (b) shows the oligonucleotides incubated with 20 µg/mL HeLa S-100 extracts. Note that degradation products are observed for PO-CT in (a) because exonucleases are predominant in serum, whereas a complete disappearance of the band is seen in (b), because endonucleases are present in S-100 extracts.

3. UV-visible melting and annealing curves are obtained by placing the oligonucleotide in a stoppered quartz cuvette (almost full) and monitoring absorbance at 295 nm as the temperature changes 20–90–20°C at a rate of 0.5°C/min. Alternative techniques to determine melting and annealing curves include monitoring absorbance at 260 nm or, if the CD spectropolarimeter is equipped with a thermal module, monitoring ellipticity at 264 nm or 295 nm. 4. Analysis of results: CD spectra of G-quadruplexes can consist of the “parallel” signature, which is characterized by a positive peak at 264 nm with a negative at 240 nm, or the “antiparallel” signature, which has a positive peak at 295 nm and a negative at 265 nm. Often, multiple conformations can coexist in solution and a combination of the two types of spectra is

390


Fig. 4. CD spectra of quadruplex-forming oligonucleotides. As described in the text, the spectra shown are indicative of G-quadruplex formation. The names for the various types of spectra relate to the supposed orientation of the strands in different conformations; however, caution must be used when interpreting data, because strand polarity cannot necessarily be inferred from CD spectra (see (14) for discussion). By contrast, a positive peak at 275–280 nm (not shown) is characteristic of nonquadruplex oligonucleotides and double-stranded DNA (B-form).

observed. By contrast, a predominant positive peak at 275–280 nm is characteristic of nonquadruplex single-stranded oligonucleotides. Examples of CD spectra are shown in Fig. 4. For thermal denaturation–renaturation using a UV-visible spectrophotometer, there is a decrease in A295 (or an increase in A260) corresponding to denaturation of the quadruplex. The melting (or annealing) temperature is defined as the midpoint of the transition. The reversibility of the transition can also be instructive with regard to the relative kinetics of quadruplex formation (14).

4. Notes 1. Ultrapure (Milli-Q) water is used for most applications. Regular distilled water is used to dilute 10X TBE as the running buffer for electrophoresis and for the fixing solution. 2. We generally store stock solutions of oligonucleotides in water at −20°C and have not observed any loss of activity after several months. However, 10 mM Tris-HCl or 10 mM phosphate buffer in pH range 7–8 can be used, if preferred. 3. Many other methods for measuring cell viability, apart from the MTT assay, are also available.


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4. Siliconized tubes, which can be purchased from many suppliers or prepared using Sigmacote (Sigma), are recommended for radiolabelling, EMSA, and stability assays. 5. Poly(dI-dC) can be added to EMSA reactions (1 µg/reaction) to reduce nonspecific binding. It was not used for competitive EMSAs, in this case, because we were comparing relative binding. 6. The annealing procedure described in Subheading 3.7.1 is sufficient to induce quadruplex formation in most G-rich sequences we have examined. However, a protocol with longer annealing at a higher temperature (as described in (14)) may be required to for some sequences, especially those that form tetramolecular structures. Note also that structure can be highly dependent on the concentration and identity of the salt used (e.g., KCl or NaCl).

Acknowledgments and Disclosures The work described herein was funded in part by the US National Cancer Institute and the US Department of Defense Prostate Cancer Program. In addition to these sources, the authors currently receive or have previously received funding from the Kentucky Lung Cancer Research Program, the Komen Breast Cancer Foundation, the University of Louisville, and Antisoma PLC (London, England). Paula J. Bates owns shares in Antisoma.

References 1. Opalinska JB, Gewirtz AM. (2002) Nucleicacid therapeutics: basic principles and recent applications. Nat Rev Drug Discov. 1:503–14. 2. Gleave ME, Monia BP. (2005) Antisense therapy for cancer. Nat Rev Cancer 5:468–79. 3. Scherer LJ, Rossi JJ. (2003) Approaches for the sequence-specific knockdown of mRNA. Nat Biotechnol 21:1457–65. 4. Mahato RI, Cheng K, Guntaka RV. (2005) Modulation of gene expression by antisense and antigene oligodeoxynucleotides and small interfering RNA. Expert Opin Drug Deliv 2:3–28. 5. Devi GR. (2006) siRNA-based approaches in cancer therapy. Cancer Gene Ther 13:819–29. 6. Tomita N, Ogihara T, Morishita R. (2003) Transcription factors as molecular targets: molecular mechanisms of decoy ODN and their design. Curr Drug Targets 4:603–8.

7. Krieg AM. (2008) Toll-like receptor 9 (TLR9) agonists in the treatment of cancer. Oncogene 27:161–7. 8. Nimjee SM, Rusconi CP, Sullenger BA. (2005) Aptamers: an emerging class of therapeutics. Annu Rev Med 56:555–83. 9. Chu T, Ebright J, Ellington AD. (2007) Using aptamers to identify and enter cells. Curr Opin Mol Ther 9:137–44. 10. Ireson CR, Kelland LR. (2006) Discovery and development of anticancer aptamers. Mol Cancer Ther 5:2957–62. 11. Bates PJ, Kahlon JB, Thomas SD, Trent JO, Miller DM. (1999) Antiproliferative activity of G-rich oligonucleotides correlates with protein binding. J Biol Chem 274:26369–77. 12. Xu X, Hamhouyia F, Thomas SD, Burke TJ, Girvan AC , McGregor WG , Trent JO ,

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13.

14.

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16.

Bates, Choi, and Nayak Miller DM, Bates PJ. (2001) Inhibition of DNA replication and induction of S phase cell cycle arrest by G-rich oligonucleotides. J Biol Chem 276:43221–30. Dapic' V, Bates PJ, Trent JO, Rodger A, Thomas SD, Miller DM. (2002) Antiproliferative activity of G-quartet-forming oligonucleotides with backbone and sugar modifications. Biochemistry 41:3676–85. Dapic' V, Abdomerovi V, Marrington R, Peberdy J, Rodger A, Trent JO, Bates PJ. (2003) Biophysical and biological properties of quadruplex oligodeoxyribonucleotides. Nucleic Acids Res 31:2097–107. McMicken HW, Bates PJ, Chen Y. (2003) Antiproliferative activity of G-quartet-containing oligonucleotides generated by a novel single-stranded DNA expression system. Cancer Gene Ther 10:867–9. Girvan AC, Teng Y, Casson LK, Thomas SD, Jüliger S, Ball MW, Klein JB, Pierce WM Jr, Barve SS, Bates PJ. (2006) AGRO100 inhibits

17.

18.

19.

20.

activation of nuclear factor-kappaB (NF-kappaB) by forming a complex with NF-kappaB essential modulator (NEMO) and nucleolin. Mol Cancer Ther 5:1790–9. Teng Y, Girvan AC, Casson LK, Pierce WM Jr, Qian M, Thomas SD, Bates PJ. (2007) AS1411 alters the localization of a complex containing protein arginine methyltransferase 5 and nucleolin. Cancer Res 67:10491–500. Laber D, Bates P, Trent J, Barnhart K, Taft B, Miller D. (2005) Long term clinical response in renal cell carcinoma patients treated with quadruplex forming oligonucleotides. Clin Cancer Res 11:9088S Miller D, Laber D, Bates P, Trent J, Taft B, Kloecker GH. (2006) Extended phase I study of AS1411 in renal and non-small cell lung cancers. Ann Oncol 17(Suppl 8):ix147–8. Dignam JD, Lebovitz RM, Roeder RG. (1983) Accurate transcription initiation by RNA polymerase II in a soluble extract from isolated mammalian nuclei. Nucleic Acids Res 11:1475–89.

Section II Clinical Approaches in Cancer Gene Therapy

Subsection A Requirements for Clinical Gene Therapy Trials

Chapter 22 Regulatory Aspects for Translating Gene Therapy Research into the Clinic Carolyn M. Laurencot and Sheryl Ruppel Summary Gene therapy products are highly regulated, therefore moving a promising candidate from the laboratory into the clinic can present unique challenges. Success can only be achieved by proper planning and communication within the clinical development team, as well as consultation with the regulatory scientists who will eventually review the clinical plan. Regulators should not be considered as obstacles but rather as collaborators whose advice can significantly expedite the product development. Sound scientific data is required and reviewed by the regulatory agencies to determine whether the potential benefit to the patient population outweighs the risk. Therefore, compliance with Good Manufacturing Practice (GMP) and Good Laboratory Practice (GLP) principles to ensure quality, safety, purity, and potency of the product, and to establish “proof of concept” for efficacy, and for safety information, respectively, is essential. The design and conduct of the clinical trial must adhere to Good Clinical Practice (GCP) principals. The clinical protocol should contain adequate rationale, supported by nonclinical data, to justify the starting dose and regimen, and adequate safety monitoring based on the patient population and the anticipated toxicities. Proper review and approval of gene therapy clinical studies by numerous committees, and regulatory agencies before and throughout the study allows for ongoing risk assessment of these novel and innovative products. The ethical conduct of clinical trials must be a priority for all clinical investigators and sponsors. As history has shown us, only a few fatal mistakes can dramatically alter the regulation of investigational products for all individuals involved in gene therapy clinical research, and further delay the advancement of gene therapy to licensed medicinal products. Key words: Clinical trial, FDA, gene therapy, GLP, GMP, regulatory guidelines.

1. Introduction Clinical research involves the testing of investigational products with unknown safety and efficacy in human volunteers, and, therefore, it is highly regulated. However, research scientists often become Wolfgang Walther and Ulrike S. Stein (eds.), Methods in Molecular Biology, Gene Therapy of Cancer, vol. 542 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-561-9_22

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Laurencot and Ruppel

frustrated and overwhelmed with regulatory requirements for products intended for human use. To comprehend the perspectives of regulatory organizations, it is often helpful to explore the evolution of laws that govern the use of these products. In most countries, drug regulation is partly due to a response to a human health crisis. For example, the regulation of gene therapy products was intensified in 2000 after the first death of a human subject attributed to gene therapy in a clinical trial conducted at the University of Pennsylvania. This example also illustrates how the calamitous errors of a few individuals can have dramatic consequences for the regulation of novel investigational products. Even though the specifics of gene therapy regulation may vary between countries throughout the world, the premise of assuring patient safety remains the same in all countries. The focus of this chapter will be a discussion of general product development and regulatory principals applicable to gene therapy clinical research, which will expedite the translation of gene therapy products into the clinic. In the European Union (EU), the procedures for harmonizing clinical trial research have been implemented, however, review by National Agencies is required and the details of gene therapy regulation varies among EU countries (1). The US Food and Drug Administration (FDA) is responsible for the regulation of clinical trial research with investigational agents in the USA. FDA scientists analyze data and conduct a risk assessment on investigational products to determine whether the benefit to the intended patient population outweighs the risks. Gene therapy products are defined by the FDA as “all products that mediate their effects by transcription and/or translation of transferred genetic material and/or by integrating into the host genome, and that are administered as nucleic acids, viruses, or genetically engineered microorganisms” (2). Gene therapy products can be characterized as either in vivo or ex vivo products depending on whether they are directly administered to the patient or are administered as a component of a cellular therapy product. The US Code of Federal Regulations (CFR) Title 21 contains all of the FDA regulations and is updated annually (http://www.gpoaccess.gov/cfr/index. html). Regulations are the interpretation of the laws by the executive branch of the US government, and are binding until they are revised or withdrawn. The principle regulations for drug and biologic products are listed in Table 1. Part 312 of Title 21, referred to as 21CFR312, contains the regulations for Investigational New Drug (IND) Applications. In the USA, to conduct clinical trials with investigational agents, an IND application must be submitted to and reviewed by the FDA. The roles and responsibilities of individuals involved in clinical trial research are also described in 21CFR312 Subpart D. Such individuals include the sponsor and the investigator. The regulations define sponsor as a person

Regulatory Aspects for Translating Gene Therapy Research into the Clinic

399

Table 1 Principle US regulations for drug and biological products Code of Federal Regulations: Title 21: Food and Drugs Chapter I: Food and Drug Administration, Department of Health and Human Services Part 312

Investigational New Drugs

Part 50

Protection of Human Subjects

Part 56

Institutional Review Boards

Part 58

Good Laboratory Practices for Nonclinical Laboratory Studies

Part 314

New Drug Applications

Parts 600–680

Biologics

Part 54

Financial Disclosure by Clinical Investigators

Part 25

Environmental Impact Considerations

Parts 201 & 202

Labeling and Advertising

Parts 210 & 211

Current Good Manufacturing Practices

Parts 800–861

Devices and In Vitro Diagnostics

Parts 1270 & 1271

Human Tissues

who takes responsibility for and initiates a clinical investigation. The sponsor may be an individual or pharmaceutical company, governmental agency, academic institution, private organization, or other organization. The sponsor does not actually conduct the investigation unless the sponsor is a sponsor–investigator. An investigator, also referred to as a principal investigator, is the person who actually conducts a clinical investigation (3).

2. Planning for Clinical Trials Clinical development is a labor- and resource-intensive endeavor. Proper planning is imperative to conduct a cost-effective clinical study that meets the intended goals. This can only be accomplished by clear communication of the goals and timelines within the clinical development team. The clinical development team should include at a minimum experts in chemistry and manufacturing processes, nonclinical animal studies (pharmacology and toxicology), clinical studies, statistics, data management, and regulatory affairs. Because the overall clinical development plan drives the nonclinical development, the amount of product required,

400


and the regulatory strategy, changing the clinical plan at any point during development can have dramatic effects on the cost and timelines for the clinical study. Input from the entire clinical development team should be obtained from the initial concept so experts from each discipline can contribute to the design of the plan. Starting with knowledge of the requirements for demonstrating safety and efficacy for the product in a specific indication, the clinical plan can be formulated. It is not uncommon for inexperienced investigators to change the clinical plan after product has been manufactured, resulting in inadequate supplies of material needed for the new nonclinical and clinical studies, resulting in substantial delays for trial initiation. Information regarding regulatory expectations during the development of gene therapy products can be obtained by consulting guidance documents published by various regulatory agencies, including the FDA, the International Conference on Harmonisation (ICH), and the Committee for Proprietary Medicinal Products (CPMP) within the European Agency for the Evaluation of Medicinal Products (EMEA), as well as journal articles and other publications by experts in the field. The ICH has established the Gene Therapy Discussion Group to foster communication and harmonize regulations, and they have also sponsored workshops and published articles on emerging topics in the gene therapy field. Another source of information in the USA is the Federal Register. Table 2 provides a list of selected guidance documents pertaining to the development of gene therapy products. It is important to remember that all of these documents provide guidance only, and are subject to interpretation. Guidance documents are not legally binding, and represent the current thinking of the regulatory authorities at the time they were published. Alternative approaches to those mentioned in guidance documents may be used if they satisfy the requirements in the regulations. The field of gene therapy is rapidly evolving, and even though guidance documents are a useful and necessary resource, they cannot take the place of an experienced clinical development team. In addition, regulatory authorities should be contacted early in development of the gene therapy product to ensure that the regulatory requirements will be addressed appropriately. The Office of Cellular, Tissue and Gene Therapies (OCTGT), in the Center for Biologics Evaluation and Research (CBER) at the FDA regulates gene therapy products in the USA, and encourages sponsors to contact them early and often during development. A discussion of manufacturing, preclinical, and clinical plans can be conducted with OCTGT staff via a pre-IND meeting. The sponsor can present the development plan to the FDA, and ask questions regarding manufacturing processes and product testing, and nonclinical and clinical study designs. Even though the FDA’s comments during a pre-IND meeting are not


401

Table 2 Selected guidances for the development of gene therapy products Sourcea

Date

Title

ICH

May 1996

Guideline for Good Clinical Practice (E6)

FDA

November 2006

Gene Therapy Clinical Trials—Observing Subjects for Delayed Adverse Events

FDA

April 2007

Guidance for Clinical Investigators, Sponsors, and IRBs Adverse Event Reporting—Improving Human Subject Protection (Draft)

FDA

May 2007

Protecting the Rights, Safety, and Welfare of Study Subjects—Supervisory Responsibilities of Investigators (Draft)

FDA

May 2007

Clinical Trial Endpoints for the Approval of Cancer Drugs and Biologics

Clinical

Nonclinical ICH

July 1997

Preclinical Safety Evaluation of Biotechnology-Derived Pharmaceuticals (S6)

EMEA

July 1998

Note for Guidance on Preclinical Evaluation of Anticancer Medicinal Products (CPMP/SWP/997/96).

FDA

1998

Regulatory considerations for preclinical development of anticancer drugs. Cancer Chemother Pharmacol. 1998; 41:173–185

FDA

1999

Pilaro AM, Serabian MA. Preclinical development strategies for novel gene therapeutic products. Toxicol Pathol 1999; 27:4–7.

ICH

November 2000

Nonclinical Safety Studies for the Conduct of Human Clinical Trials for Pharmaceuticals (M3)

EMEA

December 2006

Non-Clinical Testing for Inadvertent Germline Transmission of Gene Transfer Vectors (EMEA/273974/05)

Manufacturing FDA

July 1993

Points to Consider in the Characterization of Cell Lines Used to Produce Biologicals

ICH

September 1998

Quality of Biotechnological/Biological Products: Derivation and Characterization of Cell Substrates Used for Production of Biotechnological/ Biological Products (Q5D)

FDA

May 2001

IND Meetings for Human Drugs and Biologics Chemistry, Manufacturing, and Controls Information

FDA

August 2001

Submitting Type V Drug Master Files to the Center for Biologics Evaluation and Research (Draft)

FDA

August 2003

Instructions and Template for Chemistry, Manufacturing, and Control (CMC) Reviewers of Human Somatic Cell Therapy Investigational New Drug Applications

FDA

June 2008

Guidance for Industry: CGMP for Phase 1 Investigational Agents

(continued)

402


Table 2 (continued) Sourcea

Date

Title

FDA

April 2008

Guidance for FDA Reviewers and Sponsors: Content and Review of Chemistry, Manufacturing, and Control (CMC) Information for Human Gene Therapy Investigational New Drug Applications (INDs)

FDA

April 2008

Guidance for FDA Reviewers and Sponsors: Content and Review of Chemistry, Manufacturing, and Control (CMC) Information for Human Somatic Cell Therapy Investigational New Drug Applications (INDs)

Multidisciplinary FDA

November 1995

Content and Format of Investigational New Drug Applications (INDs) for Phase 1 Studies of Drugs, Including Well-Characterized, Therapeutic, Biotechnology-Derived Products

FDA

March 1998

FDA Guidance for Industry: Guidance for Human Somatic Cell Therapy and Gene Therapy

FDA

November 1999

Dear Gene Therapy IND Sponsor/Principal Investigator Letter

FDA

February 2000

Formal Meetings With Sponsors and Applicants for PDUFA Products

FDA

March 2000

Dear Gene Therapy IND or Master File Sponsor Letter

EMEA

April 2001

Note for Guidance on the Quality, Preclinical, and Clinical Aspects of Gene Transfer Medicinal Products (CPMP/BWP/3088/99)

NIH

April 2002

Appendix M of the NIH Guidelines “Points to Consider in the Design and Submission of Protocols for the Transfer of Recombinant DNA Molecules into One or More Human Research Participants”

2003

Grilley, BJ, Gee, AP. Gene Transfer: regulatory issues and their impact on the clinical investigator and the good manufacturing production facility. Cytotherapy 2003; 5:197–207

NCI

March 2005

The Sponsor’s Guide to Regulatory Submissions for an Investigational New Drug (http://wwwbdp.ncifcrf.gov/pdf/GuidetoRegSubs.pdf)

FDA

November 2006

Supplemental Guidance on Testing for Replication Competent Retrovirus in Retroviral Vector Based Gene therapy Products and During Follow-up of Patients in Clinical Trials using Retroviral Vectors

a FDA guidances on drugs can be found at http://www.fda.gov/cder/guidance/index.htm. FDA guidances on biologics can be found at http://www.fda.gov/cber/guidelines.htm. FDA gene therapy guidances can be found at http://www.fda.gov/cber/genetherapy/gtpubs.htm. EMEA guidances can be found at http://www.emea.eu.int/index/indexh1.htm. ICH guidances can be found at all three of these sites or http://www.ich.org

binding, they provide valuable information for the sponsor. The FDA has published a guidance document regarding meetings that should be consulted for information on requesting a meeting, expectations for information submitted to the FDA for review before the meeting, and the pre-IND meeting process (4).


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3. Clinical Development It is expected that Good Clinical Practice (GCP) principals are adhered to in the development, implementation, and oversight of clinical studies in humans (5). GCP is an ethical and scientific quality standard for the design, conduct, monitoring, auditing, recording, analysis, and reporting of clinical trials based on the ethical principles established in accordance with the Declaration of Helsinki (5). The clinical site must maintain complete and accurate records of observations and activities occurring during the clinical trial. This documentation provides evidence that the clinical trial was conducted in compliance with the protocol, and according to GCPs and applicable regulatory agency requirements. Ultimately, this documentation will be used to provide evidence of the safety and effectiveness of the treatment. The success of clinical trials depend on the quality of the data, therefore, the case report forms (CRF) and the data management plan should be developed in conjunction with the clinical development team, and finalized before protocol implementation. Investigators and site personnel should be qualified to conduct clinical trials, and sponsors must provide oversight to ensure quality standards are followed. For studies in which the investigator is also the sponsor (referred to as sponsor– investigator), it is imperative that a separate and independent monitoring group is used to prevent a conflict of interest and ensure the quality of the clinical trial. In March 2006, the FDA issued a letter to sponsors requesting information regarding the quality assurance plan for clinical trials (6). This plan is now required in all new gene therapy IND applications. Even though there are numerous types of clinical trials, all clinical protocols include a hypothesis and the plan to address the hypothesis. The clinical protocol states the objectives and design of the study, describes the intervention in detail, the measurement of the variables that indicate a beneficial or a detrimental response to treatment, and the monitoring and reporting of the responses to sponsors and regulatory groups. The informed consent document accompanies each protocol and is part of the informed consent process, which informs the clinical trial participants of all aspects of the clinical study in a noncoercive manner. The informed consent document is written in a language understandable to the subject and also outlines the rights of the clinical trial participant. Trial participants are required to sign the informed consent before enrolling in the clinical trial (5). The study must be designed and conducted in a fashion to minimize the risks to the clinical trial participants while optimizing the potential benefit. The required elements of a protocol are outlined in Table 3. Because of concerns regarding long-term consequences of gene therapy, studies to include the collection of data on delayed adverse events in subjects treated with gene therapy products

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Table 3 Elements of a clinical protocol • Background/justification ° Name and description of investigational product ° Justification for route of administration and dose ° Where we are in the field ° Summary of findings of nonclinical and clinical studies that are relevant to trial ° What the study will add that is important ° Summary of potential risks, if any • Names and address of investigators, facilities, and IRB • Objectives ° Primary hypothesis ° Secondary hypotheses • Eligibility criteria • Study design and methods ° Type of study, comparison ° Description of intervention (what, how much, how long) ° Concomitant therapy ° Examination procedures (baseline, follow-up, outcome assessment) ° Intervention assignment procedure • Monitoring and management ° Data and safety monitoring ° Adverse event assessment, reporting ° Contingency procedures ° Withdrawal criteria ° Testing for gene therapy product specific safety issue (when applicable) ° Long-term follow-up for patient safety (when applicable) ° Data handling and review • Statistics ° Sample size ° Stopping guidelines ° Analysis plans (continued)


405

Table 3 (continued) • Patient protection issues ° Informed consent process ° IRB review and approval of informed consent document

may be necessary. The risk of long-term consequences varies between gene therapy products, therefore the FDA’s guidance “Gene Therapy Clinical Trials—Observing Subjects for Delayed Adverse Events” should be consulted to determine which products are exempt from this additional monitoring. Product-specific safety concerns, such as replication-competent retrovirus (RCR) formation, and immunogenicity of specific vectors (i.e., adenoassociated virus vectors), may require additional patient samples collection and testing (7). If additional data is required, information regarding the collection and analysis of the data should be included in the protocol and informed consent. As required by the US Food and Drug Modernization Act and the Best Pharmaceuticals for Children Act, US IND sponsors are responsible for registering clinical trials involving investigational agents that are intended for the treatment of a serious or life-threatening disease or condition in the Clinical Trials Data Bank (http://clinicaltrials. gov/ and http://prsinfo.clinicaltrials.gov/). Additional information on registering clinical trials, including the required and optional data elements and the FDA guidance (8) is available at the Protocol Registration System (PRS) Information Site listed above. Review and initial approval of a gene therapy clinical protocol and the informed consent document may take several months upwards to a year depending on the review process at the clinical site, the sponsor, and the quality of the clinical protocol. The clinical protocol review process often involves scientific reviews by the clinical site and the sponsor, human subjects protection review by the Institutional Review Board/Independent Ethics Committee (IRB/IEC), and regulatory review by the applicable regulatory agency. In the USA, in addition to the FDA, research involving recombinant DNA molecules is regulated by the National Institutes of Health (NIH). Therefore, a gene therapy clinical protocol must also be submitted to the Office of Biotechnology (OBA), NIH and the Institutional Biosafety Committee (IBC) for review. The Recombinant DNA Advisory Committee (RAC) is the review panel for OBA, and consists of experts in clinical

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trials research, gene therapy, manufacturing, and ethics. Whereas submission of documentation to the FDA is the responsibility of the clinical trial sponsor, submissions to the IRB/IEC, IBC, and OBA are the responsibility of the principal investigator of the gene therapy study. The NIH Guidelines for Research Involving Recombinant DNA Molecules should be consulted before initiating any gene therapy studies in animals or humans (available at http://www. nih.gov/od/oba/). The NIH Guidelines are applicable to all recombinant DNA research within the USA or its territories that is conducted or sponsored by an institution that receives any support for recombinant DNA research from the NIH. The NIH guidelines are also applicable to research that involves testing in humans of materials containing recombinant DNA developed with NIH funds, if the institution that developed those materials sponsors or participates in those projects. Privately funded projects using recombinant DNA must also adhere to the NIH Guidelines if these projects are being carried out or funded by an organization that has any NIH contracts, grants, or other support for this kind of research. Numerous sponsors of privately funded recombinant DNA research who do not receive NIH support voluntarily follow the NIH Guidelines. Nonclinical and clinical studies investigating recombinant DNA molecules will require submission of the protocols to the Institutional Biosafety Committee (IBC) before study initiation. Appendix M of the NIH Guidelines “Points to Consider in the Design and Submission of Protocols for the Transfer of Recombinant DNA Molecules into One or More Human Research Participants” outlines the requirements for conducting human gene therapy clinical trials. In addition to submitting the protocol to OBA for RAC review, the investigator will need to submit responses to Appendices M-II through M-V of the NIH guidelines, which pose questions concerning the objective and rationale of the proposed research, and informed consent and privacy issues. Once the protocol is reviewed, revised, and approved by the sponsor, the scientific reviewers, the IRB/IEC, the IBC, the FDA, and the RAC, the clinical trial can then be initiated. Within 20 working days of enrollment of the first patient, OBA requires submission of the following additional information: the IRB and IBC approval, the IRB- and IBC-approved protocol and informed consent, responses to RAC recommendations, modifications to the protocol and consent required by the FDA, the IND number, the applicable NIH grant numbers, and the date of initiation of the study. During the conduct of the clinical trial, all protocol amendments, adverse event safety reports, and annual updates on safety and response outcomes must be submitted to the sponsor, IRB/IEC, IBC, OBA, and the FDA according to the federal regulations (9) and the NIH Guidelines for Research


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Involving Recombinant DNA Molecules. In addition to semiannual or annual updates to the IRB/IEC, and annual updates to the FDA, IBC, and OBA, gene therapy clinical trials often undergo an additional safety review by a Safety Monitoring Committee (SMC) (also called Data Safety and Monitoring Committees [DMSB] because of the perceived increased risk of gene therapy products. Effective October 2000, the NIH issued Guidance on Data and Safety Monitoring for Phase I and II trials. The NIH policies can be found at http://grants.nih.gov/grants/ guide/notice-files/not98-084.html and http://grants.nih.gov/ grants/guide/notice-files/NOT-OD-00-038.html.

4. Nonclinical Development The objectives of nonclinical animal studies are to establish “proof of concept” for efficacy, and provide safety information for products intended for use in a clinical trial. The initial nonclinical toxicology studies should be designed to provide information that can be used to select a starting dose and dose escalation plan for the clinical trial, as well as identify the target tissues of toxicity and the reversibility of these toxicities. The toxicity profile in animals will be used to establish the clinical parameters to monitor for safety and provide information needed to determine the eligibility criteria. In general, animal safety studies should be conducted according to the Good Laboratory Practice (GLP) regulations as defined in 21 CFR part 58. The GLP regulations set forth the rules for conducting and reporting nonclinical safety studies that support or are intended to support clinical research or marketing approval as regulated by the FDA. The regulations cover corporate organization and personnel, facilities, equipment, testing facilities operation, test and control article documentation, the study protocol, record keeping and reporting, and procedures for disqualifying testing facilities. The purpose of the GLP regulations is to assure the quality and integrity of the nonclinical safety data submitted to regulatory agencies. However, not all of the GLP regulations apply to all studies. For example, GLP compliance may not be required or possible when disease model systems are used for testing the safety of gene therapy products. Sponsors should consult the FDA to determine whether certain key studies supporting safety can be conducted without full compliance with GLP provisions. Nonclinical pharmacology and toxicology information is required by regulatory agencies not only before, but throughout clinical testing of products. Risks that need to be evaluated in animals before exposing humans include those related to duration

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of administration, formulation, and route and schedule of administration. Bioactivity studies provide the rationale for the introduction of the gene therapy product into human clinical trials. These studies should be designed to determine the duration and level of gene expression, the dose–response relationship, and the optimal route of administration and dosing regimen to be used in the clinical trail. Early stage studies may include dose range finding and short-term toxicology studies, vector biodistribution and persistence studies, and metabolism studies. As clinical development of the gene therapy product progresses, late stage toxicology studies may be required to provide additional information on the safety of the medicinal product after extended exposure (e.g., carcinogenicity studies) or in special populations (e.g., pregnant animals). Factors such as the properties of the vector, the transgene product, the delivery system, and the clinical indication will need to be considered in the design and implementation of these studies (10). The nonclinical animal studies should be conducted in a similar fashion as the proposed clinical study in regards to the product tested, formulation of the product, route of administration, and schedule of treatment. Changing the formulation (e.g., by adding liposomes, altering pH, or adjusting salt concentration), or the route of administration may change the biodistribution of the product and thereby affect bioactivity or toxicity. Key principles for gene therapy nonclinical studies include species specificity of the transduced gene, permissiveness for infection by viral vectors, and comparative physiology, therefore, studies should be conducted in a relevant animal model (11). Because of the strict species specificity of many gene therapy products, OCTGT at the FDA should be consulted for concurrence on nonclinical study designs before initiating any studies. Safety issues of great concern with gene therapy products include distribution of the vector from the site of injection, and genomic integration of vector sequences. Biodistribution studies are necessary to determine the distribution and persistence of the vector in nontarget organs (12). Of particular importance is the vector distribution to the germ cells; therefore, testicular and ovarian tissues must be analyzed in nonclinical studies for the presence of the gene therapy product. Other tissues that should be evaluated in the biodistribution study include peripheral blood, tissue at the injection site, highly perfused organs such as brain, liver, kidneys, heart, and spleen, as well as tissues which would potentially be affected due to the route of administration or the toxicity of the transgene. The presence of the vector sequence in tissues can be evaluated via DNA polymerase chain reaction (PCR) methodology designed to detect a sequence unique to the product. If vector sequences are observed in nontarget tissues, studies should be conducted to determine whether the gene is


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expressed and/or associated with any toxicity (11). Often the safety and biodistribution of vectors can be evaluated in the same toxicology study. Toxicology testing should obtain information regarding the toxicities related to the vector delivery system and the safety of the expressed gene. Vector persistence, in vivo expression of the transgene, identification of the target organs, and the reversibility of toxicities should be determined.

5. Manufacturing Gene therapy products present numerous manufacturing challenges to provide a safe and effective product for use in clinical trials. These products involve the modification of genetic material of living cells (13). They are complex biologic products due to the components and manufacturing processes used to generate them, and they are not easily defined. Due to the biologic nature of these products, there is an inherent risk for variability and introduction of adventitious agents. Therefore, appropriate controls must be in place to ensure manufacturing consistency and product quality. These controls include qualification of source materials as well as appropriate quality oversight. Initial clinical trials must focus on ensuring the safety of the patients receiving the investigational gene therapy product. Demonstration that a gene therapy product is safe involves documentation of the components, process, and testing involved in its manufacture. 5.1. Good Manufacturing Practices

It is essential that investigational products are manufactured in a manner that ensures they are safe for human use. Good Manufacturing Practices (GMP) were developed for this purpose, and are required for the manufacture of a product to be administered in the USA (14). Similarly, Europe also has a set of GMPs that must be followed for manufacturing products. GMPs are a set of current, scientifically sound methods, practices, or principles that are implemented and documented during product development and production to ensure consistent manufacture of safe, pure, and potent products. GMPs play an important role in control and regulation of not only the final product, but all steps of the manufacturing process. Adherence to GMPs provides for quality and safety throughout the process and will lead to consistent performance of product lots. Any product not manufactured under GMPs in the USA is considered adulterated per the US Food, Drug, and Cosmetic Act, even if the product meets its final specifications. In July 2008, the US Food and Drug Administration (FDA) issued the Guidance for Industry entitled “CGMP for Phase 1

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Investigational Drugs” (15). This guidance document discusses the concept that product manufacturing is recognized as a dynamic process that will change as more information is gathered about the product and the manufacturing process during development. A frequent misconception is that the FDA requires full GMP compliance for all investigational products irrespective of the phase of development. In reality, the FDA expects an incremental approach to manufacturing investigational drugs/biologics during the various stages of clinical development. This is often referred to as the “sliding scale” of GMP. During product development, it is essential to maintain the quality and safety of the investigational product. As manufacturing experience is gained throughout development, more controls should be put in place, taking into consideration the product and process information that has been obtained. Each individual manufacturer must ensure that their procedures, facilities, and testing provide a product that meets appropriate standards of safety, identity, quality, strength, and purity. The basic principles of GMPs involve traceability and documentation. Careful control of the entire manufacturing process is critical to ensure safety of the gene therapy product to be administered to a patient. Manufacturing of a product for initial clinical trials, known as Phase I, involves adherence to written procedures, having equipment that is adequately controlled, and having records of all data from the production and testing of the product (14, 15). Many small-scale investigators or manufacturers will document procedures in laboratory notebooks, although the use of batch production records is also a common method. Records should be in sufficient detail that the process could be repeated. Traceability is the ability to track all materials used in the manufacturing process. Because materials used in production can impact the safety of the product, traceability provides regulators with some assurance of final product safety. All of the work done in the laboratory, raw materials, cell lines, vectors, and reagents that were used to make the product need to be recorded. Documentation of all raw materials should include vendor names, catalog numbers, lot numbers, expiration dates, and certificates of analysis. Methods for selection of the final gene construct, the method of transfer of the gene construct into the host cell, and the selection and characterization of the recombinant host cell clone need to be described (16). Details of the expansion of the recombinant host cell clone, seed stock, or any cell or virus banking information need to be recorded. For autologous and allogenic cell components, the cell source (tissue type and cell type) needs to be identified, and donor safety testing should be performed (16, 17). In addition, the method of collection, processing, and culture conditions should be described for these products. This information is critical for establishing that a product is safe and pure.


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The source and country of origin needs to be determined for all raw materials of animal origin. Assurance from animal viruses needs to be provided, so animal derived materials should be avoided when possible. Any materials of bovine or porcine origin, like fetal bovine serum or trypsin, that can harbor adventitious viruses should be appropriately tested (16, 18). Bovine serum should be from countries that are considered free of bovine spongiform encephalopathy (BSE). Because of the risk of BSE in ruminant animals, all ruminant-source raw materials should be obtained from BSE-free countries. Certificates of origin for animal-derived materials can be obtained from most vendors and should be available and submitted with regulatory submissions, along with certificates of analysis. GMPs are about controlling the manufacturing process and the environment, therefore, manufacturing and Quality Control personnel should have adequate education, training, and experience for the particular function that they perform. They should be familiar with GMP requirements and know the importance of adhering to them to produce a safe investigational product. Regulations usually refer to a Quality Control unit that can be made up of both a Quality Assurance group and a Quality Control group. The Quality Control group is usually responsible for the in-process and release testing of the product. The Quality Assurance group is responsible for approving and rejecting all components, in-process material, packaging materials, labeling, and drug products. They also review documentation of the production records to assure that no errors have occurred or, if they have, that they have been thoroughly investigated. The facility or laboratory used in manufacturing and Quality Control testing should be properly maintained and have work areas sufficient for the intended functions to be performed. The environment should be clean and have appropriate lighting, ventilation, heating, cooling, and plumbing. The air-handling systems should help prevent any contamination of the product. Laminar flow hoods are recommended. Equipment used should not contaminate the product or be reactive or absorptive to it. Equipment must be calibrated, cleaned, and sanitized at appropriate intervals. Dedicated equipment and or disposable parts are recommended when possible. Special precautions need to be considered for the manufacture of sterile products (15). Aseptic manipulations should be conducted under laminar flow conditions (air classification of Class 100 or ISO 5). The work area should be disinfected before any manufacturing, and between different operations. Attention should be given to make sure the airflow of the laminar flow workstation is not interrupted. All procedures should be documented and followed. Media fills are usually performed to demonstrate a process consistently produces a sterile product. The product should be

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tested for sterility, and it is critical that the test article does not interfere with the sterility test. The Quality Assurance group should also perform a review of the production records to show that the aseptic procedures were followed, and should not release the final product until acceptable results of sterility are known. There are special exceptions to release of a product before receiving sterility results when a product must be administered before the final results are available. However, these exceptions must be discussed with a regulatory agency and agreed upon. The best situation is for only one product to be produced in a separate area or room at a time. However, if appropriate cleaning and control procedures are in place to ensure no product mix-ups or cross contamination, then the same areas can be used for multiple purposes (15). In the case of multiuse facilities, careful consideration should be given to the design of the room and flow of the procedures to prevent any cross contamination. Component and product separation should be considered, and change-over procedures can be put in place to clear the room or areas before use. Appropriate cleaning and testing procedures must be in place to ensure prevention of contamination by any adventitious agents especially if live viral or vector processing has occurred. Other factors that can facilitate conformance with GMPs during manufacturing include the use of closed-process equipment (not open to the environment) and disposable equipment, which can reduce the cleaning needed (15). The risks of the environment where the product is being manufactured should be carefully considered in terms of how it might adversely affect the quality of the investigational product. One area of particular concern is cross contamination with other substances that may have been used or are being used in the manufacturing environment. Manufacturers must perform their own risk assessment for their product and manufacturing process and follow good scientific principles to ensure GMP procedures are met. 5.2. Manufacturing and Quality Control Testing Issues

The product’s attributes are maintained in part by having appropriate Quality Control procedures in place. Analytical tests used during production and for product release should be scientifically sound and reproducible. The tests should follow written procedures with the appropriate controls and standards. Laboratory equipment used for analytical procedures should be properly maintained and calibrated according to written procedures, and personnel should verify the equipment is in good working condition before use. An investigational product must be evaluated before release for its identity, strength, potency, purity, and quality (19, 20). Acceptance criteria should be established with the information known for this stage of development. Not all product attributes may have acceptance criteria determined at this early stage.


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Characterization and release testing of your gene therapy product must be completed to demonstrate the product is acceptable for administration to patients. Testing is performed at various stages, including cell banks, virus banks, in-process testing, bulk purified material, and final vialed product. Testing can also be performed on pooled harvest material if appropriate. Specific testing is determined on a case-by-case basis dependent on the product and process by which it is manufactured. However, products should be tested for sterility (21CFR610.12, USP), endotoxin content, mycoplasma, identity, adventitious viruses, concentration, purity, and potency (13, 14, 16). Mycoplasma testing should be performed on the product at a stage such as pooling of culture harvests because it is a step that is most likely to detect contamination. Identity testing can be performed by restriction enzyme mapping, sequencing, or PCR (13). For early potency assays, the level of gene expression is acceptable, but whenever possible the biological activity of the expressed gene product should be measured. Purity testing for DNA plasmids can include measures of DNA quality that include linear and supercoiled content. Impurities should be assessed as part of product characterization (16, 18). Product impurities such as cellular DNA or protein should be considered, as well as process related impurities. The FDA recommends that specifications are in place for residual host cell protein and DNA. For residual DNA, a specification of less than 10 ng/dose and less than 200 bp in size is suggested. Processrelated impurities can include testing for the removal of reagents used during the process such as column chromatography materials, fetal bovine serum, or solvents used. All release testing for cell banks, virus banks, and bulk and final vialed product lots should be summarized on certificates of analysis. When gene therapy vectors will be continually made from the same cell source, cell banking is important. A formal cell banking system should be used, which usually includes the use of a Master Cell Bank (MCB) and Working Cell Bank (WCB) (13, 16, 17, 21). A Working Cell Bank is derived from the Master Cell Bank and is used for production runs. The WCB only requires limited testing because it is derived from the MCB, which is tested thoroughly. The origin and history of each cell bank should be well documented. Cell banks are fully tested and characterized, and provide a consistent source of qualified cells for future production runs. The specific tests required will depend on the origin of the cells and is assessed on a case-by-case basis. For example, if the cell line is of human origin, tests should be performed for a variety of human viruses. Cell banks must be free of adventitious viruses and mycoplasma, and must be sterile. Adventitious viruses are generally assessed using two assays, an in vitro assay and an in vivo assay. Both assays screen for a variety of different nonspecific viruses. In addition to the nonspecific virus assays, many

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tests for specific viruses must be performed (16, 17). Methods for cell culturing also commonly use fetal bovine serum or porcine trypsin. Therefore, if these materials are not certified free of viruses, then the cell banks would need to be tested for porcine parvovirus and bovine viruses. The identity and stability of the banks should be confirmed. It should be noted that it is possible to manufacture a product without the use of a cell-banking system; however, the final product would generally need to be tested for all the required cell bank tests in addition to the final product release tests. This testing would need to happen every time a lot is manufactured, whereas if the lot was produced from a qualified cell bank source, the final product specific tests would only be needed for each production run. It is therefore recommended that a cell bank system be used when possible so that a well-characterized cell source is available for future production runs. Virus banks are also important and should be well characterized and tested (13). A master virus bank (MVB) provides a stock of virus that is used as the inoculum for all production runs. A Working Virus Bank (WVB) can also be prepared and it requires less testing because it was derived from the MVB (16). An MVB is tested similar to a master cell bank. Tests are performed for specific viruses, along with in vivo and in vitro adventitious viruses, sterility, and mycoplasma. An adenovirus MVB should be tested for replication-competent adenovirus (RCA) because RCA is a potential byproduct of adenovirus vector production. The target specification for RCA is less than one RCA in 3 × 1010 virus particles (16). Other characteristics to test for include identity (nucleotide sequence), activity of the transgene, number of vector particles, and infectious titer. The FDA currently recommends a ratio of viral particle to infectious units of less than or equal to 30:1 (16). A stability program should also be put in place for investigational products to ensure the quality of the final formulated vialed product in use during the clinical trial (16). A subset of the release tests can be performed at various intervals on the investigational product after storage at its current storage temperature/conditions. A written stability plan should be followed that includes storage conditions, tests, specifications, and time points to be tested. Stability analysis should include evaluation of product potency, integrity, and sterility. Typical testing intervals used are 0, 3, 6, 9, 12, 18, and 24 months. To conserve on product, sterility testing can be performed at time zero, an intermediate time point, and at the end of the stability study (16). Testing the final product should continue throughout the duration of the clinical trial to indicate the product meets specifications during its use. When only small lot sizes are available for a study, reduced testing and frequency can be negotiated with the appropriate regulatory agency.


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5.3. Quality Assurance

There should always be appropriate documentation of the manufacturing and testing, with quality oversight. Quality oversight involves both the Quality Control and Quality Assurance units. The Quality Control unit is usually the department that does the analytical testing needed for the in-process manufacturing and release of the final product. Quality Assurance has the responsibility to approve and reject all components, in-process material, packaging materials, labeling, and drug products. They should also review documentation of the production records to assure no errors have occurred or if errors have occurred, that they have been thoroughly investigated. It is important to have separate quality assurance oversight from manufacturing personnel to ensure quality of your product (15, 18). This way the person responsible for assurance that the manufacturing and testing have been performed properly and met acceptance criteria are separate from the personnel responsible for conducting the manufacturing and testing, and this prevents a potential conflict of interest. This separation provides extra assurance that the product was made per written documentation and when tested for its identity, strength, quality, purity, and potency, met its intended release specifications.

5.4. Gene Therapy Product-Specific Issues

There are additional manufacturing issues for gene therapy products that require consideration. Because of the presence of unexpected genetic material commonly observed in vectors, tests for identity now include vector sequencing before the initiation of Phase I clinical studies. This assures that the manufactured vector has the appropriate characteristics and that production did not alter the structure of the vector. Complete sequencing of plasmid vectors should be performed to check for sequence accuracy. The FDA requires complete sequencing of vectors that are 40 kb or smaller (16). Larger vectors should have the transgene sequenced along with the 3´ and 5´ flanking regions, the gene insert, and any regions of the vector that are modified. Vector construction including the source materials used need to be documented and characterized. Known regulatory elements such as promoters or enhancers need to be identified. Genetic stability needs to be examined because it can lead to coding errors and changes in protein expression. Use of penicillin-like (beta-lactam) antibiotics for selection is not recommended because of the possibility of allergic reactions (13, 16) and possible interference with microbial sterility tests. For viral vectors, genetic stability needs to be studied with respect to the possibility of recombination resulting in the generation of replication-competent viruses (RCV). In the case of replication-defective or selective vectors, the virus seeds should be free of RCV. Testing for replication-competent retrovirus (RCR) is recommended during the various stages of retrovirus vector production (7, 16). The history and derivation of the vector,

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including detailed descriptions of all constructs used to generate a final vector and raw materials used for construction should be documented. Another important issue is stability of the gene therapy product (16, 17, 22). As mentioned previously, stability of the final formulated vialed product must be demonstrated for use in the clinical trial. Establishing and following a written stability plan is highly recommended. The plan should include the storage conditions, tests, specifications, and time points to be tested. Accelerated stability studies should be considered for later stages of development. Stability should also be evaluated with the specific infusion apparatus to be used in both the toxicology study and the clinical study. Stability with these materials should be demonstrated over the time period they will be used. In addition, stability data should demonstrate that the integrity, sterility, and potency of the product will be maintained under shipping conditions. Formal shipping stability studies should also be considered, and will be needed by later stages of product development. Reference standards should also be considered (22). A laboratory reference standard made at a reduced production scale can be used as a starting reference standard. A well-characterized laboratory reference standard in sufficient quantity makes the process of showing identity and bioequivalence to subsequent product lots much easier. Again, it is important to document how the reference standard was made and tested. A commercial reference standard is available in some instances such as for adenoviruses that can be used to calibrate assays for infectivity and particle number (www.wilbio.com). Product tracking and labeling are important considerations (16, 17), especially for autologous or patient-specific products. A formal product tracking system should be set up to track the product from collection to patient administration. Procedures that segregate the product from other products need to be in place. Labeling of patient-specific products is also critical and it should include patient identifiers as well as the date of manufacture, product name, and storage conditions.

6. US Regulatory Submissions As previously mentioned, in the USA, to conduct clinical trials with investigational agents, an IND application must be submitted to and reviewed by the FDA. The IND process provides federal oversight of clinical studies, and the main focus of the FDA in reviewing all types of INDs is to assure the safety and rights of clinical trial participants. The FDA can provide nonbinding advice


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on information that will be included in an IND at a pre-IND meeting. This is a formal meeting (4) that must be requested by the sponsor. The FDA will review the information and questions submitted before the meeting, and draft responses are communicated to the sponsor before the meeting. This enables cancellation of the meeting if no further discussion of the material is warranted. If the meeting is conducted, the FDA is required to send a copy of the meeting minutes within 30 days of the meeting. Pre-IND meetings provide sponsors with valuable advice on clinical, nonclinical, and manufacturing issues, which can expedite drug development. They also provide the advantage of introducing the FDA to new products that will shortly be coming to them for review. The IND application consists of ten sections as outlined in Table 4 (19, 20). The main information required in an IND includes: manufacturing information (composition, manufacture, stability, controls), nonclinical pharmacology and toxicology information, clinical protocols and investigator information (including FDA Form 1572), and previous human experience with the investigational drug or related compounds, if available. Item 1 of the IND is the FDA Form 1571, which must accompany every IND submission to the FDA including IND amendments. By signing the 1571 form, the sponsor agrees to abide by all the applicable regulations including not initiating the study until 30 days after the FDA’s receipt of the IND and/or not to begin or

Table 4 Content of an investigational new drug applicationa Cover Letter Item 1: Form 1571 Item 2: Table of Contents Item 3: Introductory Statement and General Investigational Plan Item 4: [Reserved] Item 5: Investigator’s Brochure Item 6: Protocol Item 7: Chemistry, Manufacturing, and Control Data Item 8: Pharmacology and Toxicology Item 9: Previous Human Experience Item 10: Additional Information a

21CFR312.23

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continue studies that are on hold. The FDA does not approve an IND. Unless the FDA indicates otherwise, an IND can be activated and the clinical trial initiated 30 days after the FDA’s receipt of the IND. The IND receipt date is included in the IND acknowledgement letter from the FDA, which is sent to the sponsor within 2 weeks of IND submission. This letter also contains the IND number, and other requirements for the investigational agent. The amount of information included in an IND depends on the novelty of the drug, the extent to which it has been studied previously, the known or suspected risks, and the developmental phase of the drug (i.e., Phase I–IV). It is expected that sufficient manufacturing and control information is submitted to assure the proper identification, quality, purity, and strength of the investigational drug. Nonclinical pharmacological and toxicological studies should be included from which the sponsor has concluded that it is reasonably safe to conduct the proposed clinical investigations. The central focus of the initial IND submission should be on the general investigational plan and the protocols for specific human studies. The FDA will carefully review the clinical monitoring plan, testing for product specific safety issues, and the long-term follow-up plan. For gene therapy INDs, the sponsor must include in the IND a response to the FDA’s March 6, 2000 Dear Gene Therapy IND or Master File Sponsor Letter (http:// www.fda.gov/cber/ltr/gt030600.htm) detailing manufacturing Quality Assurance/Quality Control and clinical trial oversight and monitoring. As previously mentioned, long-term patient follow-up, if required, and patient sample collection and testing for product-specific safety concerns, such as RCR or immunogenicity, should be included in the IND protocol. Additional guidance on regulatory submissions, which includes an IND template, can be found in the National Cancer Institute publication “The Sponsor’s Guide to Regulatory Submissions for an Investigational New Drug” (http://wwwbdp.ncifcrf.gov/pdf/GuidetoRegSubs.pdf). Gene therapy INDs should be sent to the Office of Cellular, Tissues and Gene Therapy, CBER, FDA (www.fda.gov/cber/ genadmin/octgtprocess.htm). Upon receipt of the IND, the IND is processed and scientific experts in manufacturing, nonclinical pharmacology/toxicology, clinical, and statistics review the IND sections as appropriate. Approximately 1 week before the 30-day review due date, the FDA review team meets and discusses the need for clarifications from the sponsor, including whether a clinical hold is necessary. Clarifications from the sponsor are usually necessary, therefore, it is prudent for the sponsor to contact the FDA before the due date to resolve any issues so that initiation of the clinical study is not delayed. Once the IND study is initiated, amendments to the IND can include protocol amendments, new protocols, manufacturing


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information, nonclinical study reports, IND safety reports, updated Investigator’s Brochures, IND annual reports, and other information amendments. Subsequent amendments to the IND that contain new or revised protocols should build logically on previous submissions and should be supported by additional information, including the results of animal toxicology studies or other human studies as appropriate. Annual reports to the IND should serve as the focus for reporting the status of studies being conducted under the IND and should update the general investigational plan for the coming year. Annual report requirements are outlined in 21CFR312.33 and include adverse event and response outcome summaries, patient sample testing results (i.e., RCR testing), and long-term follow-up data, if applicable. The annual report should also contain confirmation that manufacturing Quality Assurance/ Quality Control and clinical trial oversight and monitoring have been conducted as indicated in the original IND submission and, if not, modifications should be provided.

7. Summary Gene therapy products are highly regulated, therefore, moving a promising candidate from the laboratory into the clinic can present unique challenges. Success can only be achieved by proper planning and communication within the clinical development team, as well as consultation with the regulatory scientists who will eventually review the clinical plan. Regulators should not be considered as obstacles but rather as collaborators whose advice can significantly expedite the product development. Sound scientific data is required and reviewed by the regulatory agencies to determine whether the potential benefit to the patient population outweighs the risk. Therefore, compliance with GMP and GLP principles to ensure quality, safety, purity, and potency of the product, and to establish “proof of concept” for efficacy, and for safety information, respectively, is essential. The design and conduct of the clinical trial must adhere to Good Clinical Practice principals. The clinical protocol should contain adequate rationale, supported by nonclinical data, to justify the starting dose and regimen, and adequate safety monitoring based on the patient population and the anticipated toxicities. Proper review and approval of gene therapy clinical studies by numerous committees, and regulatory agencies before and throughout the study allows for ongoing risk assessment of these novel and innovative products. The ethical conduct of clinical trials must be a priority for all clinical investigators and sponsors. As history has shown us, only a few fatal mistakes can dramatically alter the regulation of

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investigational products for all individuals involved in gene therapy clinical research, and further delay the advancement of gene therapy to licensed medicinal products.

References 1. Gonin, P, Buchholz, CJ, Pallardy, M, Mezzina, M. Gene therapy bio-safety: scientific and regulatory issues. Gene Therapy 2005; 12:S146–S152. 2. Guidance for Industry: Gene Therapy Clinical Trials- Observing Subjects for Delayed Adverse Events (draft), US Department of Health and Human Services, Food and Drug Administration, Center for Biologics Evaluation and Research, August 2005. 3. US Code of Federal Regulations, Title 21, Part 312, Section 3 (21CFR312.3) 4. Guidance for Industry: Formal Meetings with Sponsors and Applicants for PDUFA Products, US Department of Health and Human Services, Food and Drug Administration, Center for Drug Evaluation and Research, Center for Biologics Evaluation and Research, February 2000. 5. Good Clinical Practice: Consolidated Guidance (ICH E6), International Conference of Harmonization, June 10, 1996. 6. Dear Gene Therapy IND or Master File Sponsor Letter, CBER, FDA, March 2006 (http:// www.fda.gov/cber/ltr/gt030600.htm) 7. Supplemental Guidance on Testing for Replication Competent Retrovirus in Retroviral Vector Based Gene therapy Products and During Follow-up of Patients in Clinical Trials using Retroviral Vectors, US Department of Health and Human Services, Food and Drug Administration, Center for Biologics Evaluation and Research, November 2006. 8. Guidance for Industry: Information Program on Clinical Trials for Serious or Life-Threatening Diseases and Conditions (draft), US Department of Health and Human Services, Food and Drug Administration, Center for Drug Evaluation and Research, Center for Biologics Evaluation and Research, January 2004. 9. US Code of Federal Regulations, Title 21, Part 312, Sections 30–33 (21CFR312.30-33). 10. Frederickson RM , Carter BJ , Pilaro AM . Nonclinical toxicology in support of licensure of gene therapies. Mol Ther. 2003; 8:8–10 .

11. Pilaro AM, Serabian MA. Preclinical development strategies for novel gene therapeutic products. Toxicol Pathol 1999;27:4–7. 12. Gonin P, Gaillard C. Gene transfer vector biodistribution: pivotal safety studies in clinical gene therapy development. Gene Therapy 2004, 11:S98–S108. 13. Guidance for Industry: Guidance for Human Somatic Cell Therapy and Gene Therapy, US Department of Health and Human Services, Food and Drug Administration, Center for Biologics Evaluation and Research, March 1998. 14. US Code of Federal Regulations, Title 21, Parts 210 and 211 (21CFR210–211). 15. Guidance for Industry: CGMP for Phase 1 investigational Drugs US Department of Health and Human Services, Food and Drug Administration, Center for Drug Evaluation and Research, Center for Biologics Evaluation and Research, June 2008. 16. Guidance for FDA Reviewers and Sponsors: Content and Review of Chemistry, Manufacturing, and Control (CMC) Information for Human Gene Therapy Investigational New Drug Applications (INDs), US Department of Health and Human Services, Food and Drug Administration, Center for Biologics Evaluation and Research, April 2008. 17. Guidance for FDA Reviewers and Sponsors: Content and Review of Chemistry, Manufacturing, and Control (CMC) Information for Human Somatic Cell Therapy Investigational New Drug Applications (INDs), US Department of Health and Human Services, Food and Drug Administration, Center for Biologics Evaluation and Research, April 2008. 18. Bauer SR, Pilaro AM, Weiss KD. Testing of Adenoviral Vector Gene Transfer Products: FDA Expectations. Adenoviral Vectors for Gene Therapy, 2002, 21:615–654. 19. US Code of Federal Regulations, Title 21, Part 312, Section 23 (CFR312.23). 20. Guidance for Industry: Content and Format of Investigational New Drug Applications (INDs) for Phase 1 Studies of Drugs, Including Well-Characterized, Therapeutic, Biotechnology-

Regulatory Aspects for Translating Gene Therapy Research into the Clinic derived Products, US Department of Health and Human Services, Food and Drug Administration, Center for Drug Evaluation and Research, Center for Biologics Evaluation and Research, 1995 21. Guidance on Quality of Biotechnology/Biological Products; Derivation and Characterization of Cell Substrates Used for Production of

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Biotechnological/Biological Products (Q5D), International Conference of Harmonization, July 16, 1997. 22. Aurigemma R., Tomaszewki JE, Ruppel S, et al. Regulatory Aspects in the Development of Gene Therapies. Contemporary Cancer Research, Cancer Gene Therapy. September 2004, 29: 441–472.

Chapter 23 Ethics of Cancer Gene Transfer Clinical Research Jonathan Kimmelman Summary Cancer gene transfer is a relatively novel intervention strategy. In part because of this novelty, trials often present greater uncertainties than those investigating more conventional approaches. In the following review, I examine how this greater uncertainty might affect how clinical studies are designed, when they are initiated, their degree of risk, and whether such risk can be justified in terms of therapeutic benefit. The review also discusses two other ethical issues presented by gene transfer clinical research: fairness in subject selection and communications with the public. I conclude with a series of recommendations directed toward researchers, policymakers, and ethics committee members. Key words: Cancer, correlative studies, ethics, gene transfer, informed consent, phase 1, risk uncertainty, value.

1. Introduction Why an entire chapter on the ethics of gene transfer (GT) in the oncology research setting? Many readers of this volume will probably wonder whether the ethics of cancer gene transfer is distinguishable from, say, that for cytotoxic or targeted cancer therapies. If cancer gene transfer research presents no distinctive challenges, then all that is needed is a review of the basic tenets of research ethics. Instead, I hope to persuade the reader that there are certain aspects of GT clinical research in oncology that diverge from classic cancer trials. Although the same broad principles of research ethics apply—namely, respect for persons, beneficence, and justice (1–3)—continuities with more familiar realms of clinical research potentially obscure ethically important differences. Yet in making this argument, I also emphasize that, strictly speaking, Wolfgang Walther and Ulrike S. Stein (eds.), Methods in Molecular Biology, Gene Therapy of Cancer, vol. 542 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-561-9_23

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few of the ethical issues encountered in GT oncology research are unique, and many of these are paradigmatic for a series of emergent ethical challenges in other areas of translational oncology clinical research. The central thesis is that GT cancer trials (like other cuttingedge experimental interventions) involve a very high degree of scientific and clinical uncertainty. This has implications for when such studies are initiated, how they are designed, the assessment of their risks, and the conduct of informed consent. I also briefly discuss issues like justice and public expectations. The chapter closes with recommendations for designing and implementing cancer GT studies (Table 1). The reader is directed to other thoughtful sources for a discussion of related ethical and policy matters, such as how GT research is regulated (4–7), animal welfare and protections (8–11), and a broader account of the ethics of early phase oncology studies (12–16).

Table 1 Recommendations for researchers contemplating cancer GT clinical studies Value: Investigators should design studies such that “negative” outcomes—like adverse events or observing no biological effect—will lead to new insights and stimulate further research (rather than simply closing off one research avenue). When to Begin: Researchers should only begin studies when there is a modest translational distance between preclinical and clinical studies. Doing so requires that internal and external validity in preclinical studies be maximized. The Justification of Risk: Investigators and ethics committees should generally avoid using the therapeutic value of a study drug to justify risks for such studies. Instead, risks should be justified by the value of the knowledge gained. Assessing Risk: Risk estimates of study interventions in translational trials should generally be based on the agent’s “upper bound” estimate rather than an investigator’s “best estimate.” Consent and Therapeutic Misestimation: Investigators should provide specific information on the probability, nature, and duration of benefits. In general, they should disclose that translational phase 1 studies are unlikely to lead to dramatic clinical improvements and that the primary beneficiaries of the study are future patients. Consent and Therapeutic Misconception: Investigators should focus their consent discussion on elements of the study design that frustrate a volunteer’s therapeutic objectives. Thus, they should discuss subtherapeutic dosing, procedures like tissue biopsies, rigidity of dosing, or posttrial access to the study intervention (if an agent involves chronic application). Justice: Researchers should pursue studies in economically disadvantaged populations only when interventions seem likely to expand the capacity of existing health systems to address the urgent health needs of these populations. Interactions with the Public: Researchers should monitor public expectations, and consider the effects of their communications. In general, researchers should avoid presenting patients to the media to convey the significance of their research.

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2. Cancer GT and Uncertainty All human studies present uncertainties; the very goal of the research enterprise is the conquest of uncertainty. Yet, the nature and degree of uncertainty in clinical studies will vary. Consider a side-by-side trial aimed at comparing the efficacy of a single preoperative dose of a frequently used antibiotic combination against a three-dose regimen of the same antibiotic combination. Here, the clinical experience for both regimens is very extensive, and investigators have a reasonably clear picture of what kinds of outcomes to expect—which side effects to look for, when these will become manifest, which outcomes to measure, how much of each drug to use and how to administer it, etc. The trial is aimed at asking a narrow question: whether the simpler regime performs as well in controlling postoperative infection as the standard, three-dose regimen. Either it will prove equivalent or inferior. Contrast this scenario to a first-in-human phase 1 study. The safety profile for the drug in human beings is unknown, much less its efficacy. We might use preclinical studies to make certain predictions about side effects, but we have no bank of clinical experience from which to draw. Investigators do not yet know which types of adverse effects to expect, how long the drug will stay in the volunteers’ system, and whether it will have the desired biological effects. The study does not have a well-defined hypothesis; its future is much more open-ended (17). Obviously, the degree and nature of uncertainties between the antibiotic randomized controlled trial (RCT) and the phase 1 study varies. Yet even within categories, uncertainties will vary in character and degree. For example, a dose escalation trial combining two licensed agents presents a more narrow range of uncertainty than, say, a trial testing a monoclonal antibody that targets a recently discovered biological pathway. The former might demonstrate toxicities that do not occur with the single agents, or that occur at much lower doses than expected. However, the probability of something unexpected occurring is lower, and the range of uncertainty is more bounded by clinical experience than with the novel monoclonal antibody, which has never been administered to human beings. As a class, cancer GT interventions (indeed, all GT interventions) are relatively novel. That is, GT interventions apply agents for which clinical experience is relatively limited. For a variety of reasons, our ability to predict the properties and behavior of gene transfer agents has important limitations (18). First, they frequently involve active agents: vectors might be capable of executing biological programs (like conditional replication for oncolytic vectors), or they might recombine with wild-type virus

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to produce novel pathogens. In contrast, conventional drugs are passive compositions of matter. Second, GT interventions are simultaneously delivery devices as well as pharmacologic agents. Both vectors and transgenes can trigger toxic responses. Furthermore, the toxicities of each component cannot always be assumed to be additive. Third, GT agents often demonstrate a high degree of species specificity. For example, the main organism used to validate cancer GT, the mouse, does not develop infections when exposed to adenoviruses. As such, the predictive value of the mouse for modeling immunoreactivity is limited (indeed, most mice used in cancer preclinical research lack functional immune systems altogether). Similarly, transgenes often have speciesspecific properties. Indeed, the trend toward targeted and biological interventions in cancer translational research presents a safety paradox. On the one hand, targeted interventions should be safer because they are less likely to have off-target effects. On the other hand, their high degree of target specificity makes animal modeling of toxicity extremely difficult. Fourth, toxicities are often mediated through the immune system. In addition to the difficulty of modeling immune responses, any volunteer’s immunological reaction to a given vector will depend on their previous exposures to epitopes like those presented by the vector. In contrast, an individual’s response to a small molecule drug is less likely to be influenced by medical history. Fifth, some gene transfer vectors involve nonlinear dose–response curves (19). As such, the safety of dose escalation in a traditional phase 1 study is subject to greater uncertainty. Obviously, not all gene transfer interventions present the same degree and character of uncertainty. For instance, as of 2008, a protocol investigating the safety of intratumoral injection of oncolytic measles virus presents greater uncertainty than one testing intratumoral administration of Ad-p53. Nevertheless, this review will, on the one hand, focus on the former type of studies, while presuming that even the best-characterized gene transfer agents still present enough uncertainties as to make the central arguments largely applicable.

3. The Value of Early Phase GT Trials What are the ethical implications of human clinical testing pursued under conditions of radical indeterminacy? One place to begin an examination of how biomedical novelty might affect the trial process is with the seemingly simple question: why perform a phase 1 cancer GT study? Clinical trials are ethical only insofar as they produce valuable knowledge. The question might thus be


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reframed as “what kind of valuable knowledge should be sought in a phase 1 cancer GT study? Moreover, how might the knowledge sought in such studies differ from that sought in a more conventional cancer drug study? The conventional answer to the former questions is that phase 1 studies are medically valuable insofar as they provide adequate information for designing a phase 2 study (20). They do this by identifying major toxicities and determining the appropriate dose for studying efficacy in a subsequent study (21). This framework for defining the value of a phase 1 study might be termed a “regulatory model.” According to recent data, the likelihood that a phase 1 cancer GT study will actually accrue this type of value is small compared with more conventional cancer drugs. For example, the probability that a cancer drug will advance from phase 1 into phase 2 testing is high (approximately 77%); the figure for advancement from phase 1 through to licensure is about 27% (22). Although the study from which these figures are drawn did not stratify according to drug class, one can venture very crude comparable estimates for GT using public databases. According to a National Institutes of Health (NIH) database, 314 phase 1 cancer GT protocols have been submitted to the Office of Biotechnology Activities; 65 phase 2 studies have also been submitted. No cancer GT agents have yet been licensed in North America or Europe. Based on historical data, then, the probability that a GT cancer agent entering phase 1 studies will progress immediately into phase 2 studies is on the order of 20%; the probability of licensure is presently zero (23). Similar figures can be obtained using a worldwide database of GT clinical trials (24). This comparatively low number probably reflects that GT agents are relatively novel, and researchers still have much to learn to turn them into effective interventions. Phase 1 cancer GT studies have a much better shot at producing medically valuable knowledge—and, in the process, advancing application of GT—if they are designed along an alternative model of the value of a phase 1 study—what I will term the “translational model.” According to this model, phase 1 studies can produce knowledge about the properties of an agent that can stimulate refinement at the preclinical level, leading to another phase 1 study. Alternatively, they can produce findings that stimulate entirely different lines of research altogether. In their 1995 report to the NIH on the status of gene transfer, Stuart Orkin and Arno Motulsky tacitly endorsed this “scientific model.” According to them, gene transfer studies (like any translational study) “…need to address specific hypotheses, enabling investigators to interpret negative as well as positive findings” (25). A well-designed phase 1 cancer GT study, then, should aim to collect information that can enable future investigators to

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troubleshoot and refine their application, as well as other related applications. This might include a determination of whether the vector reaches appropriate targets, whether it has effectively transferred genes to target tissues, whether transgenes are expressed and at what levels, whether tumor response correlates with transgene expression, whether the intervention effectuates tumor response by the intended mechanism, or whether the vector causes subclinical toxicities or has undesirable biodistribution properties (26). Applying a scientific model of value to cancer GT studies often necessitates trial designs that might be different than for typical phase 1 cancer studies. The ethical analysis of when and how to initiate phase 1 testing, acceptable risk, and informed consent derives in no small part from a recognition that the objectives of phase 1 GT translational studies should have a more scientific and iterative character.

4. The Ethics of Translation: When to Initiate Testing One of the most persistent and unresolved ethical challenges for trials testing highly innovative approaches is the decision to initiate a first-in-human trial. Pursuing a study too early exposes volunteers to gratuitous risk and burden, and squanders resources for research. Too cautious an approach, however, potentially frustrates the autonomy of otherwise willing volunteers or leads to inefficiencies like delay and duplicative research. For later-stage clinical trials, research ethics has a readymade solution to the question of when an investigator is ethically justified in initiating a drug trial: clinical equipoise. This refers to a state of uncertainty among the expert community regarding the comparative merits of agents used in a controlled clinical trial (27, 28). Proponents of clinical equipoise argue that a trial is ethical only where it is designed in a way that can perturb equipoise at the trial’s completion, and thus change clinical practice. Equipoise is an attractive concept, in part because it simultaneously captures the obligations to promote the welfare of volunteers and maximize scientific value in a single concept that can be readily grasped and implemented by investigators, trial designers, referring clinicians, and Institutional Review Boards (IRBs). Unfortunately, the concept has limited utility for first-inhuman studies. For example, first-in-human trials are statistically underpowered and hence unable to perturb clinical consensus. If equipoise is to guide decisions about initiating first-in-human studies, at the very least it requires considerable interpretation and revision.


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I have elsewhere advanced a position, the principle of modest translational distance, which holds that researchers should not initiate human studies unless the number and character of assumptions linking a preclinical study to a clinical study are modest. The principle derives from the view that preclinical studies are performed to project what will occur in the clinical setting. This process of projection requires that the researcher hold in place a number of assumptions about similarities between the (laboratory) model and the (clinical) modeled. If these assumptions are numerous or extravagant in character (i.e., if the translational distance is great), the predictive value of the preclinical studies is more limited. Clinical studies that attempt to cross a large translational distance (i.e., that are launched on a thin evidentiary base) have two related ethical drawbacks. First, they expose human subjects to the risks of an agent for which evidence of safety or clinical utility is severely limited. Even if the risks are modest, the clinical investigator is literally wasting the time of the trial participant (time may be particularly scarce and scheduling complicated for the types of treatment refractory volunteers enrolled in translational cancer studies (29)) unless the study intervention is supported by a scientifically compelling rationale. Second, studies crossing a more than modest translational distance are less likely to fulfill scientific objectives. As noted already, most phase 1 studies in novel translational research arenas fail to progress smoothly toward licensure. Such “negative studies” should enable to investigator to troubleshoot their intervention or gain biological insights that lead to further inquiry. When translational distance is great, however, a “negative result” will typically have numerous plausible and/or trivial explanations. The latter, which might include observations that the agent never reached its target, might have easily been anticipated with less risk for volunteers had smaller-scale feasibility studies been performed. As such, translational studies should generally not be conceptualized as screening studies, but rather as experiments aimed at testing various hypotheses that, if refuted, likely invalidate an intervention strategy. 4.1. Internal Validity

One major factor that contributes translational distance is internal validity of preclinical experiments. Numerous studies from other translational research arenas have documented sporadic adherence to sound methodological practice (30–32). I am not aware of similar studies assessing the methodological rigor of GT preclinical studies. Nevertheless, a scan through the methods sections in journals like Molecular Therapy, Cancer Gene Therapy, Cancer Research, or Nature Medicine does not lead one to think that the situation is any different in cancer GT research. Consider, for example, the main outcome measure for cancer drugs: death. One might be tempted to view death as a “hard

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endpoint” whose measure does not require blinded assessment. Yet, many animal welfare policies specifically discourage researchers from using death as an endpoint (33–35) (this author strongly endorses such policies). Instead, they urge killing moribund animals to limit suffering. However, without an explicit and objective criterion for determining moribund status, or allocation blinding of laboratory personnel, the decision to kill an animal—and the resulting survival curves—are susceptible to bias. On the other hand, consider randomization. Because cancer studies typically use inbred animals that are housed under identical conditions, many researchers consider pointless the random treatment allocation of animals. Nevertheless, injury states like tumor grafting are subject to variation, which potentially introduces another source of bias. A third and common weakness in preclinical research is failure to perform a priori power calculations. Corner a researcher at a scientific conference and she will confess that researchers often “expand” their sample sizes until a treatment effect crosses the threshold of statistical significance. Unless such studies are followed by others using a priori power calculations, such methodological deficiencies can lead to the needless launch of clinical testing. 4.2. External Validity

Another major source of “translational distance” is external validity: the degree to which the preclinical experiment recapitulates the clinical setting. In translational research, one of the biggest challenges for external validity is the quality of animal models. The main model used in cancer preclinical research, the murine xenograft of human tumor tissue, suffers numerous weaknesses as a model of human cancers. First, murine xenograft models are immunodeficient. Given the role of the immune system in effecting response and/or causing toxicity for gene transfer interventions, this represents a major shortcoming. Mice are also typically inbred; human patients are outbred. Murine tumors are at least an order of magnitude smaller than those in human patients; their morphology is different, and they are less prone to hypoxia or genetic heterogeneity. Tumor tissue in animal models is passaged for many generations before grafting, whereas human tumors are always “fresh” (36). Transgenic models show promise for alleviating these problems. Nevertheless, they too are subject to limitations like size, physiological differences with human beings, inbreeding, and intellectual property (37).

4.3. Maintaining Modest Translational Distance

How might a translational researcher establish a sufficiently firm evidentiary basis for launching a human study involving a novel GT strategy? My recommendations divide into three categories. The first is that the researcher should ensure (and the review committee should demand) methodological rigor in preclinical studies by applying practices that are routine for clinical research. These include the use of a priori power calculations, randomized


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treatment allocation, blinded treatment allocation, and blinded (or automated) assessment. Departures from such practices might be justified, but they should always be explained (on the other hand, the use of such procedures should not be taken as a guarantor of validity!). The second set of recommendations would be for research teams to use alternative models where strategies are highly innovative or expected to involve pathways that are poorly recapitulated by xenograft models. One promising model is the companion animal (38–40). Dogs develop cancer at roughly the same rate as human beings; at about 60 million in the USA (41), they also represent a reasonably large population of potential study subjects. Unlike many murine models, dogs are outbred and immunocompetent; their tumors are also larger and spontaneous. They are ethically advantageous in that companion animal studies are less likely to deliberately impose nontherapeutic suffering on an animal, and more likely to merge therapeutic objectives for the animal subject (42). In numerous instances, researchers have tested novel cancer drugs in companion animals (43–45), and the US National Cancer Institute recently established a comparative oncology program to promote such studies (46). A third recommendation would be to perform smaller-scale, hypothesis-testing trials before launching a classic phase 1 dose escalation study. Such feasibility studies have been used profitably in cystic fibrosis GT (47, 48) and muscular dystrophy cell therapy (49). Within cancer, phase 0 studies typically administer very small doses of a study agent for a short period of time to study an agent’s distribution, pharmacokinetics, pharmacodynamics, and biological effects. Phase 0 studies have the advantage of limiting a volunteer’s systemic exposure to a potentially toxic agent, minimizing the commitment of resources toward a particular investigational agent, and strengthening the evidentiary base for a phase 1 translational trial; the ethics of phase 0 studies has been reviewed elsewhere (50).

5. Risk and Cancer GT Studies Thus far, this review has emphasized that GT cancer trials have scientific dimensions that necessitate greater emphasis on correlative studies and/or feasibility studies. Because such studies often involve burdens without offering volunteers any compensatory medical benefit, they invite obvious ethical questions about risk and benefit.

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5.1. Correlative Studies and Risk

As noted in the previous section, cancer GT studies will often collect tissue specimens to determine whether an agent has the expected biological effects on target tissues. This can necessitate the use of invasive tissue biopsies that have no conceivable medical benefit for volunteers. Is it ethical for oncologists to ask cancer patients who are already debilitated by their illness to submit to painful organ biopsies that have no medical justification? Is it ethical to impose such procedures as a condition for study access? Will cancer patients, who often feel a debt of gratitude toward their oncologists, feel at liberty to decline painful biopsy procedures? Indeed, is it realistic to expect that emotionally debilitated cancer patients will understand that such procedures are aimed at benefiting society, not them? Unfortunately, there are no simple answers to these questions. Evidence that cancer patients are often willing to endure significantly greater risk for nontherapeutic procedures than their oncologists find acceptable suggests grounds for concern (51). At the same time, critics of such procedures should bear in mind that they are ethically very similar to the use of subtherapeutic dosing in phase 1 dose escalation studies. That is, the first few volunteers in a phase 1 study typically receive levels of a drug that are well below those expected to induce a therapeutic response. As such, initial dose cohorts involve risk without expectation of compensating benefit. There are several possible courses of action for enhancing the ethical strength of a proposal to conduct correlative or “nontherapeutic” feasibility studies. The first is to pursue initial studies in cancers where tissues are readily accessible—either because of tumor location (e.g., head and neck cancers) (52) or because of planned and therapeutic surgical procedures (e.g., tumor resection). A second, emerging possibility is the use of noninvasive imaging techniques to monitor vector transduction, biodistribution, or gene expression (53–55). Third is a more rigorous approach to informed consent, to assure that volunteers fully understand the nontherapeutic nature of such procedures (this will be discussed below). Last, investigators should request autopsy from volunteers before they enroll in a study. In other contexts, some have argued that willingness to undergo autopsy should be an inclusion criterion (although, of course, volunteers should always be informed of their right to withdraw consent) (56).

5.2. Ethics and the Risk of Cancer Gene Transfer Agents

Closely related to the uncertainty that underwrites my argument about value, trial design, and trial initiation is the issue of risk for GT agents. Almost 2 decades of GT clinical trials in cancer provide some assurance that, when compared with classic cytotoxic drugs, GT agents have a significantly better safety profile. Several years ago, two teams of researchers performed meta-analyses of phase 1 oncology studies with the aim of characterizing their


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risks and benefits. The studies are imperfect—the samples are potentially biased, clinical benefit is inferred from tumor response, and, obviously, the data are drawn from open-label studies. Nevertheless, these best estimates show a death rate in GT studies significantly below that for cytotoxic drug studies (0.67–0.80% vs 0–0.19%). However, they also indicate a much lower response rate: in one of the meta-analyses, partial and complete response rates in gene transfer trials were 3.3%; the figure for cytotoxic drugs was on the order of 11%. Although serious adverse events have occurred in cancer GT studies, (57) by comparison with the types of poisons tested in traditional oncology trials, GT studies seem homeopathic—in both the best and worst senses of the term (58). 5.3. Risk and Uncertainty

Nevertheless, there are two important respects in which GT trials can be said to present challenges for the ethical management of risk. The first concerns uncertainty and complexity (18). For reasons specified in the second section (Cancer GT and Uncertainity) above, risks in cancer GT studies may be difficult to predict with confidence. The conventional regulatory response to uncertainty is generally to assume greater risk, even if best risk estimates suggest relative safety. Thus, for example, in deciding starting doses for healthy volunteer first-in-human trials involving novel agents, the US Food and Drug Administration (FDA) typically recommends that investigators start at doses ten times lower than the predicted safe starting dose (59). Therefore, although available evidence would suggest a favorable safety profile for phase 1 cancer studies, the uncertainties surrounding GT generally support a position that regards such studies as presenting nontrivial risk. The uncertainties surrounding such studies have at least three other implications with respect to how phase 1 trials are conducted. First, researchers should execute trials that escalate risk in a cautious and incremental fashion. For example, cohorts should be observed carefully for adverse events before moving to a higher dose cohort. In addition, researchers should monitor volunteers for subclinical toxicities that might provide some indication of whether toxicity thresholds are near. Another incremental strategy for minimizing risk is a “staged” approach to clinical testing, in which the safety of novel agents intended for systemic administration is first confirmed by intratumoral injection, then by administration to a body cavity (e.g., intraperitoneal), and then arterially, and finally, intravenously (60).

5.4. Risk and Bystanders

The second way cancer GT studies present challenges to ethical management of risk is that they can present risks to parties other than the study volunteer. Although this includes the possibility of vertical transmission, I will focus my discussion around the possibility of vector transmission to a volunteer’s social contacts

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(what I have elsewhere called “bystander risks”) (61). This is a particular concern where protocols involve living or conditionally replicating agents such as genetically modified bacteria (62, 63) or oncolytic viruses. Bystander risk implicates an additional layer of complexity in managing risk in that the risks will depend, in part, on the behavior of study volunteers—much less the behavior of microorganisms outside the controlled clinical setting. According to one 2004 review, the extent to which conditionally replicating vectors present transmission risks is not well studied; vector shedding has been detected in some studies (64), but I am not aware of any documented transmission to third parties. Existing ethics codes do not effectively address bystander risks, and current management practices, like Institutional Biosafety Committee (IBC) review, do not necessarily safeguard all ethical interests of bystanders (e.g., there are no provisions for informing or obtaining consent from bystanders). Nevertheless, there are sound ethical reasons to protect bystanders by establishing formal risk disclosure mechanisms where studies involve potentially transmissible agents (65).

6. The Research– Treatment Distinction: Phase 1 Trials and Direct Benefit

Do Phase 1 GT Trials Have Therapeutic Utility? Another ethical issue extending from this discussion of risk and uncertainty is whether GT agents in translational phase 1 trials should count as therapeutic interventions. On the one hand, proponents of this view argue, variously, that their risks and benefits are probably comparable to those for off-label treatments (66), and that there are instances of Lazarus-like responses in translational phase 1 studies (67, 68). Countering these views are those who argue that benefits are too improbable to count as therapy, and above all, that the primary objective of research is to answer biomedical questions rather than serve an individual patients’ needs. Proponents of the latter view have coined a term to diagnose the failure to distinguish clinical care (which tends to be individualized and flexible) with research (which involves randomization, masking, and protocol rigidity): the therapeutic misconception (69). The resolution of this controversy has important ramifications for informed consent. A large body of evidence indicates that the vast majority of patients enter phase 1 oncology studies with therapeutic objectives (13, 70). Indeed, some investigators conceive phase 1 studies as embodying a legitimate therapeutic modality (71, 68). Moreover, several surveys show that phase 1


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study volunteers overestimate the probability of benefit when compared with historic estimates of benefits (72) (this has been called “therapeutic misestimation” (73)), and overestimate the probability of benefit compared with estimates of clinical investigators (74) (in addition, there is some suggestion in these studies that oncologists themselves often overestimate the probability of benefit when compared with historic levels). If phase 1 trials are a species of research rather than care, then three concerns follow. First, research subjects who enter phase 1 trials are generally not providing valid informed consent. Second, clinicians who knowingly enroll patients harboring therapeutic misconceptions are abetting misinformed consent. Third, clinicians who regard phase 1 trials as therapeutic are, at best, conceptually muddled and, at worst, deceiving their desperately ill subjects. The problem with this debate is that partisans on both sides have tended to view the research–treatment distinction—and, in particular, the function of phase 1 studies—as categorical and fixed (75). However, phase 1 trials are a type of technology, and technologies can be interpreted and applied in a wide variety of ways. The internet, for example, was originally designed by the Defense Advanced Research Projects Agency to provide a communication network in the event of a nuclear missile attack. It has since been “interpreted” as a system for ordering books on Amazon. A similar argument can be made about phase 1 trials: reinterpretation of phase 1 studies as serving therapeutic objectives has led to reforms, like accelerated dose escalation regimes, that are designed to enhance their therapeutic value. Of course, this reinterpretation is not without important hazards: one recent study indicated that novel dose escalation regimes might increase risk without compensatory therapeutic benefits gains (76). However, what such reforms sacrifice in terms of physical risks and benefits, they gain by better aligning the objectives of clinical investigators with those of research volunteers. For these reasons, it seems inappropriate to dismiss the notion that phase 1 cancer trials have therapeutic value, or that volunteers who enter phase 1 trials seeking treatment are somehow misguided. Nevertheless, there are compelling ethical reasons for clinical investigators, ethicists, and policymakers to avoid using therapeutic value to justify risks in studies of novel interventions like GT (unless, of course, the agent is tested in combination with another validated therapy). For one, based on cumulative experience with first-in-human GT trials, probability of cure or significant clinical improvement is less than 1%—a figure that several commentators have advanced as a threshold below which medical interventions should be deemed futile, and, hence, unethical to offer as treatment (77, 78). Although this threshold has many critics, regulatory agencies do not typically grant licensure for

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interventions for which fewer than 5% of patients show efficacious responses (unless, of course, the drug had major quality of life benefits). Perhaps one of the most extreme examples is the licensure of interleukin (IL)-2 for the treatment of stage IV renal cell carcinoma, where complete responses were observed in only 7% of volunteers (79). Contrast such low probabilities of clinical benefit against the very high probability that such trials perform a valuable service for the biomedical community and future patients, much less sponsors, investigators, and institutions, and it should be apparent that the utility of phase 1 trials of novel therapeutics is overwhelmingly weighted toward the interests of others (80), and that contemporary policy is generally inconsistent with such distributions of risk and benefit. One further argument against justifying phase 1 studies on therapeutic grounds is this: should the agent prove promising enough to test in randomized controlled trials, how will the investigators then justify randomizing half of their volunteers to control groups? The position advanced above has two important implications for the assessment of risk in phase 1 GT cancer studies. All major research ethics codes demand that risks of clinical studies be balanced favorably against benefits. Benefits for clinical trials divide into those for volunteers (direct benefits) and those flowing to society through the production of generalizable knowledge (what some commentators have termed aspirational benefits). The foregoing discussion suggests that the therapeutic value of phase 1 GT cancer studies should generally not be used as a justification for risk.

7. Consent Practices and Cancer GT Studies

A second implication of the position argued above is that informed consent practices should clearly aim at thwarting therapeutic misestimations and misconceptions by reflecting the improbability of direct benefit and describing trial design features that might interfere with a volunteer’s therapeutic objectives. Regarding the former, studies of consent documents used in phase 1 GT oncology trials show that fewer than a quarter state that any direct benefit—for example, cure or major clinical improvement—is improbable (81, 82). Instead, consent documents tend to offer vague descriptions, like the statement “we can not guarantee this drug will shrink your tumors.” This statement contains virtually no information about the probability or magnitude of benefits. In contrast, the statement “participating in this study is unlikely to result in major clinical improvement” is far more informative and accurately represents current knowledge about benefits for phase 1 studies. Along a similar


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vein, consent forms used in oncology tend toward therapeutically loaded language by using terms like “treatment” and “gene therapy” instead of “study agent” or “gene transfer.” With respect to therapeutic misconceptions, consent forms and discussions have a long way to go in terms of describing how phase 1 studies might frustrate certain therapeutic objectives. Although consent forms used in phase 3 studies routinely disclose blinding and randomization, it is far less common for consent forms used in phase 1 studies to state clearly that initial volunteers will receive doses of a study agent that are well below those expected to induce a clinical response, or that volunteers receiving subtherapeutic doses are ineligible to receive doses that are more promising in subsequent treatment cycles. Indeed, in one anthropological study of consent discussions surrounding a cancer GT phase 1 study, volunteers in the lowest dose cohort were consistently told that low doses were likely to be safer; however, they were never told that higher doses were more likely to induce a response. The relationship was reversed when high-dose cohorts were consented: volunteers were told that they would receive a dose that was more likely to induce a response; however, they were never told that they would also receive doses that were less likely to be safe (83). Similarly, cancer GT protocols occasionally are designed in ways that categorically thwart therapeutic objectives. In one example, researchers pursuing a combination therapy trial used a volunteer’s uninjected tumors as a control in measuring response rates for tumors injected with an oncolytic virus (84). Although the standard agents in this study likely had therapeutic value, the GT component did not.

8. Justice Major statements on clinical research ethics require that research subjects be selected fairly. On the one hand, this is generally interpreted as meaning that subject selection should be based on scientific rationales rather than convenience, easy availability, or dependence. On the other hand, disadvantaged populations should not be excluded from access to potentially beneficial research. Recent reinterpretations of the principle of justice have furthermore urged greater inclusiveness of women, minorities, and children in drug studies to assure that medical knowledge has relevance for these historically disadvantaged or underrepresented populations (85). Concerns about justice in GT research—as in other research realms—have generally been overshadowed by discussions about risk, consent, and conflict of interest. However, they have long

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been present in cancer research. In 1963, Chester Southam (who conducted some of the earliest experiments in viral oncolytic therapy (86)) performed a series of experiments in which he injected a suspension of cancer cells into indigent, aged patients at the Jewish Chronic Disease Hospital in Brooklyn to study the role of the immune system in cancer (87). This controversial experiment, along with others involving powerless human subjects, motivated a recognition of justice as an important consideration for human protections (88). Another episode highlighting justice concerns, more squarely within cancer GT, occurred in 1993, when a patient with glioblastoma used political connections (89) to secure “compassionate use” access to an experimental gene transfer intervention (90, 91). Presently, the major justice concern in cancer GT is what seems to be an incipient trend toward outsourcing translational clinical trials to low- and middle-income countries like China. Several factors appear to be behind this phenomenon. First, trials in low- and middle-income countries are often less costly than in the USA and Europe, where GT is bound by an elaborate regulations and production specifications (92). Second, volunteers are easier to recruit, in part because they lack access to standard care (93). In general, major research ethics codes like the Declaration of Helsinki (3) and Council for International Organizations of Medical Sciences (CIOMS) (94), as well as statements of the National Bioethics Advisory Commission (95) and the Nuffield Council (96), articulate two criteria for ensuring that trials conducted in resource-poor settings are ethical. The first, providing volunteers posttrial access to an intervention, has limited applicability to phase 1 trials, because these are not aimed at vindicating the efficacy of a study drug. The second is “responsiveness.” For example, the CIOMS states in its section on general ethical principles that “research project[s conducted in] low-resource countries or communities… should be responsive to their health needs and priorities in that any product developed is made reasonably available to them, and as far as possible leave the population in a better position to obtain effective health care and protect its own health” (97). The Belmont Report, although it did not address international research specifically, stated that “…it seems unfair that populations dependent on public health care constitute a pool of preferred research subjects if more advantaged populations are likely to be the recipients of the benefits”(1). One reasonable interpretation of responsiveness, then, is that an intervention should not be tested in a population unless there is a reasonable likelihood that it will expand the capacity of a community to meet its urgent healthcare needs (98). Experience with Gendicine, a GT intervention licensed for the treatment in head and neck cancer in China, points to some of the challenges for fulfilling responsiveness should researchers


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from high-income countries pursue clinical testing in low- and middle-income countries. A typical course of treatment with Gendicine reportedly costs over US$3,000—a figure that exceeds per capita yearly income in Beijing (99), one of the country’s wealthiest cities. The treatment is also not presently covered by health insurance. As a result, many patients receiving Gendicine are wealthy foreigners (100, 101). In general, trials pursued in resource-poor settings with the objective of obtaining regulatory relief, or expediting patient accrual, will have a more difficult time making the case that the study embodies sincere objectives of responding to the health deprivations of the host community.

9. Managing Public Expectations

With rare exception, research ethics scholarship and policy has centered on the obligations of the clinical investigators toward study volunteers. This focus is hardly surprising, given the abuses that have occurred in human experimentation. Nevertheless, basic and clinical investigators have other ethical responsibilities as well. One important set concerns how GT researchers interact with members of the public, the media, and patient advocacy organizations. Mishandled communications matter ethically for several reasons. First, there is reason to think that they may influence the consent process. For instance, some studies suggest that many volunteers decide on enrollment before consent discussions have even occurred (102). If true, enrollment decisions are likely to be shaped by expectations that volunteers bring to these studies. Through their interaction with various publics, researchers play an important role in shaping this environment of expectation. Second, expectations have consequences in determining whether and to what extent scientific research programs are funded and supported. One episode that starkly illustrates some of the problems encountered in bringing translational research to the public occurred in 1998, when the Sunday New York Times ran a front page story on Judah Folkman’s discovery of a class of cancer drugs called angiogenesis inhibitors (103). The report contained several choice quotes from Nobelist James Watson and then National Cancer Institute director Richard Klausner underscoring the immense promise of this new strategy. The next morning, shares of Entremed, a biotechnology firm founded by Folkman, increased sixfold (104). When Harvard initiated trials, more than 1,400 cancer patients sought entry for three slots; one

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wealthy study candidate even offered to buy a controlling stake in Entremed (105). Notwithstanding a general inattentiveness to such concerns in standard ethics commentary, there are ethics codes that address these issues. For example, the American College of Physicians states that researchers should “use… precise and measured language” when communicating with the public, and “avoid raising false public expectations or providing misleading expectations” (106). The Medical Research Council of India also states “Researchers have a responsibility to make sure that the public is accurately informed about results without raising false hopes or expectations” (107). And the Association of Medical Research Charities states that “it is important that researchers consider the impact any publication of research findings may have on patients with the condition, those involved in their care, those involved in the research and consumer groups” (108). There is insufficient space here for a full analysis of management expectation. Nevertheless, I offer the following recommendations, and direct readers elsewhere for a lengthier analysis (109). First, public expectation management activities are an ethically appropriate practice, because they enable a better public understanding of, on the one hand, the medical potential of a research undertaking, and, on the other hand, the glacial pace of translational research. Second, researchers should carefully monitor public expectations, lest these outrun the ability of researchers to deliver. Third, researchers—and research leaders—should be as energetic in countering exuberant expectation as they might be in trying to neutralize adverse expectations brought on by major mishaps or disappointments. Fourth, researchers should consider vetting their press releases before ethics committees or patient advocates. Last, researchers should be extremely cautious about using individual patients to promote their research, for reasons that are entirely independent of protecting the privacy of patients. However many earnest disclaimers a researcher might offer about the precariousness of their research findings, the visual impact of patients is powerful, and often conveys two potentially misleading messages: that cures are imminent, and that early phase trials of novel agents are a viable form of therapy.

10. Conclusions: Different, but the Same? In summary, early phase cancer GT trials present ethical challenges that are somewhat distinct from those of testing conventional cancer drugs. These challenges derive from the high degree


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of scientific uncertainty surrounding such studies, the complexity of GT interventions, and the often basic character of clinical investigations. Obviously, where GT studies involve simple or well-characterized agents, their ethical properties become more akin to those for conventional cancer drugs. On the other hand, where non-GT interventions—such as targeted interventions or immunotherapies—involve high levels of novelty and complexity, ethical challenges in testing these become very similar to those for GT. In this way, working out the ethics of gene transfer is an occasion to explore issues that will be confronted with increasing frequency as cancer translational clinical research extends its frontiers.

Acknowledgments This work was funded by the Canadian Institutes of Health Research.

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Chapter 24 Virus Production for Clinical Gene Therapy Tiago Vicente, Cristina Peixoto, Manuel J.T. Carrondo, and Paula M. Alves Summary Gene therapy is becoming increasingly relevant for the treatment of prominent human diseases. Viral vectors are currently used in more than 50% of the gene therapy clinical trials, most of them aimed at cancer diseases. Clearly, the increasing needs of high-quality viral preparations require the elimination of process bottlenecks, streamlining the development of a viral vector into a real-world clinical tool. Virus production for clinical gene therapy can be a limiting step because many virus generation protocols rely on labor-intensive, bench-scale methods; robust, cost-effective strategies for the delivery of clinical-grade viruses are thus essential for the future of gene therapy. A comprehensive picture of key aspects on the integration of upstream and downstream processing is addressed in this chapter, by describing the case study of recombinant budded baculoviruses for gene therapy; scalable methods are described in detail as well as mandatory characterization techniques for a proper and complete quality assessment of the viral vectors. Key words: Analytical tools, characterization, recombinant baculoviruses, upstream and downstream processing, viral vectors.

1. Introduction Viral vector manufacturing for preclinical and clinical gene therapy requires both understanding of virus biology and expertise on animal cell technology/bioprocessing to conveniently generate batches without impairing the biological activity of such complex particles. Indeed, in vitro cultures of animal cells (mammalian or not) currently constitute the best alternative for gene therapy viral vector production (1); the proper use of the cell machinery allows the generation/replication of the viral protein components, assembly, and, in most of the cases, delivery of the virus to the extracellular medium. Subsequent separation steps Wolfgang Walther and Ulrike S. Stein (eds.), Methods in Molecular Biology, Gene Therapy of Cancer, vol. 542 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-561-9_24

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are normally facilitated in contrast to the use of living organisms or eggs as producer systems, where a large amount of intrinsic biological impurities and adventitious agents raise safety concerns (e.g., Influenza A virus vaccine production (2)). In these cases, more expensive and rather poorly reproducible separation processes are required to ensure product quality, typically resulting in low yields. However, optimization of recombinant virus production using cell lines can be a difficult task because viral replication is typically destructive for the host cell. Bioengineering of cell cultures for virus production leads to processes that are more cost-effective and that can be further implemented in the pilot/ process scale thus meeting the demands for clinical trials. Currently, clinical-grade viral preparations are obtained based on a classic bioprocess basis: an upstream phase, wherein the bioreaction takes place; and a downstream phase, wherein the generated bulk containing the viruses (harvested at its optimum1) undergoes a sequence of concentration/purification/polishing steps allowing the separation of the desired product from cell debris, host cell protein contaminants, DNA, and/or damaged/ empty viral vectors (e.g., transgene deficient vectors) (3). Analytical measurements throughout the bioprocess are critical not only for a proper documentation (in agreement with current good manufacturing process [cGMP] guidelines) but, most importantly, for identification of critical issues needing further investigation and development. Moreover, viral vector preparations for clinical trials must meet the quality requirements with respect to the allowed contaminant limits (4). Therefore, special attention should be paid to the analytical tools used to monitor the bioreaction operational conditions (pH, dissolved oxygen, dissolved carbon dioxide) and to performing viral vector characterization; these can be either biological assays (viral titration methods) or molecular/cell biology techniques such as sodium dodecyl sulfate (SDS) polyacrylamide gel electrophoresis (PAGE), total protein content determination, immunoblotting for specific viral protein(s), DNA content determination, and viral genomes (vg) number assessment by quantitative polymerase chain reaction (q-PCR) (5). This chapter focuses on state-of-the-art and day-to-day methodologies for the production of gene therapy viral vectors. Among the most widely used in clinical trials are retroviruses, adenoviruses, and adeno-associated viruses (AAVs) (6). Methods for retrovirus production for clinical gene therapy have been reviewed recently (5, 7, 8). These vectors, derived from wild-type, singlestranded RNA (ssRNA) viruses belonging to the Retroviridae 1

“Viral quality” (total viruses/infective viruses ratio) or “viral titer” can be used to define the harvest optimum, depending on the process specificities and the strategy of optimization available.

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family, have been used very efficiently as gene transfer vehicles (9); they have the intrinsic capacity of delivering a specific transgene into a variety of cell hosts and, most importantly, by using their own and host cell machinery, effectively integrate a corrective gene in the genome thereby allowing long-term gene expression. Within the retrovirus taxonomic group, lentiviruses (e.g., human immunodeficiency virus [HIV]-1]) are also being paid special attention for their ability to infect both dividing and nondividing cells; however, security and ethical concerns arise with their use because of eventual appearance of replication-competent lentiviruses (RCLs) and random insertional mutagenesis (10). Apart from its paramount potentialities, this biological system still faces several technological challenges, e.g., low-titer supernatants (up to 107 infective particles [IP]/mL when a dose may require as much as 1010 IP/patient (11)) produced using engineered packaging cell lines2, and inherent retroviral vector instability due to physical envelope disruption, RNA polymerase and/or reverse transcriptase activity decay, and possibly other mechanisms (5, 12). Inevitably, these drawbacks pose new challenges to the production and purification tasks. As such, it is not surprising that, over the last decades, much effort has been directed into retrovirus bioprocess development seeking technological solutions for both vector stability improvement and for refined purification steps preserving the product biological quality (5). Adenoviruses are technologically interesting vectors because they replicate in cell culture to very high titers (up to >1011 infective viruses/mL) and can be readily produced in large-scale bioreactors in suspension conditions (1, 13). Unlike retroviruses, adenoviruses are double-stranded DNA (dsDNA), non-enveloped viruses, with spike protrusions covering the virion capsid, thus serving as a protective shield; they efficiently infect a variety of hosts and are typically safer than integrative viruses, allowing more controlled, time-spanned therapeutics. However, as an important disadvantage, these viruses elicit strong immune responses during infection, which can be detrimental in specific clinical applications (14). An extensive knowledge is now available in the scientific literature concerning the understanding, use, and development of adenoviral vectors for clinical gene therapy (15, 16); different modifications and molecular-based enhancements pursuing ideal vectors have been proposed as well as technological improvements in terms of scalable bioprocesses aiming at optimized productions of clinical-grade adenoviral preparations (14, 17, 18). AAVs hold great promise as permanent-expression viral vectors and are well covered in the literature (19, 20). AAVs are

2

Cells previously transfected thereby becoming capable of permanently generating a designed viral vector.

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non-enveloped, ssDNA viruses that belong to the Dependovirus genus of the Parvoviridae family. These viral vectors have the capability of integrating into a specific site of the host cell genome, opening wide perspectives for permanent and safe correction of a specific defective gene (20). However, these viruses need the presence of a helper virus for their replication and expression. As such, a variety of producer systems have been developed: multiple plasmid transfection systems using a suitable cell line; mammalian packaging cell lines expressing some of the AAV components plus a helper virus (e.g., adenovirus Ad5); or baculovirus expression vector systems (BEVS) by means of a coinfection with recombinant baculoviruses expressing the different components of the AAVs (20). Upstream scalability is presently feasible, because producer systems involving suspension cultures in bioreactors fed with protein-free media (for instance, the BEVS system) are easy to implement. Nevertheless, whenever a helper virus or recombinant baculovirus in the BEVS is needed, a mixture of different viral entities (i.e., AAV, Ad5, baculovirus) is obtained during the production process. This requires highly efficient, scalable purification methods, and the development of intricate characterization assays ensuring the absence of the process auxiliary viruses, in agreement with the regulatory agencies strict guidelines (20). Recombinant Autographa californica multicapsid nucleopolyhedrosis viruses (AcMNPV), classically referred to as recombinant baculoviruses, are dsDNA, enveloped viruses derived from the Baculoviridae family. Beyond the handiness of the baculovirus expression vector system for the production of recombinant proteins (21, 22), these viruses have more recently been recognized as potential clinical gene therapy vectors given their advantages over other viruses (23–25). Indeed, scientific interest has been redirected toward the development of the molecular biology of these viruses improving their effectiveness as gene therapy tools (25–27). Major advantages of this system include: remarkable handling ease of insect cell suspension cultures (for the virus infection/replication), bioreaction upscale feasibility, serumfree media cell culture, capability of accommodating very large transgene DNA (>38 kbp), and very low cytotoxicity (24, 25). However, not enough knowledge is available and there are not enough downstream processing (DSP) and characterization methods mandatory for clinical-grade baculovirus preparations, although occasional reports on the use of chromatography have been released (28–30). An overview of current processing issues and challenges inherent to different viral vectors used for clinical gene therapy is summarized in Table 1. As briefly reviewed here, there are specific issues and challenges inherent to the selected viral vector system for a given clinical application. Upstream and downstream processes for the


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Table 1 Viral vectors envisaged for application in clinical gene therapy, inherent advantages, issues, and production process challenges Major drawbacks/technological challenges

Viral vector

Key advantages

Retroviruses (enveloped, ssRNA)

- Very efficient transgene integration - Infect dividing (and nondividinga) cells - Low immunogenicity - Long-term gene expression

- Inherent instability - Low titer supernatants - Difficult production in stirred systems - Short shelf-life: very low virus halflives at 37°C (8 h) - Virus reverse transcriptase enzyme activity decay

(5, 8, 9, 12)

Adenoviruses (nonenveloped, dsDNA)

- Very efficient transient expression - Large-scale production and purification available

- Empty capsid vectors, increasing downstream processing hurdles and cost - Immunogenic and potentially toxic leading to strong immune responses; safety concerns - Aggregation at high virus titers

(14, 35)

AAVs (nonenveloped, ssDNA)

- Very efficient transgene integration into specific genome locus - Replication-defective, thus safer vectors - Low immunogenicity

- Empty capsid viruses, increasing downstream processing hurdles and cost (36) - Low packaging capacity (up to 4.5 kbp) - Mixture of different viruses on the produced preparations - Complex virus production strategies using different molecular contributors

(19, 20)

Baculoviruses (enveloped, dsDNA)

- Large packaging capacity (>38 kbp) - Large scale bioreaction possible - Typically high stability - Lack of cytotoxicity and replication

- No clinical trial experience - Inefficient transfection efficiency on some target cells/tissues - Large, rod-shaped virus complicating downstream processing - Aggregation at high virus titers (>1010 pfu/mL), storage compromised

(25)

a

Reviewed in

Lentiviral vectors

delivery of clinical-grade viral vector preparations use somewhat similar strategies. The current paradigm is based on tightly controlled and modeled upstream processes using stirred vessels or disposable bioreactors alongside robust downstream purification technologies. Chromatographic and membrane processes are regarded as a strategy of choice as new, state-of-the-art materials

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and matrices are being used and optimized for the DSP of viruses. The removal of the traditional ultracentrifugation-based purification methods for a wide variety of viruses is becoming a must-do shift toward the establishment of a robust and effective strategy. In the context of this chapter, methodologies and procedures will be presented for the production, harvesting, concentration, purification, and characterization of non-occluded, cell-budded recombinant baculoviruses as an example of (emerging) vectors for application in a clinical setting. The versatility and horizontality of most of the methodologies presented here will hopefully provide key instructions/hints/guidance applying to other major viral vector systems, where such processes are more advanced than for recombinant baculoviruses. Whenever pertinent, references to special methodologies used for other vectors will be included. Additionally, several notes are presented at the end of the chapter emphasizing critical and specific issues.

2. Materials 2.1. Cell Culture and Viral Propagation

1. Spodoptera frugiperda Sf9 cell line purchased from collections commercially available (e.g., ATCC, ECACC, DSMZ). 2. GibcoTM Sf-900 II SFM medium (Invitrogen, Paisley, UK), sterile liquid formulation available. 3. 500-mL Erlenmeyer shake flasks (Schott, Mainz, Germany), autoclaved. 4. Stirred tank bioreactor (5-L working volume) (Sartorius Stedim Biotech, Göttingen, Germany). 5. 100-, 250-, 500-, 2,000-, and 5,000-mL flasks (Schott), with (at least) two glass connections for tubing, 0.2-μm vent filters connected to the Schott flasks, all autoclaved. 6. 1.5-mL Eppendorf sterile tubes (Nunc, Roskilde, Denmark). 7. 15-mL, 50-mL polypropylene tubes (Nunc).

2.2. Buffers

1. Phosphate-buffered saline (PBS), sterile: 154 mM NaCl (Merck, Darmstadt, Germany), 1.34 mM Na2HPO4 (Merck), 1.54 mM KH2PO4 (Merck) in water, adjust pH to 7.2 (see Note 1). 2. GibcoTM Dulbecco’s-PBS (with Ca and Mg) (D-PBS) (Invitrogen), sterile solution available (Buffer 1). 3. D-PBS containing 1,500 mM NaCl, sterile solution available (Buffer 2).


2.3. Downstream Processing 2.3.1. Harvest/Clarification

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1. 3-μm retention Sartopure PP2 filter capsule(s) (Sartorius Stedim Biotech) (see Note 2), 0.25-inch inside diameter Master Flex PharmedTM tubing (Cole-Parmer, Vernon Hills, IL, USA), stainless steel connections and sealings, two or more 2-L and one 5-L Schott flasks, all autoclaved. 2. Optionally, 0.65-μm retention Sartopure PP2 can be included in series for improved cell debris removal.

2.3.2. Ultrafiltration/ Diafiltration

1. 500-, 2,000-, and 5,000-mL flasks (Schott), with (at least) two glass connections for tubing, 0.2-μm vent filters connected to the Schott flasks, all autoclaved. 2. Sartoflow Slice 200 Benchtop apparatus including reservoir, magnetic stirrer, scale, 0.19-inch and 0.12-inch inside diameter MasterFlex PharmedTM tubing (Cole-Parmer) and connections/software for spreadsheet data collection (all Sartorius Stedim Biotech) (see Fig. 1); tubings sanitized with a 0.1 M NaOH solution; reservoir and pressure sensors should only be sanitized with 20% (v/v) ethanol. 3. Sartocon Slice 200 ultrafiltration cassettes, HydrosartTM membranes, 30-kDa nominal molecular weight cutoff (MWCO) and/or 0.45-μm pore size (all Sartorius Stedim Biotech), sanitized with a 0.1 M NaOH solution (mounted on the holder).

2.3.3. Chromatography

1. 500-, 2,000-, and 5,000-mL flasks (Schott). 2. Matrix 1: SartobindTM D 75 MA membrane adsorber units (Sartorius Stedim Biotech). 3. Matrix 2: HiPrepTM 26/10 Desalting column (GE Healthcare, Uppsala, Sweden).

Fig. 1. Ultradiafiltration system design. P1, P2, and P3 refer to inlet pressure sensor, retentate pressure sensor, and permeate pressure sensor, respectively; the pressure sensors, scale, and peristaltic pump are connected to the Sartoflow Slice 200 Benchtop controlling unit; online data are transmitted to a MicrosoftTM Excel spreadsheet.

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4. 0.5 M NaOH solution. 5. 20% (v/v) absolute ethanol in water. 2.4. Recombinant Baculovirus Quantitation

1. 96-well flat-bottomed sterile plates (Nunc). 2. 3.5-mL polystyrene tubes (Nunc).

2.4.1. Infective Particles (Endpoint Dilution Method Using Green Fluorescent Protein [GFP] as Transgene) 2.4.2. Quantitation of Genome Containing Baculoviruses (q-PCR)

1. 1.5-mL Eppendorf sterile tubes. 2. High Pure Viral Nucleic Acid kit (Roche, Mannheim, Germany). 3. DNAse I (Sigma, Steinheim, Germany). 4. Nuclease-free water (Promega, Madison, WI, USA). 5. LightCycler-DNA Master SYBR Green I (Roche). 6. 20-μL LightCycler capillaries (Roche). 7. Forward primer: CCC GTA ACG GAC CTC GTA CTT, Tm= 59.8°C, 20 μM (TIB MOLBIOL, Berlin, Germany). 8. Reverse primer: TTA TCG AGA TTT ATT TGC ATA CAA CAA G, Tm= 56.4°C, 20 μM (TIB MOLBIOL).

2.5. Recombinant Baculovirus Characterization 2.5.1. Total Protein (Bicinchoninic Acid Method) 2.5.2. Protein Profile (SDS-PAGE) and Immunoblotting (for Recombinant Baculovirus Envelope gp64 Glycoprotein)

1. 96-well plates (Nunc). 2. Bicinchoninic acid (BCA) kit (Pierce, Rockford, IL, USA). 3. D-PBS. 1. 1.5-mL Eppendorf sterile tubes. 2. NuPAGE Novex 4–12% Bis-Tris precast polyacrylamide gel, 1.0 × 10 well (Invitrogen). 3. SimplyBlueTM SafeStain (Invitrogen). 4. SilverXpressTM Silver Staining kit (Invitrogen). 5. BenchMarkTM Pre-Stained Protein Ladder (Invitrogen). 6. Running buffer: NuPAGE MES buffer (20×) (Invitrogen) diluted to 1× in water. 7. Transfer buffer: NuPAGE transfer buffer (20×) (Invitrogen) diluted to 1× in water and supplemented with 20% (v/v) methanol (Merck). 8. Nitrocellulose HybondTM-C Extra (GE Healthcare) and 3MM chromatography-grade filter paper (Whatman, Maidstone, UK).


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9. Washing buffer (TTBS): Tris-buffered saline (TBS) (Sigma) supplemented with 0.05% (v/v) Tween 20 (Roche) solution in water. 10. Antibody dilution buffer: TTBS supplemented with 1% (w/v) bovine serum albumin (Roche). 11. Blocking buffer: 5% (w/v) skim milk (Roche) dissolved in TTBS. 12. Primary antibody: mouse monoclonal antibody (mAb) against gp64 baculovirus glycoprotein, AcV5 clone, affinity purified (eBioscience, San Diego, CA, USA). 13. Secondary antibody: Goat anti-mouse antibody, γ-chain specific, immunopure (Sigma). 14. 1-stepTM NBT/BCIP solution (Pierce).

3. Methods 3.1. Cell Culture and Viral Propagation

Recombinant baculovirus replication is routinely performed using Sf9 insect cells infected with a previously propagated, titrated baculovirus stock. Upon viral replication, de novo baculovirus capsids are produced in the cell nucleus; these are further released by budding out from the host cell wherein the double lipid layer envelope is incorporated (21, 22, 31). However, after two or more rounds of replication (the previously budded baculoviruses may infect the same cell), the host insect cells cease growth and suffer lysis due to the infection burden (21–23). A good indicator of the baculovirus infection and its replication during the bioreaction is therefore the determination of cell viability. The multiplicity of infection (MOI) (number of infective virus added per cell in culture), the time of infection (TOI) for the cell culture, the cell concentration at infection (CCI), and the time of harvest (TOH) needs to be defined by the operator. These parameters strongly impact the infection and virus propagation process in this biological system and a significant amount of literature related to this subject is available (32, 33). The materials and methods presented herein allow for the production of a 5-L recombinant, enhanced green fluorescent protein (eGFP)-encoding baculovirus in a stirred batch bioreactor. In other virus production systems, it is worthwhile remarking that packaging cell lines (e.g., for retroviral vectors production (5)) consequently avoid the infection process; the upstream optimization in these cases is more focused in the implementation of suspension cultures improving virus titer and stability.

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3.1.1. Recombinant Baculovirus Stock Propagation

1. Sf9 insect cells are grown in serum-free medium up to a CCI of 1–1.5 × 106 cells/mL (see Note 3) at 27°C and 150 rpm (~3–4 days) using a 125-mL (working volume) spinner vessel (Wheaton Instruments, Millville, NJ, USA). The cell inoculum used must be withdrawn from a mid- to late-exponential routine culture (i.e., 2 to >4 × 106 cells/mL). 2. Infect the culture with a secondary recombinant baculovirus stock (see Note 4) with an MOI of 0.1 pfu/cell. Low MOIs (especially lower than 1 pfu/cell) are preferable to prevent the production of “defective” baculovirus generations (lacking the transgene). This genomic instability is promoted when using heavier viral loads that will increase the number of rounds of replication (33). 3. Monitor the infected spinner culture daily, assessing cell viability (see Note 5). 4. (Optional) For a more accurate quantification of the infected cells during the process, run a Fluorescent Activated Cell Sorting (FACS) analysis for GFP-positive cell counting: transfer ~1 mL of cell suspension (diluted 1:2 if the cell concentration is >>106 cells/mL) to 3.5-mL polystyrene tubes (Nunc); count up to 20,000 events (4 × 106 cells/mL to inoculate 4 × 125-mL spinner flasks (all working volumes); let these cultures grow up to >4 × 106 cells/mL and inoculate a 2-L working volume culture in the 5-L vessel bioreactor (Sartorius Stedim Biotech). 2. Alongside with the previous step, prepare the bioreactor by following these procedures in the specified order: (i) pH electrode calibration; (ii) assembly of all the necessary tubing and flasks needed, glass male (for outflow) and female (for inflow) connections (two flasks for medium, one for cell inoculum, one for viral inoculum, one for eventual NaOH addition, two spare inlets for safety reasons, an outlet 100-mL Schott flask for sterile sampling, an outlet nonsterile tubing and glass


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tube); (iii) sterilization of all the material by autoclaving at 121°C for 1 h; (iv) pressure testing of the vessel (for sterility assurance reasons); (v) culture medium transfer into the bioreactor (in sterile conditions [see Note 6]); (vi) dissolved oxygen (pO2) electrode calibration, 0% (saturating with N2), and 100% (saturating with normal air inlet), at the culture temperature (27°C), stirring frequency and aeration rate (70 rpm, 20 mL/min (1 atm, 20°C, normal temperature and pressure [NTP]). 3. Inoculate the bioreactor by seeding 0.5 × 106 cells/mL in a total 2 L starting working volume. 4. Set the digital controller unit set point to maintain the pO2 at 30% using a cascade action control strategy with the following order: (i) agitation from 70–200 rpm; (ii) aeration from 20–50 mL/min (NTP); (iii) gas mixture with pure O2 enrichment from 0–100% (v/v). 5. Withdraw bulk samples throughout cell growth for cell concentration and viability assessment. 6. At a cell concentration of around 2 × 106 cells/mL, add the remaining medium to fulfill ~5 L working volume. Let cell growth proceed until infection with the working viral seed stock. 7. Use a CCI of 1–1.5 × 106cells/mL and an MOI of 0.1 pfu/ cell by adding the corresponding volume of the working viral seed stock previously titrated (see Note 7). 8. Allow the viral infection/replication to occur until the cell viability decreases down to ~50%; monitor the bioreaction withdrawing samples and perform FACS analysis to observe the infection progression; the bioreaction may last up to 4–7 days after infection. 3.2. Downstream Processing 3.2.1. Harvest/Clarification

The bulk clarification can be performed classically by centrifugation at 1,500g for 20 min at 4°C, discarding the pellets and collecting the supernatants containing the recombinant baculoviruses; alternatively, an integrated process designed to warrant easiness of operation and use a sterile, disposable process using depth-filters is desirable. A scalable protocol is thus presented herein: 1. Set-up the depth-filter apparatus (see Fig. 2) and test it for pressure maintenance using the bioreactor aeration system (sterility assurance); rinse the whole tubing and filter with water, and sterilize by autoclaving. The apparatus is connected by setting its female glass inlet connection to a glass male bioreactor outlet connection (in sterile conditions). 2. Using a peristaltic pump (Watson-Marlow, Wilmington, MA, USA), let the bulk flow through the depth-filter unit(s) to

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Storing flask

Peristaltic pump

Storing flask

Depth-filter

Bypass stream

Fig. 2. Depth-filter system design integrated with the bioreactor outlet stream.

control overpressure conditions; if such happens, immediately use the last resource alternative, i.e., flow the remaining bulk through the bypass tubing and proceed to the clarification of this amount later on; if needed during this step, mildly pressurize the bioreactor vessel to facilitate the process. 3. Collect the flasks with the clarified viral preparation and move them into a laminar flow chamber to aliquot the material into freshly autoclaved Schott flasks in the desirable volumes. 4. Clarified viral preparation aliquots must be stored at 4°C in the dark until further processing. 3.2.2. Ultrafiltration/ Diafiltration

Intermediate purification and concentration is done in an ultrafiltration step, where the recombinant baculoviruses are retained in the retentate. The bulk medium is exchanged to the selected working buffer, D-PBS, performing a diafiltration as soon as a chosen concentration factor (CF) is achieved in the retentate during ultrafiltration. This step yields an outlet stream suitable to be processed in an ion-exchange chromatographic step with enhanced yields. 1. Install the ultrafiltration cassette in the holder and perform a washing step with water. 2. Test the cleanliness of the cassette by performing a calibration curve (permeate flux vs transmembrane pressure [TMP]) with water: set the controlling unit to fix the operator-chosen TMP (see Eq. 1) by automatically changing the recirculation/retentate crossflow rate (within a range of 50–500 mL/min); fix a number of TMPs spanning from 0 to 40 psig; data should be autonomously transmitted to the Excel spreadsheet (see Note 8). TMP =

p feed + pretentate 2

− p permeate

(1)

3. Rinse the whole system with the D-PBS (to be used subsequently for diafiltration).


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4. Transfer a selected amount of clarified bulk (e.g., 500 mL) into the apparatus reservoir and start the procedure defining a mild TMP—e.g., 12.5 psig—setting a low retentate pressure (pretentate), up to 5 psig (controlling the tightening of the tubing pressure valve), and a null permeate gauge pressure (pperme) (atmospheric pressure) (see Note 9). ate 5. When the CF approaches 5, note down the remaining reservoir volume (from now on defined as diafiltration volume [DV]) and feed four DVs into the reservoir (letting the retentate reach the DV between each buffer volume addition). 6. Recover the ultradiafiltered retentate into a sterile container and store at 4°C in the dark until subsequent processing; collect a sample for titration of infective viruses (see Subheading 2.4.1). 3.2.3. Chromatography Preparation of the Liquid Chromatography System

1. Connect one Matrix 1 unit (see Subheading 2) or more in series (alternatively, use one or more Sartobind D 100 MA) to an ÄKTAexplorer 100 system coupled with the UNICORNTM software (all GE Healthcare) or other “fast protein liquid chromatography (FPLC)” system with a UV absorbance cell detector, conductivity meter, pH sensor, and a fraction collector; ensure that the inlet streams are purged and no air is imprisoned in tubing or valves; the whole system must be washed before any operation first with 20% (v/v) ethanol and next with Buffer 1 to be used as running/equilibration buffer. 2. Equilibrate the Matrix 1 unit(s) with Buffer 1 with 10 membrane volumes (MVs) at a flow rate of 20 mL/min; the system back pressure should not exceed the Matrix 1 default pressure limit, 6 bar. 3. Equilibrate the Matrix 2 with Buffer 1 with 10 column volumes (CVs) at a flow rate of 5 mL/min; the system back pressure should not exceed the Matrix 2 default pressure limit, 1.5 bar. 4. Sanitize both matrices with 0.5 M NaOH for 10 MVs or 10 CVs at the corresponding flow rates mentioned above in steps 2 and 3. 5. Sanitize the whole system with 0.5 M NaOH including all valves and inlet and outlet streams at a high flow rate (e.g., 15 mL/min). 6. Equilibrate the system with Buffer 1 at a high flow rate (e.g., 25 mL/min) until the conductivity and pH of the outlet stream reaches the Buffer 1 values (pH 7.2 and ~16.7 mS/cm). 7. Re-equilibrate both Matrices 1 and 2 as described in steps 2 and 3 until the conductivity and pH of the outlet stream reaches the Buffer 1 values (pH 7.2 and ~16.7 mS/cm).

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Anion-Exchange Membrane Chromatography

1. With the system and Matrix 1 equilibrated with Buffer 1, reset the UV detector. 2. Load the ultradiafiltered viral preparation volume onto Matrix 1 at a flow rate of 10 mL/min (working flow rate). 3. Wash the Matrix 1 with five MVs of Buffer 1. 4. Start an isocratic elution of the bound material with Buffer 1. 5. Start a second isocratic elution of the remaining bound material (recombinant baculoviruses) with 60% (v/v) of Buffer 2 in Buffer 1 using the mixing inlet feature of the ÄKTA system (if not available in the FPLC system used, switch the inlet buffer to the mentioned Buffer 1 and 2 mixture). An example of an elution chromatogram profile is shown in Fig. 3.

Desalting of Purified Recombinant Baculoviruses

1. With the system and Matrix 2 previously equilibrated with Buffer 1, reset the UV detector. 2. By means of a sample loop (previously sanitized and washed with Buffer 1), inject the high-salt content recovered recombinant baculovirus peak fraction coming from step 3.2.3.2 onto Matrix 2 at 5 mL/min (working flow rate). 3. Flow Buffer 1 through the Matrix 2 to recover the viral peak fraction. This peak, more or less broad depending on the

Flowthrough pool

Elution peak

90 80

1200

70

1000

60 800 50 600 40 400

30

200

Conductivity (mS/cm)

280 nm Absorbance (mAU)

1400

20

0 20

30

40

50

60

70

80

90

100

110

10 120

Volume (mL) 280 nm mAU

Conductivity

Fig. 3. Elution profile of an anion-exchange chromatography run designed for capture/purification of recombinant baculovirus; the flow-through pool mostly contains process impurities; the “elution peak” corresponds to the product peak, concomitantly with the increase in salt concentration detected in the conductivity meter; process description: Sartobind D 75 loaded with ~35 mL of concentrated, ultradiafiltered (to D-PBS) recombinant baculovirus stock (starting upstream bulk volume: 500 mL); elution performed with a step dilution of 60% of Buffer 2 in Buffer 1 (as described in Subheading 2). The peak tailing observed is the consequence of the chromatographic system dispersion in combination with a heavy load of material desorbing from the matrix upon elution.


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amount loaded, should elute before the salt peak; as such, the injection volume should not exceed 10% of the CV. Although some loss is to be expected (up to >50%) (baculoviruses are rod-shaped with 50–90 × 100–450 nm, filter the viral preparation using 0.22-μm sterile filters to assure sterility (31). 4. Store the purified recombinant baculoviruses at 4°C in the dark. In a typical situation, these conditions allow long shelflives with a good biological activity preservation; however, in some cases (as yet not clearly elucidated why), freezing of the recombinant baculovirus stocks down to − 80°C is preferable, because vector activity decay through time at 4°C is more significant. 3.3. Recombinant Baculovirus Quantitation 3.3.1. Infective Particles (Endpoint Dilution Method Using GFP as Transgene)

1. Sf9 cell density must be between 1–1.8 × 106 cells/mL; seed 100 µL of cell suspension in 96-well plates (duplicate plates) to yield a concentration of 5 × 104 cells/well; allow the cells to settle and adhere to the bottom of the plate for approximately 1 h at 27°C see Note 10. 2. Prepare, in 3.5-mL polystyrene tubes, serial dilutions of tempered (30 min at 27°C) viral samples. 3. Virus dilution range: from 10—1 to 10—12 (300 µL of the virus stock + 2,700 µL of medium for the 10—1 dilution, 30 µL from the previous virus solution + 2,770 µL of medium for 10—3 dilution, 30 µL from the previous virus solution + 2,770 µL of medium for 10—5 dilution, 300 µL of the previous solution + 2,700 µL of medium for a 10—6 dilution, and repeat this last step until a dilution of 10—11 is reached) (10—1 can be considered as a positive control [P], if analyzing a nonconcentrated, supernatant-derived virus sample that has a titer of approximately 108 pfu/mL). 4. Remove the supernatant after cell attachment to the plate and replace it with 100 µL of each virus dilution for each well. Typically, dilutions 10—4 (1st row) (n = 10), 10—5 (2nd row) (n = 10), …, 10—11 (8th row) (n = 10); use a column of negative control wells (11th column only with cells) and a column of positive control wells (12th column), considering also a final volume of 100 µL see Note 10. 5. Incubate the plate at 27°C for 7 days inside a closed plastic container with dampened paper towels in the bottom to prevent dehydration. 6. Analyze the plates under a regular fluorescence microscope: detection of the green signal (even if few cells are expressing GFP) is counted as a positive well (+). 7. Count the total number of positive and negative wells for each virus dilution; the 50% tissue culture infection dose (TCID)50 is determined as corresponding to the dilution that gave rise to 50% positive wells (+) and 50% negative wells, as interpolated

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from the experimental data obtained. In an accurate titration assay, the dilutions used should range from 100% positive to 100% negative wells (see Note 11). 8. Do the following calculations (22): ⎡(+% on dilution next to and above 50% + ) − 50% + ⎤⎦ PD = ⎣ ⎡(+% on dilution next to and above 50% + ) − ⎤ ⎢ ⎥ ⎣⎢(+% on dilution next to and below 50% + ) ⎦⎥ log10 TCID50 = log10 (dilution next to and above 50% + )+ (− PD ) TCID50 = 10 log10 TCID50 Titer (TCID50 / mL )=

1 TCID50 value × virus volume added per well (mL )

Titer ( pfu / mL ) = TCID50 / mL × 0.69 pfu / mL

3.3.2. Quantitation of Genome Containing Baculoviruses (q-PCR)

In this method, the quantification of recombinant baculoviruses by quantitative real time PCR is described. The wild-type baculovirus, i.e., one gene sequence is used as template for the amplification of a chosen amplicon of 150 bp (protocol adapted from (34)). 1. Contaminant DNA removal can be performed incubating the samples with DNAse I (1 U/μL) for 30 min at room temperature. 2. Inactivate the DNAse I enzyme, incubating the mixture at 75°C for 10 min. 3. Extract the viral DNA from each sample using the High Pure Viral Nucleic Acid kit following exactly the detailed manufacturer instructions; sample volume for extraction: 200 μL. 4. Mix the reagents from the SYBR green PCR kit in a sterile Eppendorf tube (Master mix), according to the volumes and order shown in Table 2 (see Note 12). 5. Mix and centrifuge the mixture and add 10 μL to the necessary glass capillaries. 6. Add 10 μL of PCR-grade water to the control capillary. 7. Do the sample dilutions and add 10 μL to the respective capillaries; depending on the sample viral titer, 1:100 or 1:1,000 sample dilutions in PCR-grade water should be made so that the DNA copies number stays within the standard calibration curve (see step 8). 8. Dilute the standard baculovirus DNA (that can be previously quantitated with another q-PCR based method (e.g., other template gene) (1:10 serial dilutions down to 102 or 101 copies/10 μL) and add 10 μL of diluted standards to the respective capillaries.


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9. Centrifuge the capillaries and start the q-PCR run using the LightCycler method indicated in Table 3. 10. Analyze the data using the “Fit Points Method” in the LightCycler software; examine the melting curves to check the presence of specific amplification (amplicon peak in the expected melting temperature) and the presence or not of

Table 2 Volume per sample and final concentrations of reagents used (Master Mix) Reagent

Volume (mL)/sample

Final concentration

PCR-grade water

4.4

—

25 mM MgCl2

3.2

4 mM

Forward primer (stock solution at 50 μM)

0.2

0.5 μM

Reverse primer (stock solution at 50 μM)

0.2

0.5 μM

SYBR green master mix

2

1×

Table 3 Quantitative real-time PCR program Program

Temperature Hold time Slope Acquisition target (°C) (s) (°C/s) Cycles mode

Pretreatment

95

600

20

1

None

Denaturation

95

30

20

45

None

Annealing

63

5

20

None

Elongation

72

6

20

Single

Denaturation

95

0

20

Annealing

70

15

20

None

Elongation

95

0

0.1

Continuous

Cooling

40

30

20

Amplification:

Melting: 1

1

None

None

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primer dimers (presence of peaks at temperatures lower than the amplicon’s). The quality of water is of critical importance; however, it is usual that some primer dimers may naturally occur. 11. The calibration curve should present a slope between −3.3 and −3.9 cycle number/log dilution and an error lower than 0.1 (calculated by the LightCycler software). 12. The samples initial DNA copy concentration is calculated according to: Titer (vg / mL) = PCR copies × dilution × 25

where “titer” is the initial sample titer expressed in viral genomes per milliliter of stock (before the DNA extraction step); “PCRcop” is the sample amplicon copies in the capillary, calculated by ies the LightCycler program; “dilution” is the corresponding sample dilution made in step 7; and “25” is a constant value that takes into account the concentration of the sample during DNA extraction (four times) and the extrapolation of the value to milliliters (because the concentration was obtained in 10 μL of sample and the titer is given in viral genomes per milliliter). 3.4. Characterization of the Recombinant Baculoviruses

The characterization of the recombinant baculoviruses throughout the process is crucial. It allows for a proper quality control and determines eventual pitfalls for future rectification and improvement. Herein, we focus on key characterization procedures for the baculovirus system. Total protein determination is essential for quantification of remaining host cell protein; SDS-PAGE profiling enables one to check the coexistence of the expected baculovirus proteins and foreign/host cell proteins; and immunoblotting serves as a product detection method by using an envelope baculovirus protein as a target. Other methods are available and must be used (as applicable) for more detailed characterization, such as DNA quantitation (using commercially available kits) or particle integrity assessment using electron microscopy. Retroviruses, adenoviruses and adeno-associated viruses have their own specificities in the available characterization methods (7, 19, 35); however, their methodologies are very similar to the protocols described herein.

3.4.1. Total Protein (BCA Method)

1. Follow the instructions of the BCA kit from Pierce using 96-well plates. 2. Prepare a set of serial dilutions in duplicate of the standard bovine serum albumin included in the kit (e.g., from 1 mg/mL to 7.82 μg/mL, by serially diluting 1:2 from well to well eight times); use D-PBS as dilution buffer. 3. Assay the samples in duplicate and, if necessary, in different dilutions; use negative control wells with D-PBS. 4. Let the plate incubate for at least 30 min at 37°C.


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5. Check the color formation of the calibration curve samples; if the color of the most-diluted samples is too similar to the negative control samples, let the plate incubate for an extra 10–30 min at 37°C. 6. Measure the absorbance at 562 nm in a 96-well plate spectrophotometer. 7. Subtract the background from all absorbance data using the buffer measurement values, and plot the standard data points to generate a linear regression curve; in some cases, the most concentrated standards could show a somewhat saturated signal thus deviating from the linear trend; these data points may be either neglected or a quadratic (or cubic) fitting could be used instead. 3.4.2. Protein Profile (SDS-PAGE)

1. Run the viral samples by SDS-PAGE using NuPAGE 4–12% Bis-Tris gels. 2. The gels can be stained for protein profile visualization using SimplyBlueTM SafeStain or performing silver staining with SilverXpressTM kit following the manufacturer instructions.

3.4.3. Immunoblotting (for Recombinant Baculovirus Envelope gp64 Glycoprotein)

1. Run the viral samples by SDS-PAGE using NuPAGE 4–12% Bis-Tris gels (see Note 13). 2. After electrophoresis, start the transfer protocol using a semidry transfer cell system Hoefer TE70 (GE Healthcare). 3. Briefly immerse one nitrocellulose membrane (or more if more than one gel has been run; up to four gels can be simultaneously processed in this transfer cell unit) in transfer buffer. 4. Briefly immerse six filter papers/membrane in transfer buffer. 5. Place three filter papers/gel on the transfer unit and above it the membrane. 6. Place the gel(s) over the membrane carefully (preventing gel disintegration) without moving as soon as a contact is established with the membrane to avoid protein transfer and later band smears after immunodetection. 7. Stack three filter papers above each gel and remove the imprisoned air bubbles from the sandwich. 8. Start the transfer, setting the electric source to 0.8 mA/(cm2 of membrane) during 75 min; use a transparency film canvas to block the remaining transfer unit area. 9. Immerse the membrane(s) in blocking buffer in a rocking plate at room temperature for 1 h (or leave at 4°C overnight). 10. Wash the membranes with washing buffer 3 × 3 min. 11. Dilute primary antibody 1:2,000 in antibody dilution buffer (for a final volume of ~10 mL/membrane) (see Note 14).

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12. Incubate the membrane(s) immersed in the primary antibody diluted solution for 1–2 h at room temperature in a rocking plate (or leave overnight at 4°C if the viral content is expected to be low) (see Note 15). 13. Wash the membranes with washing buffer 3 × 3 min. 14. Dilute secondary antibody 1:5,000 in antibody dilution buffer (for a final volume of ~10 mL/membrane). 15. Incubate the membrane(s) immersed in the secondary antibody diluted solution for 1 h at room temperature in a rocking plate. 16. Wash the membranes with washing buffer 3 × 3 min. 17. Immerse the membrane(s) in cold detection solution (1-stepTM NBT/BCIP) until visible protein bands appear (this can take up to 15–30 min); change to fresh NBT/BCIP solution to accelerate the detection reaction. As an example, an immunoblot analysis obtained for the study of the adsorption kinetics of the recombinant baculoviruses onto Matrix 1 (see Subheading 2) is shown in Fig. 4.

a 1

2

3

4

5

6

7

8

9

10

gp64

b 1 0.9

Arbitrary Units

0.8 0.7 0.6 0.5 0.4 0.3 0.2 0.1 0

2

3

4

5

6

Lane

7

8

9

10

Fig. 4. a: Immunoblot for recombinant baculovirus envelope glycoprotein gp64. Samples were withdrawn from an adsorption test using a recirculation loading mode of a Sartobind D 75 membrane adsorber unit. Lanes: 1, molecular weight marker; 2, loading recombinant baculovirus stock; 3, 4, 5, 6, 7, and 8, samples collected during loading at times 0’, 2’, 4’, 8’, 15’, and 45’, respectively; 9 and 10, washing step sample and eluted peak sample, respectively. These results show that the recombinant baculoviruses should have been captured onto this matrix. b: Plot demonstrating band intensities (estimated by densitometry analysis using ImageJ software); numbers here refer to the lanes described in a.


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18. Wash the membrane(s) with water and let dry with absorbent paper.

4. Notes 1. All the water used during the bioreaction and DSP should be, unless otherwise mentioned, of high purity with a resistivity of 18.2 MΩ/cm. 2. Depth-filter capsules are previously rinsed with water and sterilized with a glass female connection (to be later connected to an outlet stream of the bioreactor vessel). These filters are mounted in appropriate stainless steel apparatus provided by the manufacturer, suitable for autoclaving for at least 20 min at 121°C. Bypass tubing should be included for security reasons, because depth-filters may become clogged in the case of high cell density bulks; preferentially, oversized capsules or two smaller ones in parallel should be used to ensure process completion. 3. Cell passage routine is performed using 50-mL working volume suspension cultures in 500-mL previously autoclaved Erlenmeyer shake flasks (Schott); culture conditions: 27°C incubator (without CO2) and 90–110 rpm on an orbital shaker (IKA-Werke, Staufen, Germany); cell seeding is 0.3– 0.5 × 106 cells/mL; allow cell growth up to 3–8 × 106 cells/ mL as desired and assuring good cell viability (>90%); cell passage should be below 40. 4. Primary virus stock is the virus-containing supernatant derived from a transfection of Sf9 cells grown in a static culture. This stock is labeled as the “master viral seed”. 5. Cell concentration and viability can be assessed using the trypan blue exclusion method, i.e., diluting the cell samples 1:2 (previously diluted in PBS or not depending on the original suspension cell concentration) in 0.4% trypan blue exclusion dye (Merck) in PBS and counting the cells on a hemocytometer (Brand, Wertheim, Germany). 6. Sterile conditions are granted by using a flame and working close to the bioreactor in the sterile air environment created; additionally, glass materials must be flamed before and after an opening/switching operation of glass connections or Schott flask caps; laboratory air conditioning should be switched off (if close to the working space) or, e.g., windows must be completely closed. 7. The working viral seed stock must have an infective titer of >8 × 107 pfu/mL to allow moderate viral stock volumes for

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infection in large-scale cultures (moderate MOIs, i.e., 0.01–1 pfu/cell); ultracentrifuge to concentrate (21), or prepare new viral stock if needed. 8. Should the calibration curve for water yield a good linear regression fitting (r2 > 0.98), the cassette can be considered clean; otherwise, use a 0.1 M NaOH solution to get rid of any impurities clogged in the membrane. 9. Maintaining the retentate at low pressure allows higher retentate cross-flow velocities that will prevent sharp drops in the permeate fluxes caused by gel layer formation over the membrane pores. 10. A calibrated multichannel pipette and dispenser should be used for the transfer of viral dilutions and cell seeding to minimize experimental error. 11. If the recombinant baculovirus does not contain a GFPencoding transgene, the recognition of the positive and negative wells should be performed by an operator experienced in cell culture. This can be achieved through comparison between the cells in each well and the cells in the positive and negative control wells with respect to cell morphological changes (cell death/cell lysis demonstrating viral infection). As opposed to a noninfected well (negative control well), the positive wells present lower cell concentrations as a result of cell growth ceasing after virus replication; furthermore, the positive wells typically contain a larger amount of cell debris in suspension. 12. Before starting the preparation for q-PCR, prechill the LightCycler capillary block at –20°C; this will prevent the capillaries from warming up and yielding unexpected annealing and primer dimer formation during sample transfer to the capillaries before the LightCycler run; all material used in this protocol should be disposable and kept strictly sterile, namely pipette tips (with protecting filters) and capillaries. 13. The sample volumes are chosen depending on the expected concentration of recombinant baculoviruses, typically 10 μL of original sample (e.g., crude bioreaction bulk) are diluted in 10 μL of loading buffer included in the NuPAGE kit (Invitrogen) (and then loading the gel with the whole mixture); however, up to 30–40 μL total volume can be loaded onto the specified gel lanes. 14. The primary and secondary antibody incubations are assumed here in small, membrane area-sized lidded boxes; antibody diluted solutions can be reused for up to three times with minor signal loss (stored at –20°C); nonetheless, smaller antibody dilution volumes can be used using in, e.g., sealed plastic bags. 15. If the immunoblotting assay results in nondetectable baculovirus gp64 protein bands, either the primary antibody


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incubation can be prolonged, e.g., overnight at room temperature in a rocking plate; or a larger amount of sample (200 μL or more) can be precipitated before SDS-PAGE using 70% ethanol during 2 h or longer at –20°C, centrifuging at 13,000g, drying the pellet, and dissolving in the chosen volume, for instance, 10 μL of loading buffer.

Acknowledgments We thank Marcos Sousa for the expertise on the upstream processing and Dr. Uwe Gottschalk from Sartorius Stedim Biotech (Göttingen, Germany) for providing the Sartobind membrane adsorber units and the ultrafiltration device. Financial support from the European Commission (Baculogenes, LSH-2005– 1.4.4.6) and the Portuguese Fundação para a Ciência e Tecnologia (SFRH/BD/31257/2006) is gratefully acknowledged.

References 1. Wu, N., and Ataai, M. M. (2000) Production of viral vectors for gene therapy applications. Curr Opin Biotechnol 11, 205–8. 2. Ozaki, H., Govorkova, E. A., Li, C., Xiong, X., Webster, R. G., and Webby, R. J. (2004) Generation of high-yielding influenza A viruses in African green monkey kidney (Vero) cells by reverse genetics. J Virol 78, (4), 1851–7. 3. Cruz, P. E., Maranga, L., and Carrondo, M. J. T. (2002) Integrated process optimization: lessons from retrovirus and virus-like particle production. J Biotechnol 99, 199–214. 4. CBER guidelines (2007), http://www.fda. gov/cber/guidelines.htm 5. Rodrigues, T., Carrondo, M. J. T., Alves, P. M., and Cruz, P. E. (2007) Purification of retroviral vectors for clinical application: Biological implications and technological challenges. J Biotechnol 127, 520–41. 6. Gene Therapy Clinical Trials Worldwide. Charts and Tables, Vectors (2007), http:// www.wiley.co.uk/genmed/clinical/. 7. Cruz, P., Carmo, M., Rodrigues, T., and Alves, P. (2007) Retrovirus Production and Characterization. In Animal Cell Biotechnology: Methods and Protocols, 2nd ed.; Pörtner, R., Ed. Humana Press: Totowa, NJ, pp 475–87.

8. Segura, M. M., Kamen, A., and Garnier, A. (2006) Downstream processing of oncoretroviral and lentiviral gene therapy vectors. Biotechnol Adv 24, (3), 321–37. 9. McTaggart, S., and Al-Rubeai, M. (2002) Retroviral vectors for human gene delivery. Biotechnol Adv 20, (1), 1–31. 10. Cockrell, A. S., and Kafri, T. (2007) Gene delivery by lentivirus vectors. Mol Biotechnol 36, (3), 184–204. 11. Lenz, H. J., Anderson, W. F., Hall, F. L., and Gordon, E. M. (2002) Clinical protocol. Tumor site specific phase I evaluation of safety and efficacy of hepatic arterial infusion of a matrix-targeted retroviral vector bearing a dominant negative cyclin G1 construct as intervention for colorectal carcinoma metastatic to liver. Hum Gene Ther 13, (12), 1515–37. 12. Merten, O. W. (2004) State-of-the-art of the production of retroviral vectors. J Gene Med 6, (Suppl 1), S105–24. 13. Ferreira, T. B., Alves, P. M., Aunins, J. G., and Carrondo, M. J. T. (2005) Use of adenoviral vectors as veterinary vaccines. Gene Ther 12, S73–S83. 14. Volpers, C., and Kochanek, S. (2004) Adenoviral vectors for gene transfer and therapy. J Gene Med 6, (Suppl 1), S164–71.

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15. Graham, F. L., and Prevec, L. (1991) Manipulation of adenovirus vectors. In Methods in Molecular Biology: Gene Transfer and Expression Protocols 7, Murray, Ed. Humana Press: Clifton, NJ. 16. Wold, W. S. M., and Tollefson, A. E. (2006), Adenovirus Methods and Protocols, Adenoviruses, Ad Vectors, Quantitation, and Animal Models. Humana Press: Totowa, NJ. 17. Morenweiser, R. (2005) Downstream processing of viral vectors and vaccines. Gene Ther 12, (Suppl 1), S103–10. 18. Peixoto, C., Ferreira, T. B., Carrondo, M. J. T., Cruz, P. E., and Alves, P. M. (2006) Purification of adenoviral vectors using expanded bed chromatography. J Virol Methods 132, 121–6. 19. Grieger, J. C., and Samulski, R. J. (2005) Adeno-associated virus as a gene therapy vector: vector development, production and clinical applications. Adv Biochem Eng Biotechnol 99, 119–45. 20. Merten, O. W., Geny-Fiamma, C., and Douar, A. M. (2005) Current issues in adeno-associated viral vector production. Gene Ther 12, (Suppl 1), S51–61. 21. O’Reilly, D. R., Miller, L. K., and Verne, A. L. (1994), Baculovirus Expression Vectors: A Laboratory Manual. Freeman: New York. 22. King, L. A., and Possee, R. D. (1992), The Baculovirus Expression Vector System: A Laboratory Guide. Chapman & Hall: London. 23. Kost, T., Condreay, J., and Jarvis, D. (2005) Baculovirus as versatile vectors for protein expression in insect and mammalian cells. Nat Biotechnol 23, (5), 567–75. 24. Kost, T. A., and Condreay, J. P. (2002) Recombinant baculoviruses as mammalian cell gene-delivery vectors. Trends Biotechnol 20, (4), 173–180. 25. Hu, Y. C. (2006) Baculovirus vectors for gene therapy. Adv Virus Res 68, 287–320. 26. Kaikkonen, M. U., Raty, J. K., Airenne, K. J., Wirth, T., Heikura, T., and Yla-Herttuala, S. (2006) Truncated vesicular stomatitis virus G protein improves baculovirus transduction efficiency in vitro and in vivo. Gene Ther 13, 304–12. 27. Raty, J. K., Airenne, K. J., Marttila, A. T., Marjomaki, V., Hytonen, V. P., Lehtolainen,

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P., Laitinen, O. H., Mahonen, A. J., Kulomaa, M. S., and Yla-Herttuala, S. (2004) Enhanced Gene Delivery by Avidin-Displaying Baculovirus. Mol Ther 9, 282–291. Barsoum, J. (1999) Concentration of recombinant baculovirus by cation-exchange chromatography. Biotechniques 26, (5), 834–6, 838, 840. Wu, C., Soh, K. Y., and Wang, S. (2007) Ionexchange membrane chromatography method for rapid and efficient purification of recombinant baculovirus and baculovirus gp64 protein. Hum Gene Ther 18, (7), 665–72. Transfiguracion, J., Jorio, H., Meghrous, J., Jacob, D., and Kamen, A. (2007) High yield purification of functional baculovirus vectors by size exclusion chromatography. J Virol Methods 142, (1–2), 21–8. Blissard, G. W., and Rohrmann, G. F. (1990) Baculovirus Diversity and Molecular Biology. Annu Rev Entomol 35, 127–55. Roldao, A., Vieira, H. L., Charpilienne, A., Poncet, D., Roy, P., Carrondo, M. J., Alves, P. M., and Oliveira, R. (2007) Modeling rotavirus-like particles production in a baculovirus expression vector system: Infection kinetics, baculovirus DNA replication, mRNA synthesis and protein production. J Biotechnol 128, (4), 875–94. Palomares, L. A., Estrada-Mondaca, S., and Ramírez, O. (2006) Principles and Applications of the Insect Cell-Baculovirus Expression Vector System. In Cell culture technology for pharmaceutical and cell-based therapies, Ozturk, S. S.; Hu, W.-S., Eds. Taylor & Francis: New York, pp. 417–431. Lo, H. R., and Chao, Y. C. (2004) Rapid titer determination of baculovirus by quantitative real-time polymerase chain reaction. Biotechnol Prog 20, 354–60. Altaras, N. E., Aunins, J. G., Evans, R. K., Kamen, A., Konz, J. O., and Wolf, J. J. (2005) Production and formulation of adenovirus vectors. Adv Biochem Eng Biotechnol 99, 193–260. Urabe, M., Xin, K., Obara, Y., Nakakura, T., Mizukami, H., Kume, A., Okuda, K., and Ozawa, K. (2006) Removal of Empty Capsids from Type 1 Adeno-Associated Virus Vector Stocks by Anion-Exchange Chromatography Potentiates Transgene Expression. Mol Ther 13, 823–828.

Chapter 25 Production of Plasmid DNA as a Pharmaceutical Martin Schleef and Markus Blaesen Summary Developments in gene therapy, cell therapy, and DNA vaccination require a pharmaceutical gene vector that, on one hand, fulfils the properties to express the encoded information—preferably at the right place, time, and level and, on the other hand, is safe and productive under good manufacturing practices (GMP). Here we summarize the features of producing and modifying these nonviral gene vectors and ensuring the required quality to treat cells and humans or animals. Key words: CGE analysis, contract manufacturing, fermentation, gene therapy, miniplasmid, minicircle, plasmid DNA, pharmaceutical scale, plasmid topology, quality assurance, quality control, vaccination, vector production.

1. Introduction Curing diseases at the level of the specific gene defects rather than at the conventional phenotype level is the first “therapy” as opposed to just working on the modification of a certain disease-correlated phenotype (1, 2). In addition, preventive or curative vaccination against pathogens such as bacteria or viruses or for immunotherapy turns out to at least work in animal studies (3–10). Guidelines for clinical DNA vaccine, cell therapy, or gene therapy trials were available early and the fields of application were reviewed (11–17). Ethical aspects of gene therapy and cell therapy were recently reviewed by King and Cohen-Haguenauer (18). The economic perspectives of this fast-developing field underline the need for the development of industrial-scale processes for the production of plasmid DNA of adequate quality. Such processes have to conform with current GMP guidelines Wolfgang Walther and Ulrike S. Stein (eds.), Methods in Molecular Biology, Gene Therapy of Cancer, vol. 542 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-561-9_25

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and to be acceptable to the regulatory agencies. The production of an active pharmaceutical ingredient (API) requires not only fulfilment of the respective regulatory guidelines, the production also needs to be of a sufficient scale. The term “large scale” was used for productions between 10 and 1,000 mg until the end of the 1990s, but recent large-scale technology for plasmid DNA refers to 100 g up to kilogram scales (19–21). The first promising application might be genetic vaccination with DNA molecules. A major advantage of vaccination with circular plasmid molecules (rather than linear plasmid vectors or viral vaccines) is that there is thought to be no risk of genome integration. Traditional methods of purifying plasmids usually require sophisticated methodology if the DNA is to be separated from contaminating organic components. Plasmids have been produced as drug substances for some years now (19). In the meantime, significant progress has been made on purifying plasmid DNA (22, 23). Compared with proteins, these products require less time for development and for validation of different individual plasmids. Process design can make use of a technology that is generic, at least for plasmids of up to 10-kbp capacity. The requirement for producing plasmids with higher capacities led to the design of different process types—either for large vectors or those considered to be a challenge: these plasmids tend to instability due to their size or sequence structure. Physical stability is distinguished from segregative stability. Progress in gene design and synthesis allows the redesigning of such vectors to avoid problems in manufacturing, which increase the costs of these pharmaceuticals. Any process development has to be accompanied by a powerful in-process control (IPC) system to generate data on the characteristics of the plasmid molecules. In particular, methods are required for obtaining supercoiled covalently closed circular (ccc) plasmid DNA in pure form—the form considered to be the one with lowest risk of chromosomal integration. Commonly, other plasmid topologies appear as well, which have to be separated from the desired product. In addition, the ccc form is the only intact molecule and, hence, is potentially expressed completely and efficiently. From a pharmaceutical point of view, these are the only API molecules that are considered to be homogenous. The established technology of capillary gel electrophoresis (CGE) is the most powerful tool to quantify different plasmid topologies, and additional analytic tests are established to ensure that the new class of drug substances may be safely used for clinical applications (24, 25). A list of applicable quality control (QC) assays for clinical-grade plasmid DNA, manufacturing technology, and examples of process evaluation will be presented here. Recent developments show that a new type of nonviral but not even plasmid DNA vector may successfully enter preclinical

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studies: the “minicircle” DNA. This circular, nonviral DNA vector derives from plasmid DNA but lacks those elements of plasmids not useful or even in doubt for clinical applications. The bacterial origin of replication (ori) and the selection marker used to propagate the plasmid are removed, and the first in vitro data show at least identical results to those obtained with standard plasmids. The minicircle system was developed in a microgram-scale initially and has been scaled up in cultivation to obtain milligram amounts (26).

2. Nonviral DNA Vectors 2.1. Plasmid DNA Vectors

Plasmid DNA has become quite important as a pharmaceutical substance because it was shown that naked DNA injected into muscle tissue was expressed in vivo. Thus, the introduction of immunogenic sequences may result in vaccination against an encoded peptide in an animal model (8, 27–29). Plasmids are circular duplex molecules, which may be stably maintained as episomal genetic information within bacteria (30–32). Their size ranges from 0.8 to ∼120 kbp, and the plasmid copy number per bacterial cell may vary considerably (33). In the case of small plasmids, copy numbers as high as 1,000 copies per cell have been reported. The replication (amplification) does not depend on any plasmid-encoded protein, and is not synchronized with the replication of the bacterial host chromosome (33). The plasmid dimension depends on its form. A linear plasmid of 3 kbp has a molecular mass of 2 × 106 Da and a length of 1 μm (33). The exact form of a plasmid molecule depends on its integrity. Although the ccc form is in a supercoiled state, the “open circular” (oc) form is in a relaxed or nicked state. In addition, monomeric forms are distinguishable from multimeric forms (24, 34). Figure 1 shows the schematic supercoiled (ccc) form, the nicked (oc) form, and dimers of both topologies, as well as linear forms. A potential influence of the plasmid form on the efficacy of DNA vaccines, transfection, co-transfection, and virus production is under investigation (35). The effect of plasmid size was analyzed with different plasmids carrying the same transgene or recently with the comparison of monomeric and dimeric plasmids (35–38). The success of a gene transfer is not only driven by these parameters, but is also and especially driven in combination with the plasmal form by the gene transfer system applied because there are, e.g., transfection, electrogenetransfer (also see Chapter “Electroporative Gene Transfer”), aerosolization, hydrodynamic delivery, ultrasound, or laser beam driven gene transfer (39–46).

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Fig. 1. Plasmid DNA exists in different forms.

Plasmid DNA may be used as a novel vaccine. The major difference between classic vaccines and DNA vaccines is the opportunity of saving time during the development of different vaccines. Classic approaches need years each time—about 10 years for the development of a drug; a substantial part of which is spent on process development (22, 47, 48). No generic process is suitable for manufacturing different protein vaccines, because they are usually different and, hence, require different processing conditions. In contrast, the characteristics of plasmids are not very dependant on the particular target sequence. Therefore, a generic process may be developed for the manufacturing of different plasmid DNA vaccines. During the last 20 years, significant progress was made on the process technology for generating material needed for clinical trials in a fast, economic, and consistent way. The common use of animal-derived and complex raw materials like bovine RNase or fermentation media based on meat peptones could be excluded recently (49). The formulation and stabilization of plasmid DNA for long-term applications is under investigation (additional studies are ongoing), and improvements of the vector system for preventive applications were successfully reached (see “minicircle” within Section 2.4 below and aspects of removing immune stimulatory sequence motifs such as CpG or the optimization of the codon usage) to avoid potential side effects caused by the presence of nonrelevant sequences of the vector system. These CpG motifs that are typically present within prokaryotic (and eukaryotic) sequences within plasmids have a dramatic influence on the expression level of the respective vectors transfected into cells (51–54). One very important advantage of plasmid DNA vaccines is the opportunity of combining vaccine-encoding sequences


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within one vaccine formulation. This would not be possible in the majority of cases with classic vaccines based on proteins or pathogen-inactivated antigens because of, e.g., different storage buffer or formulation buffer requirements (55). Especially in veterinary DNA vaccine applications, the costs for the application per animal are substantial and add up quickly because of the number of different vaccinations required. It would represent a great advantage if all vaccines could be applied in one single injection. No difficulty of this type is expected in mixing different plasmids within one cocktail. However, such an approach may be the result of individual studies performed with the different vectors in individual series of tests. With respect to manufacturing and development costs, it makes sense to design vectors containing more than one epitope for vaccination (56). 2.2. Plasmid Manufacturing: From Vector Design to Fill and Finish

A plasmid manufacturing process must start at the laboratory or pilot scale. Any component used is required to be of a quality that is also available later on for GMP processes. The scaling-up process has to be investigated, and it is not just a multiplication of relevant factors but a time-consuming evaluation of steps to be performed. For example, modification of temperature within a large fermenter requires a fast temperature transfer to all parts of the vessel. In another example, alkaline lysis is not scaled up merely shaking a very large bottle instead of a small one. In line lysis and mixing devices may help to overcome such bottlenecks. Figure 2 gives an overview of a typical large-scale manufacturing process for plasmid DNA.

2.2.1. Production Strain, Cultivation, and Harvest

The manufacturing of plasmid DNA is divided into two major phases. The first phase starts with the transformation of the fully characterized vector plasmid into appropriate and characterized host cells. The resulting genetically modified organism (GMO) is to be checked carefully for the expected characteristics. Subsequently, the GMO will be transferred into the GMP environment for GMP-conforming processing. This processing includes the generation of a master cell bank (MCB) and a working cell bank (WCB), which are required for reproducible large-scale cultivation of the bacterial biomass. The cell banks have to be fully characterized to be of sufficient quality for further manufacturing. Table 1 summarizes the QC assays to be performed with this cell bank material for lot release. The cultivation of Escherichia coli cells is a fast process compared with other strains. Typically, growth overnight is sufficient to obtain large amounts of cell paste with a high amount of plasmid. However, this depends on the proper cultivation parameters (49). The productivity is very important with respect to the subsequent scaling up of any of such processes to keep the product economically possible. Productivity and quality with respect to

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Fig. 2. Overview of a typical large-scale manufacturing process for plasmid DNA. (a) Construction and transformation of the plasmid vector is performed under research and development conditions (R&D). The host cells used for transformation are from a host cell master stock and all further work is performed under GMP; cultivation of E. coli cells in shaker flasks (preculture) up to fermentation and harvesting; (b) purification of plasmid DNA in downstream processing: alkaline lysis followed by filtration and chromatography to remove components other than plasmid DNA. Ultrafiltration or other buffer-exchange procedures are followed by formulation and aseptic filling of the API.

the content of plasmid forms depends on the host strain used. Figure 3 demonstrates the use of different host strains. Most of the host strains show further differences if cultivation conditions change. This happens if the process is modified—starting with scaling up the fermenter size.


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Table 1 QC assays performed with cell bank material for plasmid manufacturing Test

Analytical method

Plasmid identity and integrity Plasmid structure and identity ( 3 enzymes)

Restriction digestion, agarose gel electrophoresis (AGE)

Plasmid yield (μg/mg biomass)

Small-scale preparation and UV absorption (260 nm)

General QC Test of containers

Visual inspection

Plasmid identity and integrity Plasmid identity (sequence)

Sequencing (double stranded)

Host cell identity and integrity Host identity (K12)

Valine sensitivity and phage u3 specificity

Host identity (API-20E)

API-20E identification system (biochemical tests)

Segregative plasmid stability

Replica plating (e.g., 100 colonies) on LB medium with and without antibiotic

Host identity (16S rDNA)

Sequencing (partial) of 5¢ end of 16S rDNA

Host identity (genomic structure)

Riboprinting

Purity Bioburden (microbiological contamination)

Ph. Eur., USP or DAB or comparable

Purity (bacteriophages)

Plating of supernatant on host cells UV induction and plating on host cells

Productivity and performance Pilot cultivation (yield)

Cultivation (5£ L fermenter)

Cultivation is a semigeneric process. With some plasmids, the same process type may be used, with other plasmids (typically those used for virus production likely caused by their repetitive elements such as invested terminal reports (ITR) for adeno-associated viruses or long terminal reports (LTR) for lentiviral transfer plasmids), for each new plasmid, a cultivation process development is required. Figure 4 shows the typical pilot run for GMP process evaluation to determine the cultivation parameters. The biomass of a cultivation needs to be separated from the supernatant liquid

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Fig. 3. DNA from different Escherichia coli host strains transformed with a 6.5-kb plasmid pCMV-luc (PlasmidFactory, Bielefeld, Germany, Art. No. PF461). Lane M = 1 kb DNA ladder (PlasmidFactory, Bielefeld, Germany, Art. No. MSM-865-50), lanes 1–7: E. coli DH5a, Top 10, XL1 blue, DH10B, GT115, TB1, and JM109, respectively. The prominent lower band is the supercoiled (ccc) plasmid form, the upper band shows the open circular (oc) or the dimeric band (may not be distinguishable by AGE). The higher bands represent either multimers or chrDNA of the host cells copurified with the analytic plasmid preparation (Nucleobond PC20, Macherey-Nagel, Düren, Germany). The DNA signal above the lane numbers is a high amount of bacterial chromosomal DNA within the gel loading slots—a major reason to investigate this for the evaluation of plasmid DNA quality rather then cutting this portion off the picture.

of the culture. In laboratory scale, this can be performed by batch centrifugation, in large-scale processing, it can be performed by flow-through centrifugation or cell concentration with tangential flow filtration processes. The biomass is subject to QC tests for product content and absence of any contamination and the biomass will be processed, if it is released for manufacturing. In addition to the use of qualified and well-documented production strains for the microbiological amplification of the

Macherey-Nagel, Düren, Germany) to determine the plasmid yield. The optical density was measured against water. Glycerol determination was carried out on a cation-exchange column (Nucleogel ION 300 OA, Macherey & Nagel) and detection by an RI detector (ERC-7515 A, ERMA CR. INC., Tokyo, Japan). The column was tempered at 70°C (ERC Gecko 2000, ERMA CR. INC., Tokyo, Japan). Sulfuric acid (2.5 mM) was used as eluent at a flow rate of 0.4 cm3/min. Quick plasmid purification was accomplished by means of a standard purification kit with alkaline lysis, anion-exchange chromatography, followed by a precipitation and washing step. After suspension of the DNA in TE buffer, the concentration was determined by spectroscopy at a wavelength of 260 nm (UV mini 1240, Shimadzu). After threefold analysis, the plasmid yield was calculated with the net weight and the DNA concentration.

Fig. 4. (a) Typical pilot run for GMP process evaluation to determine the cultivation parameters. Batch cultivation of E. coli DH5α pF565 on a semisynthetic medium at a 5/dm3 scale. (b) Batch cultivation of E. coli DH5α pF565 on a semisynthetic medium at a 5 dm3 scale: carbon dioxide concentration in the exhausted air of three independent cultivations. The results obtained from three cultivations (a, b, c) were consistent. (c) AGE in-process control (IPC) of bacterial fermentation at different times (cultivation operating time): Lane 1 = 10 h, lane 2 = 11 h, lane 3 = 12 h, lane 4 = 13 h, lane 5 = 15.5 h, lane 6 = 16.5 h, and Lane M = 1 kb ladder (see Fig. 3). Large-scale cultivation of E. coli is performed by fermentation. The fermentations of plasmid pF565, as an example, were carried out in a MBR bioreactor (MBR BIO REACTOR, Switzerland) with a total volume of 7/dm3 operated with 5/dm3 at an operating temperature of 37°C and a pH of 7.0. Adjustment of pH was carried out with a 2 M sodium hydroxide solution and 2M phosphoric acid. The flow rate of air was fixed at 5 L/min. The oxygen concentration (60%) was controlled by varying the stirrer speed in the range of 500 to 2,000/min. The bioreactor was inoculated with 20 cm3 of a shaking flask culture grown in the same medium at 37°C for a period of 4 h. For preculture, 300/cm3 shaking flasks with baffles were used. The filling volume was approximately 60/cm3. The shaking flask cultivations were carried out under selective conditions by adding 0.045/cm3 of a kanamycin stock solution to obtain a final concentration of 75 mg/dm3. The biomass concentration in the inocula was about 0.5–1 g/dm3 (corresponds to an optical density of 2.0–2.5). Before inoculation, the culture was examined under the microscope for possible contaminations. Cells were harvested by centrifugation for 6 min at 9039g, filled in plastic bags, frozen, and stored at —20°C before use for alkaline lysis. During bioreactor cultivations, samples were taken at 1-h intervals by an automatic sample collector and stored at 4°C before determination of the optical density at a wavelength of 600 nm (UV mini 1240, Shimadzu), the glycerol concentration by high-performance liquid chromatography (HPLC) and for quick analytical plasmid purification (NucleoBond® PC 100,

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required plasmid DNA, the cultivation of biomass in fully defined media has become a safety issue with respect to recent discussions on the use of animal-derived raw materials (49). Today’s technology for the generation of complex bacterial growth media uses soy bean peptones to avoid animal-derived protein sources (57) in the face of problems caused by bovine spongiform encephalopathy (BSE) or transmissible spongiform encephalopathy (TSE). Generally, to avoid BSE risk materials, the use of synthetic growth media should be favored, as recommended by regulatory guidelines (58). To ensure high productivity in cultivation, a large biomass concentration with a high plasmid content has to be produced. Generally, these high-biomass concentrations are achieved by fed-batch techniques. Such high cell density cultures have been described for a variety of products derived from E. coli, including recombinant proteins, antibodies, or polyhydroxybutyric acid (59–61). Feed of concentrated medium may be controlled by monitoring different operating variables in the bioreactor like pH, or dissolved oxygen, indirect determination of the specific growth rate, or by online monitoring of a limiting substrate (62–66). Several processes for plasmid DNA production have been described, most of them aimed at producing high biomass and product concentrations only. The homogeneity of the plasmid at the cultivation stage is rarely addressed. Reinikainen et al. 1989 examined the influence of pH and temperature on plasmid copy number in cultivations in a semidefined medium (67). However, no statement was made regarding plasmid homogeneity. Lahijani et al. 1996 described the cultivation of a pBR322-derived plasmid. The copy number of the plasmid was increased by introducing a temperature-sensitive point mutation (68). Setting the cultivation temperature to 42°C in the growth phase resulted in a plasmid concentration of 37 mg/ dm3 in batch experiments on semidefined medium, and 220 mg/dm3 in fed-batch experiments. However, the isolated DNA was a nonhomogenous product comprising several multimeric plasmid forms and chromosomal DNA. Additionally, segregative plasmid stability was maintained by supplementation of antibiotics. Schmidt et al. 1999 described a dissolved oxygencontrolled fed-batch cultivation on a defined glycerol medium (24). A product concentration of 100 mg/dm3 and a dry biomass concentration of 48 g/L were achieved, resulting in a yield of 2.1 mg/g. The cultivation of E. coli to high cell densities for plasmid DNA production in a batch mode was described by Voss et al. 2004 (69). By using a fully defined synthetic glycerol medium,


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45 mg/dm3 plasmid DNA could be produced. Recent developments allow even higher productivity (21). For subsequent purification, the produced biomass is separated from the culture medium by centrifugation or microfiltration and stored at low temperatures (–20°C). A cultivation process needs to be performed with identical results at least three times to ensure constant quality of biomass. In scaling up, the quality of biomass should fulfil identical specifications. Because the scale of a fermenter might be limited by economic or physical parameters, it is worth thinking about feeding strategies for plasmid manufacturing. This would allow keeping a small volume for cultivation while reaching high productivity. All of these developments require solid process development and can lead to a multiplication of cultivation capacity. 2.2.2. Downstream Processing: Cell Lysis, Chromatography, and Filling

Plasmid molecules are usually released from host cells by alkaline lysis. This process step is critical because the major contamination source of plasmid DNA productions is the bacterial chromosomal DNA (chrDNA) also released at this step (19). If this high molecular weight material should be subject to degradation, either by enzymatic or physical matters (e.g., shear force), the resulting fragments will be present within the subsequent purification steps and will need to be removed there. Because removal of these fragments is a major task in plasmid manufacturing, it is worth avoiding this by keeping the bacterial high molecular weight DNA intact and removing it together with bacterial debris within the lysis separation and filtration. Because the cleared lysate contains a large amount of potentially contaminating substances (Table 2), the removal has to be evaluated on the basis of these contaminants.

Table 2 Constituents of Escherichia coli lysates (70) Content of bacterial cell lysates Proteins

55%

RNA

21%

Host chromosomal DNA

3%

Lipopolysaccharides

3%

Plasmid DNA

3%

Others

15%

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Cleared lysates contain only about 3% plasmid DNA. The separation of plasmid DNA from host cell impurities comparable to the product in their physical and chemical characteristics is a major challenge for bioprocess engineering. The requirement of also scaling up the lysis step was earlier approached by thermal alkaline lysis in combination with in-line processing. Significant progress in performing the alkaline lysis in line was described previously (71–74). The resulting lysate is separated from insoluble matter and cell debris. This “cleared lysate” is sterilized by filtration for further downstream processing (DSP). This DSP comprises the second phase of plasmid manufacturing (Fig. 2b). Here the plasmid molecules should be separated from soluble biomolecules (e.g., host chromosomal DNA, RNA, nucleotides, lipids, residual proteins, amino acids, and saccharides), salts, and buffer components. A typical approach is chromatography. For clinical applications, no other way is possible at present. The design of the chromatographic process depends on the required purity of the plasmid product. So far, only a few techniques are used in pharmaceutical-grade processing of plasmids. Anion-exchange chromatographic techniques were initially described for certain processes (19, 74–79). Alternative chromatographic technologies make use of size exclusion chromatography for small scale, and of reverse phase chromatography or hydrophobic interaction chromatography. Other approaches describe plasmid DNA purification systems for the selective removal of contaminants or improved binding capacity for the intended plasmid product (e.g., monolithic stationary phase) (60, 80–82). The second phase of plasmid manufacturing ends with bulk purified plasmid DNA being formulated within the appropriate buffer or solution for further processing or storage and application. Recent manufacturing technology makes use of removing the undesired open circular (oc) and linear plasmid forms as well as of fragmented chrDNA from the bacterial chromosome. The removal of damaged plasmid forms can be demonstrated by agarose gel electrophoresis (AGE) and CGE, the removal of chrDNA by quantitative polymerase chain reaction (qPCR) ((49), PlasmidFactory, unpublished). 2.3. Improvements in Plasmid-DNA: Quality Control (QC) and Quality Assurance (QA) 2.3.1. Quality Assurance in Manufacturing Plasmid DNA

The type of production process used for plasmid DNA purification is responsible for the quality profile of the product. This profile can be monitored within the finished product but, ideally (and conforming to GMP), this is already done during evaluation and validation, and, in all cases, by IPC while production is running. Table 3 summarizes all QC assays to be performed for the API and, if possible, in parallel with any manufacturing. The extent of performing these assays depends on different aspects:


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Table 3 Test assays and required quality controls for the quality assurance and product release of plasmid DNA Test

Method

DNA concentration

UV absorption (260 nm)

Appearance

Visual inspection

DNA homogeneity (ccc content)

Densitometry

DNA homogeneity (ccc content)

CGE (capillary gel electrophoresis)

DNA purity

UV scan (220–320 nm)

RNA

Agarose gel (visual)

Endotoxin (LPS)

Limulus amebocyte lysate (LAL) test

Bacterial chromosomal DNA

Quantitative PCR

Protein

Bicinchoninic acid (BCA) test

Purity (microorganism)

Bioburden

Plasmid identity (different restriction enzymes)

Restriction digestion and agarose gel

Identity

Sequencing (double strand)

the intended use of the product, the decision of any regulatory body involved, and the respective rules, guidelines, and regulations, including GMP. Recommended references are summarized by the European Medicines Evaluation Agency (EMEA) and the World Health Organisation (WHO) (83, 84). Plasmids of identical nucleotide sequence isolated from E. coli may exist in different shapes and forms. Previous literature used a nomenclature that was significantly consolidated by Schmidt and coworkers in 1999 (24, 31, 85–91). The ccc isoform exhibits the most compact structure. If one strand is broken (nicked), the oc form results, and coiling is lost. Linear forms are generated when both strands are cleaved once at approximately the same position. In addition, plasmids may appear as oligomeric forms, e.g., concatemers, which may, in turn, exist as different isoforms. All forms may be isolated in plasmid DNA manufacturing processes and should be considered as contaminations as long as their mode of action is unknown (92). Furthermore, the appearance of such forms requires ensuring that they appear reproducibly if they cannot be excluded completely. The reason for this is that the plasmid product should be as homogeneous as possible because it is intended for use as a drug. Further forms are catenanes, which are oligomeric plasmids

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connected like chain links. However, these forms appear mainly in kinetoplasts and rarely in bacterial plasmid preparations (90). The usual method applied to determine the plasmid form distribution was by agarose gel electrophoresis (AGE). Different bands in AGE from an in-process control sample or a plasmid product sample may be assigned to different plasmid isoforms. The assignment of bands to forms, however, is not easy because the electrophoretic mobility of plasmids of different shape changes with the electrophoretic operating conditions (86, 91, 93, 94). In addition, the quantification of isoforms based on the signal intensity of stained bands in AGE may not be reliable due to no linear responses. Adequate equipment is required to obtain reproducible results. In cases where only some restriction enzyme digestions are performed to evaluate the fragment sizes of a plasmid, the isoforms cannot be distinguished. We recommend an AGE assay with undigested plasmid samples at least. It is well known that typically only one band, the ccc form, is observed when only a small amount of a plasmid sample is applied to an agarose gel. Standard AGE usually reveals two prominent bands: the ccc form and another slower-migrating form, commonly thought to be the oc form (see Fig. 1). Schmidt and coworkers showed that this is not always the case (24). It could be demonstrated that the slower migrating upper band within AGE is the oc form and, at nearly the same position, also the ccc dimer form of the plasmid. Earlier publications reported the separation of monomeric forms only (95–97). CGE enables identification of the other forms mentioned, as well as the ability to individually quantify them ((24, 98); see also www.CGEservice.com). CGE analysis was used to perform long-term real-time stability studies of plasmid DNA where the storage of DNA could be monitored (50). Another aspect that makes CGE outstanding is that the characterization of a plasmid sample can be performed in less than 50 min and requires only 50–100 ng of plasmid sample. Plasmid sizes of up to 37 kb were analyzed so far (PlasmidFactory, unpublished). Additional studies were carried out to determine whether the DNA concentration, the buffer, the containment, or the plasmid size are responsible for any damage in storage. In addition, the way of freezing or thawing DNA was considered to be of influence and analyzed (PlasmidFactory, not published). Figure 5 demonstrates first results from lyophilization and subsequent reconstitution after storage at room temperature of plasmid DNA (pCMV-luc) carrying a reporter gene function. 2.4. Improvements of Nonviral Vectors: Minicircle Technology

The general structure of plasmids consists of the following elements: origin of replication (ori), selection marker (typically antibiotic resistance element like, e.g., kanamycin resistance [kan], or ampicillin resistance [bla]), and the element containing the gene of interest (GOI).

Fig. 5. (a) AGE and (b) CGE analysis of the 6.5-kb plasmid pCMV-luc (PlasmidFactory, Bielefeld, Germany, Art. No. PF461) before lyophilization and after lyophilization and reconstitution. The DNA was resuspended in water for injection (WFI). a: Lane M = 1 kb DNA ladder (PlasmidFactory, Bielefeld, Germany, Art. No. MSM-865–50), lanes 1–4 = before lyophilization, lanes 5–8 = after lyophilization and reconstitution. DNA in lanes 3 and 4 as well as in 7 and 8 were restriction enzyme digested with Eco RI. b: Electropherograms before and after lyophilization and reconstitution. The plasmid DNA shows 79.3% ccc monomer, 16.7% ccc dimer, and 4% oc forms before lyophilization. After lyophilization and reconstitution, the results were 79.6% ccc monomer, 16.4% ccc dimer, and 4% oc, demonstrating no damage of plasmids in this process step.

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Fig. 6. Scheme of the MC recombination system (26).

In general, the optimal vector would contain only (!) the gene cassette with the GOI and no other elements. The antibiotic resistance elements are especially considered redundant and a safety problem. Within the chapter use of minicircle plasmids for same theory, we show the most significant improvement of the last years on nonviral vectorology: minicircle (MC) DNA vectors (26). These vectors derive from parental plasmids with antibiotic resistance markers, GOI, and ori, as well as two special recombination sequences right and left of the GOI. Through an intramolecular recombination process, the GOI (plus one of the signal sequence elements) is cut out of that parental plasmid, circularized, and finally consists of only the GOI and the signal sequence in a circular molecule. The residual part of the parental plasmid contains the ori, the selection marker, and is, by definition, still a plasmid and hence called a miniplasmid (Fig. 6). We developed a process technology to produce large amounts of this minicircle vector by fermentation (PlasmidFactory, unpublished).

3. Future Perspectives Approximately 25% of all gene therapy protocols performed so far in clinical studies were directly based on plasmid DNA vectors (99). The market share of plasmid DNA vectors with respect to all


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vaccines was initially expected to rise to about 60% (100). This also includes the plasmid DNA used to produce viral vectors—e.g., by transient transfection of producer cells. However, diverse aspects of producing plasmid DNA need still to be addressed to develop processes for the safe and economic production of large amounts of plasmid DNA. As soon as minicircle vectors enter the large-scale market, they will become a major part of those applications where either the direct application into human or animal requires an improved safety profile or where the manufacturing of viral particles or antibodies requires the specific features of those patients. Further improving nonviral plasmid or minicircle vectors— e.g., by adding scaffold/matris attachment region (S/MAR) elements—might increase the efficacy of the vector system, although recent studies with a first-generation minicircle system demonstrated that it did not show episomal replication in liver cells (101–106). Recent studies of the first scalable “plasmid factory minicircle system” demonstrate a significant improvement of transfection efficacy with minicircle DNA (107). In addition, the following aspects will influence the use of gene vectors on their way from research and development into clinics. 3.1. Subcontracting Plasmid Manufacturing and Availability of Large Plasmid DNA Amounts

Outsourcing research, development, and biopharmaceutical manufacturing for preclinical and the first steps in clinical research has become an attractive way around the risk of time consuming and expensive construction of manufacturing plants. Small start-up biotech and pharmaceutical companies with a strong scientific background but limited financial resources especially benefit from this development within their first years in developing innovative biotech-based therapeutics. Limiting the economic risk in drug development is a necessary aspect for larger pharmaceutical companies. Hence, these large companies also benefit from the availability of subcontractors for the manufacturing of their products for clinical phases I through II, maybe even III. If successful until this point, commercial manufacturing may require dedicated facilities. The increasing amounts of plasmid DNA—namely for DNA vaccination—will lead to a requirement of very large-scale manufacturing technology in combination with an effect on the costof-goods for the pharmaceutical. However, scaling up is not only a question of fermenter size but also of cultivation conditions and aspects of harvesting and processing (see above).

3.2. Critical Aspects: Animal-Derived Growth Media

Culture media for the growth of microorganisms were developed in the 19th and 20th centuries. Liebig extracted beef and concentrated the essence for storage and later use in media preparation for the cultivation of microorganisms. The history

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of development, manufacture, and control of microbiological culture media was summarized by Bridson 1994 (57). One major improvement for such media was the addition of peptones and salts, which led to the increased supplementation with amino acids and an enhanced osmolarity. These peptones were generated by enzymatic digest of meat. Other protein sources were also digested to generate peptones. If it is indicated not to have any potential source of animalderived components within a plasmid manufacturing process, complex media should be completely avoided. Data is available on cultivation processes with the intent to manufacture plasmid DNA using synthetic growth media (see www.PlasmidFactory. com) and able to ensure at least comparable product yields. High cell density cultivation processes for plasmid production based on synthetic media are available (21). If vegetable peptones shall be used, it should be considered that no enzymes of animal origin are applied (e.g., pepsin, pancreatin, or trypsin). Hydrolysis with acids avoids the use of such enzymes but leads to higher salt content. A further alternative is the use of yeast extract. If no enzymatic digestion was performed in production, this may be a complex component of choice. However, even this yeast may have been grown on culture medium containing components of animal origin. A promising approach to make use of complex components while excluding any external contamination is the use of a homologous complex component for cultivation (108). 3.3. Critical Aspects: Animal-Derived Processing Aids

Ribonucleases (RNases) are able to hydrolyze phosphodiester bonds within RNA molecules. These enzymes are needed in cells to modify and process RNA molecules. In plasmid production processes, one common major contaminant is RNA from E. coli. In general, RNA has a short half-life, but is a substantial contamination in plasmid preparations. This RNA may block the binding capacity in, e.g., anion-exchange chromatography. Therefore, an enzymatic digestion of RNA before the chromatographic step is usually applied (75, 77, 78). If this is carried out in pharmaceutical manufacturing processes, the question must be raised whether the quality of such processing aids, which normally are derived from animal sources, is sufficient. Because RNase A is typically prepared from bovine pancreas, the origin of these glands has to be assessed and certified. Some plasmid manufacturing processes completely avoid the use of RNase. In these cases, the RNA is removed by specific precipitation techniques (80, 109, 110). However, this was only applicable in a laboratory scale so far. We developed a recombinant RNase to replace the traditional one. The manufacturing process is not trivial because the RNase is also active within the producer cell. If the process for plasmid


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manufacturing does not allow avoidance of this processing aid, it can be used in large amounts. 3.4. Critical Aspects: Bacterial Chromosomal DNA

A major contaminant of plasmid preparations commonly is the bacterial chromosomal DNA (chrDNA). It has to be reduced significantly or, ideally, be removed by the process steps applied. The most critical step is the alkaline lysis, where the chrDNA as well as the plasmid DNA are denatured by a pH shift to alkaline buffer conditions. A subsequent neutralization step allows the reannealing of plasmid DNA within a short time. The chrDNA, however, does not reanneal completely to a DNA double strand. Therefore, the major part of the chrDNA will be a component of the flaky material that is generated by the alkaline lysis. It mainly consists of potassium dodecyl sulfate, insoluble proteins, cell debris, lipopolysaccharides (LPS), and DNA. Chromosomal DNA is extremely sensitive to shear, which may easily result in DNA fragmentation. A removal of DNA by subsequently applied process steps is difficult. Some chrDNA fragments are large enough to migrate in one distinct band in AGE or be stuck within the gel slot when being loaded. Smaller fragments can be detected as an undefined smear by overloading the AGE gel. Below a certain size, these fragments leave the gel without being detected. More sensitive assays like Southern blot hybridization can demonstrate this—depending on the hybridization and washing conditions applied (111). A most sensitive assay was recently presented by Smith et al. 1999, for which a kinetic PCR method using a TaqMan probe was used to quantify chrDNA contaminations (112). This technique makes use of the detection of the seven-copy 23S-rDNA gene in E. coli. Other systems are also available today. The removal and quantification of chrDNA is of high importance, because this contamination will be present within a plasmid DNA product even when a CpG-free plasmid is manufactured. In such cases, the effort to obtain a so called “zero-CpG plasmid” is foiled because the chrDNA contains a lot of CpG (113). Levy et al. claim to be able to remove chromosomal E. coli DNA from plasmid-containing process liquids to a residual content of less than 1% (114). These authors describe how cleared lysate, resuspended PEG precipitates, or anion-exchange chromatography eluates were filtered through nitrocellulose membranes to selectively remove chromosomal DNA. From our observation, the removal has to start with a biomass of high quality (low chrDNA) to end up with a sufficient quality rather than removing this in a time-consuming and costly process. A general safety aspect is the potential integration of gene vectors into the chromosome of the transfected cells. Studies analyzing the fate of the administered DNA were performed (115, 116). The integration of a plasmid could be monitored

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through the knowledge on its exact sequence. In contrast, detecting the integration of a piece of contaminating bacterial chromosomal DNA is a lot more difficult and, hence, needs to be excluded on the level of plasmid DNA quality. 3.5. Next Goals in DNA Manufacturing

The clinical development of DNA vaccines, gene therapeutics, or even viral vectors produced through transient transfection of producer cells requires a significant development of: (a) Validated QA assays to ensure highest quality of gene transfer pharmaceuticals (even for initial clinical trials or preclinical work) (b) Processes to obtain plasmid qualities significantly increased with respect to critical contaminants such as bacterial chromosomal DNA (c) More economic ways of manufacturing plasmid DNA Additionally, improvements such as the lyophilization of DNA without causing damage when performing the lyophilization, storage, or reconstitution of the DNA; the new types of efficient and vector systems with improved safety without bacterial ori and antibiotic resistance marker sequences; as well as increased efficacy through codon usage or, in general, sequence optimization, will increase the development of DNA pharmaceuticals.

Acknowledgments We thank the German Federal Ministry of Education and Research (BMBF) for grants BioChancePLUS (0313749) and Nano-4-Life (13N9063), the research team of PlasmidFactory, Bielefeld, Germany for contributing to the work and the whole manufacturing team of PlasmidFactory for their discussion. We also thank M. Schmeer and our partners within CLINIGEN (FP6) for critical discussion. References 1. Alton, E. W. F. W., Middleton, P. G., Caplen, N. J., Smith, S. N., Steel, D. M., Munkonge, F. M., Jeffery, P. K., Geddes, D. M., Hart, S. L., Williamson, R., Fasold, K. I., Miller, A. D., Dickinsons, P., Stevenson, B. J., McLachlan, G., Dorin, J. R., and Porteous, D. J. (1993) Non-invasive liposome-mediated gene delivery can correct the ion transport defect in cystic fibrosis mutant mice. Nat. Genet. 5, 135–142. 2. Caplen, N. J., Gao, X., Hayes, P., Elaswarapu, R., Fisher, G., Kinrade, E., Chakera, A., Schorr,

J., Hughes, B., Dorin, J. R., Porteous, D. J., Alton, E. W. F. W., Geddes, D. M., Coutelle, C., Williamson, R., Huang, L., and Gilchrist, C. (1994) Gene therapy for cystic fibrosis in humans by liposome-mediated DNA transfer: U.K. regulatory process and production of resources. Gene Ther. 1, 139–147. 3. Michel, M. L., Davis, H. L., Schleef, M., Mancini, M., Tiollais, P., and Whalen, R. G. (1995) DNA-mediated immunization to the hepatitis B surface antigen in mice: Aspects of


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Subsection B Clinical Protocols in Cancer Gene Therapy

Chapter 26 Gene Immunotherapy for Non-Small Cell Lung Cancer John J. Nemunaitis Summary Historically, limited results have been observed with immunity in non-small cell lung cancer (NSCLC). In the last 5 years, however, several immune-stimulating products have demonstrated enhancement of tumor antigen recognition through activation of dendritic cell-involved processes. Moreover, clinical benefit has been demonstrated in subsets of patients, justifying ongoing phase III investigation. Results of key gene immunotherapies being tested in NSCLC are reviewed. Preliminary results in advanced NSCLC suggest evidence of well-tolerated immune activation with suggested evidence of clinical benefit with respect to survival and response. Key words: Gene immunotherapy, non-small cell lung cancer.

1. Introduction For decades, immune-based therapies have been investigated in many types of cancer, with a focus related to observed activity in melanoma, prostate, renal cell, non-Hodgkin’s lymphoma, bladder cancer, and renal cell carcinoma (1–6). Lung cancer, however, has historically not been considered an immunogenic tumor. Historically, there have been several hypotheses to explain the potential lack of anticancer immune activity in NSCLC. These include ineffective priming of tumor-specific T cells, lack of high avidity of primed tumor-specific T cells, and physical or functional disabling of primed tumor-specific T cells by the primary host and or tumor-related mechanism. For example, in NSCLC, a high proportion of the tumor-infiltrating lymphocytes are immunosuppressive T regulatory cells (CD4+ CD25+) that secrete transforming growth factor (TGF)-β and express a high Wolfgang Walther and Ulrike S. Stein (eds.), Methods in Molecular Biology, Gene Therapy of Cancer, vol. 542 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-561-9_26

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level of cytotoxic T lymphocyte (CTL) antigen-4 (7). These cells have been shown to impede immune activation by facilitating T cell tolerance to tumor-associated antigens rather than cross-priming CD8+ T cells, resulting in the nonproliferation of killer T cells that recognize the tumor without attacking it (7–9). Elevated levels of immune inhibitors such as interleukin (IL)-10 and TGF-β are found in circulation in patients with NSCLC, and animal models have shown that immune suppression is mediated by these cytokines serving as a defense for malignant cells against the body’s immune system (10–12). Recently, however, several advances have been made with respect to mechanisms by which the immune system can actually be manipulated to enhance tumor antigen recognition and provide anticancer activity in NSCLC. Within the next 5 years, it is possible that clinical trials involving gene-based vaccines such as a Belagenpumatucel-L, GVAX®, B7.1, L523S, and α(1,3)-galactosyltransferase will progress and expand our knowledge of the immune system’s role against cancer, potentially further improving our clinical experience with the use of various vaccines involving different stages of disease in NSCLC. This chapter reviews clinical aspects of gene-based vaccines in NSCLC. Our understanding of gene vaccine activity in NSCLC may also provide evidence to successful application in management of other solid tumors.

2. Summary of Current Therapy in NSCLC

The 5-year survival rates for patients with early stage (£IIIA) NSCLC range from 13% to 61% with effective local regional management (surgery, radiation) (13–20), while patients with advanced disease, Stage IIIB or IV, have a 5-year survival rate of less than 5% (19). Standard systemic therapy in advanced stage lung cancer involves combination of cytotoxic agents, angiogenesis inhibitors, and small molecule tyrosine kinase inhibitors. Cytotoxic chemotherapies are based on platinum doublet regimens and have been the standard of care for first-line treatment of advanced NSCLC for several years (21). Recently, the potential therapeutic advantages of combining cytotoxic therapies with agents targeting the tumor vasculature have been discovered (22). In the phase III Eastern Cooperative Oncology Group (ECOG) 4599 trial in previously untreated nonsquamous NSCLC, the addition of bevacizumab to paclitaxel/carboplatin yielded significant improvements in overall response rate (ORR) (35% vs 15%; P < 0.001), median progression-free survival (PFS) (6.2 months vs 4.5 months; P < 0.001), and median overall survival (OS) (12.3 months vs 10.3

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months; P = 0.003) (23). Although the addition of bevacizumab yielded a significant increase in efficacy, paclitaxel/carboplatin/ bevacizumab was associated with significant increases in hematologic and nonhematologic toxicities compared with paclitaxel/ carboplatin alone, including grade 4 neutropenia (25.5% vs 17%; P = 0.002), febrile neutropenia (grade 3: 4% vs 2% and grade 5: 1.2% vs 0.2%; P = 0.02), grade 3/4 hypertension (7% vs 1%; P < 0.001), grade 3/4 proteinuria (3% vs 0; P < 0.001), and bleeding events (all grades) (4% vs 1%; P < 0.001) (23). Moreover, treatment-related deaths were more frequent with the addition of bevacizumab. Docetaxel, Alimta, and erlotinib are approved for secondline NSCLC (24–29). Response rates of 10% or less are demonstrated and the median survival rates achieved range from 7 to 8 months. Despite improvement of 2–3 months in median survival compared with palliative management with no anticancer therapy, the success of cytotoxic and targeted therapy in second-line treatment of NSCLC is far from optimal. Recently, studies have been performed to evaluate the effect of gene-based vaccines in NSCLC. Encouraging results have been observed and are described in subsequent sections.

3. Immune Function Capacity in Cancer Patients

Tumor cells often inappropriately express normal cellular antigens and neo-antigens that are recognized by the immune system of the tumor-bearing host. The production of tumor-specific antibodies by B cells can be elicited by the immunization of animals with tumor cells, and tumor-associated determinants have been defined by their ability to stimulate an antibody response in association with their presence on tumor cells. Detectable titers of antibodies that bind to tumor-associated antigens have been demonstrated in cancer patients, but there is no compelling evidence that administration of exogenous antibodies to tumor-associated determinants cause solid tumors to regress in humans. However, there is an accumulating body of evidence to suggest that many tumors in experimental model systems and from cancer patients express molecules that are recognized by the T cell arm of the immune system. Rejection of antigen-expressing tumor cells has been shown to be mediated by specific host cytolytic T cells (30, 31). Tumor infiltrating lymphocytes (TIL) have also been shown to mediate durable regression of established pulmonary tumors in mice with advanced tumor burdens (32, 33). In patients bearing metastatic tumors, a number of groups have demonstrated the existence of

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antitumor CTL responses. Peripheral blood mononuclear cells as well as TIL contain populations of cells and individual clones that demonstrate tumor specificity; they lyse autologous tumor cells, but not natural killer targets, allogeneic tumors cells, or autologous fibroblasts (34–38). Therefore, tumor-specific antigens exist on metastatic human tumors, and are capable of stimulating a specific T cell response that can be expanded ex vivo to achieve clinical objective responses in tumor-bearing patients. Endogenously synthesized antigens of virtually all mammalian cells are processed in an endoplasmic reticulum compartment and converted to small epitope peptides that are subsequently displayed on the cell surface in association with Class I major histocompatability complex (MHC) molecules (39–41). However, specialized antigen processing cells (dendritic cells) are capable of presenting very small numbers of exogenous peptides in association with Class I molecules to stimulate T cell recognition (42, 43). The role of dendritic cells (DCs) in cell-mediated immunity has been extensively investigated (44–48). DCs have been found to play a central role in the induction of antitumor immunity in tumor-bearing hosts by a process of antigenic cross-presentation and have displayed activity in NSCLC (49). They efficiently display antigens on MHC II, ultimately stimulating proliferation and activation of CD4+ and CD8+ T cells. CD4+ cells further augment the activity of natural killer cells and macrophages, in addition to amplifying antigen-specific immunity by local secretion of cytokines (50–54). Recent studies in animal models have shown that immunization with dendritic cells loaded with defined tumor antigens in the form of peptides, proteins, or RNA are capable of priming CTL responses and inducing tumor immunity (55–58). Dendritic cells mixed in vivo with irradiated tumor cells have been shown to produce a protective immune response against a challenge with autologous tumor cells, using a nonimmunogenic breast cancer cell line, 4T1 (59). Tumor-specific CTL response was also detectable. Clinically, infusion of dendritic cells has shown evidence of antitumor activity in patients with NSCLC (49) and other solid tumors (60, 61). These attributes make DCs a pivotal component in therapeutic strategies of many current immune-based therapies in NSCLC.

4. Vaccine Therapies 4.1. TGF-b2 Antisense Gene Vaccine

LucanixTM is a nonviral gene-based allogeneic vaccine that incorporates the TGF-β2 antisense gene into a cocktail of four different NSCLC cell lines (62). Elevated levels of TGF-β2 are linked to immunosuppression in cancer patients (63–68). Systemic levels of TGF-β are inversely correlated with prognosis in patients with


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NSCLC (69). TGF-β2 has an antagonistic effect on natural killer cells, lymphokine-activated killer cells, and DCs (11, 12, 70–73). Using an antisense gene to inhibit TGF-β2, several researchers have demonstrated an inhibition of cellular TGF-β2 expression resulting in an increased immunogenicity of gene-modified cancer cells (74–82). In a recent phase II study involving 75 early stage (n = 14) and late stage (n = 61) advanced NSCLC patients, a dose-related effect of LucanixTM was defined (62) (Tables 1 and 2). Patients were randomized to one of the three dose cohorts (1.25 × 107 cells/injection, 2.5 × 107 cells/injection, or 5 × 107 cells/injection; each, 16 injections). Injections were administered one time each month or every other month until progressive disease criteria were fulfilled. Treatment was well tolerated with only one grade 3 event attributed to the vaccine (arm swelling). A significant survival advantage at dose levels ³ 2.5 × 107 cells/injection, with an estimated 2-year survival of 47% in response to LucanixTM in 41 advanced Stage IIIB or IV patients, was found (Tables 1 and 2). This compared favorably with the historical 2-year survival rate of 75 years old) or in patients with bulky or rapidly progressive disease. As such, appropriate trial design with combination or sequential use of gene vaccines with other standard approaches needs to be further studied. It is not known whether combination with other targeted therapies or cytotoxic therapy will enhance any potential effect of the vaccine. Furthermore, debulking surgical management followed by vaccination may provide additional opportunity for investigation. Completion of phase III investigation will address these important questions.


4.2. Granulocyte Macrophage Colony-Stimulating Factor Gene-Based Vaccines in NSCLC

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Given the histological heterogeneity of NSCLC and the relative absence of information on the relevant immunodominant antigens in this disease, initial trials with GVAX® used autologous tumor cells as the source of tumor antigens in NSCLC (83) (Tables 1 and 2). The first pilot study of autologous GVAX® in NSCLC involved harvest from 35 patients. Thirty-three patients received vaccine treatment at three different dose levels (1 × 106 cells/injection, 4 × 106 cells/injection, and 1 × 107 cells/injection). The vaccine was administered weekly for 2 weeks then biweekly until the supply was exhausted. Vaccines were well tolerated, with the most common toxicity being local, self-limited vaccine site reactions and mild flu-like symptoms in a minority of patients. Antitumor immunity was demonstrated by induction of delayed-type hypersensitivity (DTH) reaction to injections of irradiated, genetically unmodified autologous tumor cells in 82% of patients as well as the presence of inflammatory infiltrates in metastatic tumor biopsies. In addition, one patient demonstrated evidence of tumor regression (mixed response), five patients achieved stable disease for a median of 12 months, and two other patients have remained recurrence-free for more than 5 years after resection of isolated metastatic sites harvested for vaccine preparation. Subsequently, a multicenter phase I/II trial investigating another autologous NSCLC tissue vaccine was performed. Manufacturing processes were modified to enable potential commercial development. This study also involved patients with both early stage and advanced stage disease (84). Patients were enrolled in two cohorts. Cohort A included patients with Stage IB or II NSCLC with planned primary surgical resection and no preoperative or postoperative chemotherapy or radiotherapy. Patients in cohort B had surgically nonresectable Stage III or IV NSCLC and accessible tumor to harvest for autologous vaccine processing. Vaccines were administered subcutaneously every 2 weeks for a total of three to six vaccinations. The vaccine dose was individualized based on yield. Dosing fell into three categories or cohorts: (1) 5 × 106 to 10 × 106 cells per vaccination; (2) 10 × 106 to 30 × 106 cells per vaccination; and (3) 30 × 106 to 100 × 106 cells per vaccination (Tables 1 and 2). Eighty-three patients underwent tumor harvest (20 in cohort A, 63 in cohort B) and 43 patients initiated vaccine treatment (10 in cohort A, 33 in cohort B). All ten patients in cohort A completed vaccine treatment. The median number of vaccinations in cohort B was five. The median number of days from tumor harvest to vaccine release was 31 days and from harvest to initiation of vaccine treatment was 49 days. Vaccines were successfully manufactured in 80% of patients in cohort A and 81% of patients in cohort B. The majority of manufacturing failures resulted from an insufficient number of tumor cells.

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The most common vaccine-related adverse events were local vaccine injection site reactions (93%), followed by fatigue (16%), nausea (12%), and pain, arthralgia, and upper respiratory infection (each at 5%). Two grade 4 (pericardial effusion) and six grade 3 (dyspnea, fatigue, injection site reaction, hypokalemia, malignant ascites, and pulmonary embolism) possibly related events were reported. There was no association between vaccine dose and the total number of adverse events or grade 3 and 4 adverse events. Vaccine injection site reaction size (skin induration) was positively associated with level granulocyte macrophage colonystimulating factor (GM-CSF) secretion from the transfected autologous malignant cells used as the product. Analysis of vaccine site biopsy specimen showed dense infiltration with CD4+ and CD8+ T cells, CD1a+ dendritic cells, and eosinophils. Three patients in cohort B achieved durable, complete tumor regressions. Two of these three patients remain in continuous complete remission now longer than 6 years. The other patient had tumor recurrence after 2 years, responded temporarily to a second GVAX® treatment, but recurred and passed away nearly 3 years after initial treatment. In addition, there was one minor response (30% decrease in a lung nodule) and two mixed responses; seven patients had stable disease with a mean duration of 7.7 months. Correlation of dose to survival was demonstrated to be significant at a threshold of 40 ng GM-CSF/106 cells/24 h expressed from an aliquot of the vaccine before the first injection. Long-term follow-up of two of the patients (Stage IV refractory disease to previous cytotoxic therapy) achieving complete response reveals continued disease-free survival now more than 8 years after initial GVAX® vaccination (85). In an effort to remove the requirement for genetic transduction of individual tumors and to optimize GM-CSF transgene expression (given that this correlated with improved survival), a second approach was developed called “bystander” GVAX®, which is a vaccine composed of autologous tumor cells mixed with an allogeneic GM-CSF-secreting cell line (K562 cells) (86). A phase I/II trial of this vaccine in advanced stage NSCLC was conducted. Tumors were harvested from 86 patients, tumor cell processing was successful in 76 patients, and 49 patients proceeded to vaccination. Serum GM-CSF pharmacokinetics were consistent with secretion of GM-CSF from vaccine cells for 4 days, with associated transient leukocytosis confirming the bioactivity of vaccine-secreted GM-CSF. Evidence of vaccine-induced immune activation was demonstrated. However, objective tumor responses were not seen despite a 25-fold higher GM-CSF secretion concentration with the bystander GVAX® vaccine (Tables 1 and 2). The frequency of vaccine site reactions, tumor response, time to progression, and survival were all less favorable to autologous GVAX®, although results were similar to historical cytotoxic


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therapy for second-line NSCLC. Overall, these results suggest that autologous malignant tissue transfection with adenovirusdelivered GM-CSF is optimal over the bystander approach, despite variability of GM-CSF expression levels and practical limitations inherent to surgically harvested tumor tissue. 4.3. L523S Gene Vaccine

L523S is a lung cancer antigen originally identified through screening of genes differentially expressed in cancer cells versus normal tissue (87, 88). L523S is expressed in approximately 80% of NSCLC cell lines (87, 88). In preclinical studies, the immunogenicity of L523S in humans was initially shown by detecting the presence of existent antibody and CD4+ T cell responses to L523S in patients with lung cancer. Subsequent studies further validated the immunogenicity of L523S, demonstrating that human CTLs could specifically recognize and kill cells that express L523S. It has demonstrated preclinical safety when the gene is injected intramuscularly as an expressive plasmid (pVAX/L523S) and when delivered by E1B-deleted adenovirus (Ad/L523S). A phase I clinical trial of 13 Stage IB, IIA, and IIB NSCLC patients was conducted using a prime/boost vaccination strategy first with pVAX/ L523S at a dose of 8 mg on days 0 and 14 then Ad/ L523S at three dosing cohorts of 1, 20, and 400 × 109 viral particles on days 28 and 56 (Tables 1 and 2) (89). No significant toxic effects related to the vaccination were reported. Although all but one patient demonstrated at least a twofold increase in anti-adenovirus antibodies, only one patient demonstrated a significant immune response to L523S. The reasons for the minimal detection of immune response are unknown, but suggest that alternate formulations and/or regimens need to be considered in addition to other surrogate immune function parameters. Two patients developed disease recurrence and all patients were alive after the 290-day follow-up. The significance of the disease-free survival cannot be assessed because of the small sample size; however, one cannot exclude the possibility that the vaccine may induce a T cell response that is below the threshold of detection in peripheral blood. The results of this trial suggest an excellent safety profile, but limited evidence of L523S-directed immune activation.

4.4. B7.1. Gene Vaccine

B7.1 (CD80+ ) is a co-stimulator molecule associated with induction of a T and natural killer (NK) cell response (90–93). Tumor cells transfected with B7.1 and HLA molecules have been shown to stimulate an avid immune response by direct antigen presentation and direct activation of T cells, in addition to allowing crosspresentation (94–96). In a phase I trial, Raez et al. (97) used an allogenic NSCLC tumor cell line (AD100) transfected with B7.1 and HLA-A1 or HLA-A2 to generate CD8+ CTL responses. Patients who were HLA-A1 or HLA-A2 allotype received the

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corresponding HLA-matched vaccine. A total of 19 patients with Stage IIIb or IV NSCLC were treated and most had received previous chemotherapy. Patients who were neither HLA-A1 nor HLA-A2 received the HLA-A1 transfected vaccine. Each patient received three intradermal vaccinations of 5 × 107 cells every 2 weeks. If the disease remained stable and toxicity was low, treatment was continued. A total of 18 patients received at least one full course (three vaccinations) of treatment. One patient was removed before the completion of the first course because of a serious adverse event not associated with the vaccine. Three more patients experienced serious adverse events, which were also not associated with the vaccine. Side effects associated with the vaccine included minimal skin erythema in four of the patients (Tables 1 and 2). All but one patient had a measurable CD8+ response after three vaccinations. There was no statistically significant difference in CD8+ response dependent on whether or not the patients were HLA matched. One patient showed a partial response for 13 months and five patients had stable disease ranging from 1.6 to >52 months (97, 98). Based on the six surviving patients, the tumor vaccine appears to elevate immune response for at least 150 weeks. Overall, the Kaplan–Meier estimate for the survival of 19 patients was 18 months. One-year survival was estimated at 52% (Tables 1 and 2). The low toxicity and good survival in this study suggested benefit from clinical vaccination. Further clinical investigation is ongoing. 4.5. a(1,3)-Galactosyltransferase

α(1,3)-Galactosyltransferase (aGal) epitopes present on the surface of most nonhuman mammalian cells are the primary antigen source inductive of hyperactive xenograft rejection. aGal directs the addition of aGal to N-acetyl glucosamine residues in humans. Expression of aGal epitopes after gene transfer of aGal (using retroviral vector) in human A375 melanoma cells prevented tumor formation in nude mice (99). Preliminary results by Morris et al. (100), using three irradiated lung cancer cell lines genetically altered to express xenotransplantation antigens by retroviral transfer of the murine aGal gene, were recently described in seven patients with Stage IV, recurrent or refractory NSCLC. Intradermal injections were given at doses of 3 × 106 cells/vaccine, 10 × 106 cells/vaccine, 30 × 106 cells/ vaccine, or 100 × 106 cells/vaccine once every 4 weeks, spanning a total of four doses. Only four patients received all four vaccinations, two patients received three vaccinations, and one patient received two vaccinations at the time of publication of the abstract. Toxicity involved grade 1 and 2 pain at the injection site, local skin reactions, fatigue, and hypertension. Four patients had stable disease for >16 months (Tables 1 and 2). Morris et al.


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concluded that the aGal vaccine was feasible and safe. Full analysis is awaiting completion of this trial.

5. Conclusion The poor overall survival of patients with lung cancer combined with mounting evidence of multistep carcinogenesis (cancer is the result of multiple molecular defects) (101–103), which may, in part, provide a mechanism for malignant tissue’s remarkable ability to evade many treatment modalities, mandates novel approaches to clinical management of lung cancer. Traditional cytotoxic approaches for management of advanced stage lung cancer have likely reached a plateau with respect to survival and response. Recent data on combinations of cytotoxic therapies with angiogenesis inhibitors and/or epidermal growth factor receptor (EGFR) inhibitors appear encouraging in subsets of patients. Results summarized in this chapter suggest that immune-based therapies may also soon provide sufficient validation to be considered as part of the therapeutic armamentarium for lung cancer. Yet more remains to be discovered. Uncloaking the mechanisms of molecular-based and immune-based therapeutics as we study their effect in cancer patients may one day enable nontoxic synergistic combinations of such therapies, potentially reducing our dependence on cytotoxic approaches, with their associated adverse effects. Molecular- and immune-based therapies operate by mechanisms that may not be immediately evident to investigators. For example, the autologous GVAX® lung vaccine, a patient-specific vaccine taking advantage of paracrine secretion of GM-CSF and patient-specific tumor cells, in one trial reported three complete responses. This strategy showed more favorable results than the allogeneic GVAX® bystander vaccine in similar patient populations. Comparison of these two relatively similar vaccines reveals countless logistic and mechanistic variables, some unknown, which contribute to the many hypotheses regarding why GVAX® bystander was less effective and serves to illustrate the challenges facing researchers in evaluating these therapies. However, it is clear that the opportunity to activate the immune system is a real possibility; although the method and mechanism by which this can be optimally achieved is unknown. Through enhancement of tumor antigen recognition and immune activation, these vaccines may one day provide patients with a highly tolerable therapy to use in combination with traditional and other novel approaches.

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Gene Immunotherapy for Non-Small Cell Lung Cancer 77. Fakhrai H, Dorigo O, Shawler DL, et al. Eradication of established intracranial rat gliomas by transforming growth factor beta antisense gene therapy. Proc Natl Acad Sci USA 1996;93(7):2909–14. 78. Dorigo O, Shawler DL, Royston I, Sobol RE, Berek JS, Fakhrai H. Combination of transforming growth factor beta antisense and interleukin-2 gene therapy in the murine ovarian teratoma model. Gynecol Oncol 1998;71(2):204–10. 79. Tzai TS, Shiau AL, Liu LL, Wu CL. Immunization with TGF-beta antisense oligonucleotide-modified autologous tumor vaccine enhances the antitumor immunity of MBT-2 tumor-bearing mice through upregulation of MHC class I and Fas expressions. Anticancer Res 2000;20(3A):1557–62. 80. Tzai TS, Lin CI, Shiau AL, Wu CL. Antisense oligonucleotide specific for transforming growth factor-beta 1 inhibit both in vitro and in vivo growth of MBT-2 murine bladder cancer. Anticancer Res 1998; 18(3A):1585–9. 81. Marzo AL, Fitzpatrick DR, Robinson BW, Scott B. Antisense oligonucleotides specific for transforming growth factor beta2 inhibit the growth of malignant mesothelioma both in vitro and in vivo. Cancer research 1997;57(15):3200–7. 82. Park JA, Wang E, Kurt RA, Schluter SF, Hersh EM, Akporiaye ET. Expression of an antisense transforming growth factor-beta1 transgene reduces tumorigenicity of EMT6 mammary tumor cells. Cancer Gene Therapy 1997;4(1):42–50. 83. Hege KM, Carbone DP. Lung cancer vaccines and gene therapy. Lung Cancer 2003;41(Suppl 1):S103–13. 84. Nemunaitis J, Sterman D, Jablons D, et al. Granulocyte-macrophage colony-stimulating factor gene-modified autologous tumor vaccines in non-small-cell lung cancer. J Natl Cancer Inst 2004;96(4):326–31. 85. Nemunaitis J, Vorhies JS, Pappen B, Senzer N. 10-year follow-up of gene-modified adenoviral-based therapy in 146 non-small-cell lung cancer patients. Cancer Gene Therapy 2007;14(8):762–3. 86. Nemunaitis J, Jahan T, Ross H, et al. Phase 1/2 trial of autologous tumor mixed with an allogeneic GVAX vaccine in advanced-stage non-small-cell lung cancer. Cancer Gene Therapy 2006;13(6):555–62. 87. Mueller-Pillasch F, Pohl B, Wilda M, et al. Expression of the highly conserved RNA binding protein KOC in embryogenesis. Mech Dev 1999;88(1):95–9.

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101. Ten Have-Opbroek AA, Benfield JR, Hammond WG, Teplitz RL, Dijkman JH. In favour of an oncofoetal concept of bronchogenic carcinoma development. Histol Histopathol 1994;9(2):375–84. 102. Toloza EM, Roth JA, Swisher SG. Molecular events in bronchogenic carcinoma and their implications for therapy. Semin Surg Oncol 2000;18(2):91–9. 103. Ihde DC, Minna JD. Non-small cell lung cancer. Part I: Biology, diagnosis, and staging. Curr Probl Cancer 1991;15(2):61–104. 104. Salgia R, Lynch T, Skarin A, et al. Vaccination with irradiated autologous tumor cells engineered to secrete granulocyte-macrophage colony-stimulating factor augments antitumor immunity in some patients with metastatic non-small-cell lung carcinoma J Clin Oncol 2003;21(4):624–30.

Chapter 27 Gene Therapy for Antitumor Vaccination Seunghee Kim-Schulze and Howard L. Kaufman Summary Tumor immunotherapy depends on the interactions between the host, the tumor, and the immune system. Recent data suggests that priming of antigen-specific T cells alone may not be adequate for mediating regression of established tumors because of the immune inhibitory influences within the tumor microenvironment. Thus, we developed a recombinant vaccinia virus vector to express single or multiple T cell costimulatory molecules as a vector for local gene therapy in patients with malignant melanoma. This approach is feasible and generated local and systemic tumor immunity and induced objective clinical responses in patients with metastatic disease. This chapter reviews the details and major issues related to using live, replicating, recombinant poxviruses for gene delivery and antitumor vaccination within the tumor microenvironment. Key words: Antitumor vaccination, mmunotherapy, melanoma, tumor antigen, vaccinia virus.

1. Introduction The successful implementation of immunotherapy for cancer depends on the complex interaction between host, tumor, and immune system. These elements intersect within the tumor microenvironment and recently there has been significant emphasis placed on understanding how the immune system mediates tumor recognition and rejection within the milieu of an established tumor. The importance of the local tumor microenvironment has been nicely demonstrated in patients with melanoma in whom functional antigen-specific T cells can frequently be detected in peripheral blood but not within established tumors or regional tumor-draining lymph nodes (1). This has led to speculation that the microenvironment of successfully proliferating


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tumors can actively suppress effective tumor immunity. In fact, there are now several well characterized molecular and cellular pathways that are known to mediate local tumor immunosuppression and these likely represent major obstacles to successful tumor immunotherapy (2). Thus, the therapeutic effectiveness of tumor immunotherapy will depend on not only activating innate and adaptive antigen-specific immune responses but will also require overcoming the local suppressive factors within the tumor microenvironment. The identification of tumor-associated antigens suggested that tumors could be recognized by the host immune system in a highly specific manner (3). To prime antigen-specific T cells, we developed a recombinant poxvirus platform wherein live, replicating viruses were engineered to express specific tumor antigens (4). These vectors were initially administered systemically through subcutaneous or intramuscular injections and demonstrated an excellent safety profile but few, if any, clinical responses were seen (5). To improve the potency of the vaccines, they were reengineered to coexpress tumor antigens and T cell costimulatory molecules based on the two-signal hypothesis of T cell activation (6). This hypothesis states that activation of T cells requires recognition of antigen and stimulation by a set of surface molecules collectively termed “costimulatory.” The prototypical example is B7.1 (CD80), which signals via T cell CD28 to induce T cell proliferation, survival, and cytokine release. The addition of B7.1 to recombinant poxviruses did indeed improve therapeutic activity of the vaccines in murine models and resulted in enhanced T cell activation in early clinical trials (7, 8). Subsequently, expression of multiple costimulatory molecules in the recombinant poxvirus vectors demonstrated incremental improvements in T cell activation in vitro, in vivo, and in preclinical therapeutic tumor models (9–11). In studies pioneered by Jeffrey Schlom and his colleagues, the inclusion of B7.1, ICAM-1 (CD54), and LFA-3 (CD58) were found to be superior to B7.1 alone for T cell activation, and this triad of costimulatory molecules was termed TRICOM (12). Although the TRICOM vectors have shown significant promise in early phase clinical trials, the functional status of vaccine-primed T cells within the tumor microenvironment remained largely unexplored (11, 13). Thus, we hypothesized that activated T cells might not be able to mediate complete tumor regression because of the inhibitory influences of the tumor microenvironment. This possibility was supported by data suggesting that tumors with low levels of B7.1 expression could block rejection of established tumors in murine models of colorectal cancer (14). To replace local costimulatory molecules and provide a more favorable inflammatory cytokine profile within established tumors, we sought to use a recombinant vaccinia virus expressing B7.1 as a direct intratumoral delivery vector. Because melanoma is often associated with

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tumor-infiltrating lymphocytes and many lesions are often accessible for injection, we focused these studies on patients with metastatic melanoma (15). The initial Phase I clinical trial resulted in objective regression in 3 of 12 patients, with one patient achieving an objective complete response that is ongoing for longer than 65 months and was associated with the appearance of autoimmune vitiligo. An additional four patients had stable disease after local vaccination. Based on these initial results, we sought to evaluate the effects of a recombinant vaccinia virus expressing the TRICOM combination in a similar patient population. A Phase I clinical trial found that 30.7% of patients had an objective clinical response, with one patient achieving a complete response for longer than 22 months (16). This chapter reviews the major issues related to gene therapy using live, replicating, recombinant poxviral vectors in patients with cancer (see Note 9).

2. Materials 2.1. Vaccine Formulation

1. The recombinant vaccinia virus expressing B7.1 (rV-B7.1) was generated from the Wyeth vaccinia virus (New York City Board of Health) strain by homologous recombination. The B7.1 gene was derived from a human complementary DNA (cDNA) library and was inserted into a recombinant vaccinia DNA plasmid before in vitro recombination (15). Recombinant virus was selected through plaque purification, and high titers were generated by subsequent tissue culture using chick embryo dermal (CED) cells. The rV-TRICOM vector was generated in a similar fashion to rV-B7.1, except that the recombinant plasmid was generated to include the genes encoding human B7.1, ICAM-1, and LFA-3. 2. Gene expression was confirmed by infecting a monolayer of CV1 cells with various multiplicities of infection (MOI) of each virus (range, 10–1,000) for 24 h. Cells were then collected and subjected to flow cytometry with anti-B7.1, anti-ICAM-1, and anti-LFA-3 monoclonal antibodies (and IgG isotype-matched controls). 3. Sterility and quality control were performed as previously described (17, 18). 4. Vaccines were vialed with the maximum concentration of virus per vial. For rV-B7.1, each vial contained 2.13 × 109 plaque-forming units (pfu) per milliliter in a total volume of 300 μL; for rV-TRICOM, there was 2.54 × 109 pfu/mL in a total volume of 200 μL.

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2.2. Vaccine Storage and Preparation

1. All vaccines were stored in a locked 70°C freezer in a Biosafety Level 2 laboratory approved for virus storage and preparation (see Note 1). 2. Strict drug accountability records were maintained during vaccine storage, preparation, and disposal. 3. When patients were vaccinated, individual vials were thawed completely at room temperature. The thawed contents were gently vortexed for at least 10 s before dose preparation. 4. The vials were swabbed with alcohol before withdrawing the necessary amount of vaccine or diluent. Vials allowed easy recovery of 0.2 mL (4.26 × 108 pfu rV-B7.1 and 5.1 × 108 pfu rV-TRICOM). Lower doses were achieved by tenfold serial dilutions in 0.9% sodium chloride solution. 5. The vaccine was prepared by removing 0.2 mL from the vial (or after dilution for lower doses) with a 1-cc tuberculin syringe and needle under sterile conditions in a biohazard safety cabinet. 6. The tuberculin syringe containing 0.2 mL total volume was used to directly inject the index lesion (after alcohol wiping) as close to the center as possible and injecting outwards in a circumferential manner (see Notes 2 and 3).

3. Methods 3.1. Clinical Protocol

1. The clinical protocols were written and approved by the Cancer Therapy Evaluation Program (CTEP) of the National Cancer Institute (NCI), Office of Biotechnology Activities (OBA) of the National Institutes of Health, local institutional review board (IRB), and biosafety committee (IBC) (see Note 4). 2. The primary endpoints of the study were to: (a) Determine the maximum tolerated dose (MTD) of vaccine that elicits a host immune response and is associated with acceptable toxicity in patients with metastatic melanoma. (b) Evaluate the safety and clinical toxicity associated with intralesional vaccine administration. 3. The secondary endpoints of the study were to: (a) Establish evidence of host antimelanoma immune reactivity after intralesional vaccine administration. (b) Evaluate the effect of vaccination on T cell immunity. (c) Evaluate the clinical response to intralesional vaccine administration. (d) Evaluate quality of life during treatment.


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Based on the replicating nature of the vaccinia virus, the use of recombinant DNA, and the limited experience with direct intratumoral injections at the time these studies were initiated, patient selection was strict and all patients were required to meet these eligibility criteria (see Note 5). 1. Age 18 years or older. 2. Presence of histologically proven metastatic, nonresectable cutaneous, subcutaneous, or lymph node melanoma accessible for percutaneous injection. 3. Lesions for injection must be measurable and measure at least 1.0 cm. 4. Patients must have a good performance status defined as ECOG 0–1 or Karnofsky 80–90%. 5. Women of childbearing potential must have a negative serum β-HCG and were instructed in the use of and agreed to use adequate contraception. 6. Patients with a history of surgery for management of primary or metastatic lesions were eligible to enter the study if longer than 4 weeks had elapsed from the time of surgery, the patient was fully recovered, and there was measurable disease still present. Patients with metastatic brain lesions, leptomeningeal disease, or seizure disorders were ineligible. Patients with a history of successfully treated brain metastases were eligible provided they were at least 6 months from definitive therapy (surgery or radiation) and had no evidence of disease or edema on brain imaging. 7. Patients with a history of chemotherapy were eligible for inclusion if the following criteria were met: (a) No more than two previous chemotherapy regimens were administered. (b) More than 4 weeks had elapsed since the last cycle was administered. (c) The patient had fully recovered from any toxicity associated with previous chemotherapy. 8. Patients with a history of radiation therapy were eligible for inclusion, if the following criteria were met: (a) More than 2 weeks had elapsed since receiving radiation. (b) The patient was fully recovered from any toxicity associated with previous radiation therapy. (c) The patient had no evidence of bone marrow toxicity associated with previous radiation treatment. 9. Patients with a history of immunotherapy were eligible for inclusion, provided that:

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(a) More than 8 weeks had elapsed since previous therapy. (b) No toxicity related to previous therapy was evident. (c) Previous therapy was not with a live poxvirus vector. 10. Patient must have had a life expectancy longer than 3 months at the start of the trial. 11. Patients must have had adequate organ function as defined by: (a) White blood cell count greater than 4,000 cells/mm3, platelet count greater than 100,000 cells/mm3, and hemoglobin level greater than 10 g/100 mL. (b) Serum creatinine less than 2.0 mg/dL or creatinine clearance greater than 60 mL/min. (c) Direct bilirubin less than 1.5 mg/dL. (d) No evidence of congestive heart failure, serious cardiac arrhythmia, recent myocardial infarction, or signs of coronary artery disease. (e) No other clinical or laboratory data suggestive of an underlying immunosuppressive disorder. 12. Patients were required to have proof of previous smallpox immunization with vaccinia virus as evidenced by: (a) Physician certification of previous vaccinia immunization or (b) Patient recollection and appropriate vaccination-site scar 13. There must have been no history of eczema or other contraindications to vaccinia virus administration. Patients must also have been able to avoid other household contacts that had active or a history of eczema. 14. Patients were ineligible if they required systemic corticosteroids less than 4 weeks before registration or were deemed likely to require corticosteroid use for intercurrent illness during the study period. 15. Patients with significant allergy or hypersensitivity to eggs were excluded because the vaccine was prepared in embryonic chicken cells. 16. Patients must not have had any active or chronic infections or concurrent medical illnesses that could interfere with the ability to receive vaccination with a live virus. 17. There must have been no evidence of encephalitis, cerebral metastasis, or other structural brain lesions by clinical or radiological evaluations. 18. Patients with any significant medical disease other than the malignancy (e.g., chronic obstructive pulmonary disease [COPD], ascites, or pleural effusions) that, in the opinion of the investigator, would significantly increase the risk for immunotherapy were not eligible.


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19. Patients with a history of another malignancy within the last 5 years, other than stage I carcinoma of the cervix or basal cell carcinoma, were not eligible. 20. Patients must have been able and willing to sign an informed consent form indicating that they understood the nature of their disease, treatment options, the experimental nature of the study, the risks, the possible side effects, any discomforts, and the potential benefit of entering the study as outlined in the protocols. 3.3. Patient Treatment

1. Initial concerns about the possibility of inducing significant autoimmunity prompted a study design using two doses for rV-B7.1 and three doses for rV-TRICOM escalations. Thus, patients were sequentially enrolled at lower doses of vaccine (each dose was adjusted tenfold) and, provided no serious adverse events were seen at the lower doses, subsequent patients could receive higher doses of vaccine (17, 18). 2. Vaccine was administered directly into an established melanoma in a cutaneous, subcutaneous, or nodal location using the appropriate dose in a tuberculin syringe at 4-week intervals for a total of three vaccinations. The injected sites were covered with a dry gauze and Tegaderm dressing. 3. Patients were required to have a baseline computed tomography (CT) scan of the chest, abdomen, and pelvis and a magnetic resonance imaging (MRI) scan of the brain within 4 weeks of receiving the first vaccination. A re-staging set of CT scans was obtained within 4 weeks of completing the third vaccination for comparison. Patients without evidence of objective disease progression could receive an additional three vaccinations at 4-week intervals. 4. Before each vaccination, a fine needle aspiration was obtained of the injected lesion for cytology and analysis of the tumor microenvironment (see below). Peripheral blood samples were also obtained at baseline and 2 weeks after each injection for immune analysis (see below). Blood was also collected every 4 weeks for routine blood counts, chemistry, and liver function analysis to monitor toxicity. 5. Patients were given a standardized quality of life questionnaire to complete before the first vaccination and at each subsequent clinic visit (17, 18).

3.4. Immune Monitoring

A major goal of these trials was to determine the effect of local gene delivery into established tumors on local and systemic immunity. Thus, a series of correlative laboratory studies were conducted in association with the clinical trial (see Note 8).

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1. Tissue samples were obtained by fine needle aspiration before each vaccine injection and these were used for cytologic analysis of melanoma cells and amplification of RNA for quantitative real-time polymerase chain reaction (PCR) analysis. 2. Cytology was performed by a certified cytopathologist on site after cytospin preparation and acetone fixation onto glass slides. The slides were stained with hematoxylin and eosin, anti-MART-1 murine IgG2b (M2-7C10) and anti-Pmel17/ gp100 mAB (HMB45). Secondary staining was performed with biotinylated goat antimouse IgG and developed with avidin–biotin–peroxidase, as described elsewhere (15). 3. The HLA genotype of individual patients was determined by PCR with sequence-specific primers (SSP) using commercially available kits (One Lambda, Los Angeles, CA). 4. Enzyme-linked immunosorbent assay (ELISA) was used to detect antivaccinia antibodies before treatment and after each vaccination. Briefly, 96-well plates were coated with lysate of the wild-type vaccinia virus. Serial serum dilutions from patients were applied to the coated wells and incubated overnight. The wells were then washed and then incubated with an antihuman immunoglobulin conjugated to horseradish peroxidase, and developed using a horseradish peroxidasespecific substrate. A standard control curve was established with serum containing known antibody concentrations. The plates were read at an optical density of 450 nm. 5. An interferon (IFN)-γ enzyme-linked immuno spot (ELISPOT) assay was performed to detect gp100-, tyrosinase-, and MART1-specific T cell responses before and after vaccination in HLAA2+ patients where defined HLA-A2-restricted peptide epitopes were available. This assay was performed using peripheral blood mononuclear cells (PBMC) collected before and 2 weeks after each vaccination. The PBMC were Ficoll separated and frozen in human AB serum and 10% DMSO until testing. On the day of assay, PBMC were thawed and resuspended in complete T cell medium (10% AB serum) and incubated overnight. The following day, cells were cocultured with antigen-presenting cells (T2-A2) in the presence of the HLA A2-restricted melanoma peptides (Melan-A/Mart-131–39, gp10013–21, and tyrosinase487–495 at each peptide concentration of 2 μg/mL) for 24 h on nitrocellulose ELISPOT (Millipore) plates coated with the capture anti-IFN-γ monoclonal antibody (mAb) (1 μg/mL). After incubation, cells were lysed and washed. Detection antibody was added, and the spots were developed and counted by a blinded observer using an automated ELISPOT counter. The number of spots per plate correlated with the frequency of T cells responding to the specific melanoma antigens. Specificity was confirmed by adding anti-major histocompatability complex (MHC) class


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I or class II mAbs during the assay to show the inhibition of IFN-γ production. 6. T cells were further characterized for cell surface marker expression using flow cytometry. T cell phenotyping was performed to evaluate the ratio of helper T cells, cytotoxic T cells, regulatory T cells, and natural killer (NK) cells in a 100% cell population defined by CD45 bright-CD14– lymphocytes. The activation state of T cells was also tested by FACS analysis (HLA-DR, CD25, and CD69). Human CD4+ and CD8+ T cell subsets were defined by staining with phycoerythrin (PE)-conjugated mAbs to CD4 and Cy-Chrome (CYC)-conjugated mAbs to CD8 in the CD3+ lymphocyte population. To detect cytotoxic T cells, the level of granzyme B and perforin expression in combination with CD28 was documented by staining CD8+ T cells. NK cells were defined as double positive for FITC–CD16 and PE-CY5–CD56. NK cell cytotoxicity was determined by the level of expression of granzyme B and perforin. Regulatory CD4+ T cells were detected by cell surface staining with FITC–CD4 and PE–CD25, and intracellular APC–FOXP3 staining. Regulatory CD8+ T cells were detected by staining with cell surface FITC–CD28 and PE–Cy5-CD8, and intracellular APC–Foxp3 staining. Dendritic cells were characterized by staining with the following mAbs: PE–CD40, PE–CD54, PE–CD62E, PE–CD83, PE–DC-SIGN, CYC–CD11c, CYC– HLA ABC, and CYC–HLA DR and ILT3–PC5. Rat IgG was added as a blocking agent before staining with antibodies. For each cell surface or intracellular marker, a corresponding isotype-matched control antibody conjugated with the same fluorescent dye was used. Six-parameter analyses (forward scatter, side scatter, and four fluorescence channels) was used for list mode data analysis. 7. Quantitative real-time PCR analysis was performed using fine needle aspirate samples obtained before each monthly vaccine injection (see Notes 6 and 7). This approach allowed the ongoing monitoring of the local microenvironment during treatment with minimal disruption of the vaccination site and could be done with miniscule amounts of material, provided RNA was amplified before PCR assay. Because these initial studies were designed for feasibility, we selected only a few gene transcripts for analysis in the initial clinical trials, focusing on interferon-γ, CD8, and interleukin (IL)10, based on previous studies suggesting that alterations in local IFN-γ and IL-10 correlated with clinical responses in melanoma patients treated with high-dose IL-2 (19). Briefly, total RNA was isolated from tissue samples using RNeasy mini kits (Qiagen, Santa Clarita, CA). RNA was eluted with water and stored at 70°C. For cDNA synthesis, about 1 μg of total

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RNA was transcribed with cDNA Transcription Reagents (Perkin-Elmer, Foster City, CA) using random hexamers. cDNA was then stored at 30°C until quantitative RT-PCR could be performed. Quantitation of gene expression was performed using the ABI prism 7700 Sequence Detection System as previously described (20). Primers and TaqMan probes (Custom Oligonucleotide Factory, Foster City, CA) were designed to span exon–intron junctions to prevent amplification of genomic DNA and resulted in amplicons 2.5 × 106 cfu/mL. 3. MCB and WCB vials were labeled with name of product, clone, number of cells, vector titer, and date. 4. RVPCs of the MCB and WCB were grown at 37°C with 5% CO2 in DMEM supplemented with 10% (v/v) heat-inactivated FBS, penicillin (P; 100 U/mL, Sigma-Aldrich, St. Louis, MO, USA) and streptomycin (S; 100 µg/mL, Sigma-Aldrich), and the antibiotic G418 (800 µg/mL, Sigma-Aldrich) for selection of transduced cells. All antibiotics were resuspended in water at the desired concentration, filter-sterilized (0.22-µm filters, Millipore, Bedford, MA), and stored in aliquots at −20°C. 5. All materials in contact with cells were sterile, pyrogen-free, and were stored and used according to the manufacturer’s directions. 6. Production of the MCB and WCB was carried out in a biosafety level 3 GMP facility according to the European Community (EC) Guide to GMP. MCB and WCB cells underwent the following sterility and identity testing: (a) Viability: Trypan blue exclusion dye (Sigma-Aldrich) was used to determine cell viability. Briefly, the cell suspension was diluted 1:2 with trypan blue (0.4% w/v), and approximately 20 µL of cells were loaded into a hemocytometer chamber. Living cells exclude the dye, whereas dead cells take up the blue dye. The blue stain is easily visible, and cells can be counted using a light microscope. (b) NeoR transgene activity: To determine neomycin resistance gene (NeoR) activity, cells were grown in medium supplemented with G418 at 800 µg/mL final concentration. The synthetic antibiotic G418, which is an effective inhibitor of protein synthesis in both eukaryotic and prokaryotic cells, is used as a selective agent for transformants that express the bacterial NeoR gene. Using NeoR as a positive selectable marker, selection and isolation of recombinants containing the transgene is facilitated. This system is particularly powerful because no equivalent enzymatic activity exists in eukaryotic cells and therefore there is no background of spontaneous G418-resistant mutants. (c) HSV-TK activity: To assess HSV-TK activity, transduced cells were seeded in triplicate at a concentration of 5 × 103 cells/well in 96-well flat bottom microplates and treated with serial dilutions of GCV, ranging from 0.01–100 µM. After incubation for 5 days, GCV cytotoxicity was measured by the MTT Cell Proliferation Kit I colorimetric

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assay (Roche Applied Science, Monza, Italy), following the manufacturer’s protocol. This assay allows calculation of the IC50, which is the concentration of drug that inhibits cell growth by 50%, by plotting the percentage of surviving cells (ordinate) versus log10 GCV concentration (abscissa). (d) hIL-2 activity: hIL-2 activity was measured by a bioassay using the CTLL cell line, as reported (13). Briefly, the CTLL mouse cell line (kindly provided by Dr. K Smith, New York Hospital, Cornell Medical Center, NY, USA) was used as an indicator cell system for determination of IL-2 activity in supernatants of RVPCs. CTLL cells are known to proliferate in response to hIL-2. IL-2 activity was assessed by 3H-thymidine uptake by CTLL after a 6 h pulse with 0.5 mCi at the end of a 24 h incubation, with serial 1:2 dilutions of supernatants from the RVPCs. Serial dilutions of human recombinant IL-2 (Eurocetus, Amsterdam, The Netherlands), containing a known amount of international units (IU) were used as standards. (e) Sterility testing: Bacterial and fungal contamination was tested by bacterial and fungal cultures, gram staining, and Hoechst staining and polymerase chain reaction (PCR) to detect mycoplasma contamination. (i) Bacterial and fungal cultures were performed by inoculating BACTEC™ bottles (BD Diagnostic Systems, Cockeysville, MD, USA) with 2 mL supernatant medium and incubation for at least 7 days. (ii) Gram staining on cell culture samples was performed to evaluate bacterial contamination. Traditional Gram staining was carried out with the three-step Gram stain procedure kit (BD Microbiology Systems), in strict accordance with the manufacturers instructions. (iii) Detection of mycoplasma contamination in cell cultures was performed by the Hoechst DNA staining method. After treatment with Hoechst dye (Hoechst 33258 stain solution; prod. no. H6024, Sigma-Aldrich), cells exhibit fluorescent nuclei against a dark background under observation at ×1,000 magnification using epifluorescence. Cell cultures infected with mycoplasma exhibit both fluorescent nuclei and extranuclear fluorescence attributable to mycoplasma DNA. (iv) PCR analysis of mycoplasma sequences was also used to detect mycoplasma contamination in cell cultures. The PCR method is considered the method of choice for its high sensitivity in the detection of mycoplasma contamination in cell cultures and other cell-culture

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derived biologicals. Mycoplasma PCR analysis was performed with the LookOutTM Mycoplasma PCR Detection Kit (Code MP0035, Sigma-Aldrich) according to the manufacturer’s instructions. The primer set and probe are specific to a highly conserved 16S ribosomal RNA (rRNA) coding region in the mycoplasma genome. This allows for detection of most Mycoplasma species tested and usually encountered as contaminants in cell cultures. (f) Screening for adventitious viral infections: (i) Detection of porcine viruses (Porcine parvovirus [PPV]) in trypsin and cell products was performed by inoculation of test articles onto monolayers of PPK cells, a primary porcine kidney cells line that is permissive for porcine viruses. PPK cells are monitored for 21 days after inoculation and positive results are confirmed with specific assays of hemadsorption and immunofluorescence at the end of the culture period. (ii) Detection of bovine viruses (bovine viral diarrhea virus, bovine parvovirus, blue tongue virus, bovine adenovirus, bovine respiratory syncytial virus, reovirus, and rabies virus) in bovine serum and cell products by inoculation of test articles onto a simian kidney cell line (Vero) and a bovine cell line (bovine turbinate [BT] cells), which allow the detection of a wide range of viruses. The detector cells are monitored for 21 days after inoculation for the development of characteristic changes in morphology attributable to replication of viral agents. A blind passage is made on day 21 and the cells are isolated, lysates are prepared, and fresh Vero and BT cells are inoculated and observed for additional 14 days for changes in morphology. Positive results are confirmed by hemadsorption and immunofluorescence. (g) Screening for RCR in cells and supernatant: Screening for RCR in cells and supernatant was performed by the provirus mobilization assay, as detailed in the Materials (paragraph 3.3.) and Methods (paragraph 4.3.3) sections. (h) Endotoxin production: Endotoxin production was analyzed by the chromogenic limulus amebocyte lysate (LAL) assay (Cambrex Bio Science Walkersville, Inc., Walkersville, MD, USA), according to the protocol provided by the manufacturer. The LAL assay is a quantitative test for rapid, chromogenic quantitation of Gram-negative bacterial endotoxin. The assay uses the initial part of the LAL


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endotoxin reaction to activate an enzyme that, in turn, releases p-nitroaniline (pNA) from a synthetic substrate, producing a yellow color. Gram-negative bacterial endotoxin catalyzes the activation of the proenzyme in the LAL. The initial rate of activation is determined by the concentration of endotoxin present. The concentration of endotoxin in a sample is calculated from the absorbance values of solutions containing known amounts of endotoxin standard. 2.2.1. WCB Lot Release Criteria

Release criteria for WCB lots to be administered to patients include: 1. Cell viability >70%. 2. Negative for RCR. 3. Negative culture for bacteria and fungi after 7 days and negative Gram stain. 4. Endotoxin testing 5 EU/mL. 5. Negative result for mycoplasma by Hoechst staining and PCR. 6. Negative screening for adventitious viral infections.

2.3. Detection of ReplicationCompetent Retroviral Particles

1. NIH3T3-LacZ, a cell line harboring the LacZ-containing retroviral vector MFGnlsLacZ (kindly provided by Y. Takeuchi). 2. DMEM + 10% FBS + P/S culture medium. 3. 8 µg/mL Polybrene (Sigma-Aldrich) in water, filter-sterilized. 4. Phosphate-buffered saline (PBS): 120 mM NaCl, 25 mM Na–phosphate, pH 7.3. 5. 5-bromo-4-chloro-3-indolyl-β-D-Galactopyranoside (X-gal) staining reagent: (a) staining solution: 0.01% (w/v) sodium deoxycholic acid sodium salt, 0.02% NP40, 2 mM MgCl2 (Sigma-Aldrich), 5 mM potassium ferricyanide (Sigma-Aldrich), 5 mM potassium ferrocyanide (Sigma-Aldrich), in PBS; (b) 1 mg/mL X-gal (Sigma-Aldrich). The staining solution is light sensitive, and stable at room temperature. Aliquots of 40 mg/mL X-gal (in DMSO) are stored at −20°C; X-gal is added to the staining solution just before use. 6. Six-well tissue culture plates (flat-bottom wells).

2.4. PCR Detection of Vector Sequences in Biological Samples

1. QIAamp® DNA mini kit (Qiagen GmbH, Hilden, Germany). 2. Real-time PCR: TaqMan Master Mix 2X (Applied Biosystems, Foster City, CA, USA); ABI-PRISM 7900HT Real Time PCR System (Applied Biosystems). 3. Oligonucleotide primers (Sigma-Aldrich) and probes (Applied Biosystems) for real-time PCR detection of HSV-TK: forward

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primer, 5′ CCA ACG GCG ACC TGT ACA A 3′; reverse primer, 5′ CAT CCC GGA GGT AAG TTG CA 3′; TaqMan probe, FAM–5′ CTG GGC CTT GGA CGT CTT GGC C 3′–TAMRA. 4. Oligonucleotide primers (Sigma-Aldrich) and probes (Applied Biosystems) for real-time PCR detection of hIL-2: forward primer, 5′ CCA GGA TGC TCA CAT TTA AGT TTT AC 3′; reverse primer, 5′ GAG GTT TGA GTT CTT CTT CTA GAC ACT GA 3′; TaqMan probe, FAM–5′ TGC CCA AGA AGG CCA CAG AAC TGA A 3′–TAMRA. 5. PCR: AmpliTaq DNA polymerase 5 U/µL (Applied Biosystems), 10X PCR buffer (Applied Biosystems), 25 mM MgCl2 (Applied Biosystems), 10 mM dNTPs (Applied Biosystems), 15 µM sense and antisense primers. 6. Oligonucleotide primers (Sigma-Aldrich) for PCR detection of NeoR: forward primer, 5′ ACT GAA GCG GGA AGG GAC TGG 3′; reverse primer, 5′ AGA AGC CGA TAG AAG GCG ATG 3′. 7. Oligonucleotide primers (Sigma-Aldrich) for PCR detection of gag: forward primer, 5′ CTT CCT AGA GAG ACT TAA GG 3′; reverse primer, 5′ GTT GGG ACC TCC TTC GTT CTC 3′. 2.5. Real-Time RT-PCR Analysis of Therapeutic Gene Expression and Cytokine Gene Expression

1. RNeasy® Mini Kit (Qiagen GmbH) 2. DNaseI (DNA-freeTM, Ambion, Applied Biosystems). 3. pGEM®-T Easy Vector System vectors (Promega Corp., Madison, WI, USA). 4. Real-time reverse transcriptase PCR (RT-PCR): SuperScript TM II Reverse Transcriptase (Invitrogen Life Technologies); RNase OUT TM 40 U/μL (Invitrogen), 5X First Strand Buffer (Invitrogen Life Technologies), 25 mM MgCl2 (Invitrogen Life Technologies), 100 mM dNTPs (Invitrogen Life Technologies), 200 ng random examers (Invitrogen Life Technologies). 5. Oligonucleotide primers (Sigma-Aldrich) and probes (Applied Biosystems) for quantitative real-time RT-PCR analysis of the following transcripts: • Human GAPDH (housekeeping gene): forward primer, 5′ GAA GGT GAA GGT CGG AGT C 3′; reverse primer, 5′ GAA GAT GGT GAT GGG ATT TC 3′; TaqMan probe, FAM–3′ CAA GCT TCC CGT TCT CAG CC 5′–TAMRA. • HSV-TK (see Subheading 3.4). • hIL-2 (see Subheading 3.4). • Tumor necrosis factor (TNF)-α: forward primer, 5′ CCC AGG GAC CTC TCT CTA ATC 3′; reverse primer, 5′ ATG


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GGC TAC AGG CTT GTC ACT 3′; TaqMan probe, FAM– 5′ TGG CCC AGG CAG TCA GAT CAT C 3′–TAMRA; • Interferon (IFN)-γ: forward primer, 5′ AGC GGA TAA TGG AAC TCT TTT CTT AG 3′; reverse primers, 5′ AAG TTT GAA GTA AAA GGA GAC AAT TTG G 3′; TaqMan probe FAM–5′ TCT GTC ACT CTC CTC TTT CCA ATT CTT CAA AAT G 3′–TAMRA; • IL-1β: forward primer, 5′ ACA GAT GAA GTG CTC CTT CCA 3′; reverse primer, 5′ GTC GGA GAT TCG TAG CTG GAT 3′; TaqMan probe, FAM–5′ CTC TGC CCT CTG GAT GGC GG 3′–TAMRA; • IL-10: forward primer, 5′ ACG GCG CTG TCA TCG ATT 3′; reverse primer, 5′ TTG GAG CTT ATT AAA GGC ATT CTT C 3′; TaqMan probe, FAM–5′ CTT CCC TGT GAA AAC AAG AGC AAG GCC 3′–TAMRA. 2.6. HSV-TK Immunofluorescence Staining

1. 4% Paraformaldehyde in PBS for fixation. 2. 0.2% Triton X-100 in PBS for permeabilization. 3. PBS containing 5% bovine serum albumin (BSA) as blocking reagent. 4. Donkey serum (Sigma-Aldrich) to block nonspecific antibody binding. 5. Polyclonal rabbit antibody anti-HSV-TK (kindly provided by Professor A Cavaggioni, Department of Human Anatomy and Physiology, University of Padova, Italy). 6. FITC-conjugated secondary antibody (BioSource, Europe S.A., Nivelles, Belgium).

2.7. Immunohistochemistry

The following antibodies were used to evaluate the cellular components of the tumor mass and the inflammatory infiltrate: 1. Anti-glial fibrillary acidic protein (GFAP), anti-CD68, Mac387, CD57, CD20, CD79a, CD8, CD3, CD15 (Dako, Glostrup, Denmark). 2. Anti-CD1 (Immunotech, Marseille, France). 3. Anti-CD56/N-CAM (Neomarkers, Fremont, CA, USA). 4. Granzyme-B (Monosan, Uden, The Netherlands).

2.8. Detection of Cytokine Production by ELISA

Detection of cytokine in plasma and cerebrospinal fluid (CSF) samples was determined by the use of highly specific and sensitive enzyme-linked immunosorbent assays (ELISA): 1. IL-2 ELISA kit (BioSource, Europe S.A.). 2. IL-6 ELISA kit (BioSource, Europe S.A.). 3. TNFα ELISA kit (BioSource, Europe S.A.). 4. IFNγ ELISA kit (BioSource, Europe S.A.).

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3. Methods

3.1. Patient Selection

Patients with recurrent glioblastoma multiforme and fulfilling the following inclusion and exclusion criteria were selected for gene treatment after they had given their informed consent.

3.1.1. Eligibility Criteria

Patients required the following characteristics to be included in the protocol: 1. Age ≥ 18 and < 75 years. 2. Performance Status according to the World Health Organisation (WHO) 2. 4. Severe impairment of renal function (i.e., serum creatinine > 1.5 mg/dL or creatinine clearance < 80 mL/min/m2). 5. Severe impairment of liver function (i.e., alanine aminotransferase [ALT] and alkaline phosphotransferase levels > two times normal range). 6. Severe anemia, leukopenia, or thrombocytopenia (Hb < 10 g/dL; granulocyte count < 1,000 cells/mm3; platelet count 106 of the 50% tissue culture infective dose (TCID50) of a particular strain of reovirus. Within the group that received T1L, three individuals developed clinical illnesses with the symptoms of headache, pharyngitis, sneezing, rhinorrhea, cough, and malaise, which began 24–48 h after the viral challenge and lasted for a maximum of 7 days. A fourth individual was reported to have loose stools but was otherwise asymptomatic. The groups that received strains T2J or T3D had no reported clinical illness, except for one individual inoculated with T3D who developed a “mild rhinitis.” Interestingly, the majority of these patients seroconverted and virus was able to be isolated from anal swabs. Since this study, there have been several reports of clinical illnesses such as meningitis and encephalitis in association with the isolation of reovirus-like particles (24, 25); however, it has never been established that reovirus is the causative agent.

4. Molecular Basis of Reovirus Oncolysis

Although viral tropism is often linked to specific cell surface receptors capable of interacting with the virus, the ubiquitous sialic acid receptor and the junction adhesion molecule that permit reovirus binding and entry do not account for the differences observed in reovirus infectivity in transformed versus normal cells (26). Because aberrant cellular signalling pathways are often linked to cellular transformation, investigation was then directed towards these pathways being involved in viral oncolytic sensitivity/restriction. Strong et al. (26) showed two murine cell lines that were epidermal growth factor receptor (EGFR) negative and resistant to reovirus to be reovirus sensitive after transfection with EGFR-containing constructs. In addition, Tang et al. (27) demonstrated the capability of reovirus binding to the N-terminal ectodomain of the EGFR. The significance of intracellular signalling rather than receptor binding for reovirus infection of transformed cells was subsequently demonstrated, where reovirus-nonpermissive NIH-3T3 cells were rendered permissive after transfection with

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a v-erb B oncogene that confirmed ligand-independent constitutive tyrosine kinase activity (28). Because constitutive activation of Ras signalling has been implicated in carcinogenesis and tumour progression in a number of malignancies, the Ras pathway appeared central to EGFR downstream signalling (29, 30). When reovirus-resistant NIH3T3 cells were transfected with ras or downstream Ras pathway elements, reovirus infectivity was conferred (31). Further, reverse transcriptase (RT) polymerase chain reaction (PCR) performed for the viral S1 gene in Ras-transformed and -untransformed NIH3T3 cells indicated viral gene transcription occurring in both cell types. However, viral protein translation was confirmed only to the transformed cells (31). The translational block evidenced in untransformed cells was linked to dsRNA-activated protein kinase (PKR) activity and its antiviral defence capabilities. The ability of Ras-transformed cells to abrogate the antiviral defence mechanism had been indicated in previous studies (32, 33). In untransformed cells, dsRNA infection leads to an antiviral state in which PKR is phosphorylated. Phosphorylated PKR in turn phosphorylates eukaryotic translational initiation factor, which then terminates viral gene translation (34, 35). In Ras-transformed cells, PKR phosphorylation is downregulated, allowing a viral replication to be initiated (31) (see Fig.1). The mechanism of how Ras activity affects PKR phosphorylation has recently been investigated by Norman et al. (36). Activated Ras (GTP-bound Ras) is known to stimulate more than 18 effectors, with the most well characterized being the Raf kinases, phosphatidylinositol 3-kinase (PI3kinase) and guanine nucleotide exchange factors (GEFs) for the small G protein, Ral (29). Signalling through these effectors in conjunction with other downstream molecules leads to tumourigenesis. The results from the study by Norman et al. suggest that reovirus sensitivity is linked to the Ras/Ral guanine exchange factor (RalGEF)/p38 pathway (36). These investigations have contributed significantly to understanding the links between Ras signalling and reovirus oncolysis. Initial clues to reovirus’s potential as an anti-cancer therapeutic originated from the pioneering work by Hashiro et al. in 1977 (37). The preferential replication of reovirus in transformed cells was first demonstrated by this group. Further to this, Duncan et al. demonstrated that transformation of WI 38 cells with SV-40 large T antigen sensitizes cells to reovirus (38). Although these early studies provided clues to reovirus’s potential as an anti-cancer therapeutic, it was not until the late 1990s that the molecular basis of reovirus oncolysis was evaluated. The concept of using reovirus for the treatment of cancer was first tested by Coffey et al. (39). In this study, 25 human cancer cell lines were tested for in vitro reovirus oncolytic activity and 20 of these demonstrated susceptibility (39). The oncolytic potential of

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EGFR ISVP

1. Attachment

EGFR ligands

3. ISVP penetration

2. Uncoating

Core 4. 1° Transcription

mRNA

?

5. Translation, Viral core assembly

6. (−) strand synthesis and 2° Transcription

7.Translation and outer capsid synthesis

8. Cell lysis

Fig. 1. Reovirus replication in transformed (epidermal growth factor [EGF] overexpressing) tumour cells.1.Reovirus attaches to sialic acid and/or junction adhesion molecule receptors followed by receptor-mediated endocytosis.2. Shedding of outer capsid proteins σ1 and σ3 and proteolytic cleavage of μ1 C occurs within the endosome.3.Intermediate sub-viral particles (ISVPs) that have already under gone extracellular proteolytic cleavage of outer capsid proteins enter cells through direct penetration of cell membrane.4.Primary transcription takes place within the cores after endosomal membrane penetration and capped transcripts are released.5.Translation of viral messenger RNA (mRNA) is blocked in normal cells via phosphorylation of PKR and subsequently eIF2α. In cells with aberrant oncogenic signalling, this translational block is released through signalling of Ral GEF, P38, and likely through other uncharacterized molecules.6.Secondary transcription and negative strand synthesis takes place in the newly synthesized cores.7.Viral proteins are amplified through secondary transcription/translation, and the outer capsid is assembled.8.Viral progeny is released and cell death occurs.

reovirus was then confirmed in vivo in severe combined immune deficient non-obese diabetic (SCID/NOD) murine xenograft models bearing v-erb B-transformed NIH-3T3 or U87 glioblastoma multiforme tumours. In addition, in a syngeneic C3H murine model representing Ras-transformed C3H-10T1/2 tumours, reovirus treatment resulted in complete tumour regression in 6 out of 9 mice (39). It is well known that many viruses have the capabilities of both inducing and inhibiting apoptosis after infection. Apoptosis resulting from cellular viral entry and replication may facilitate release of viral particles from cells that allow targeting of bystander


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cells. Alternatively, apoptosis may be the result of the host’s antiviral defence mechanism where early cell death may attenuate further spread of the virus (lack of mature virion production). There is evidence to suggest that reovirus oncolysis of cancer cells is mediated mainly via apoptotic mechanisms (40–45). Transcription factor nuclear factor-kappa B (NF-kB) has been shown to be a pivotal molecule in apoptosis induction by reovirus in a cervical carcinoma cell line, Hela (40). It has also been shown that in ovarian (OVCAR3, PA-1, and SKOV-3), breast (ZR75–1), and lung (H157) cancer cell lines, reovirus infection results in sensitizing these cells to tumor necrosis factor (TNF)-related apoptosis-inducing ligand (TRAIL) and acts synergistically to enhance apoptosis (41, 42). In contrast, in colorectal cancer cell lines (C26 and HCT116), a direct interaction between reovirus and TRAIL has not been observed (43). In colorectal cells, reovirus-mediated apoptosis has been shown to be dependent on the ras mutation status of the cell and not on reovirus replication capabilities (43). To elucidate the apoptotic signalling pathways during reovirus oncolysis of breast cancer, our laboratory has performed microarray analyses. Reovirus treatment of MCF7 and HTB 133 (T47D) breast cells resulted in 2- to 12-fold increases in several receptor-associated genes such as TNFα-induced protein (TNFαI-P), TRAIL receptor 2, TNF receptor 6, TNF member 1, and TNF receptor superfamily member 6-associated factor, as well as 2to 27-fold increases in NF-kB, signal transducer and activator of transcription (STAT-5), and p53-upregulated modulator of apoptosis (PUMA) (44). The significance of these molecules in reovirus breast cancer oncolysis is currently being investigated. Our preliminary work using pharmacologic as well as molecular (vector based as well as short interfering RNA [siRNA]) inhibitors indicate both NF-kB and PUMA to be important molecules in reovirus oncolysis of breast cancer (45).

5. Preclinical Studies of Reovirus as a Cancer Therapeutic

During the past 7 years, reovirus has been tested for its in vitro and in vivo oncolytic activity in a myriad of cancers such as colorectal, ovarian, brain, breast, bladder, pancreatic, prostate, and haematological malignancies (22, 46–52). The majority of these investigations have used a SCID/NOD murine xenograft model system. A single intratumoural injection of reovirus containing 1 × 107 plaque forming units (PFUs) has proven sufficient to cure the majority of tumours tested. These pioneering studies opened up a new chapter in oncolytic viral therapy that has lead to several ongoing phase I and II clinical trials.

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5.1. Brain Tumours

The prevalence of aberrant Ras signalling in malignant gliomas (53, 54) and the lack of suitable therapy for this disease have prompted intense investigation pertaining to reovirus as a therapeutic for this malignancy. Early in vitro studies conducted by Wilcox et al. (46) have shown 20 (83%) of 24 malignant glioma cell lines as well as 9 (100%) of 9 primary malignant glioma cultures tested to be reovirus sensitive. In this study, the majority of reovirus-susceptible cell lines showed high Ras pathway activity as assessed by mitogen activated protein kinase (MAPK) activity, a downstream effector of Ras, whereas the less susceptible cell lines showed no MAPK activity. In an in vivo setting, subcutaneous or intracerebral tumours of U251N and U87 cells established in SCID/NOD mice showed striking and frequently complete tumour regression after reovirus treatment. Ten of 10 mice showed significant regression of U251 and U87 subcutaneous tumours at 11 days after live reovirus injection, in comparison with the UVinactivated virus-treated controls, and the viral oncolytic effect lasted up to day 21, or 28 days until the animals were killed. Additionally, reovirus oncolysis was confirmed in intracerebral U251N and U87lacZ glioma tumours, where reovirus treatment lead to significant tumour regression. The tumour burden was 2.4–2.5% with live virus treatment versus 38.3–40.8% with dead (UVinactivated) virus treatment. Intracerebral tumours implanted in nude mice showed significantly prolonged survival rates with intratumoural live reovirus treatment in comparison with dead virus treatment (46). Mice bearing U251N or U87lacZ tumours treated with dead virus showed a median survival of 42 and 48 days, respectively. In comparison, 8 of 12 mice bearing U251N and 9 of 11 mice harbouring U87lacz tumours treated with live reovirus had not reached median survival at 90 days (46). More recently, reovirus sensitivity of gliomas in other immunecompetent systems has also been established. Yang et al. (55) showed that racine glioma cells 9L and RG2 implanted subcutaneously or intracranially in Fisher 344 rats showed significant tumour regression and prolonged survival with reovirus treatment. In 16 of 16 rats, reovirus treatment dramatically inhibited subcutaneous tumours of RG2 origin compared with dead virustreated tumours. In a separate experiment, ipsilateral reovirus treatment of RG2 tumours did not result in tumour shrinkage of bilateral tumours. Interestingly, viral titres assessed in these tumours indicated the viral load to be 3–5 logs lower in the contralateral tumours when compared with the ipsilateral tumours, partially explaining the viral oncolytic inefficacy. 9L or RG2 tumours implanted in the putamen of Fisher rats and intratumoural treatment of live virus showed significant tumour shrinkage, as well as improved survival. Dead virus-treated tumours occupied 28.8% of the brain compared with 3.8% tumour in the live virus-treated brains. In survival experiments, nine of nine animals treated with dead virus were killed (ethical euthanasia) by


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day 32, whereas only seven of ten live virus-treated animals had to be killed by day 41. Interestingly, three (30%) of ten live virustreated animals were long-term survivors (alive until they were killed at day 169) and were devoid of tumour (55). In addition, reovirus pre-immunization of animals before tumour implantation did not abrogate reovirus tumour oncolysis (55). These findings have been translated to a phase I glioma clinical trial that has just been completed (56). In addition to gliomas, reovirus has also been evaluated in medulloblastoma (MB) a tumour that is prevalent among children and young adults. Because of its highly invasive nature, it has the ability to metastasize throughout the cerebral spinal axis (57). MB are resistant to current treatment modalities and overall outcome is poor. In vitro studies with medulloblastoma cell lines of human and mouse origin and surgical specimens have shown to be reovirus sensitive (58). Although little is known about the Ras status of MB, this study found a good correlation between reovirus susceptibility and ras activity, indicating that aberrant Ras signalling may be a critical factor in reovirus permissibility of this disease as well. Reovirus’s ability to treat leptomeningeal metastasis from medulloblastoma (MB) has also been investigated (58). To determine whether multiple intratumoural (i.t.) injections of reovirus would prolong survival and reduce the incidence of MB dissemination, Yang et al. (58) used a Daoy cell orthotopic model system. In this study, green fluorescent protein (GFP)-labelled Daoy cells were inoculated to the right putamen of nude mice and 20 days later, live or UV-inactivated virus was administered i.t. A whole-body fluorescence imaging system (WBFIS) was used to monitor tumour growth and dissemination in vivo. Detected fluorescent tumours were given multiple reovirus injections, to a maximum of seven injections (1 × 107 PFUs/injection). Interestingly, live virus treatment resulted in significantly long-term survival and a dramatic reduction in spinal and leptomeningeal metastasis of MB. All of the dead virus-treated animals died of recurrent tumour (median survival = 78 days), whereas four of seven animals with live virus treatment were long-term survivors (median survival not reached at the time of publication). These results support future clinical trials in which reovirus could be used to control metastasis of MB to the spinal cord and thereby allow a reduction in the dose/field of total neuroaxis cerebrospinal radiotherapy currently used to prevent or treat cerebrospinal fluid dissemination. 5.2. Breast Cancer

Work conducted by Norman et al. has shown reovirus’s oncolytic potential against five breast tumour-derived cell lines as well as its ability to replicate in human ex vivo breast tumour specimens (47). Reovirus infection, multiplication, and lysis of HTB 133 (T47D) breast tumour cells are depicted in Fig. 2.

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a

b

c

Fig. 2. Reovirus oncolysis of breast cancer: viral protein detection by immunofluorescence. HTB 133 breast cancer cells that have been challenged with 40 multiplicities of infection (MOI) of reovirus for 2 h (a), 24 h (b), and 72 h (c) have been stained with anti-reovirus antibody and FITC-conjugated secondary antibody, and photographed. Note the increase in viral protein synthesis (fluorescence) at 24 h and cell disruption at 72 h of incubation. (Adapted from (82) with permission) (see Color Plates).


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Reovirus efficacy evaluated in vivo, in an MDA-MB-43S mammary fat pad model in SCID mice, has shown significant tumour regression after intratumoural reovirus therapy. Reovirus’s systemic ability to control tumours distal from its site of injection has also been demonstrated in a breast MDA-MB-468 SCID/NOD bilateral tumour model (47). In this study, ipsilateral reovirus treatment of tumour resulted in shrinkage of the contralateral tumour, suggesting a role for future systemic reovirus therapy for breast cancer. In conjunction with these laboratory findings, four breast cancer patients recruited in the phase I reovirus intratumoural clinical trial has shown encouraging results after reovirus treatment (59). Although mutations in the ras proto-oncogene are rare in breast cancer, this malignancy harbours several other mutations, resulting in deviant signalling pathways that may originate upstream or downstream of Ras. For instance, 30% of breast carcinomas overexpress Her-2, and upregulated Src family kinases are common in breast cancer cell lines and primary tumours (60– 65), thereby possibly enhancing the Ras signalling pathway to the advantage of reovirus. In addition, the ability of reovirus to control brain and leptomeningeal metastasis of breast cancer has been evaluated by Yang et al. (66). In this study, intracranially implanted breast tumours (HTB 129 and HTB 132) in nude mice have exhibited tumour shrinkage, absence of metastasis, and prolonged survival with reovirus treatment, whereas dead virus-treated animals showed spread of disease to other areas of the brain. In a separate study involving immunocompetent animals, breast tumours originating from 13762MATBIII cells were implanted intrathecally in Fisher 344 rats and reovirus (2 × 109 PFU/rat) was administered via the same route on day 4 and day 8 –after tumour inoculation. As expected, the median survival of animals with live reovirus treatment was significantly longer (16 days, n = 15) in comparison with the UV-inactivated virus-treated group (10 days, n = 14). A similar outcome was noted when the experiment was repeated with GFP-labelled 13762MATBIII breast cancer cells. All animals in this study however, succumbed to metastatic disease, likely secondary to immune clearance of virus (66). 5.3. Colon Cancer

Because mutations in the K-ras proto-oncogene are found in approximately 50% of colorectal cancers (67), Hirasawa et al. studied reovirus’s oncolytic potential against this malignancy (22). Five colon cancer cell lines but not a normal colon cell line were shown to be reovirus sensitive in vitro, and augmented Ras activity was detected in all of these sensitive cancer cells in contrast to a normal cell line (22). Subcutaneous (s.c.) tumours of colon cancer cell lines HCT-116, HT-29, and DLD-1 established in SCID/NOD mice showed significant tumour regression when

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reovirus therapy was administered intratumourally (n = 6). Intravenous reovirus therapy of s.c. HCT-116 tumours in nude mice also demonstrated significant tumour growth inhibition in comparison with dead virus-treated controls (n = 6) (22). A more recent study used a syngeneic mouse model to examine the role of immunosuppression on reovirus therapy of colorectal liver metastases (68). In this study, murine colorectal C26 tumours were established in livers of BALB/c mice and intratumoural reovirus therapy was given in the presence or absence of cyclosporine A (to dampen down the viral immune response). Significant, but short-term (up to 5 days) suppression of tumour metastases was observed in a group of 9 mice that received a single injection of reovirus with no immunosuppression. In comparison, mice that received cyclosporine A and reovirus showed markedly enhanced and prolonged anti-tumour activity at days 5, 7, 9, 11, and 13 with 16 mice in each group. Complete tumour eradication was seen in two mice with reovirus treatment and cyclosporine A treatment (68). 5.4. Ovarian Cancer

Efficient reovirus infection and cytopathic effects have been demonstrated against four ovarian cancer cell lines but not in a normal ovarian cell line (22). Intratumoural injections of live reovirus (1 × 1010 PFU, given at days 0 and 14 after tumour establishment) to SKOV3 ovarian tumour-bearing CD-1 nude mice lead to marked tumour regression in comparison with dead virus-treated animals (n = 6 in each group). To assess reovirus capability of invading metastatic tumours, an intraperitoneal human ovarian cancer xenograft nude mouse model has also been studied (22). In this study, intraperitoneal tumours of MDAH2774 cells established in nude mice were given 5 × 108 PFU of live or UVinactivated reovirus at days 5 and 19 after tumour establishment. Live reovirus treatment significantly inhibited ascites tumour formation and prolonged animal survival in comparison with the controls (n = 10 in each group) emphasizing the oncolytic potential of reovirus against this tumour type as well.

5.5. Prostate Cancer

Reovirus’s oncolytic potential against prostate cancer has also been investigated in detail. As shown by Nodwell et al. (50), prostate carcinoma cell lines LN-Cap, DU-145, and PC-3 were exquisitely sensitive to reovirus as indicated by cytopathic effects, viral protein synthesis evaluated by metabolic labelling, immunofluorescence staining for reovirus antibodies, and viral progeny production (50). Hind flank subcutaneous xenograft SCID/NOD model systems harbouring tumours of these cell types showed dramatic tumour regression with intratumoural treatment of reovirus in comparison with the controls (50). These interesting findings have also been translated to a phase I prostate clinical trial with indications of efficacy (PSA reduction) and minimal toxicity.


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5.6. Bladder Cancer

Kilani et al. (48) have tested reovirus susceptibility of transitional cell carcinoma (TCC) of the bladder cancer. MGH-U3 and RT-112 TCC cells of human origin, as well as a rat TCC cell line AY-27 demonstrated susceptibility to reovirus at doses ranging from 320 to 2 × 108 PFUs. An in vitro spheroid model system representing homogeneous TCC cells, or co-culture spheroids representing a mixture of these TCC cell lines and normal fibroblasts, showed specific killing of TCC tumour with reovirus treatment as assessed by cell viability tests (SYTO 16/propidium iodide) and immunohistochemical staining (cytokeratin 13 for TCC cells and anti-vimentin specific for fibroblasts) as well as time-lapse imaging techniques (48). Furthermore, reovirus’s ability to control superficial bladder cancer has also been demonstrated in an in vivo rodent model system (69). Although intravesical immunotherapy with Bacillus Calmette-Guérin (BCG) is currently used as a therapeutic adjuvant after surgery of superficial bladder cancer, recurrence is seen in 20% of patients and complications arising from BCG treatment are not infrequent. Therefore, Hanel et al. (69) have compared reovirus treatment of superficial bladder cancer with BCG treatment in Fisher rats implanted with AY-27 rat TCC cells. Interestingly, this study showed lesser side effects and higher tumour-free survival with reovirus treatment than with BCG treatment. Tumour response assessed as animal survival at 100 days after tumour implantation was 90% with reovirus treatment, whereas it was 50% with BCG treatment. These results support using reovirus for future clinical applications for this indication.

5.7. Pancreatic Cancer

The prevalence of ras mutations in pancreatic cancer (over 80%) (70) has lead Etoh et al. (49) to investigate reovirus effects in this tumour as well. Five pancreatic tumour cell lines tested showed susceptibility to reovirus and exhibited elevated Ras activity. Intratumoural reovirus treatment of hind flank xenografts of Panc1 and BxPC3 tumours in nude mice lead to significant tumour regression in all mice studied in comparison with the mice that received dead virus (n = 5). In the same study, systemic effect of unilateral reovirus injection was studied in bilateral tumours derived from Panc1 tumours. Live ipsilateral, intratumoural virus treatment resulted in significant tumour shrinkage bilaterally (n = 5) and these effects were not observed with dead virus tumour treatment. More recently, Himeno et al. (71) tested reovirus’s ability to control liver metastasis of pancreatic cancer. Using a syngeneic hamster pancreatic model, this group was able to demonstrate that administration of reovirus via the portal vein significantly decreased the number and size of liver metastasis in comparison with the controls. Six of six animals that received phosphate-buffered saline as treatment showed no response, whereas in reovirus-treated hamsters, only four of six showed tumour response.

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5.8. Lung Cancer

Systemic reovirus therapy for metastatic lung cancer and routes of preventing immune invasion of reovirus has been addressed by Hirasawa et al. (72). In this study, the Lewis lung carcinoma C57BL mouse model was adopted in which removal of established primary tumours leads to metastatic lung cancer. Intravenous reovirus treatment of 5 × 108 PFUs on days 1 and 7 lead to dramatic reductions in the mean number of metastatic foci (2 metastatic foci with reovirus treatment and 39 in controls) as well as more than threefold reduction in total lung weight, which was comparable to healthy animals. Of note however, tumour shrinkage seen in these immunocompetent animal models after systemic reovirus therapy appeared to be transitory, because after 3 weeks of reovirus administration, tumours resumed growth. Rising anti-reovirus antibodies within the animals indicated that the oncolytic effect of reovirus was circumvented by the host immune system. When systemic reovirus therapy was combined with immunosuppressive drugs such as cyclosporin A or anti-CD4 or -CD8 monoclonal antibodies, the anti-tumour efficacy of reovirus was restored.

5.9. Haematological Malignancies

A variety of haematological cancers have been tested for reovirus sensitivity in vitro, in vivo, and ex vivo in our laboratory (52) and by Alain et al. (51). In vitro reovirus oncolysis evaluated in five Burkitt’s lymphoma cell lines confirmed sensitivity in only two cell lines, Raji and CA46. The other three cell lines, Daudi, Ramos, and ST486, were reovirus resistant. This in vitro resistance was also observed in vivo. Reovirus treatment of Raji and Daudi orthotopic tumour xenografts in SCID/NOD mice showed regression of only Raji but not Daudi tumours. Although an ex vivo human specimen of Burkitt’s lymphoma appeared reovirus sensitive (51), another tested in our laboratory showed resistance to reovirus (52). In contrast, results that were more promising were seen with diffuse large B cell lymphoma (DLBCL). Four established cell lines and three ex vivo specimens of this malignancy tested so for have shown sensitivity to reovirus (51, 52). Initial experiments conducted with B cell chronic lymphocytic leukemia (B-CLL) has shown significant activity. Nineteen of 19 ex vivo human specimens exposed to reovirus to date have shown sensitivity (51, 52), indicating reovirus to be a possible therapeutic for this malignancy. Among other non-Hodgkin’s lymphomas, two ex vivo patient samples of small cell lymphocytic lymphoma have proved positive for reovirus infection and oncolysis. In contrast, follicular lymphoma appears to be refractory for reovirus treatment because only one of seven ex vivo samples tested so far has shown sensitivity (51, 52). Multiple myeloma is a haematological malignancy that often shows failure to conventional therapy and reovirus has shown promise towards this disease as well. RPMI 8226 multiple myeloma


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and U937 monocytic cell lines tested to date have shown exquisite sensitivity to reovirus as confirmed by cytopathic effect and viral protein synthesis and propidium iodide exclusion assessed by flow cytometry (52). The possibility of using reovirus as a therapeutic against mantle cell lymphoma is currently underway in our laboratory. Two of five established cell lines tested were sensitive to reovirus as demonstrated by cytopathic effect and metabolic labelling data (73). Treatment of reovirus virions with extracellular proteases results in the removal of the outer capsid protein σ3 and cleavage of the µ1 protein. This “uncoating” process results in an advantage for reovirus where it can now directly penetrate the cell membrane and bypass receptor binding and receptormediated endocytosis (see Fig. 1). By using this strategy, Bebb et al. (73) has shown that infection of these three resistant cells lines with intermediate subviral particles (ISVPs) rendered two of three resistant cell lines sensitive. A unique characteristic of reovirus is its oncolytic restriction to tumour cells and its inability to replicate in normal cells. Indeed, this has been demonstrated in a majority of the abovementioned studies, when the resected tumours were tested for the reovirus infectivity by means of immunohistochemistry, only the tumour areas and not the adjoining normal tissue showed apparent staining for reovirus, demonstrating that its oncolytic ability is restricted to transformed cells. 5.10. Other Tumours

6. Reovirus in Combination with Chemotherapy or Radiation Therapy

The ability of reovirus to replicate and kill mouse and human melanoma cell lines has been investigated by Errington et al. (74). Viral plaque assays and Western blot analysis against viral proteins have indicated reovirus to replicate in all cell lines tested. In vitro reovirus efficacy determined by trypan blue exclusion and propidium iodide incorporation has shown virus replication to directly correlate with oncolysis. In vitro efficacy of reovirus has also been demonstrated in cell lines of childhood sarcomas such as Ewing’s sarcoma, rhabdomyosarcoma, synovial sarcoma, and osteosarcoma origin (75). In vivo, 1 × 109 PFUs of reovirus was well tolerated in a nude osteosarcoma xenograft model. All other tumour lines tested in mouse xenografts exhibited reovirus anti-tumour activity, with a rhabdomyosarcoma cell line showing complete tumour regression (75).

When the combination of reovirus with either chemotherapy or radiation therapy was examined for synergy, there appeared to be some local benefit in animal models. These pre-clinical data have

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now been translated into early phase clinical trials looking at radiation therapy timed with intralesional REOLYSIN. One of the first chemotherapy/REOLYSIN combination studies involved the use of the HCT 116 (a human colon carcinoma) cell line and dose escalations of both REOLYSIN and the antimetabolite DNA synthesis inhibitor, gemcitabine. At all doses tested, there appeared to be synergy between the two agents (76, 77). Interestingly, this same group showed an additive effect when cisplatin and doxorubicin were used individually with REOLYSIN, but no such additive effect with paclitaxel (77). In contrast, Sei et al. found that in all NSCLC cell lines that were sensitive to cisplatin, gemcitabine, mitomycin, or vinblastine, synergy was seen with REOLYSIN (78). Further, in all six NSCLC cell lines tested, synergy was seen with paclitaxel and REOLYSIN, unlike the colon carcinoma cell line data. Cisplatin was also shown to be synergistic with REOLYSIN in a malignant melanoma (B16.F10) model both in vitro and in vivo (79). When radiation was looked at for possible synergy with REOLYSIN in multiple cancer cell lines, it was determined that in cell lines that exhibited only moderate REOLYSIN sensitivity when treated with radiation there was an increase tumour response when compared with radiation alone (80). This was also seen in SW480, HCT 116, and B16 in vivo murine models (80).

7. Other Possible Clinical Indications for Reovirus 7.1. Purging of Minimal Residual Disease During Autologous Stem Cell Transplants (ASCT)

In addition to its oncolytic efficacy of multiple tumour types, reovirus has also been evaluated as a novel purging strategy during ASCT. Autograft contamination by clonogenic malignant cells is a frequent occurrence, and gene-marking studies have provided evidence that contaminating tumour cells within the autograft could lead to relapse (81). An ideal purging technique should preserve the number and function of stem cells while selectively destroying the contaminating tumour cells. With this in mind, we tested reovirus’s ability to infect stem cells, its propensity to affect long-term colony forming abilities of stem cells, and finally its therapeutic use as a successful purging agent. In our laboratory, positively selected (CD34+) stem cells exposed to reovirus in culture medium for 5 days did not show any adverse virus effect (52). In addition, these stem cells preserved their clonogenic potential as revealed by long-term colony-forming assays in methylcellulose (52). Lastly, by using metabolic labelling experiments, we also confirmed that reovirus does not replicate in growth factor (granulocyte colony-stimulating factor)-stimulated


623

haematopoietic progenitor cells (52). These findings implied that reovirus could purge autografts without affecting the number and function of stem cells. Using an ex vivo purging model system, we have demonstrated that apheresis product contaminated with haematological tumour cell lines could be successfully purged with reovirus. Purging efficacy assessed by flow cytometry and tumour re-growth assays indicated complete purging of U937 at 0.01% tumour burden and RPMI 8226 even at 1% tumour burden. Admixing apheresis products with primary tumour cells (1–10%) of transplantable diseases such as B cell chronic lymphocytic leukemia (B-CLL), diffuse large B cell lymphoma (DLBCL), small lymphocytic lymphoma (SLL), or Waldenstrom macroglobulinemia and purging with reovirus resulted in successful elimination of malignant cells, as detected by flow cytometry. Successful reovirus purging of DLBCL was seen at 10% tumour burden at 2 days. Four to 5 days of reovirus purging of CLL tumour at 10% admixed tumour showed complete purging of two patient specimens and significant purging with another two patient samples. Complete purging was also seen at 5 days with a Waldenstrom macroglobulinemia patient specimen at 10% tumour burden. In addition, tumour of SLL also showed complete purging at 5 days with reovirus at 10% tumour burden (see Fig. 3). More than 50% purging was seen with a patient-derived multiple myeloma specimen at 5% tumour burden at 5 days. In contrast, we found incomplete purging of primary tumour cells of a Burkitt’s lymphoma and a follicular lymphoma by reovirus at 1% tumour contamination (52). More recently, we have demonstrated that reovirus’s purging abilities can be extended to solid tumours such as breast cancer (82). Our group and others have demonstrated that breast tumour micrometastases found in patients’ apheresis product predict shorter progression-free survival and overall survival. These findings imply that either the contaminating tumour cells within the apheresis product reflect a higher systemic disease burden and/or that re-infused contaminating tumour cells contribute to relapse. The ex vivo breast purging model used in our study demonstrated complete purging of up to 1% tumour burden. The tumour burden encountered in a clinical setting is usually less than 0.01% and this falls well within the range of successful purging seen in our experiments. Our group is also attempting to improve the quantitation of reovirus purging efficacy and human stem cell repopulating ability in vivo, using a murine myeloma transplant model system. A variety of haematological ex vivo tumour specimens are currently being tested for reovirus sensitivity, to increase the number of potential cancers that may benefit from this purging procedure, i.e., chronic lymphocytic leukemia. The aim of these purging

624


a

Small lymphocytic lymphoma NV

LV

b

% small lymphocytic lymphoma cells

CD19 PC7

c

Kappa FITC

Malignant Monoclonal B Cells

Lambda PE Kappa FITC

CD20 ECD

CD5 PC5

CD5 PC5

CD19 PC7

LV

AB

CD5 PC5

CD5 PC5

NV

A

Normal Polyclonal B Cells

A

CD20 ECD

Lambda PE

120 100 80 60 40 20 0 NV/LVD0

NV D5

LV D5

Treatment and days post purging

Fig. 3. Reovirus-purging effects on patient-derived small lymphocytic lymphoma (SLL). (a) Cytopathic effect of reovirus on human SLL cells 72 h after infection. Purified cells were infected with reovirus (40 MOI) and cells were photographed 72 h after infection. Cytopathic effect was seen in reovirus infected but not in uninfected cells. (b) Flow cytometric analysis of SLL after reovirus purging. Apheresis product cells were mixed with SLL cells (10%) and purged for 5 days with reovirus. Samples were analyzed using flow cytometry. Dim CD5+/CD19+/20+ B cells were gated using two regions (A and B) and assessed for clonality. The lambda-positive SLL was clearly distinguished from the normal polyclonal B cells. (c) Representative histograms of viable SLL cells before and after purging with reovirus. The arrow indicates that SLL cells were not detected. (Adapted from (52) with permission).


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experiments is to generate pre-clinical data that will eventually be translated to a phase I clinical trial in the near future. 7.2. Reovirus and the Immune System

8. Persistently Infected Tumour Cells and Acquired Resistance

It was recognized early on using immunocompetent xenograft murine models that immune clearance of virus was going to be a significant barrier to systemic administration of reovirus. Initial studies by Hirasawa et al. using a ras-expressing C3 syngeneic immunocompetent murine model demonstrated that preimmunization with reovirus diminished reovirus systemic efficacy, and the use of immunosuppressive agents such as cyclosporine A or cyclophosphamide augmented tumour response (72). The original phase I intratumoural human trial involving 18 patients revealed a robust immune response early on after the first dose of REOLYSIN, and the response was dose dependent (59). Further, in this trial, only local responses were seen and no viral culture or PCR evidence of virus could be isolated from serum, urine, stool, or cerebrospinal fluid (59). Several studies have confirmed the significance of immune recognition and subsequent clearance of reovirus as deleterious to potential efficacy. Vile et al. used cyclophosphamide as an immune modulator in a melanoma model and found that the addition of cyclophosphamide enhanced reovirus tumour efficacy and increased the amount of reovirus that replicated in the tumour (83). Smackman et al. used a murine model of colon cancer (C26 cells) to demonstrate the potentiation of reovirus with the addition of cyclosporine A (68). Several laboratories including our own have taken a different approach to the immune recognition of reovirus. We and others have attempted to use reovirus as a tumour targeting agent as well as a non-specific immune stimulant. Errington et al. have provided data supporting the finding that reovirus may activate dendritic cells and thereby promote innate anti-tumour immunity (84). This group has shown that reovirus caused dendritic cell maturation, cytokine production, and increased natural killer and T cell cytotoxicity in colorectal, melanoma, and lymphoma model systems (84–86).

Although a myriad of tumour types have been studied with regard to the oncolytic efficacy of reovirus, it has been suggested that a subpopulation of cancer cells may survive the initial viral challenge and become persistently infected with the virus. Although persistently infected cells with reovirus have been established in vitro (87), it is not known whether these cells exist in vivo. The presence of viral particles in persistently infected cells may

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present a potential challenge to viral therapy in a clinical setting. This issue has been addressed by Alain et al. (88) by using persistently infected cells as well as cured cells (persistently infected cells treated with anti-reovirus antibodies) developed from the reovirus-susceptible Burkitt’s lymphoma cell line, Raji. Reovirus infection and oncolysis of these cells along with the parental cells were examined both under in vitro and in vivo conditions. Persistently infected cells were found to be non-tumourigenic in vivo, however, in contrast, anti-reovirus antibody-treated cells were highly tumourigenic in NOD/SCID mice. Reovirus antibody-treated Raji cells were resistant to reovirus in vitro but were highly susceptible in vivo. A single injection of reovirus showed significant tumour regression in eight of eight mice. Additional experiments showed that proteases within the tumour microenvironment of cured Raji tumours enhanced the viral uncoating process and its subsequent oncolytic activity in vivo (n = 5). The lack of tumourigenicity of persistently infected cells and in vivo reovirus susceptibility of cured cells as evidenced by these experiments indicate that such conditions should not pose a potential obstacle for reovirus oncolytic therapy in vivo. Kim et al. (89) also showed acquired resistance to reovirus therapy in the HT1080 human fibrosarcoma cell line. Prolonged exposure to reovirus yielded highly resistant HTR1 cells that still exhibited high Ras activity, as high as the parental cell line. As in the Alain et al. (88) study, persistently infected cells were not tumourigenic but immunologically cured virus-free cells were highly tumourigenic in vivo in SCID/NOD mice and even grew faster than the parental cells as seen in five of five mice. Reovirus resistant HTR1 cells still maintained their apoptotic potential when induced with chemotherapy or adenovirus (89), suggesting that alternative therapies may still be a solution for such virusresistant sub-populations of cells.

9. Reovirus Toxicity Studies Toxicities attributed to reovirus have been observed in severely immunocompromised SCID/NOD animals. A fatal vasculitic injury to the animal’s extremities (toes, ears, and tail) and myocardium is commonly seen approximately 30 days after reovirus injection (21) that is never detected in athymic nude mice even with repeated injections via intratumoural and intravenous administration of reovirus (22). Pre-clinical toxicology experiments with laboratory-grade T3D reovirus have demonstrated that intratumoural (including intracerebral) and subcutaneous routes of administration were


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not associated with toxicity in immunocompetent racine models (55). In addition, a contracted company was used to investigate subcutaneous and intravenous administration of REOLYSIN (good manufacturing practise [GMP]-produced clinical-grade T3D reovirus) in rats, beagle dogs, and cynomolgus primates and the studies revealed no significant toxicity (International Toxicity Research [ITR], Montreal, Quebec, Canada). The results from the 32-animal primate study using daily doses of 5.0 × 109 PFU/day administered intravenously for 28 consecutive days did not reveal any REOLYSIN-related clinical signs or changes in food consumption, body weight, electrocardiograms, haematological, or coagulation parameters. The only finding compared with control animals was evidence of a mild polyarteritis manifested as effacement of the glomerular basement membrane and heavy kidney/spleen weights. These findings were thought to be caused by REOLYSIN-induced immune stimulation as opposed to a direct REOLYSIN effect. Yang et al. (55) evaluated reovirus (laboratory grade) neurotoxicity in nude mice. When doses of at least 108 PFUs were administered intracranially 12 (100%) of 12 mice survived, whereas doses of 109 PFUs/mouse were lethal in three of four mice tested, within 2–3 days after virus administration. A transient (10%) loss of body weight was seen in mice receiving 108 PFUs, the maximum tolerated dose (MTD). In another study conducted in Fisher 344 rats by the same group, the MTD was found to be 109 PFUs with intracranial administration of reovirus. Rats treated with less than 109 PFUs of reovirus did not show significant loss of body weight and appeared healthy. Animals treated with 4 × 109 PFUs or multiple injections (five injections) of 109 PFUs died or showed transient loss of body weight (15–25%) 2–4 weeks after viral administration. Fisher rats given three 1 × 109 PFU doses at 4-day intervals showed no behavioural changes as determined by the Morris water maze model (n = 8). Histology of the brains of these animals revealed mild inflammation, reactions in the injection site, and communicating hydrocephalus (55). Yang et al. (55) also studied REOLYSIN toxicology in cynomolgus monkeys. Doses ranging from 1.51 × 106 to 1.51 × 109 PFUs were administered intracerebrally to male and female monkeys with a monkey in each sex receiving a designated dose level. Female and male animals were killed at 2 and 6 months, respectively, and no unplanned deaths were seen during this period. A transient event was seen on day 6 in a male monkey that received the highest dose (i.e., 1.51 × 109 PFUs), a selflimiting convulsion. The group that received the highest dose of REOLYSIN showed potential treatment-related increases in rectal temperatures (39.6–40.3°C). However, no treatment-related effect on body weight or food intake was observed during the

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course of the study. No REOLYSIN effects were seen in haematological, biochemical, or urinary parameters. Reactive changes at the site of injection were seen in a dose-related manner, with the highest dose causing a microscopic area of subacute infarction at the injection site. To date, upwards of 100 patients have been treated with REOLYIN intratumourally, intracerebrally, or intravenously during early clinical testing in cancer patients, with minimal toxicity attributed to REOLYSIN.

10. Reovirus Clinical Trials REOLYSIN (GMP reovirus) is currently manufactured under the auspices of Oncolytic Biotech, Inc. (Calgary, Alberta, Canada). This biotechnology company has been responsible for all of the clinical trial activity to date using this product. Upwards of 15 different clinical trials have either been completed, are ongoing, or are currently undergoing regulatory approval in Canada, the USA, and/or the UK (see Table 1). It should be pointed out that more than 100 patients have now been treated with REOLYSIN world wide and, to date, the MTD has not yet been reached in either intralesional or systemically administered studies. The first-in-human cancer clinical trial involved 19 patients who had exhausted standard treatments and was of a single-institution intralesional dose-escalation phase I design. Eighteen patients were evaluable and received intralesional REOLYSIN (dose escalation from a single dose of 1 × 107 PFUs to 15 injections of 1 × 1010 PFUs [injections given three times per week]). Virus was tolerated with no grade 3 toxicity reported that could be attributed to REOLYSIN. Interesting, even though local responses were seen in patients with melanoma, head and neck, and Kaposi’s sarcoma cancers, little distant systemic evidence of activity was seen. Neutralizing anti-reovirus antibody titres were measured weekly and a REOLYSIN dose-to-titre relationship was documented, suggesting an immune clearance mechanism that may have limited systemic efficacy. Further, virus was unable to be cultured from serum, urine, stool, or cerebrospinal fluid in any of the patients, suggesting immune clearance. A second intralesional clinical trial involved patients with organ-confined prostate cancer. A single injection of 1 × 107 REOLYSIN was given transrectally into an ultrasound echoic prostatic lesion previously confirmed as cancerous. Patients were followed weekly for prostate-specific antigen (PSA) level and toxicity, and they underwent radical prostatectomy 3 weeks after the injection. Within 24 h after receiving

Phase Phase I Phase I Phase I Phase I Phase I Phase I Phase I Phase II

Phase I

Phase I

Phase I

Phase II

Name/type

Reo 001Intralesional

Reo 002Intralesional prostate

Reo 003Intracerebral

Reo 004Systemic

Reo 005Systemic

Reo 006Reovirus with concurrent radiation

Reo 007Infusional reovirus

Reo 008Reovirus with concurrent radiation

Reo 009Reovirus with concurrent chemotherapy (gemcitabine)

Reo 010Reovirus with concurrent chemotherapy (docetaxel)

Reo 011Reovirus with concurrent chemotherapy (paclitaxel and carboplatin)

Reo 014Intravenous monotherapy

Table 1 Reovirus clinical trials

Sarcomas

Melanoma, lung, ovarian

Bladder, prostate, lung, upper gastrointestinal

Pancreatic, lung, ovarian

Various metastatic tumours including head and neck

Recurrent malignant gliomas

Various metastatic tumours



Recurrent malignant glioma

Organ-confined prostate cancer

Solid tumours

Indication

Ongoing

Ongoing

Approximately 21 patients Approximately 52 patients

Ongoing

Ongoing

Ongoing

Ongoing

Ongoing

Closed

Closed

Closed

Closed

Closed

Status

Approximately 21 patients

Approximately 21 patients

N = 40

Unknown

N = 23

N = 33

N = 18

N = 12

N=6

N = 19

N

United States

United Kingdom

United Kingdom

United Kingdom

United Kingdom

United States

United Kingdom

United Kingdom

United States

Canada

Canada

Canada

Location

—

—

—

—

—

—

(91)

(90)

—

(56)

—

(59)

Reference

Oncolytic Viral Therapy Using Reovirus 629

630


the injection, a flu-like illness (pyrexia, myalgias, arthralgias) was documented. If prophylaxis with acetaminophen was administered, this toxicity was significantly ameliorated. Of the six patients who received REOLYSIN, two had a reduction in baseline PSA values and three had evidence of apoptosis and/or necrosis seen at the time of pathological review of the prostatectomy specimen. A third intralesional phase I study involved patients with recurrent malignant glioma that had previously been treated with surgical resection, radiation, and in several cases chemotherapy. Dose escalation in cohorts of three ranged from 1 × 107 to 1 × 109 TCID50. MTD was not reached in this study, and 3 of the 12 patients enrolled were alive 12 months after treatment. Currently, there are nine other clinical trials either actively accruing patients or just completed accrual in the USA and the UK. Two of these studies are phase I clinical trials that involve dose escalation and repeated schedules of systemically administered REOLYSIN in patients who have exhausted standard treatment interventions and thus have been heavily pre-treated. The outcomes of these studies should be formally reported either in abstract or publication form sometime in 2008. Other trials that are ongoing include REOLYSIN and radiation as well as several trials looking at REOLYSIN in combination with several cytotoxic chemotherapeutic agents (REOLYSIN/gemcitabine in pancreatic, lung, and ovarian cancer patients; REOLYSIN/docetaxel in bladder, prostate, and lung cancer patients; REOLYSIN/ paclitaxel/carboplatin in melanoma, lung, and ovarian cancer patients).

11. Concluding Remarks Among the oncolytic viral platforms, reovirus has proved to be an efficient and successful candidate that can target a plethora of cancers under in vitro, in vivo, and ex vivo conditions. Its preclinical efficacy and minimal toxicity towards humans has lead to several phase I/II clinical trials that show indications of efficacy and are ongoing. Combining reovirus therapy with current mainstays such as chemotherapy and radiation are other approaches that are being clinically evaluated to maximize its therapeutic potential. Considerable effort is currently being focussed on either downregulating the immune system or strategies for the immunopotentiation of reovirus to minimize antiviral immunity and maximize anti-tumour effects.


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Chapter 32 Design and Testing of Novel Oncolytic Vaccinia Strains Steve H. Thorne Summary Oncolytic or replication-selective viruses have been used as powerful tools for the delivery of therapeutic genes to tumors. Because these vectors are capable of replicating within the tumor, the therapeutic gene is amplified within the target tissue itself, resulting in the spread of the virus both within the tumor, and sometimes also between tumors. Vaccinia virus holds many advantages when serving as the backbone for oncolytic viral strains, including a large cloning capacity (at least 25 kbp) (1); a short life-cycle (2, 3); extensive previous use in humans, with contraindications and adverse reactions well described and antivirals available (4); the potential for systemic (intravenous) delivery to distant tumors; and vaccinia strains have previously demonstrated antitumor benefits in clinical trials (5). Because vaccinia has no known receptor and is capable of infecting almost any cell type, tumor selectivity has to be engineered into vaccinia at steps after infection. We will therefore discuss potential viral virulence genes and metabolic targets that result in tumor-selective vaccinia strains. Because the virus has limited natural requirements for host cell proteins, and, instead, contains a large genome and multiple genes involved in virulence, a large number of possible attenuating gene deletions can result in the production of viral strains reliant on inherent properties of the host cell for replication. The protocols for producing viral gene deletions and constructing viral gene expression vectors have been well established for vaccinia and are summarized briefly in this chapter. Basic assays for testing the tumor selectivity and therapeutic index of new oncolytic constructs in vitro will be covered. In addition, we describe how bioluminescence imaging can be incorporated into preclinical testing of vaccinia gene expression strains to examine the timing, biodistribution, and kinetics of viral gene expression noninvasively after delivery of the viral agents to tumor-bearing mice via different routes. Key words: Oncolytic virus, replication-selective, tumor-targeting, vaccinia, virotherapy

1. Introduction It is evident that many of the adaptations that a virus produces within an infected cell to enhance its own replication are similar to the hallmarks of a malignant cell. These include uncontrolled Wolfgang Walther and Ulrike S. Stein (eds.), Methods in Molecular Biology, Gene Therapy of Cancer, vol. 542 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-561-9_32

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proliferation, evasion of apoptotic cell death, and an inability to produce or respond to immunomodulatory factors. It is therefore unsurprising that the root causes of some cancers are known to be chronic viral infections. Conversely, it is true that many viral virulence genes or functional domains are redundant within cancer cells and therefore their deletion results in attenuated viral strains that are still capable of normal replication within tumor tissue. It is this strategy that is most effective for creating tumortargeting strains of vaccinia. Because the virus replicates in the cytoplasm and has minimal interaction with host cell transcription and DNA replication machinery, the strategy of incorporating tumor or tissue specific promoters to drive essential viral genes or transgenes (as has been reported for other viruses, especially adenoviral strains) is unlikely to be effective in vaccinia (2). In addition, because vaccinia has no known cellular target receptor, instead entering cells via membrane fusion events, it is unlikely that the strategy of retargeting or altering viral tropism will be effective (because it is currently impossible to completely negate natural tropism, an essential component of this approach). However, vaccinia is a large virus and has an extensive array of genes involved in many aspects of viral virulence (3, 6). These can be broadly split into three categories (1) genes whose products function to disrupt or limit the immune response targeting the virus (e.g., cytokine or chemokine binding proteins); (2) genes whose function is to prevent the infected cell from undergoing apoptosis; and (3) genes whose products function to activate cell signaling pathways. Because many cancer cells already display immune dysfunction, loss of apoptotic potential and activated or unregulated cell signaling pathways, these viral genes are often redundant and their deletion produces a tumor-specific or tumor-selective vaccinia strain. Deletions in a number of these genes have been studied and shown to display tumor-targeting potential (these include viral thymidine kinase [TK], vaccinia growth factor [VGF], vaccinia type I interferon binding protein [B18R], double stranded RNA [dsRNA]-binding protein [E3L], and viral serpins [B13R, B22R]) (5, 7). However, the tumor-targeting potential of deletions in many more remain unexplored, and therefore it is possible to envisage an approach of incorporating specific viral gene deletions so as to be complemented by the common mutations within a tumor target. However, probably the most extensively studied tumor-targeting strain of vaccinia contains a double deletion of both the TK and VGF genes (often referred to as vvDD) (8). These deletions create an inherent selectivity for rapidly proliferating tumors with a high percentage of cells in S-phase, and for cells with dysfunctional epidermal growth factor receptor (EGFR) signaling pathways. This means that vvDD is broadly tumor targeting, and it

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has displayed an impressive specificity for tumor cells in vitro and in vivo. Therefore, the vvDD backbone represents a systemically deliverable, effective, and broadly applicable oncolytic vaccinia virus strain that can be used for the study of novel therapeutic transgenes.

2. Materials 2.1. Construction of Oncolytic Vaccinia Strains

1. Cloning plasmids, such as pSC-65 (9). 2. Cell lines CV-1, 143B TK-, and HeLa (ATCC, Manassas, VA). 3. 10-cm tissue culture plate. 4. Lipid vesicle transfection reagent (such as Lipofectamine (Invitrogen, Carlsbad, CA), with specialized media if necessary (e.g., Opti-MEM, Gibco, Carlsbad, CA)). 5. Cell scrapers. 6. Dry ice: methanol bath (add dry ice to an insulated bucket, then carefully add methanol until a slurry is formed). 7. Waterbath at 37°C. 8. 5 mg/mL Bromodeoxyuridine (BrdU) (made fresh and filter sterilized). 9. 96-well tissue culture plates. 10. Inverted microscope. 11. 10 mM Tris HCl, pH 9.0.

2.2. Viral Replication by Plaque Assay

1. 6-well plates. 2. Overlay medium for plaque assay (per 100 mL): 50 mL of 2X DMEM, 2 mL FBS, 48 mL of 3% carboxymethyl cellulose. 3. Crystal violet solution (0.1% crystal violet in 20% ethanol). 4. Aspiration set. 5. Branson Sonifier 250 with cup attachment (or equivalent). 6. 10 mM Tris-HCl pH 9.0.

2.3. Viral Cytotoxicity by Cell Viability Assay

1. 96-well plates.

2.4. Bioluminescence Imaging

1. 30 mg/mL d-luciferin in PBS, filter sterilized (Biosynth, Itasca, IL).

2. MTS assay reagent (Promega, Madison, WI). 3. Enzyme-linked immunosorbent assay (ELISA) plate reader.

2. 5 mg/mL Coelentrazine (Millipore Chemicon, Temecula, CA) in methanol.

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3. Mice, typically obtained from commercial vendors (e.g., Jackson Laboratories or Charles River). 4. IVIS imaging system or equivalent whole animal imaging system (Caliper Life Sciences, Hopkinton, MA) with heated (37°C) stage and appropriate image analysis software (e.g., Living Image, Caliper Life Sciences). 5. 70% Ethanol. 6. Anesthetic (ideally inhaled, such as isoflurane, but may be injectable, such as Avertin). 7. Black-walled, clear-bottom, 96-well plates. 8. Luciferase assay system (Promega).

3. Methods 3.1. Design and Construct of Oncolytic Vaccinia Virus

1. Consideration should be given to both the parental vaccinia strain as well as the choice of gene deletions when designing a backbone vector for gene delivery (see Notes 1 and 2). The location of insert of any foreign transgene is also significant, because this may affect the transgene expression levels (10). The two most common locations used for transgene insertion are (1) intragenic (with no gene deletion) or (2) into the locus of the viral thymidine kinase (TK) gene. 2. A variety of methods have been used to disrupt or delete nonessential viral genes, and vaccinia strains containing deletions in many of these virulence genes have been previously constructed by a variety of investigators. However, because methodologies are nonstandard, they will not be extensively covered in this protocol. Instead, we describe the protocol for the insertion of a foreign transgene into the viral TK gene by homologous recombination as an example (see Note 3). This approach can be adapted to target other viral genes by inserting different regions of homology into the cloning plasmid, but alternative selection procedures may need to be incorporated (e.g., reporter gene, antibiotic resistance, sugar metabolism). 3. Simple cloning strategies can be used to insert the transgene of choice into the multiple cloning site of a cloning plasmid (see Note 4). 4. Grow a CV-1 cell monolayer until 80–95% confluent in a 10-cm tissue culture dish (achieved by splitting a 90% confluent plate 1:2 overnight). 5. Infect with target virus (i.e., parental virus containing all required deletions except for the thymidine kinase deletion)


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at a multiplicity of infection (MOI) of 0.05 plaque forming units (PFU)/cell. 6. After 1.5–2 h incubation at 37°C, transfect the cell layer with the cloning plasmid (see Note 5). 7. Leave infected/transfected CV-1 cells at 37°C, 5% CO2 for 48 h, then scrape cells into media. 8. Freeze/thaw cells and media three times (by transferring between a dry ice/methanol bath and a 37°C waterbath). This crude viral lysate can then be stored at –80°C until needed. 9. Add 0.5 mL of the crude viral lysate to a 90% confluent 10-cm tissue culture dish containing 143B TK- cells (split a 90% confluent 10-cm tissue culture dish 1:3 overnight), and immediately add 1:100 volumes of 5 mg/mL BrdU (see Note 6). 10. After a 48 h incubation, again collect the cells and media and freeze/thaw three times. 11. Add 10 μL/well of second crude viral lysate to each well of the top row of a 96-well plate containing 143B TK- cells (seeded at 1 × 104 cells in 200 μL media/well overnight). Dilute virus 1:4 every row down the plate (i.e., take 50 μL from each well in top row and add to the corresponding well in the row directly below; repeat down the plate. Keep the bottom row uninfected). Add 1:100 volumes of 5 mg/mL BrdU to all wells. 12. Incubate 96-well plates for 48 h (37°C; 5% CO2), then examine the wells individually under an inverted microscope. Mark the wells with a single visible plaque (if no wells contain only one plaque, select the wells with the smallest numbers of plaques). Ideally, when a secondary marker is available for successful recombination, this can then be further applied to the wells containing a single plaque to confirm recombination (to differentiate from spontaneous TK mutation) (see Note 6). Collect the media and cells from two or three of the single-plaque wells, and repeat steps 11 and 12 for further rounds of plaque purification. 13. It is necessary to confirm correct insertion of transgene by polymerase chain reaction (PCR) of DNA collected from infected cells. Primers should be designed to flank transgene insertion site. 14. Expand the purified plaque by addition of crude viral lysate to HeLa cells. Observe the cells under an inverted microscope every 24 h for signs of cytopathic effect (i.e., cell rounding), and when this is evident, aspirate the media, add 10 mM Tris-HCl pH 9.0, and scrape cells into this buffer. Freeze/thaw three times as before; spin the crude viral lysate (300g, 5 min) to pellet the cellular debris, and retain the supernatant. Virus can be further purified if desired, and may be titered by plaque assay (see below).

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3.2. Testing Selectivity of Oncolytic Virus In Vitro

As with all new cancer drugs, a process of in vitro (cell culture) and then in vivo (animal tumor model) screening should be performed to assess the selectivity and antitumor effects of the new viral construct.

3.2.1. Viral Replication Assayed by Plaque Assay

1. A panel of tumor and normal cell lines should be grown to 80% confluence in 6-well plates (see Note 7). Preliminary experiments will need to be performed to ascertain the correct split ratios to obtain cells at the correct confluence. 2. Aspirate media and add 1 mL of the appropriate media/well with 2% FBS (heat inactivated). Then add virus at an MOI of 1.0 PFU/cell. 3. Wait 2 h, then aspirate the media again and add 3 mL/well of fresh media (with 2% FBS). 4. At 24 h intervals, scrape the cells from one well of each 6-well plate into media and collect the cells and media. Freeze/thaw three times as before, then create a dilution series of the crude viral lysate by diluting 1:10 six times (using 10 mM Tris-HCl, pH 9.0 as diluent). 5. Plate BS-C-1 cells in 6-well plates (2 × 106 cells/well) overnight. Aspirate the media and add 900 μL of media with 2% FBS per well. Sonicate dilutions of crude viral lysate for 30 s at full power. Add 100 μL of dilutions two to six of the crude viral lysate to five wells of the 6-well plate (leave the last well uninfected as a control). 6. After a 2-h incubation at 37°C, aspirate the media and add 2 mL/well of overlay media. 7. Incubate plates at 37°C for 72 h before aspirating the overlay and adding 1 mL/well of crystal violet solution. 8. Leave the plates at room temperature for 2 h before aspirating the crystal violet and inverting the plates to dry. 9. Count the plaques in all wells that display between 10 and 150 plaques, and from the dilution used, calculate the initial concentration of virus (in PFU/mL). The ratio of normal to tumor viral replication will give a value for the selectivity of the virus (Fig. 1a). The oncolytic construct should be compared with wild-type virus.

3.2.2. Determination of Viral Cytotoxicity by Cytopathic Effect

1. Seed 96-well plates with 200 μL of media with tumor or normal cell lines (see Note 7) and incubate overnight to allow cells to attach (preliminary experiments will be needed to determine the correct seeding density to give cells at 80–90% confluence). Do not seed the outer wells on the plate, instead, fill these wells with 200 μL PBS. 2. Add virus to the first column of wells containing cells at an MOI of 10,000 PFU/cell and dilute the virus across the plate


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Fig. 1. (a) Comparison of replication of two candidate viral strains in a normal and a tumor cell line. Viral titers were determined 72 h after infection, and show that strain 1 has a high level of tumor-selective replication (arrow). (b) Relative cytopathic effect of different viral stains in a tumor cell line (read by viability assay 72 h after infection). The viral titer (PFU/cell) required to reduce cell viability by 50% can be compared between strains. Strain 1 displays increased cytotoxicity in this cell line (arrow).

using 1:4 dilutions (i.e., remove 50 μL/well and add to the corresponding well in the next column). Leave the final column of cells uninfected. Keep the outer wells with PBS only. Perform each dilution series in triplicate. 3. Leave the plates at 37°C for 72 h, then aspirate the media, add fresh media with no serum to each well, and add MTS (Promega) according to manufacturer’s guidelines (earlier time points may also be examined). 4. Read the MTS color change after 2 h produced by the ATP:ADP ratio using an ELISA plate reader. 5. Calculate the plaque-forming units per cell of virus needed to kill 50% of the cell layer from the viability (MTS) assay. By defining the uninfected cells as 100% viable, and PBS control wells as 0% viable, a value for the percentage viability can be determined for all treatment wells. These can be plotted to determine the plaque-forming units per cell needed to reduce viability by 50%. Ratios of values for tumor and normal cells for different control and oncolytic viruses can be used to determine the therapeutic index of candidate oncolytic agents (Fig. 1b).

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3.3. In Vivo Screening Using Bioluminescence Imaging

Noninvasive whole animal imaging provides several advantages over traditional animal models. The ability to repeatedly image the same animals means that they become their own internal controls, and also dramatically reduces the number of animals required to run experiments. In addition, unexpected and expected signals detected during whole animal imaging can be used to define time points and tissues selected for ex vivo analysis, and therefore better define the routes of infection, timings of therapeutic responses, and any unexpected infection of nontumor tissues. 1. Viral biodistribution assays will require viral strains constructed to express luciferase as a reporter gene (see Note 8). These should be first checked in vitro to ascertain the levels of bioluminescence produced. 2. Consideration should also be given to the animal model to be used. Because of the adsorption and diffraction limitations of optical imaging, tissues greater than 1 cm deep cannot be imaged. This means that the ideal species to use will be the mouse. The choice of mouse strain may be limited by the availability of cell lines, but, in general, white mice are preferable (because the pigmentation in the skin of black or brown mice further absorbs emitted light). Some normally dark mice (e.g., C57BL/6) have been bred as albinos. Animal fur also diffracts light, and therefore shaving or use of depilatory cream may enhance a weak signal. Alternatively, athymic nu– /nu– mice may be used. Pilot experiments should always be run in vivo to first characterize the model (see Note 9). 3. Deliver the luciferin substrate to mice. Imaging of insect luciferase will typically require injection of 200 µL of 30 mg/mL luciferin intraperitoneally, 5–20 min before imaging, coelentrazine for Renilla luciferase will typically be injected intravenously (tail vein or retino-orbital injection), with 100 µL of 5 mg/mL injected no longer than 2 min before imaging. 4. Anesthetize the mice. The exact order of delivery of substrate and anesthesia will depend on the system being used. The choice of anesthetic may also be important. For instance, mice take longer to recover from injectable anesthetics and therefore fewer imaging time points can be used. Inhaled anesthetics (such as isoflurane, 3%) are therefore typically preferable, however, there is some evidence that these may produce a low level of background luminescence, and thus may not be suitable for detection of weak signal, especially from the lungs. 5. Arrange the mice in the imaging chamber so that the expected source of signal is closest to the top of the animal (e.g., ventral view for imaging major organs, dorsal view for imaging the spine, left side view for imaging the spleen). The imaging stage should be wiped with 70% ethanol before use, and a piece of black paper should be placed on the stage to protect it. Dividers may be used


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between the animals (if multiple animals are to be imaged at once) to prevent strong signal from one animal reflecting off its neighbor. All non-essential materials should be removed from the imaging chamber. The stage should be heated to 37°C, and the animals should be checked for the level of anesthesia for 30 s before closing the chamber (see Note 10). 6. Take the image. Several variables may be adjusted to improve the quality of the image, including height of stage, focus, length of exposure, binning, and aperture setting (see Note 11). To help locate the source of a signal, it is recommended that the mice are repositioned and reimaged from different angles. Recent advances in software available for image analysis may include three-dimensional (3D) reconstructions, whereby animals are imaged consecutively at different wavelengths, and the relative levels of blue and red light emitted can be used to determine the depth of the signal. 7. Recover the mice. Once imaging is complete, the mice should be recovered on a heated stage until they are capable of selfrighting. 8. Image analysis. A variety of software packages exist for image analysis, including quantification, such as Living Image (Caliper Life Science). Viral signal produced from an implanted tumor may be compared with normal regions for comparison of therapeutic index of different strains. 9. Verification of imaging results. It is important to verify patterns of gene expression and the source of any signals ex vivo in a subset of animals. The bioluminescence signal from firefly luciferase will remain strong for up to 45 min postmortem, and therefore individual organs can be imaged to verify the tissue of origin of any signal (although the signal no longer remains quantifiable). In addition, it should be remembered that the bioluminescent signal represents the biodistribution of gene expression and not viral delivery. Luciferase expression can be verified ex vivo using luciferase assay systems (on frozen, ground tissues, and normalized to protein level), and viral genomes quantified by quantitative PCR (Q-PCR) (or viral infectious units quantified by plaque assay, as above) to determine any differences between viral delivery and gene expression in different tissues.

4. Notes 1. A large number of distinct vaccinia strains have been described. These include strains that were used in different geographical regions during the smallpox eradication program (e.g., Wyeth/

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New York City Board of Health, Lister, Copenhagen, Tian Tan, USSR); these have the advantage of having been used extensively in human subjects (by intradermal delivery); other strains have been developed as laboratory strains (e.g., Western Reserve, IHD-J), the majority of our knowledge regarding the effects of gene deletions have been obtained from these; finally, some strains have been designed as highly attenuated and thus safer versions of vaccine strains (e.g., MVA, NYVAC), but because these do not replicate effectively in human cells they are not considered oncolytic. A variety of virulence genes have been found to be expressed or functional in only a subset of the vaccine and laboratory strains available, and therefore some research is necessary when choosing a parental strain. Some of the most commonly used strains (e.g., Wyeth, Western Reserve, and Copenhagen) have been completely sequenced. 2. The choice of gene deletions will depend on a number of factors, including the type of cancer being targeted (tissue or cell of origin, primary or metastatic, etc.), the route of delivery being incorporated, and the therapeutic transgene being studied. Some genes (e.g., secreted decoy receptors) may interfere with the function of some transgenes. 3. The most common approach to inserting a foreign transgene into vaccinia involves homologous recombination into the viral thymidine kinase gene. This gene was traditionally targeted because its deletion did not effect viral replication in culture, but more recently it has been shown that this deletion does attenuate the virus in vivo, and is itself tumor targeting. The deletion of viral TK creates a dependence on cellular thymidine kinase expression, which is not expressed in quiescent cells, is transiently expressed during the cell cycle of proliferating cells, but is constitutively expressed in most malignant cells. 4. A variety of different cloning plasmids have been constructed that can be used to insert a multiple cloning site (MCS) into the locus of the viral thymidine kinase gene. One of the most commonly used is pSC65. Because vaccinia carries its own transcription machinery, it uses its own promoters and promoter elements, and therefore any transgene will need to be driven by one of these. However, a number of vaccinia promoters have been well described, these may be expressed early or late during the viral replication cycle or may contain elements of both early and late promoters for constitutive transgene expression. A wide range of promoter strengths is also available. The pSC65 plasmid, for example, contains two MCS, one in front of the natural viral p7.5 promoter and the other driven by the synthetic pSE/L promoter. However, because this plasmid only inserts the foreign transgene(s) into the site


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of the thymidine kinase gene, it may not be suitable for the creation of clinical vectors, where some additional deletion of the viral virulence gene would be preferable. 5. A variety of transfection reagents and protocols are available. CaCl2 transfection will work effectively with CV-1 cells, but for ease of use and reproducibility, it is recommended that a commercial lipid vesicle reagent (e.g., Lipofectamine, Invitrogen) be used. Specialist media (e.g., Opti-MEM, Gibco) may also be required. Conduct the transfection according to the manufacturer’s guidelines. 6. Because BrdU is a known mutagen, it is possible that secondary mutations will be incorporated into the virus during selection. This is especially an issue when a virus is being constructed for future clinical use. BrdU is only appropriate for selection for loss of viral TK function. Alternative selection strategies may avoid any issues relating to the use of BrdU, or may be used after insertion of transgenes into other sites. Incorporation of a reporter gene into the cloning plasmid (e.g., luciferase, green fluorescent protein [GFP], lacZ) will allow selection of viral plaques that express these genes (but again, this will not be acceptable for clinical vectors). Alternatively, a selection or detection assay for the inserted transgene may be possible (e.g., ELISA, antibody staining, or functional assay). 7. Tumor cells to be incorporated into these assays should be selected based on the desired target for the virus (e.g., tumor type or property of tumor, for instance, loss of interferon [IFN] response; EGFR mutations, etc.); “normal” cells may include immortalized, nontransformed cell lines for ease of use, but should also include some primary cell lines, ideally to closely match the target tumor cells (e.g., normal hepatocytes to compare with hepatocellular cancers). Cells may be grown under specialized conditions to further examine methods of selectivity (e.g., pretreated with IFN, grown to confluence). 8. The choice of luciferase enzyme incorporated for viral labeling is important. To achieve maximum light output from within a living organism, a significant part of the spectrum of the light produced from any luciferase should be greater than 600 nm, because light below this wavelength will be adsorbed by hemoglobin. The most commonly used luciferase is that of the firefly (Promega), and this produces light with a spectral peak of 560 nm, however, at 37°C this shifts to 590 nm, making it suitable for in vivo use (11). Other insect luciferases, such as those of the click beetle or railroad worm produce a variety of spectra and although green luciferases are often brighter in vitro, the massive adsorption at these wavelengths in vivo makes them less useful. Several luciferases have also been cloned from marine organisms, such as Renilla and Aequorea, but these produce

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blue–green light (peak of 475 nm). However, the advantage of Renilla luciferase is that is uses a different substrate (coelentrazine) to the insect luciferases (luciferin) and therefore dual bioluminescence imaging is possible. In addition, marine luciferases do not require ATP, and therefore may also be able to function extracellularly. However, the coelentrazine substrate is unstable, meaning that a much higher background signal is produced, and the substrate only remains within an animal for a short period of time. This can be used as an advantage when dual imaging of both firefly and Renilla luciferases in a single animal is attempted, because it is possible to image the Renilla first after addition of coelentrazine, and then to image the firefly luciferase (as soon as 1 hr later) after addition of luciferin. Both substrates are nontoxic, are rapidly distributed throughout the body, and can cross most membranes, including cellular membranes and the blood–brain barrier. 9. An initial pilot experiment is always recommended. Verify the tumor growth patterns and take rates for the different injected doses of cancer cell lines. In addition, dosing by injecting limiting numbers of cells infected with a labeled virus, followed immediately by imaging, can be used to predict limits of detection or to help quantify signals. Finally, sequences of images should be taken at different times after addition of luciferin substrate to verify the optimal timing of imaging. The anesthetic to be used should also be tested to ensure the minimum required dose is used, and, if possible, to verify that the anesthetic used does not interfere with the study. Because most injectable anesthetics are metabolized by the subject, they may effect metabolic pathways, stress responses, or oxygen use. In addition, protocols for all planned experiments should first be approved by institutional review boards. 10. Many plastics and other materials (including animal bedding, chow, and dander) will produce phosphorescence, and therefore careful selection of any item used in the imaging chamber is important. In particular, the anesthesia nose cones, the paper used to protect the stage, the plastic dividers, and the animal shields should always be carefully tested; and all bedding, dander, and chow should be cleaned from the stage before imaging. 11. The height of the stage should be adjusted so that the animals to be imaged best fit the field of view with minimal empty space. It is recommended to initially try a 60-s image with medium binning if the strength of the signal is not known. If this saturates the camera, then quantification is not possible, and therefore shorter exposure times should be tried and smaller binning used. If at 1 s, minimum binning


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image still saturates the CCD camera, then the aperture may be reduced. If the original image produces weak or no signal, then increased exposure times (up to 5 min) and increased binning may be tried. Images longer than 5 min will likely not improve the signal because the background will also increase. References 1. Smith, G. L., and Moss, B. (1983) Infectious poxvirus vectors have capacity for at least 25 000 base pairs of foreign DNA. Gene 25, 21–8. 2. Moss, B. (2001) (D.M., K., Fields, B. N., and Howley, P. M., Eds.) in “Field’s Virology”, pp. Ch.84, Lippincott-Raven, Philadelphia. 3. Buller, R. M., and Palumbo, G. J. (1991) Poxvirus pathogenesis. Microbiol Rev 55, 80–122. 4. Moss, B. (1991) Vaccinia virus: a tool for research and vaccine development. Science 252, 1662–7. 5. Thorne, S. H., and Kirn, D. H. (2004) Future directions for the field of oncolytic virotherapy: a perspective on the use of vaccinia virus. Expert Opin Biol Ther 4, 1307–21. 6. Smith, G. L., Symons, J. A., Khanna, A., Vanderplasschen, A., and Alcami, A. (1997) Vaccinia virus immune evasion. Immunol Rev 159, 137–54. 7. Thorne, S. H., Hwang, T. H., and Kirn, D. H. (2005) Vaccinia virus and oncolytic virotherapy of cancer. Curr Opin Mol Ther 7, 359–65.

8. McCart, J. A., Ward, J. M., Lee, J., Hu, Y., Alexander, H. R., Libutti, S. K., Moss, B., and Bartlett, D. L. (2001) Systemic cancer therapy with a tumor-selective vaccinia virus mutant lacking thymidine kinase and vaccinia growth factor genes. Cancer Res 61, 8751–7. 9. Chakrabarti, S., Sisler, J. R., and Moss, B. (1997) Compact, synthetic, vaccinia virus early/late promoter for protein expression. Biotechniques 23, 1094–7. 10. Coupar, B. E., Oke, P. G., and Andrew, M. E. (2000) Insertion sites for recombinant vaccinia virus construction: effects on expression of a foreign protein. J Gen Virol 81, 431–9. 11. Zhao, H., Doyle, T. C., Coquoz, O., Kalish, F., Rice, B. W., and Contag, C. H. (2005) Emission spectra of bioluminescent reporters and interaction with mammalian tissue determine the sensitivity of detection in vivo. J Biomed Opt 10, 41210.

Chapter 33 Tumor-Targeted Salmonella typhimurium Overexpressing Cytosine Deaminase: A Novel, Tumor-Selective Therapy Ivan King, Martina Itterson, and David Bermudes Summary The ideal anticancer regimen is one that is specific for cancer cells with limited toxicity to normal tissues. Genetically modified, nonpathogenic Salmonella offer a potential way to induce direct tumoricidal activity or to deliver tumoricidal agents to tumors. An attenuated strain of Salmonella typhimurium, called VNP20009, and its derivative TAPET-CD (which expresses Escherichia coli cytosine deaminase) are highly selective for tumor tissue and can deliver therapeutic proteins preferentially to tumors in preclinical models. Both VNP20009 and TAPET-CD have been investigated successfully in Phase 1 clinical trials in cancer patients. Key words: cytosine deaminase, 5-fluorocytosine, 5-fluorouracil, Salmonella, TAPET-CD, tumor, VNP20009.

1. Introduction It was first reported in 1997 that live Salmonella injected systemically into mice migrates to and preferentially multiplies within implanted tumors, achieving tumor-to-normal tissue ratios of up to 1,000:1 (1). Tumor growth inhibition is observed in mice receiving the live Salmonella. This tumor-preferential accumulation offers the opportunity to engineer Salmonella to act as a tumoricidal or drug delivery agent, while sparing normal tissues from toxicity. Because wild-type Salmonella induces cytokine-mediated septic shock and death, it is advantageous for the microorganism to be attenuated for clinical use. An innocuous clinical strain of S. typhimurium, called VNP20009, was developed by deleting


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the msbB gene, responsible for adding a terminal myristoyl group to lipid A. This mutation greatly reduces the bacterium’s ability to elicit tumor necrosis factor-α, the cytokine that causes septic shock (2). As an additional safeguard, the purI gene is deleted in VNP20009, making the engineered bacterium dependent on an external source of adenine to survive and multiply. The deletion of purI insures that VNP20009 cannot survive in the environment. VNP20009 is also resistant to EGTA, a property now know to be conferred by a large spontaneous rearrangement between two IS200 elements, referred to as the Suwwan deletion (3). Like the parent strain, VNP20009 maintains the property of tumor-selective multiplication, with observed tumor-tonormal tissue ratios also reaching about 1,000:1 (4). In addition, the antitumor effects of VNP20009 are as good as or better in preclinical models than currently used anticancer agents, such as cyclophosphamide and cisplatin (5). Attenuated strains of Salmonella provide advantages as anticancer vectors. Compared with standard chemotherapy drugs, Salmonella is easily manipulated to carry foreign genes; thrives in hypoxic conditions present in tumors; moves into tumors against diffusion and pressure gradients; can be injected intravenously, intraperitoneally, or intratumorally; targets a broad range of solid tumors; and is readily eliminated from the body by antibiotics (6). Tumor amplified protein expression therapy (TAPET)cytosine deaminase (CD) is generated by incorporating the E. coli CD gene into VNP20009. TAPET-CD not only multiplies within tumors, but also expresses CD inside tumors. The enzyme produced locally in tumor tissues converts the nontoxic prodrug 5-fluorocytosine (5-FC) to cytotoxic 5-fluorouracil (5-FU). Noninvasive 19F-magnetic resonance spectroscopy of rodents, which bear human tumors and are treated with TAPET-CD, confirms the in vivo conversion of 5-FC to 5-FU (7). The selective high concentration of 5-FU in tumors makes the drug more effective and less toxic to peripheral normal tissues. Coadministration of TAPET-CD and 5-FC induces prolonged, high concentrations of 5-FU in tumors, resulting in a reduction in tumor size greater than that induced by TAPET-CD or 5-FC alone (8). A Phase 1 trial of VNP20009 in terminal cancer patients validated the safety of the attenuated Salmonella as well as its ability to colonize some tumors (9). In a pilot trial in three refractory cancer patients, intratumoral injection of TAPET-CD proved safe, and two patients showed bacterial colonization of tumors that persisted for 15 days after injection. Patients also received 5-FC, which was converted to 5-FU in the two patients with bacterial colonization (10). These early results support the concept that TAPET-CD delivers a functional gene that converts an inactive prodrug to an active cytotoxic drug in tumors. We describe here the construction of TAPET-CD and its potential for use with 5-FC as a cancer treatment.

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2. Materials 2.1. Cloning and Expression of Cytosine Deaminase (CD)

1. Salmonella strain VNP20009 is derived from a hyperinvasive strain YS72, a pur-xyl− hypersensitive mutant (11). 2. E. coli HB 101 is obtained from the American Type Culture Collection (ATCC, Rockville, MD). 3. Plasmid pBluescript II KS (Stratagene, LaJolla, CA). 4. PTrc99a (Pharmacia-Upjohn, Bridgewater, NJ). 5. MsbB medium: Modified LB containing 10 g tryptone, 5 g yeast extract, 2 mL 1 N CaCl2, and 2 mL of 1 N MgSO4 per liter, adjusted to pH 7 using 1 N NaOH. 6. NcoI, XbaI (New England BioLabs, Ipswich, MA).

2.2. CD Activity Assay

1.

14

C-5-Fluorocytosine and chemicals, Brea, CA).

14

C-5-Fluorouracil (Moravek Bio-

2. Instant Imager; Electronic Autoradiography (Perkin Elmer, Waltham, MA). 2.3. Construction of pCVD442-msbB Vector

1. AvaI, SacI, SalI, SphI (New England BioLabs).

2.4. Cloning trc-CD into pCVD442-msbB in E. coli Strains DH5 lpir and SM10 lpir and Transfer to Salmonella

1. SalI, SspI, SwaI, T4 ligase (New England BioLabs).

2.5. Transfer of the Chromosomally Integrated msbB-CD into VNP20009 to Generate the Strain VNP20029 (TAPET-CD)

1. Modified LB containing 10 g tryptone, 5 g yeast extract, 2 mL of 1 N CaCl2, and 2 mL of 1 N MgSO4 per liter, adjusted to pH 7 using 1 N NaOH, for growing msbB− strains.

2.6. Quantitation of TAPET-CD Accumulation and 5-FC/5-FU Conversion in Tissues

1. Murine B16-F10 melanoma cells are obtained from Dr. I. Fidler (M.D. Anderson Cancer Center, Houston, TX).

2. Difco MacConkey agar (VWR, West Chester, PA). 3. Carbenicillin (BioWorld, Dublin, OH).

2. Bacteriophage P22 is used for transductions in LB lacking NaCl (see Note 1) (12). 3. Carbenicillin (BioWorld).

2. C38 colon carcinoma cells are obtained from the National Cancer Institute (NCI, Frederick, MD). 3. Dulbecco’s modified Eagle’s medium (Gibco/Invitrogen, Carlsbad, CA) supplemented with 10% fetal bovine serum (FBS, Gibco/Invitrogen) 4. Trypsin-EDTA and phosphate-buffered saline (PBS, Sigma, St. Louis, MO).

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5. 5-Fluorocytosine and 5-Fluorouracil (Sigma, St. Louis, MO). 6. High-performance liquid chromatography (HPLC) column (Phenomenex, Prodigy ODS, Torrance, CA). 2.7. Antitumor Activity of TAPET-CD in Animals

1. 5-Fluorocytosine (Sigma). 2. Female C57BL/6 mice are obtained from Charles River Laboratories (Wilmington, MA). Animals used in the studies are as uniform in age and weight as possible. They are approximately 8 weeks of age, and body weights range from 19 to 21 g. All animals are kept in a well-ventilated room in which a 12-h light/12-h dark photoperiod is maintained. Room temperature is maintained at 72 ± 2°F.

3. Methods TAPET-CD was derived from VNP20009, originally modified from wild-type S. typhimurium (Fig. 1). The construction of VNP20009, an attenuated S. typhimurium with the deletion of msbB and purI genes, has been previously reported (13). 3.1. Cloning and Expression of Cytosine Deaminase (CD)

1. Total genomic DNA is prepared from E. coli HB 101 by detergent lysis, phenol/chloroform extraction, and ethanol precipitation (14). 2. CD is cloned by polymerase chain reaction (PCR) using primers based on the complete sequence for E. coli codA gene that encodes CD (15), GenBank# S56903: forward: 5¢-GCTAACCATGGCGAATAACGC-3¢ (which contains an NcoI site) and reverse 5¢-CTAGGTCTAGACCAGTCGTTCAA-3¢ (which contains a XbaI site). 3. Transformants are screened using a probe generated from the original PCR product by random priming, using [α-32P] dCTP, and positive clones sequenced to confirm that the correct DNA is cloned. 4. A CD assay described in Subheading 3.2 is used to confirm the presence of enzyme activity in the clone.

3.2. CD Activity Assay

1. A fresh overnight culture of bacteria containing pTrc99a-CD is diluted 1:5 with msbB medium and allowed to grow for 2 h at 37 ± 2°C with shaking (~250 rpm). 2. After growth, a 100-μL aliquot is removed and placed into a fresh microfuge tube.

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Strain /plasmid

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Process PCR CD gene

Contains cloned CD under control of trc promoter

pTrc99a-CD plasmid Subclone into pCVD442-msbB Suicide vector

trc CD contained in ΔmsbB

DH5α λpir PCVD442-CD Transform SM10 λpir

trc CD contained in ΔmsbB Capable of plasmid exchange for chromosomal integration

SM10 λpir PCVD442-CD Mate to Salmonella YS50102

trc CD on chromosome Sucs, carbr, tetr

YS50102 ΔmsbB-CD Transduce to VNP20009

trc CD on chromosome Sucs, carbr, tetr, ΔmsbB, purI-, xyl-,

VNP20009 ΔmsbB-CD

Suwwan, EGTAr

Sucrose resolution

trc CD on chromosome Sucr, carbs, tets, ΔmsbB, purI-, xyl-,

VNP20029 (TAPET-CD)

Suwwan, EGTAr

Fig. 1. Derivation of VNP20029 (TAPET-CD) from VNP20009 using a cloned CD under control of the trc promoter.

3. The following reaction mix is added per sample: 1.5 μL of 10 mM 5-FC (150 μM final concentration), 1.5 μL of 14 C-5-FC (0.1 μCi/μL; 0.15 μCi final per sample). 4. The reaction is incubated at 37 ± 2°C for 1 h. 5. After incubation, the reaction is centrifuged at 16,000 g for 5 min to remove the cells. 6. A 4-μL aliquot of the supernatant is spotted onto a TLC plate (silica gel) with separate lanes for 14C-5-FC and 14C-5-FU standards, and the reaction products are separated by running in 80:20 CH3Cl:ethanol for approximately 25 min until the solvent front is approximately 1 cm from the top of the TLC plate. 7. The plate is removed from the TLC chamber, allowed to air dry for about 10 min, and then quantified by autoradiography on an Instant Imager.

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3.3. Construction of pCVD442-msbB Vector

1. A plasmid capable of undergoing homologous recombination with the ΔmsbB gene in the chromosome of strain VNP20009 is constructed in the suicide vector pCVD442 (16). Primers for PCR are designed that generate portions of the 5¢ and 3¢ sections of the msbB deletion occurring in VNP20009 as two separate products: msbB-5¢: forward 5¢-GTGTGAGCTCGATCAACCAGCAAGCCGTTAACCCTCTGAC-3 ¢ and reverse 5¢-GTGTGCATGCGGGGGGCCATATAGGCCGGGGATTTAAATGCAAACGTCCGCCGAAACGCC GACGCAC-3¢; and msbB-3¢: forward 5¢-GTGTGCATGCGGGGT TAATTAAGGGGGCGGCCGCGTGGTATTGGTTGAACCGACGGTGCTCAT GACATCGC-3¢ and reverse 5¢-GTGTCTCGAGGATATCATTCTGGCCTCTGA CGTTGTG-3¢. 2. These primers also contain outer SacI (5¢) and AvaI (3¢) restriction endonuclease sites to facilitate cloning into the SacI and SalI sites of pCVD442 when these two fragments are joined via a common SphI site and generate internal Not1, PacI, SphI, SfiI, and SwaI, to facilitate cloning of DNA fragments into the ΔmsbB for stable chromosomal integration without antibiotic resistance. This vector is referred to as pCVD442-msbB in Fig. 2.

3.4. Cloning trc-CD into pCVD442-msbB in E. coli Strains DH5 lpir and SM10 lpir and Transfer to Salmonella

1. To clone trc-CD into pCVD442-msbB, the trc-CD plasmid DNA is partially digested with SspI, and the pCVD442-ΔmsbB is digested with SwaI. 2. The appropriate DNA is purified and a ligation reaction containing these two components is performed using T4 ligase. 3. The ligation reaction is then transformed to DH5 λpir and colonies are screened for the presence and orientation of the trc-CD insert by restriction digestion of purified plasmids with SalI (17). 4. The transcriptional orientation coinciding with that of the msbB gene is chosen for further analysis (see Note 1). 5. The DH5 λpir clone is transformed into the strain SM10 λpir (16) and the plasmid designated pCVD442-CD. 6. Ten colonies of SM10 λpir are screened for CD activity, as described in Subheading 3.2, and one of the SM10 λ pir clones is chosen for use as a mating donor to Salmonella strains. 7. SM10 λ pir containing the pCVD442-CD is mated to a Salmonella strain YS50102, a spontaneous derivative of the tetracycline-resistant strain YS82 with enhanced resistance to Difco MacConkey agar by standard methods and selected for on plates containing 50 μg/mL carbenicillin and 300 μg/mL streptomycin (11, 12).


1

Δ msbB

pCVD442-CD

5' DNA

bla

sacB

mobRP4

oriR6K

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Δ msbB 3' DNA

trc CD

Δ msbB 3' DNA 2

tet

disrupted chromosomal copy of msbB in YS50102 Integration forms pseudodiploid (sacB+, ampR, tetR) transducable to other strains

Δ msbB 5' DNA

P1

P2

Δ msbB

Δ msbB

3' DNA

5' DNA

trc CD

3

Δ msbB 3' DNA tet A'

A Selection for homologous recombination/excision (sacB-, ampS, tetS)

Δ msbB 5' DNA 4

P1

P2

Δ msbB 3' DNA

trc CD A A'

Fig. 2. (1) pCVD442-CD vector derived by insertion of trc-CD into pCVD44-msbB, (2) homologous recombination with the ΔmsbB chromosomal copy in Salmonella YS50102, (3) chromosomal integration in Salmonella YS50102, and after phage transduction to strain VNP20009, (4) sucrose resolution resulting in strain VNP20029 (TAPET-CD). oriR6K, the plasmid origin of replication; mobRP4, the mobilization element for the plasmid to be transferred from one strain to another; bla, the beta-lactamase gene, which confers sensitivity to β-lactam antibiotics such as carbenicillin and ampicillin; sacB, the gene that confers sensitivity to sucrose. Note: not drawn to scale.

8. The resulting YS50102-CD clones are checked for the trc-CD gene by PCR using the CD primers described in step 3.1.2 and shown as P1 and P2 in Fig. 2, respectively: forward 5¢-GCTAACCATGGCGAATAACGC-3¢ and reverse 5¢-CTAGGTCTAGACCAGTCGTTCAA-3¢. 3.5. Transfer of the Chromosomally Integrated msbB-CD into VNP20009 to Generate the Strain VNP20029 (TAPET-CD)

1. Using bacteriophage P22 (mutant HT105/1 int-201), VNP20009 is transduced to carbenicillin resistance using strain YS50102-CD as the donor. The presence of the bla and sacB genes from pCVD442 allows the selection of a carbenicillinr, sucroses strain denoted VNP20009 ΔmsbB-CD, which contains both the ΔmsbB and ΔmsbB-CD genes. 2. Strain VNP20009 msbB-CD is plated on LB sucrose to select a sucroser carbenicillins derivative, which removes the ΔmsbB gene and leaves the ΔmsbB-CD gene (16). The LB-sucrose agar plates are made without NaCl, and the plates are incubated at 30°C (see Note 2).

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3. After the growth of colonies on these plates, they are gridded to an msbB plate and replica plated to either carbenicillin- or sucrose-containing plates to detect the presence of a clone that lacks both the antibiotic and sucrose markers (see Note 3). 4. The resulting clones are checked for the presence of the CD gene by PCR. One derivative containing the chromosomally integrated trc-CD, but lacking sucrose sensitivity and carbenicillin resistance, is denoted as VNP20029, also known as TAPET-CD. 3.6. Quantitation of TAPET-CD Accumulation and 5-FC/5-FC Conversion in Tissues

1. B16-F10 cell lines are grown in Dulbecco’s modified Eagle’s medium supplemented with 10% fetal bovine serum at 37°C in a humidified atmosphere of 5% CO2. 2. At approximately 80% confluence, cells are detached from the flasks by addition of 2 mL trypsin–EDTA, resuspended in 25 mL phosphate-buffered saline (PBS), and transferred into a 50-mL Falcon conical centrifuge tube. 3. Cells are pelleted by centrifugation at 4°C for 5 min at 800 rpm in a Beckman GS-6R refrigerated centrifuge. 4. The supernatant is discarded, and the cell pellet is resuspended in PBS. The tumor cell suspension is kept on ice until implantation into mice. 5. After B16-F10 melanoma tumor implantation, animals are inoculated intravenously with 1 × 106 cfu of TAPET-CD on Day 14. 6. Mice are intraperitoneally administered 300 mg/kg of 5-FC on Day 17, 3 days after TAPET-CD injection. 7. Mice are killed by CO2 inhalation at 10, 30, 90, 270, and 360 min after 5-FC inoculation. 8. Tissues, including tumor, liver, spleen, brain, serum, and bone marrow, are weighed and homogenized in PBS. 9. Bacteria are quantitated by plating serial dilutions of the homogenates onto msbB plates, incubating overnight at 37°C, and counting bacterial colonies. 10. The conversion of 5-FC to 5-FU in tissues, including tumor, liver, spleen, brain, serum, and bone marrow, is determined by HPLC analysis (Table 1). The detection limit for 5-FC and 5-FU is 0.5 μg/g and 0.2 μg/g, respectively.

3.7. Antitumor Activity of TAPET-CD in Animals

1. C38 tumor tissue is aseptically dissected from a tumor-bearing C57BL/6 mouse. 2. The tumor is mechanically minced into 3- to 5-mm3 pieces, and transplanted subcutaneously with a 16-gauge trocar needle into the right flank of C57BL/6 mice that are under methoxyflurane anesthesia.


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Table 1 Tissue distribution of 5-FU in mice after treatment with 5-FC, TAPET-CD plus 5-FC, and 5-FU Amount of 5-FU in tissue (mg/g tissue) Serum

Liver

Tumor

Brain

Bone marrow

Spleen

5-FC alone

0

0

0

0

0

0

TAPET-CD + 5-FC

0

0

19

0

0

0

5-FU alone

10

3

4

0.6

2

4

Mice were implanted with B16-F10 melanoma and were administered intravenously with TAPETCD at 1×106 cfu/mouse 14 days after tumor implantation. Tissues were collected 30 min after mice received a single dose of 5-FU (60 mg/kg) administered intraperitoneally. For 5-FC and 5-FU groups, PBS instead of TAPET-CD was administered. Three mice were used for each group

3. Fifteen days after transplantation, when C38 tumors have grown to a volume of approximately 300 mm3, the mice are randomly divided into four groups of ten animals. 4. Groups 1 and 3 receive PBS (0.1 mL). 5. Groups 2 and 4 are injected in the tail vein with TAPET-CD at 1 × 106 cfu/mouse. 6. Groups 3 and 4 receive daily three intraperitoneal injections of 5-FC at a dose of 300 mg/kg on Days 19 through 40. 7. Tumor volume is measured in three dimensions twice weekly and calculated with the formula L × H × W/2, where L, H, and W represent length, height, and width, respectively. Tumor volume is presented as mean ± standard deviation, and the Student’s t test is performed for statistical analysis. 8. The tumors in the TAPET-CD/5-FC-treated animals are significantly smaller than in either the 5-FC or TAPET-CD groups (8).

4. Notes 1. The configuration of the upstream and downstream sections of the msbB gene, which constitute the targeting portion for homologous recombination of the pCVD-442-msbB, are oriented transcriptionally in the same direction as the other elements in the pCVD-442 plasmid. It has been noticed by us and others that the integration and/or resolution efficiency of plasmids with this type of arrangement are superior

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to non-transcriptionally coordinated plasmids. In addition, when the trc-CD is cloned into the vector using blunt-end cloning, we also select clones for uniformity of their transcriptional orientation and have greater efficiency, although we have achieved subsequent success in obtaining resolved clones when the insert is not coordinated. 2. Phage are grown on donor bacteria by diluting an overnight culture of the donor strain 1:5 in LB medium and adding P22 phage to a final concentration of 5 × 106 phage/mL. This mixture is grown overnight with aeration and treated with a few drops of chloroform for 5 min at 37°C, and debris is removed by centrifugation for 10 min at 6,000g. For transductions, this donor phage is used in a 1:20 phage-to-bacteria ratio to infect the recipient strain, consisting of a freshly growing culture of about 108/mL in LB broth. After a 20-min preadsorption period, dilutions are plated onto selective plates. 3. In the transductions and subsequent sucrose resolution of partial diploids, we sometimes are unable to obtain resolved clones, despite repeated attempts. Resolution from independently transduced clones sometimes overcomes this problem. In addition, we find that prescreening partial diploid clones is essential for identifying those with the correct properties. The prescreen involves gridding patches of independent, partial diploids to the sucrose-containing medium and growing them at 30°C. Clones with a propensity to resolve correctly show irregular “fuzzy” edges. This method allows the prescreening of large numbers of colonies.

Acknowledgements The authors thank Brooks Low, Samuel Miller, and John Pawelek for helpful suggestions and Kimberly Troy, Jeremy Pike, and Caroline Clairmont for technical assistance. This work was supported by Vion Pharmaceuticals, New Haven, CT. References 1. Pawelek, J.M., Low, K.B., and Bermudes, D. (1997) Tumor-targeted Salmonella as a novel anticancer vector. Cancer Res. 57, 4537–4544. 2. Bermudes, D., Low, B., and Pawelek, J. (2000) Tumor-targeted Salmonella: Highly selective delivery vectors. Adv. Exp. Med. Biol. 465, 57–63.

3. Murray, S.R., Suwwan de Felipe, K., Obuchowski, P.L., Pike, J., Bermudes, D. and Low, K.B. (2004) Hot spot for a large deletion in the 18- to 19-centisome region confers a multiple phenotype in Salmonella enterica Serovar Typhimurium strain ATCC 14028. J Bacteriol. 186, 8516–8523.

Tumor-Targeted Salmonella typhimurium Overexpressing Cytosine Deaminase 4. Zheng, L., Luo, X., Feng, M., Li, Z., Le, T., Ittensohn, M. et al. (2000) Tumor amplified protein expression therapy: Salmonella as a tumor-selective protein delivery vector. Oncology Res. 12, 127–135. 5. Bermudes, D., Zheng, L., and King, I. C. (2002) Live bacteria as anticancer agents and tumor-selective protein delivery vectors. Curr Opin Drug Discover Develop. 5, 194–199. 6. Bermudes, D., Low, K.B., Pawelek, J., Feng, M., Belcourt, M., Zheng, L., and King, I. (2001) Tumor-selective Salmonella-based cancer therapy. Biotechnol. Genet. Eng. Rev. 18, 219–233. 7. Dubois, L., Dresselaers, T., Landuyt, W., Paesmans, K., Mengesha, A., Wouters, B.G., et al. (2007) Brit. J. Cancer 96, 758–761. 8. King, I., Bermudes, D., Lin, S., Belcourt, M., Pike, J., Troy, K., et al. Tumor-targeted Salmonella expressing cytosine deaminase as an anticancer agent. Human Gene Therapy 13, 1225–1233. 9. Toso, J.F., Gill, V.J., Hwu, P., Marincola, F.M., Restifo, N.P., Schwartzentruber, D.J., et al. (2002) Phase I study of the intravenous administration of attenuated Salmonella typhimurium to patients with metastatic melanoma. J Clinical Oncol. 20, 142–152. 10. Nemunaitis, J., Cunningham, C., Senzer, N., Kuhn, J., Cramm, J., Litz, C., et al. (2003) Pilot trial of genetically modified, attentuated Salmonella expressing the E. coli cytosine

11.

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14.

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16.

17.

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deaminase gene in refractory cancer patients. Cancer Gene Therapy 10, 737–744. Low, K.B., Ittensohn, M., Le, T., Platt, J., Sodi, S., Amoss, M., et al. (1999) Lipid A mutant Salmonella with suppressed virulence and TNFα induction retain tumor-targeting in vivo. Nature Biotechnol. 17, 37–41. Davis, R. W., Botstein, D., and Roth, J. R. (1980) Advanced Bacterial Genetics. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York. Low, K.B., Ittensohn, M., Luo, X., Zheng, L., King, I., Pawelek, J.M., et al. (2003) Construction of VNP20009. Methods in Molecular Med. 90, 47–59. Sambrook, J., Fritsch, E.F., and Maniatis, T. (1989) Molecular Cloning: A Laboratory Manual, 2nd Ed., Cold Spring Harbor Press, Cold Spring Harbor, NY. Austin, E.A. and Huber, B.E. (1993) A first step in the development of gene therapy for colorectal carcinoma: cloning, sequencing and expression of Escherichia coli cytosine deaminase. Molecular Pharmacol. 43, 380–387. Donnenberg, M.S. and Kaper, J.B. (1991) Construction of an eae deletion mutant of enteropathogenic Escherichia coli by using a positive-selection suicide vector. Infection and Immunity 59, 4310–4317. Hanahan, D. (1983) Studies on transformation of Escherichia coli with plasmids. J. Mol. Biol. 166, 557–580.

Chapter 34 Chemoprotection by Transfer of Resistance Genes Tulin Budak-Alpdogan and Joseph R. Bertino Summary Dose-limiting toxicity of chemotherapeutic agents, i.e., myelosuppression, can limit their effectiveness. The transfer and expression of drug-resistance genes might decrease the risks associated with acute hematopoietic toxicity. Protection of hematopoietic stem/progenitor cells by transfer of drug-resistance genes provides the possibility of intensification or escalation of antitumor drug doses and consequently an improved therapeutic index. This chapter reviews drug-resistance gene transfer strategies for either myeloprotection or therapeutic gene selection. Selecting candidate drug-resistance gene(s), gene transfer methodology, evaluating the safety and the efficiency of the treatment strategy, relevant in vivo models, and oncoretroviral transduction of human hematopoietic stem/progenitor cells under clinically applicable conditions are described. Key words: CD34+ cells, drug-resistance gene, hematopoietic stem cells, murine models, myeloprotection, retroviral gene transfer, transduction

1. Introduction One of the major reasons for cancer treatment failure is specific genetic or epigenetic alterations in the cancer cells. Alterations in more than 250 genes are known to a play role in cancer drug resistance, including loss of a cell surface receptor or a drug transporter, alteration of metabolism of a drug, higher expression of energy-dependent transporters that eject anticancer drugs from cells, insensitivity to drug-induced apoptosis, induction of drugdetoxifying mechanisms, or an increase in DNA repair. De novo and acquired drug resistance are major challenges of cancer treatment, and we and others have suggested “turning the tables” and using drug-resistance genes for protecting normal tissues (1). This strategy holds the promise of not only lessening the toxicity Wolfgang Walther and Ulrike S. Stein (eds.), Methods in Molecular Biology, Gene Therapy of Cancer, vol. 542 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-561-9_34

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of the anticancer drug(s), but also increasing the efficiency of treatments. In this chapter, we summarize the process of developing gene therapy strategies for enhancing the effectiveness of anticancer chemotherapy by transferring drug-resistance genes into bone marrow stem/progenitor cells. Additionally, drug-resistance genes may be used as selectable markers when combined with a therapeutic gene for correction of a congenital hematopoietic system disorder (2, 3). The genetic chemoprotection strategy should provide evidence that (1) the choice of the myeloprotective gene(s) is effective, (2) the gene transfer method has reasonable acute and long-term safety, (3) the engraftment potential of the genetically modified cells is preserved, (4) transgene expression level and its persistence are adequate for the planned treatment period, (5) the timing for posttransplantation treatment is plausible with tumor growth kinetics, and (6) the posttransplantation treatment provides significant tumor growth control. 1.1. Selecting Drug-Resistance Gene(s)

For several malignancies, a relationship between the intensity of antineoplastic chemotherapy and tumor response has been demonstrated. However, high-dose intensity treatment has thus far had limited clinical efficiency. Salvage chemotherapy treatments after a high-dose treatment are known to have limited benefit with significant toxicity (4–8). Myeloprotection with drug-resistance gene transfer and therefore possible dose escalation may increase the clinical value of the adjuvant/or salvage chemotherapy in the posttransplantation setting. For developing a gene transfer-based myeloprotection strategy, one should consider drugs that have myelosuppression as the dose-limiting toxicity, and target tumor models should have a steep dose response to the particular drug treatment. For the last 25 years, scientists have been trying to establish safe and efficient drug-resistance gene transfer strategies, however, the process has turned out to be more complicated than originally thought. Every cancer expresses a different array of drug-resistance genes, and cells within a cancer are usually heterogeneous in respect to their drug resistance, and even the tumors that are not intrinsically resistant become selected for drug-resistant variants and then exert simultaneous resistance to many different structurally and functionally unrelated drugs. Therefore, cancers are usually treated with drug combinations that have different mechanisms of action, but the toxicities of the combinations are usually higher then the single-agent toxicity. Drug-resistance gene transfer aims to provide resistance to healthy tissue above a threshold that will spare the healthy tissue but not the cancer cells. Chemotherapeutic resistance can be caused either by general mechanisms of resistance to cell death, i.e., the expression of genes

Chemoprotection by Transfer of Resistance Genes

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that prevent cellular suicide, deficiencies of the cell to detect DNA damage and repair, changes in the cell cycle profile, or by drugspecific mechanisms. The drug-specific mechanisms are usually related to drug transport, metabolism, and/or the drug’s cellular targets. Many cancers have disturbed apoptotic, DNA damage response, and/or cell cycle regulation pathways that contribute to their resistance against a variety of anticancer drugs, however, the relevant genes usually have a role in the oncogenic process and their forced activation might alter the regulation of hematopoiesis. Figure 1 illustrates pathways that are commonly considered as good candidates for developing drug-specific resistance strategies. Table 1 summarizes the drug-resistance genes that have been explored for myeloprotection. The problems defined during developing drug-resistance gene transfer strategies are summarized in Note 1. Choosing a drug-resistance gene that will provide bone marrow protection against more than one drug is preferable. For that reason, overexpression of the multidrug-resistance gene, MDR1, that provides resistance to a variety of non-cross-resistant chemotherapeutic agents, became the first clinically applied drug-resistance gene transfer strategy (9–17). The other strategy is to transfer more than one drug-resistance gene to protect marrow stem cells against a combination of chemotherapeutic drugs. For that purpose, different drug-resistance genes are expressed

Fig. 1. Schematic representation of drug-resistance genes. The drug-resistance gene transfer strategies aim to alter drug relevant cellular pathways: inhibit the uptake of the drugs, inactivate the drugs before their interaction with cellular targets, alter the cellular proteins and make them resistant to specific inhibition with drug, or repair lesions caused by the toxic or mutagenic affects of the drugs.

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Table 1 Drug Resistance Genes Pathway

Genes

Confers resistance to

Transmembrane efflux pump

Multidrug resistance 1 (MDR-1&3) ATP-binding cassette transporter, subfamily B (ABCB1&4)

Vinca alkaloids, taxanes epipodophyllotoxins, anthracyclines

Multidrug resistance-related protein (MRP1, 3–5) ATP-binding cassette transporter, subfamily C (ABCC1, 3–5)

Vinca alkaloids, paclitaxel, antifolates, nucleoside analogs, epipodophyllotoxins, anthracyclines,

ATP-binding cassette transporter, subfamily G (ABCG2/BRCP1)

Irinotecan, topotecan, anthracyclines, flavopiridol, etoposide, methotrexate

Increased nucleoside transport

Equilibrative nucleoside transporter-2 (hENT2)

Methotrexate, trimetrexate

Decreased drug influx

Reduced folate carrier mutant

Methotrexate

Altered drug target

Dihydrofolate reductase mutant

Methotrexate, trimetrexate, pemetrexed

Thymidylate synthase mutant

5-Fluorouracil, raltitrexed, Thymitaq, pemetrexed

Ribonucleotide reductase mutant

Hydroxyurea

Tubulin mutant

Vinca alkaloids, paclitaxel

Cytidine deaminase

Cytarabine, gemcitabine

Aldehyde dehydrogenase (ALDH1)

Cyclophosphamide

Glutathione S-transferase (GST alpha, pi)

Cisplatin, cyclophosphamide, busulfan, melphalan

Altered drug metabolism

Altered detoxification/ antioxidation

Microsomal GST-2

DNA repair

Manganese superoxide dismutase

Radiation, anthracyclines, paraquat

O6–Methylguanine–DNA– methyltransferase (MGMT)

Alkylating agents, benzylguanine,

Tyrosyl–DNA–phosphodiesterase (TDP-1)

Camptothecin

from a single promoter as a fusion gene (18–20), or as two transgenes with a separate translational unit, an internal ribosomal entry site (IRES) (14, 18–28), or from two different promoters in a bidirectional vector design (29–31) (Fig. 2).


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Fig. 2. Different vector designs. Schematic representations depict: viral origin enhancer promoter (EP, solid black), R, U5 regions (LTRs), splice sites (splice donor [SD], and splice accepter [SA]) Rev-responsive element (RRE), packaging signal (Ψ), cellular origin enhancer promoter (EP, open), and posttranscriptional regulatory element (PRE). (A) LTR-driven gamma-retroviral vector with single gene; (B) LTR-driven gamma-retroviral vector with two genes separated with an internal ribosomal entry site (IRES); (C) LTR-driven gamma-retroviral vector with a fusion gene that consists of two genes; (D) self-inactivating (SIN, with a 3′ U3 deletion) gamma-retroviral vector with a cellular promoter; (E) SIN lentiviral vector with a cellular promoter; (F) SIN lentiviral vector with an intron installed 3′ of the cellular promoter; (G) SIN lentiviral bidirectional vector with two expression cassettes driven by a cellular, bidirectional promoter.

1.2. Selecting the Gene Transfer Method

In the clinical setting, effective cytotoxic therapy usually requires repeated cycles of drug administration. Therefore, sustained drug-resistance gene expression is required through the whole course of treatment. Physicochemical methods of gene delivery, i.e., electroporation, are currently emerging. Electroporation has been investigated as a tool for gene transfer in hematopoietic stem cells (HSCs), however, this gene delivery method not only had low long-term expression, but also may induce a substantial amount of cellular toxicity (32–35). Recently, targeted integration of the genes has become possible by using mobile genetic elements, namely, transposons, e.g., Sleeping Beauty, PiggBac (36–45), or by inducing double-strand breaks with engineered zinc finger nucleases that stimulate integration of long DNA stretches into a predetermined location (46, 47). However, gene transfer into long-term repopulating HSCs with these methods requires further improvements (43, 47). Among the available gene delivery systems, integrating retroviral vectors are the most commonly used delivery systems, with their stable integration and long-term expression ability. Stably integrating vectors, i.e., gamma-retroviral, lentiviral, and foamy viral vectors, have the capacity to carry different size foreign genes.

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Retroviral vector integration to host DNA was suggested to be random and expected to cause a minimal amount of harm to the target cell. Improved long-term gene marking with improved gene transfer protocols revealed the underestimated risk of retroviral gene transfer; namely, insertional mutagenesis and consequently clonal hematopoietic disorders (48–51). The French X-SCID trial data as well as carefully designed animal studies have revealed that retroviral integrations are not as random as it had been suggested. The integrations occur at varying degrees of preference for actively transcribed genes, and in distinct cis-regulatory regions that might result in abnormal activity of other genes (52–60). The semirandom insertion of transgenes into chromosomal DNA impacts the hematopoietic clonal diversity, which might induce clonal imbalance (54, 55, 57–59, 61–69). Vector backbones and the window chosen for statistical analysis influence the integrational site distributions, i.e., MLV-based vectors exhibit predilection for insertion into the immediate proximity of transcriptional start sites and gene-regulatory regions, whereas HIV-based vectors are more likely to integrate in and disrupt active transcription units (52, 53, 55, 56, 58, 70, 71). The documented risks of delayed adverse events with integrating vectors necessitate implementation of methods for detection of gene therapy-related delayed adverse events, including new malignancies, and hematological and autoimmune disorders. The nucleotide sequence adjacent to the site of the vector integration can be determined by linear amplification-mediated (LAM) polymerase chain reaction (PCR). The identified integration site sequence is compared with known human sequences in the human genome database to reveal the association of the identified sequences with any known carcinogenesis mechanisms (72–75). Because the reported leukemia cases have been observed after relatively long periods, the role of the integration sites in any clonal dominance can only be determined by repeated long-term follow-up of these vector integration sites. Oligoclonality or even monoclonality may be seen without signs of malignancy (76, 77), but high vector copies per cell, and persistent clonal dominance are known to increase the risk of insertional mutagenesis and oncogenesis, and these patients should be followed accordingly. The adverse effects relating to insertional mutagenesis have increased the importance of adding safety features to vector designs. Self-inactivating (SIN) vectors, physiologic cellular promoters, posttranscriptional regulatory elements (i.e., improving 3′ end processing with polyadenylation sequences), and insulators lessen the dysregulation of the genes in the vicinity of vector integration sites. (78–85). Figure 2 depicts different vector designs with their regulatory elements. Several investigators have reported that tissue-specific transgene expression may be accomplished with microRNA target


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site-containing vectors (86–99). Coexpression of a suicide gene may eliminate the transduced cell using systemic administration of a prodrug that is activated by the suicide gene. However, herpes simplex-derived thymidine kinase (HSV-TK), used as a suicide gene has several problems, including insufficient blood levels of the prodrug Ganciclovir, immunogenicity (100, 101), and Ganciclovir resistance related to postinsertional alterations of the transgene with cryptic splice sites (102, 103). Recently, a new suicide gene, mutant human thymidylate kinase, that provides substantial sensitivity to AZT, has shown promise in both in vitro and in vivo studies (104). This new knowledge and technology in vector design might further improve cell targeting through vector and promoter specificity and reduce the immune response to the current vectors. In vivo administration of vectors for gene therapy is currently being explored for clinical applications (29, 105–108), however, ex vivo transduction is still the most common gene transfer method for HSCs. Integration of oncoretroviral vectors into the host cell’s chromosomal DNA requires active cell cycling (109). Prestimulation with early acting cytokines increases the expression of the envelope receptors on HSCs (110–114) and the number of cells that are actively cycling, and also maintains cell viability by inhibiting HSC apoptosis (115). However, there is a fine balance between cell cycling and differentiation; prolonged ex vivo manipulation of HSCs often results in either differentiation of HSCs and/or loss of their engraftment potential (116–119). In that respect, lentivirus-, and foamy virus-derived vectors have the advantage of being less dependent on cytokines for gene transfer than oncoretroviral vectors (89, 106, 115, 120–124). Physical parameters that modulate the interactions between viral particles and target cells, i.e., high density of viral particles in the close vicinity of the cells (125), colocalization with RetroNectin® (126–130) or with polycations (131–135), spinoculation (120, 136–141), and preloading (137, 142), affect gene transfer efficiency. The benefit of adding polycations (125, 128), i.e., Polybrene or protamine sulfate, and centrifugation (125, 143) in the presence of RetroNectin® is somewhat controversial, and vector preloading might only be beneficial with highly concentrated vector supernatant batches, which might also contain inhibitory factors (144). Systematic prescreening of vector stock titers on human CD34+ cells might eliminate producer cell clones that secrete inhibitory factors. Insertional gene transfer modalities may induce deleterious stem cell behavior including stem cell exhaustion, genotoxic lesion accumulation, selection, and dominance or elimination of the transduced clones. Vector backbones as well as the target genes may determine the extent of these side effects, but certain changes in transduction design might also limit in vitro transformation

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rates and in vivo oncogenesis. For about a decade, the most commonly used transduction method was to seed the HSCs on top of adherent vector-producing cells. However, this method not only resulted in substantial loss of HSCs but also increased leukemogenesis in mice transplanted with transduced HSCs (62). Coculturing the HSCs on producer cells increased the vector copy number per cell; however, all of the animals transplanted with these HSCs eventually developed leukemia (62). The vector copy number per cell was suggested to be correlated with the expression levels of the transgene, the downside is that the higher the vector copy number per cell, the higher the chance of insertional clonal transformation (62). The correlation between integration frequency per cell and overall gene transfer efficiency has led to the suggestion that limiting the HSC’s transduction efficiency would reduce the probability of multiple integrations in single cells (145). Among a variety of cytokine combinations, the combination of stem cell factor, FLT3 ligand, and thrombopoietin has been widely accepted for HSC transductions, because this combination induces human HSC expansion that is more synchronous (146), yields higher transgene marking (143), improves survival (146), and maintains the multipotential engraftment ability of HSCs (147). For murine bone marrow progenitor cells, stem cell factor, thrombopoietin, with or without interleukin (IL)-3 and IL-6 are the commonly used cytokine combinations for ex vivo transduction protocols. The length of prestimulation, the number of transduction cycles, and the time interval between two transductions varies among various published retroviral gene transfer protocols (125, 126, 143, 148–154). The total ex vivo cell manipulation time is shorter for lentiviral vectors than for oncoretroviral vector transductions (125). 1.3. Mouse Models for Drug-Resistance Gene Transfer

The translation of successful murine studies into human therapies is questionable because large animals, including primates, had lower gene-marking levels, but nevertheless murine models answer a series of safety and efficiency concerns and are useful for demonstrating the principle of gene transfer into HSCs. In this chapter, we summarize mouse bone marrow and human CD34+ transduction protocols. Other animal models, e.g., dog, nonhuman primates, have also been used for testing the validity and safety of drug-resistance gene transfer strategies (66, 155–165), however, these models are beyond the scope of this chapter. Mouse experiments, if designed well and conducted with enough animals, may provide the preclinical data required by regulatory bodies, including proof of concept, efficacy, vector and vector-transduced cell distribution, immunogenicity, and longterm safety. Experiments should be designed for testing safety and efficacy of gene therapy, long-term gene marking, as well as biodistribution, insertional mutagenesis, and immunogenicity.


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For transducing the target cells that allow long-term hematopoiesis reconstitution, either selected hematopoietic progenitor/stem cells, sca+c-kit+lin−, or unfractionated bone marrow from mouse donors that are treated with 5-fluorouracil (5-FU) may be used. 5-FU treatment is known to eliminate the hematopoietic progenitors and stimulate stem cell proliferation (166, 167) (see Note 2). The harvested murine bone marrow cells are transduced and infused into the tail vein of lethally irradiated recipient mice. Transplant models such as nonmyeloablative preparative regimens (168–170) or infusing ex vivo expanded transduced HSCs (171, 172) may also be used for testing the selective advantage of gene transfer strategies. Retrovirus infection depends on binding of the retroviral envelope (Env) protein to specific cell surface protein receptors. The tropism of the virus is defined by the envelope protein because different envelope receptors are differentially expressed among cell lines and different hematopoietic cell subpopulations (114, 166, 173). This differential expression of envelope receptors defines the susceptibility of the specific cell line to viral transduction. Transductions with amphotropic, GALV-, FeLV-c-, RD114-, or VSV-Genveloped viruses are considered to reflect transduction parameters that are relevant to clinical applications. In ours and others experience, the longevity of gene expression in murine models is higher with ecotropic or 10A1-enveloped virus transduction than with amphotropic virus, and pretreatment of mice with 5-FU increases the ecotropic retroviral transduction efficiency (166, 174, 175). For experiments requiring proof of principle for myeloprotection and therapeutic advantage, we prefer to generate ecotropic virus by transfecting the Phoenix-Eco producer cell line (176), which provides viral titers above 106 transduction unit (tu)/mL. Methods for vector production will not be discussed in this chapter, but, briefly, selection with drug-resistance genes enables generation of stable, polyclonal high-titer producer cell lines. Further selection of single producer clones not only might allow a 1- to 2-log increase in titers (177) but also may eliminate clones that produce inhibitory factors against target cells (100, 125, 178).

2. Materials

2.1. Isolation of Mouse Lin – Hematopoietic Cells and Prestimulation (see Note 2)

1. Dulbecco’s Modified Eagle Medium (DMEM) high glucose (1X), liquid, with L-Glutamine and sodium pyruvate (DMEM-HG), 500 mL. 2. Fetal bovine serum (FBS), prescreened for hematopoietic cell growth, 500 mL.

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3. Penicillin–streptomycin (100X solution of 10,000 U/mL penicillin and 10,000 μg/mL streptomycin), 100 mL, storage at −20°C temperature. Final concentration in media is 1X (100 U/mL penicillin and 100 μg/mL streptomycin). 4. Dulbecco’s phosphate-buffered saline (D-PBS), without calcium and magnesium, 500 mL. 5. Scissors, forceps, scalpel. 6. Sterile water for injection, USP, 10-mL vial. 7. 25- and 28-gauge needles. 8. 1-, 3-, and 5-mL syringes. 9. T25, T75 vented tissue culture flasks. 10. 40-μm cell strainer (Becton Dickinson, Lincoln Park, NJ). 11. Millex-GV PVDF 0.2 μm, syringe-driven filter unit (Millipore, Bedford, MA). 12. PVDF 0.22-μm filter units for 50-mL and 250-mL volumes (Stericup, Millipore). 13. Rat stem cell factor (SCF), lyophilized 10-μg vial (PeproTech, Inc., Rocky Hill, NJ). Both the lyophilized protein and reconstituted solution are stored at −20°C. The lyophilized protein is diluted in 1 mL injectable dH2O and filtered to generate a stock solution at 10 ng/μL; 5 μL should be added for each milliliter of media for a final concentration of 50 ng/mL. 14. Recombinant murine thrombopoietin (TPO) lyophilized 10-μg vial (PeproTech, Inc.). Both the lyophilized protein and reconstituted solution are stored at −20°C. The lyophilized protein is diluted in 1 mL injectable dH2O and filtered to generate a stock solution of 10 ng/μL; 10 μL should be added for each milliliter of media for a final concentration of 100 ng/mL. 15. Mouse lineage depletion kit, including MS or LS columns, biotin-conjugated lineage antibodies, and anti-biotinconjugated magnetic beads (Cat# 130-090-858, Miltenyi Biotec, Auburn, CA). 16. D-PBS supplemented with 0.5% bovine serum albumin (BSA) and 0.6% citrate–dextrose (ACD solution from Sigma, St. Louis, MO), pH 7.2. 2.2. Cell Preparation and Prestimulation for Human CD34+ Cells

1. Human bone marrow (BM)-, mobilized peripheral blood (MPB)-, or umbilical cord blood (UCB)-derived human CD34+ cells, either fresh or frozen, are used for gene transfer. Purification of human CD34+ cells is beyond the scope of this chapter (see Note 3). 2. 5% Human serum albumin solution (HSA), 250 mL (Buminate, Baxter Healthcare, Deerfield, IL).


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3. 10% Dextran 40 in 500 mL of 0.9% sodium chloride IV, (Gentran Viaflex, Baxter Healthcare). 4. X-Vivo 10 Media without phenol red, 1 L, storage at +4°C. 5. 50 mg/mL Gentamicin, 10 mL; storage at room temperature. Final concentration in media is 50 μg/mL. 6. Recombinant human stem cell factor (SCF), lyophilized 10-μg vial (PeproTech, Inc.). Both the lyophilized protein and reconstituted solution are stored at −20°C. The lyophilized protein is diluted in 1 mL injectable dH2O and filtered to generate a stock solution at 100 ng/μL; 10 μL should be added for each milliliter of media for a final concentration of 100 ng/mL. 7. Recombinant human Fms-related tyrosine kinase 3 ligand (FLT3L), lyophilized 10-μg vial (PeproTech, Inc.). Both the lyophilized protein and reconstituted solution are stored at −20°C. The lyophilized protein is diluted in 1 mL injectable dH2O and filtered to generate a stock solution at 10 ng/μL; 10 μL should be added for each milliliter of media for a final concentration of 100 ng/mL. 8. Recombinant human thrombopoietin (TPO), lyophilized 10-μg vial (PeproTech, Inc.). Both the lyophilized protein and reconstituted solution are stored at −20°C. The lyophilized protein is diluted in 1 mL injectable dH2O and filtered to generate a stock solution at 10 ng/μL; 10 μL should be added for each milliliter of media for a final concentration of 100 ng/mL. 9. Sterile water for injection, USP, 10-mL vial. 10. Millex-GV PVDF 0.2-μm, syringe-driven filter unit (Millipore). 11. T25, T75 vented tissue culture flasks, or 6-well plates, or gas-permeable tissue culture bags. 12. Millex-GV PVDF 0.2-μm, syringe-driven filter unit (Millipore). 13. PVDF 0.22-μm filter units for 50-mL and 250-mL volumes (Stericup, Millipore). 2.3. RetroNectin® Coating

1. RetroNectin®, lyophilized 2.5 mg protein (Takara Bio, Inc., Shiga, Japan). The lyophilized protein should be stored at +4°C and the reconstituted solution (1 mg protein/mL) should be stored in aliquots at −20°C. For reconstitution, add sterilized 2.5 mL distilled water to obtain a 1 mg protein/mL solution, filter through a 0.22-μm filter and store in aliquots at −20°C. To keep the protein integrity, avoid vigorous mixing (no vortexing) and repeated freeze–thaw cycles.

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2. 30% Bovine serum albumin solution, 50 mL, should be kept at +2–8°C. 3. D-PBS with 2% BSA for mouse BM transductions, and D-PBS with 2% HSA for human CD34+ cell transductions. 4. Nontissue culture 6-well plates. 5. Sterile water for injection, USP, 10-mL vial. 6. Millex-GV PVDF 0.2-μm, syringe-driven filter unit. 2.4. Transduction of Mouse Bone Marrow Cells in RetroNectin®Coated Plates

1. Frozen vector stock, see Note 6 2. Prestimulated mouse (see Note 4) lin− bone marrow cells from Subheading 2.1 (lentiviral and foamy virus transduction protocol does not require prestimulation). 3. RetroNectin®-coated adherence-improved 6-well plates from Subheading 2.2. 4. Dulbecco’s phosphate-buffered saline (D-PBS), without calcium and magnesium, 500 mL. 5. Dulbecco’s Modified Eagle Medium (DMEM) high glucose (1X), liquid, with L-Glutamine and sodium pyruvate (DMEM-HG), 500 mL. 6. Fetal bovine serum (FBS), prescreened for hematopoietic cell growth, 500 mL. 7. Penicillin–streptomycin (100X solution). 8. PVDF 0.22-μm filter units for 50-mL and 250-mL volumes (Stericup, Millipore). 9. Rat SCF, see Subheading 2.1.1, final concentration of 50 ng/mL. 10. Murine TPO, see Subheading 2.1 final concentration of 100 ng/mL.

2.5. Transduction of Human CD34+ Cells in RetroNectin®-Coated Plates

1. Frozen vector stock with relevant envelope, see Note 6. 2. Prestimulated human CD34+ cells (see Note 5) from Subheading 2.2. 3. RetroNectin®-coated adherence-improved tissue culture vehicles. 4. X-Vivo 10 Media without phenol red, 1 L, storage at +4°C. 5. 50 mg/mL Gentamicin; final concentration in media, 50 μg/mL. 6. Recombinant human SCF, see Subheading 2.2, final concentration of 100 ng/mL. 7. Recombinant human FLT3L, see Subheading 2.2, final concentration of 100 ng/mL.


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8. Recombinant human TPO, see Subheading 2.2, final concentration of 100 ng/mL. 9. Sterile water for injection, USP, 10-mL vial. 2.6. Colony Assay for Determining Drug-Resistant Hematopoietic Progenitors

1. 0.4% Trypan blue solution for cell viability counts. 2. 3% Acetic acid with methylene blue for nucleated cell counts (Stemcell Technologies, Vancouver, BC). 3. 35-mm Tissue culture plates. 4. 100-mm Tissue culture plates. 5. Methylcellulose media supplemented with cytokines for mouse hematopoietic progenitor cell growth, MethoCult M3434 (Stemcell Technologies). Keep at −20°C frozen, repeated cycles of freezing and thawing should be avoided. After thawing, can be kept at 2–8°C for a month. 6. For human hematopoietic progenitor cell growth, methylcellulose media supplemented with cytokines, MethoCult H4434 (StemCell Technologies). 7. 16-gauge blunt-end needles for dispensing methylcellulose media. 8. DMEM supplemented with 2% FBS. 9. 3-mL syringes. 10. Sterile distilled water. 11. Drug that will be used for selection and the drug dose should be lethal for nontransduced mouse bone marrow progenitors.

2.7. Single-Colony PCR for Determining Transduction Efficiency (see Note 8)

1. Colony-forming units that are growing in methylcellulose semisolid media. 2. DNA Lysis Buffer, 5 mM Tris-HCL, 0.45% Tween 20, at pH 8.0. Store at room temperature. 3. Proteinase K, Solution (Roche) in 10 mM Tris-HCL, pH 7.5, 15.1 mg/mL. Store at 2–8°C. 4. Dulbecco’s phosphate-buffered saline (D-PBS), without calcium and magnesium, 500 mL. 5. Phenol/CHCl3/isoamyl alcohol (25:24:1) mixture. 6. Glycogen, 20 mg/mL solution (Boehringer-Mannheim, Mannheim, Germany). 7. Ammonium acetate, 7.5 M solution. 8. Absolute ethanol. 9. AmpliTaq Gold-DNA polymerase (Applied Biosystems, Foster City, CA). 10. GeneAmp 10x PCR buffer II (Applied Biosystems). 11. 25 mM MgCl2 solution (Applied Biosystems).

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12. dNTP GeneAmp Blend, 10 mM (Applied Biosystems). 13. β-Actin forward and reverse primers (relevant human or mouse-specific β-actin primers). 14. Transgene-specific forward and reverse primers. 2.8. Flow Cytometry for Peripheral Blood or Bone Marrow: Immunostaining for Mouse Cells

1. Peripheral blood or bone marrow sample. 2. CD16/CD32 FcγIII/II receptor (Mouse BD Fc Block, BD Pharmingen, San Jose, CA). 3. Lysing buffer, 10X (BD Pharma Lyse, BD Pharmingen). 4. Staining buffer, D-PBS supplemented with 0.2% BSA and 0.1% sodium azide. 5. Fluorescein isothiocyanate (FITC)-conjugated anti-mouse CD45.1 (BD Pharmingen). 6. FITC-conjugated mouse IgG2a, κ isotype control (BD Pharmingen). 7. Phycoerythrin (PE)-conjugated anti-mouse IgG2b, κ (Mac-1, BD Pharmingen). 8. PE-conjugated rat IgG2b, κ isotype control (BD Pharmingen). 9. Peridinin–chlorophyll–protein complex (PerCP)-conjugated anti-mouse CD3ε (BD Pharmingen). 10. PerCP-conjugated hamster IgG1, κ isotype control (BD Pharmingen). 11. PerCP-conjugated anti-mouse CD45R/B220 (BD Pharmingen). 12. PerCP-conjugated rat IgG2a, κ isotype control (BD Pharmingen). 13. APC-conjugated anti-mouse Ly-6G (Gr-1, BD Pharmingen). 14. APC-conjugated rat IgG2b, κ isotype control (BD Pharmingen). 15. Polystyrene 12 × 75-mm tubes (BD Falcon, San Jose, CA).

2.9. Flow Cytometry for Peripheral Blood or Bone Marrow: Immunostaining for Human Cells

1. Peripheral blood or bone marrow sample. 2. CD16/CD32 FcγIII/II receptor (Mouse BD Fc Block, BD Pharmingen). 3. Lysing buffer, 10X (BD Pharma Lyse, BD Pharmingen). 4. Staining buffer, D-PBS supplemented with 0.2% BSA and 0.1% sodium azide. 5. FITC-conjugated anti-human CD45 (BD Pharmingen). 6. FITC-conjugated mouse IgG1, κ isotype control (BD Pharmingen). 7. Polystyrene 12 × 75-mm tubes (BD Falcon).


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3. Methods 3.1. Isolation of Mouse Lin− Hematopoietic Cells, and Prestimulation

1. On Day 0, harvest bone marrow cells from femurs and tibias using 5-mL syringes and 25-gauge needles to flush bone marrow with cold DMEM supplemented with penicillin/streptomycin and 2% FBS. 2. To make a single-cell suspension, gently pass the cells through a 40-μm cell strainer, nylon mesh, to remove bone particles and cell clumps and then wash the strainer several times with D-PBS to get all the cells through. Keep the cells on ice until you finish harvesting all animals. 3. Wash bone marrow cells with D-PBS (300×g for 10 min), resuspend the cell pellet in DMEM containing 20% fetal calf serum (FCS), and count the cells both with trypan blue and 3% acetic acid with methylene blue. The total nucleated cell count harvested per mouse should be between 3 and 6 × 107. 4. Adjust the cell concentration to 1–5 × 106 cells/mL with DMEM containing 20% FCS and plate in 75-cm2 tissue culture flasks (maximum 15 mL cell suspension per flask). Place the flasks into the incubator, and allow stromal cell components to adhere for 2 h. 5. Wash the flasks three times with D-PBS (8 mL per wash) and collect all nonadherent cells. Stromal cells and monocyte/ macrophage lineage cells adhere strongly to the flasks in 2 h. 6. Centrifuge the cells at 300×g for 10 min and then proceed with lineage-negative cell depletion (Note 2) according to manufacturer’s protocol. Briefly, resuspend the cell pellet in 40 μL of D-PBS with 0.5% BSA and 0.6% ACD per 107 total cells. After adding 10 μL of biotin antibody cocktail per 107 total cells, incubate for 10 min at +2–8°C and then add 30 μL of buffer and 20 μL of anti-biotin microbeads per 107 total cells. Incubate the cells at +2–8°C for 15 min and then wash with buffer at 300×g for 10 min. Resuspend the cells (up to 108 cells in 500 μL of buffer) and pass them through the column. Collect the cells passing through the column as the negative-binding fraction. Wash the column several times with buffer and then elute the cells retained in the column by simply taking the column outside the magnetic field and then eluting with buffer. Check the purity and calculate the yield of the procedure by staining the initial, negative, and positive populations with PEconjugated anti-biotin antibody. Using a second column could increase the purity of the lineage negative cells. 7. Centrifuge the cells at 300×g for 10 min, and then resuspend the cells at a final concentration of 1 × 106 cells/mL in DMEM supplemented with 10% FBS, penicillin/streptomycin, and

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cytokine cocktail (50 ng/mL rat SCF, and 100 ng/mL murine TPO) for prestimulation. Incubate the BM cells for 36 h for prestimulation at 37°C and 5% CO2 (see Note 3). 3.2. Prestimulation of Human CD34+ Cells

1. Thaw frozen human CD34+ cells at 37°C (see Note 4). 2. Ice-cold dextran and 5% human serum albumin solution (v:v) should be mixed with the same volume of frozen human CD34+ cells and then spun at 400×g for 10 min. 3. Human CD34+ cells should be plated at a cell density of 5 × 105 cells/mL in X-Vivo 10 media supplemented with human cytokines, 100 ng/mL huSCF, huFLT3L, huTPO, and gentamicin for prestimulation. Cells can be incubated in flasks, plates, or gas-permeable bags, depending on the amount of the cells used for the experiment. 4. Incubate the human CD34+ cells for 36 h for prestimulation at 37°C and 5% CO2 (see Note 5).

3.3. RetroNectin® Coating

1. Calculate the amount of RetroNectin® according to the surface of the area; the surface area of a well for a 6-well plate is approximately 10 cm2, and the wells should be coated with RetroNectin® at a dose of 2 μg/cm2. Total amount of RetroNectin® = surface area × 2 μg/cm2 = 20 μg per well. The calculations are given for a 10-cm2 well, but one should keep in mind that total number of cells that can be transduced in one well is usually limited to 2 × 106 cells (see Note 4). According to the size of the experiment, nontissue culture T75 flasks could be coated. Calculations should be adjusted accordingly. 2. Dilute 20 μL of 1 mg/mL RetroNectin® stock solution (see Subheading 2.3) in 1 mL of D-PBS (final concentration of 20 μg/mL; for effective coating, the RetroNectin® concentration should be kept between 20 and 100 μg/mL). Incubate either at room temperature for 2 h or overnight at +4°C. 3. Prepare 1 mL blocking solution as 2% bovine serum albumin in D-PBS (for human CD34+ cell transductions, we prefer using human serum albumin instead of BSA). After aspirating the RetroNectin®-containing solution, transfer the blocking solution to the well. 4. After ½ h incubation at room temperature, aspirate the solution and wash the well with another 1 mL D-PBS. If the RetroNectin®-coated plate is not used immediately, it can be kept at +4°C for approximately 1 week.

3.4. Transduction in RetroNectin®-Coated Plates

1. Thaw vector supernatant that has been sterile filtered through a 0.45-μm filter and stored at −70°C (see Note 6). It is important that the supernatant be warmed at least to room temperature before adding to the cells. The medium should


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be warm and pH should be neutral, and the CO2 in the incubator should be 4.5–5%, or the retroviral receptor conformation will be altered and the retrovirus will not bind. 2. For oncoretroviral transduction of mouse cells, prestimulated lineage-negative bone marrow cells are washed with D-PBS and then resuspended (1 × 106 cells/mL of transduction media containing vector supernatant and cytokines: 100 ng/mL rat SCF, and 100 ng/mL human TPO). For human CD34+ cells, prestimulated human CD34+ cells from Subheading 3.2 should be washed with D-PBS and then resuspended (5 × 105 cells/mL) with transduction media containing vector supernatant and cytokines: 100 ng/mL human SCF, FLT3L, and TPO. For gamma-retroviral vectors, we prefer a multiplicity of infection (MOI) ratio of 1. Dilute the vector supernatant with complete media (either DMEM with 10% FBS and penicillin/ streptomycin, or X-Vivo 10 media with gentamicin) accordingly. The total transduction media height per well should be 2 mm/cm2 of surface area (~2 mL transduction media per well for a 6-well plate) (see Note 7). 3. After 12 h of incubation at 37°C and 5% CO2, remove supernatant and spin down the cells at 300×g for 10 min. Resuspend the cells in transduction media for the second cycle of transduction. Two cycles of 12 h of transduction usually provide 52–75% transduction efficiency with Eco-enveloped gamma-retroviral vectors. Longer transduction, 24 h/transduction cycle, or more cycles impair the engraftment potential of the cells. 4. At the end of two cycles of transduction, wash the plates several times with cold D-PBS, and then centrifuge at 300×g for 10 min. Resuspend the cells that will be used for in vitro drug toxicity in either DMEM supplemented with 10% FBS, penicillin/streptomycin, and rat SCF 50 ng/mL; or X-Vivo 10 media with gentamicin and 100 ng/mL human SCF. Incubate for another 24 h at 37°C and 5% CO2. 5. Lentiviral and foamy virus transduction protocols do not require prestimulation (see Note 4 and 6). For lentiviral transductions, resuspend the cells isolated at Subheading 3.1 in vector-containing complete media supplemented with 50 ng/ mL rat SCF and 100 ng/mL human TPO. Incubate the cells for 24 h at 37°C and 5% CO2. For human CD34+ cell transduction, use X-Vivo 10 media supplemented with gentamicin and human cytokines: 100 ng/mL of huSCF, huFLT3L, and huTPO. Remove the cells from the transduction media and wash with D-PBS. 6. For in vivo experiments, the cell pellet should be resuspended in D-PBS supplemented with 2% FBS, i.e., if 5 × 105 transduced cells are to be injected per mouse, we prepare a cell

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suspension of 5 × 10 6 cells/mL, and then inject 100 μ L of this solution per mouse. Mice, 6–8 weeks old, have an approximate blood volume of 1.1 mL, therefore, the intravenous injection volume should be at most 200 μL/day. Cells should be kept on ice and syringes should be filled one at a time after thoroughly mixing the cells before each syringe fill. Highly concentrated cells usually form aggregates and prefilled syringes get clumped with cell aggregates if left waiting without mixing during the injection process. Mixing platforms can avoid the aggregates. Aggregates could cause lung emboli and consequently cause loss of animals. Cell aggregates would also cause great graft dose variability among the animals. Cells should be injected to the irradiated recipients at a dose of 5–7.5 × 105 lineage-negative cells or 1–2 × 106 5-FU-enriched cells per mouse (see Note 2). 7. For in vivo human hematopoiesis in mice, 6–8 weeks old, sublethally irradiated NOD/SCID mice should be injected with CD 34+ cells. One may use 2 × 105 umbilical cord-derived, 1–2 × 106 bone marrow-derived, or 5 × 106 mobilized peripheral blood-derived CD34+ cells per mouse (see Note 3). 8. Incubate the cells for another 24 h in complete media for in vitro experiments. These cells will be plated to determine the efficiency of transduction. Additional incubation of the cells is required for optimal expression of the transgene. 9. The use of spinoculation, preloading, or polycations during the transduction process may increase the transduction efficiency, however, the success of these modalities depend on the vector, envelope, and the producer cell line (see Note 7). 3.5. Colony Assay for Determining Drug-Resistant Hematopoietic Progenitors (see Note 8)

1. The level of drug resistance in hematopoietic progenitors is determined by treating the cells with a dose of drug that is just 100% lethal (a dose–response curve for hematotoxicity of the drug should be established beforehand). The drug treatment schedule should be relevant to the in vivo pharmacokinetics of the specific drug, i.e., continuos exposure during the culture or short-term incubation with drug and then plating for colony-forming unit (CFU) assay. For each transduction sample, cells should be also plated without any drug treatment. 2. Thaw MethoCult overnight in the refrigerator. 3. Adjust the cell concentration to 2–5 × 104 cell/mL for mouse lin− cells, or 5 × 105 or 1 × 106 cells/mL for 5-FU-treated bone marrow samples, or 5–7.5 × 103 cells/mL for human CD34+ cells. 4. Add 0.3 mL of cells to 3 mL MethoCult medium for duplicate cultures.


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5. Vortex tubes to ensure all cells and components are thoroughly mixed, and let the tube stand for 5 min to allow bubbles to disappear. 6. Aspirate 1.1 mL to a 3-mL syringe with 16-gauge blunt needle, and plate into a 35-mm dish. 7. Distribute the methylcellulose by gently swirling the dish. Place two dishes into a 100-mm Petri dish, and add a third, uncovered dish containing 3 mL sterile water. 8. Incubate the mouse cells for 12 days or human CD34+ cells for 14 days at 37°C, 5% CO2, and 95% humidity. 3.6. Single-Colony PCR for Determining Transduction Efficiency (see Note 8)

1. Fill microcentrifuge tubes with 1 mL D-PBS. Aspirate single colonies from the methylcellulose medium in a volume of 20–50 μL with individual plugged pipette tips (P200), under visualization with a phase-contrast inverted microscope and add to the D-PBS containing tube. Place each colony into an individual tube. 2. Leave the pipette tip in the tube for about an hour at room temperature to allow the methylcellulose to dissolve. Then pipette D-PBS in and out to thoroughly resuspend the cells in D-PBS. 3. Centrifuge the tubes at 500 × g for 10 min in a microcentrifuge. Aspirate carefully the supernatant using individual pipette tips without disturbing the tiny cell pellet. It is acceptable to leave up to 10 μL D-PBS in the tube to avoid cell loss. 4. Prepare the appropriate volume of DNA lysis buffer as in Subheading 2.7 for the number of picked colonies (total volume = number of colonies picked × 50 μL + 10% of volume). Add proteinase K to DNA lysis buffer so that the final concentration is 0.1 mg/mL. 5. Transfer 50 μL of proteinase K-containing lysis buffer to each tube and resuspend the cell pellet by pipetting in and out. 6. Incubate the samples at 56°C for 90 min. 7. Inactivate proteinase K at 95°C for 5 min. 8. Add 150 μL of D-PBS to each tube. 9. Mix 200 μL phenol/CHCl3/isoamyalcohol with DNAcontaining solution from step 7. Leave at room temperature for 10 min and centrifuge at 14,000 × g in a microcentrifuge for 10 min. 10. Collect the aqueous upper phase (~150–180 μL) and precipitate by adding 2 μg glycogen, 18 μL of 7.5 M ammonium acetate, and 500 μL absolute ethanol. Incubate overnight at −20°C.

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11. Centrifuge the DNA at 14,000 × g in a microcentrifuge for 10 min. Wash the pellet with 70% ethanol. Dry the DNA pellet on the bench top; do not use vacuum drying. 12. Resuspend the DNA in 50 μL of either Tris/EDTA (TE) buffer or dH2O. Three to 5 μL of this solution should be adequate to detect the presence of most transgenes by PCR. 13. Run the PCR reaction with the transgene-specific primers. A PCR reaction with β-actin primers should also be run to confirm the presence of DNA in the sample. 3.7. Flow Cytometry for Peripheral Blood or Bone Marrow

1. Prepare a single-cell suspension from either bone marrow or peripheral blood samples. 2. Wash the cells with staining buffer and pellet the cells by centrifugation at 300×g for 5 min. Resuspend the cells with cold staining buffer to a concentration of 1 × 107 cells/mL. 3. Aliquot 100 μL of cell suspension in 12 × 75-mm polystyrene tubes (~1 × 106 cells/tube). 4. Preincubate the cell suspension with 2 μL of Mouse BD Fc Block at 4°C for 5 min (this does not need to be washed off before staining). 5. Add each antibody or isotype control at 5 μL/tube, and incubate for 20 min at 4°C, protected from light. 6. Wash the cells with 1 mL volume of staining buffer twice. Centrifuge the cells at 300×g for 5 minutes, aspirate supernatants from cell pellets, and resuspend the cells in 200 μL of staining buffer. 7. Add 2 mL of 1X lysing buffer to each tube containing 200 μL of cell suspension. 8. Gently vortex the cells after adding the lysing buffer. 9. Incubate at room temperature for 15 min, while protecting from light. 10. Centrifuge the tubes at 200×g for 5 min, aspirate the supernatant, and resuspend the cells in 0.5 mL staining buffer. 11. Acquire at least 50,000 events for mouse engraftment experiments, and 500,000 events for human engraftment experiments. Multicolor immunostaining for mouse cells: (1) IgG2a,κFITC, (2) CD45.1-FITC/IgG2b, κ-PE/IgG2a, κ-PerCP/IgG2b, κ-APC, (3) CD45.1-FITC/IgG2a, κ-PerCP, (4) CD45.1-FITC/ CD11b-PE/CD3ε-PerCP/Ly-6G-APC, and (5) CD45.1-FITC/ CD45R/B220-PerCP. Immunostaining for human cells: (1) IgG1,κFITC, and (2) CD45-FITC.


3.8. In Vivo Experimental Models

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The schematic presentation of representative murine experiments is given in Fig. 3. The murine models should be chosen according to the aim of the study: 1. Experiments evaluating the long-term expression and/ or safety, and immunogenicity of the gene transfer strategy should use either murine serial transplant models, i.e., C57BL/6, CH3, Balb/c, or known tumor-prone mouse models, i.e., Cdkn2a−/− mice (55), for testing the in vivo frequency for insertional malignant transformation. It also worth mentioning that insertional mutagenesis-unrelated tumor development has been observed, mainly defined by the genotypic background of the mouse strain, when experiments were extended to longer than 8 to 10 months. It is highly

Fig. 3. Experimental murine models for drug-resistance gene transfer. (A) A congenic or syngenic, serial transplant model for evaluating the transduction efficiency of long-term HSCs, engraftment potential of transduced HSCs, long-term transgene expression, and safety of the insertional gene transfer modality. (B) A congenic or syngenic transplant with chemotherapeutic treatment challenge for evaluating the myeloprotection and hematopoietic stem or progenitor cell selection aspects of drug-resistance gene transfer. (C) Mouse–mouse tumor or immunodeficient mouse–human tumor xenograft model for testing the therapeutic efficiency of myeloprotection strategy. (D) Immunodeficient mouse–human HSC transplant for evaluating the engraftment potential of ex vivo manipulated human HSCs, long-term gene expression in HSCs, and safety of the insertional gene transfer modality.

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recommended to assess the frequency of vector-independent tumor formation with adequate numbers of control animals, i.e., 30 mice (74). Following peripheral blood counts weekly after bone marrow transplant reveals the engraftment kinetics of manipulated cells, however, animals are usually very vulnerable to infection and bleeding during the first 2 weeks of the posttransplantation period. We limit peripheral blood withdrawal volume to less than 50 μL per week. C57BL/6 (CD45.2) and C57BL/6Ncr (CD45.1) congenic strains differ in their CD45 isomer expression on their peripheral blood leukocytes. Use of fluorophoreconjugated monoclonal antibodies against those isomers allows quantitative determination of donor engraftment with flow cytometry (see Subheading 3.7). In mouse models, long-term hematopoietic reconstitution could only be evaluated 8–10 weeks after the transplant. We recommend killing the animals 4 months after transplantation, and harvesting the bone marrow. Spare almost one third of the harvested bone marrow for the assays that include plating for colony assays with and without drug, and isolating both DNA and RNA for quantitative vector copy number and transgene expression determination (see Note 8). The rest of the harvested bone marrow is injected into the secondary transplant recipients. For secondary and tertiary transplants, we inject 10 × 106 cells per mouse. Congenic mouse models also allow determination of multilineage hematopoietic reconstitution. We use Gr-1, CD11b, CD3ε, and B220 for determining lineage-specific engraftment (see Subheading 3.7). Serial transplantation experiments aim to provide evidence for long-term hematopoiesis, and it is valuable to show donor-derived multilineage hematopoiesis in those recipients. Immune response against the gene transfer vector (179, 180), different components of transduction media (181), or the transgene (183–185) itself are a concern for HSC gene transfer. Congenic mouse models allow investigators to determine T cell clonal immunity against transduced HSCs. Induction of an immune response against transgenes expressed in HSCs correlates to the degree of immunosuppression induced by the conditioning regimen, the immunogenicity of the transgene, the cellular localization of the transgene coded protein, and the extent of gene expression in transduced cells. Because a single amino acid change within an epitope can even induce a cytotoxic T lymphocyte (CTL) response, mutant drug-resistant genes might also induce CTL responses directed to normal host proteins that only differ slightly from the wild-type form. Performing LAM-PCR with peripheral blood DNA samples requires the polyclonality of engraftment and insertional mapping of the specific vector backbone. We refer the readers to the following references for LAM-PCR methodology (75, 182).


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2. Therapeutic challenge experiments for demonstrating myeloprotection with drug-resistance gene transfer, as well as evaluating the probability of inducing high vector copy clonal dominance through the drug selection process should use either syngeneic or congenic mouse models. Before conducting murine bone marrow transplant experiments, we typically evaluate the advantage of drug-resistance gene expression with a drug dose and schedule that mimics the pharmacokinetics of the drug by applying selection pressure on unmanipulated and transduced mouse bone marrow cells. A pilot dose-finding study should be conducted using the same gender and strain mouse that will be used for in vivo myeloprotection experiments. With a clinically applicable drug(s) administration schedule, the maximal tolerated dose for the drug(s) should be defined. For estimating the first drug dose and schedule, we prefer to use the recommendations of the National Cancer Institute (NCI)–Developmental Therapeutics Program (http://dtp.nci.nih.gov), and then increase the drug dose every 2 weeks with a modified Fibonacci schedule until we see an average of 15% total body weight loss and/or more than a 70% decrease in the peripheral blood counts. Each dose increment is applied after full total body weight recovery is established, approximately 9–12 days after the treatment. We do at least one more dose increment after reaching the maximal tolerated dose. For the dose-finding experiments, we recommend using 10–12 animals per group. For murine bone marrow transplant experiments, recipients should either receive an ablative or a nonablative dose of irradiation or chemotherapeutics. A control group transplanted with a marker gene (enhanced green fluorescent protein [eGFP], Discosoma sp. red fluorescent protein [dsRed], etc.) transferred bone marrow mimics the vulnerability of the graft after transplant better than a naïve group of animals. We prefer to start drug treatment after engraftment is established at about 3 weeks, and inject the maximal tolerated drug dose. Total body weight and peripheral blood counts of the animals are followed every other day. Drug dose increments are continued for at least two more dose increments or until 50% of the animals in the drug-resistance group succumb to toxicity, whichever comes first. When an animal dies, its bone marrow is harvested and plated for colony assay, and the DNA and RNA are isolated for vector copy, expression, and LAM-PCR assays. We keep a group of animals without treatment and evaluate similar parameters immediately after engraftment, and at the end of each treatment cycle (at least 5 animals at each time point adds up to 20 animals for the untreated group). Harvesting the other tissue samples from animals might also allow the investigators to do vector distribution-related safety studies.

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This study could also answer drug selection-related safety concerns by providing the data for vector copy numbers both with and without drug treatment. 3. Therapeutic advantage of myeloprotection and posttransplantation salvage chemotherapy should be tested either with mouse/mouse tumor or immunodeficient mouse/human tumor xenograft models. In our experimental models, tumor inoculation follows bone marrow transplantation with transduced cells (186, 187). We test the number of cells and tumor growth pattern as a preliminary study, and inject tumor cells at a time and cell dose such that we could have a measurable tumor mass (tumor diameter between 2 and 4 mm) at the time of engraftment. We carry the rest of the experiments similar to the myeloprotection experiments described in the model B section. 4. Human HSCs transplanted to sublethally irradiated immunodeficient mouse are used for evaluating the engraftment ability of transduced human HSCs. This human hematopoiesis model, with or without drug challenge, could determine the frequency of clonal dominance, the longevity of transgene expression, and the transfer rate into secondary transplant recipients. The amount of transduced cells transplanted to immunodeficient mouse varies according to the HSC source, i.e., cord blood-derived CD34+ cells can be as low as 1 × 105 cells/mouse, whereas MBP-derived CD34+ cells should be as high as 3–5 × 106 cells for NOD/SCID mouse. However, NOD/SCIDγc null strain mice require a much lower amount of cells for a relatively higher amount of engraftment in both peripheral blood and bone marrow (188–190). One should also remember that the NOD/SCID model would not allow investigators to test the myeloprotection aspect of drugresistance gene transfer strategy, because human HSCs would only engraft as a small fraction of the mouse bone marrow and would not provide enough peripheral blood leukocytes for myeloprotection.

4. Notes 1. The human multidrug-resistance 1 gene (MDR1) encodes a transmembrane efflux pump, P-glycoprotein, that prevents intracellular accumulation of a variety of clinically important drugs, such as anthracyclines, vinca alkaloids, taxanes, and podophyllotoxins (191–196). Several technical and biological problems have been encountered during the development of MDR1 gene therapy. HSCs intrinsically express MDR1


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higher than their progeny and effective myeloprotection could only be provided if the transgene expression was significantly above the background levels that resulted from the activity of the cellular alleles in primitive hematopoietic cells (191, 194). The relatively large complementary DNA (cDNA) of MDR1 resulted in aberrant posttranscriptional processing of vector RNA, i.e., aberrant splice variants, defective proviral genomes, and internally deleted frame-shifted messenger RNAs (mRNAs) (197, 198). New vector designs containing partially splice-corrected MDR1 cDNA with a silent point mutation in the cryptic splice acceptor and a leader sequence that contains an artificial intron in the 5´ untranslated region eliminated aberrant reading frames for initiation of translation and improved MDR1 transgene expression (9, 16, 17, 199, 200). Additionally, a myeloproliferative disorder was observed in mice reconstituted with bone marrow cells transduced with MDR1 (201), however, this event has not been encountered in nonhuman primate studies nor in human trials (202, 203). The level of resistance and selective advantage difference determine the preference between the wild-type or mutant form of the drug-resistance gene. O6–Methylguanine–DNA–methyltransferase (MGMT) is an important DNA repair protein that protects cells against nitrosourea cytotoxicity (2, 28, 120, 204–206). HSCs and their progeny express relatively low amounts of MGMT, and myeloprotection can be obtained by transfer of either wildtype (207) or different MGMT mutants (120, 201, 205–210) into HSCs. 6-Benzylguanine (6-BG) increases the sensitivity of both tumor cells and hematopoietic cells to nitrosoureas by inactivating MGMT. The two MGMT mutants, G156A and P140K (211), have retained DNA repair activity while providing resistance to BG. These mutants have been used in animal studies, and HSCs transduced with retroviral constructs containing these mutant cDNAs have increased tolerance to BG and BCNU (212, 213). Upon comparison with wild-type MGMT, these mutant forms demonstrate substantially higher resistance to 6-BG depletion; however, high expression of MGMT P140K mediated a selective disadvantage irrespective of the vector backbone (214, 215), and expression of the P140A/G156A double mutant was associated with reduced or unstable protein in hematopoietic cells (216–218). The advantage of using mutant MGMTs in HSC gene transfer is that BG selection is actually capable of ablating endogenous wild-type MGMT carrying untransduced primitive HSCs. This advantage might increase the therapeutic index for the drugs, however, decreased survival of transduced cells without drug selection for P140A mutant-containing vectors necessitate further evaluation of gene therapy strategies with MGMT cDNA. The other disadvantage is that nitrosureas are stem cell poisons

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and carcinogenic. At an era of insertional mutagenesis, increasing the risk for treatment-associated leukemia with BCNU or temozolomide might be of concern (219). Route of administration, either intraperitoneal or intravenous, in murine models are usually similar to bolus injections in humans, and, for those kind of injections, peak plasma and bone marrow levels of the drug could be as important as plasma half-life of the drug(s). Mimicking the drug exposure time and concentrations for in vitro assays usually provide better understanding for in vivo results. In an attempt to develop resistance against fluoropyrimidines, our mutant thymidylate synthase (TSG52S) model failed to provide in vivo myeloprotection against 5-fluorouracil (5FU) intraperitoneal injections. According to the schedule of administration, 5-FU behaves as two different drugs. In bolus 5-FU injection, it is known that the dominant drug activity is RNA synthesis inhibition, whereas intravenous infusion has more of a DNA synthaie blocking effect through thymidylate synthase (TS) inhibition (220, 221). We have observed that TSG52S-carrying bone marrow progenitors can only be protected from 5-FU toxicity by continuous drug exposure, but TSG52S gene transfer does not protect from pulse high-dose 5-FU toxicity (unpublished data). 2. We harvest mouse bone marrow from both tibias and femurs, and the average cell count varies between 3 and 6 × 107 total nucleated cells per mouse. It is our experience that bone marrow cells that are not enriched cause more unpredictable cell loss during prestimulation and transduction. We limit cell purification to the lineage-negative population, about 10% of the bone marrow total nucleated cells, and our recovery from negative selection varies between 38% and 64%, with purity between 63% and 87%. Although it is possible to further purify the HSCs, it is more time consuming and adds more steps and subsequently there is more cell loss during the process. The majority of the advanced colony-forming units spleen (CFU-S) is eliminated with 5-FU treatment, whereas primitive blast colony forming cells (CFCs), and long-term culture-initiating cells (LTC-IC) are enriched. However, 5-FU treatment not only enriches preexisting, resting HSCs, but also induces phenotypical changes (168, 221), including decreased CD117 and increased CD11b expression on longterm reconstituting cells. Four days after 5-FU administration, the average total nucleated cell count harvested becomes 107 cells/mouse, at the lowest. Purification attempts from 5-FUtreated bone marrow, such as sca+/c-kithigh/lin− cells, would only result in substantial loss of long-term reconstituting HSCs. We recommend using either a purified HSC population or a 5-FU-enriched bone marrow for transduction.


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For experimental designs testing the safety and efficiency of drug-resistance gene transfer, transfusing the amount of cells that will reconstitute hematopoiesis for all of the animals is preferred. Ex vivo manipulation with cytokines alters the engraftment potential of HSCs (222), and usually increasing the graft dose threefold may provide comparable levels of engraftment. We inject 5–7.5 × 105 lineage-negative or 2–5 × 106 5-FUenriched cells per mouse after transduction. Nonmyeloablative regimens require almost a log higher amount of cells to be transplanted (223–225) for comparable levels of engraftment with myeloablative regimens. Engraftment defects caused by ex vivo manipulation can be revealed with competitive repopulation assays. For this limiting dilution assay, animals are usually divided into three or four groups, and injected with increasing numbers of cells (twofold to threefold difference for each cell dose, and 6–12 animals per group). Animals are also injected with unmanipulated 1–2 × 105 cells/mouse, to ensure that animals will be protected from the hazards of radiation until the manipulated cells engraft. We prefer two split dose total body irradiation (TBI), 3 h apart, for recipients and we usually give TBI 24 h before the transplant. The total body irradiation dose varies between mouse strains, i.e., 1100 cGy for C57B6, and 900 cGY for nu/nu mouse. Engraftment of immunodeficient NOD/SCID mice by human HSCs requires conditioning by irradiation. NODSCID mice survive sublethal irradiation doses up to 400 cGy (128, 226), and we prefer 300–350 cGy for our human HSC engraftment experiments. 3. Human CD34+ cells can be isolated from bone marrow (BM), or mobilized peripheral blood (MPB), or umbilical cord blood (UCB). The source of HSCs used for retroviral transduction affects the transduction efficiency and engraftment potential of the cells. UCB-derived CD34+ cells display the highest transduction efficiency and engraftment potential (110, 111), and adult HSCs are more dormant and less responsive to cytokine stimulation when compared with UCB. Human CD34+ cells from BM or MPB require prolonged cytokine stimulation to enter the cell cycle, to upregulate envelope receptor expression, and to be efficiently infected by oncoretroviral vectors (227, 228). Higher levels of envelope receptor mRNA are present in UCB-derived HSCs when compared with HSCs derived from BM (229). Selecting the CD34+ cells from frozen samples results in highly variable CD34+ purity and yield (112). CD34+ cells from pooled leukapheresis MPB products or fresh bone marrow samples can be selected within 24–48 h of collection.

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Thawing the frozen purified human CD34+ cells in dextran/ albumin solution increases the postthaw viability (230). Frozen/thawed UCB-derived CD34+ cells have been shown to express higher levels of amphotropic receptor mRNA but not of GaLV receptor (Pit 1) mRNA (190, 231, 232). The ability of purified CD34+ cells to respond to cytokine stimulation does not change upon freezing and thawing (190, 231–233). The level of engraftment levels of human cells varies for NOD/SCID, NOD/SCID-β2-microglobulinnull, or NOD/ SCID-γcnull (149). The numbers of human CD34+ cells mentioned in Subheading 3.4 usually provide a human CD45 engraftment level between 0.5 and 5% when injected into NOD/SCID mice. Mouse bone marrow and thymus microenvironment fail to support human stem cell proliferation and differentiation, especially T lymphopoiesis. On the other hand, human CD34+ cells injected into NOD/SCID-β2microglobulinnull or NOD/SCID-γcnull mice could establish human engraftment in much higher levels, as well as human T lymphopoiesis (124). Nevertheless, immunodeficient mouse– human HSC experiments mainly provide the evidence for engraftment potential of manipulated human CD34+ cells. 4. For gamma-retroviral transductions, prestimulation is required for stimulating the quiescent HSCs. Early cytokines, i.e., SCF, TPO, FLT3-ligand, are known to be effective for inducing cell cycling in 24–48 h (115, 124). Adding IL-3, IL-6, or granulocyte colony-stimulating factor (G-CSF) to the cytokine combination might increase the number of cells generated during the ex vivo manipulation, but, quantitatively, those cells have less potential to engraft than SCF- and TPO-stimulated cells (147, 234). Proliferation, or cell division, per se, is not necessary for efficient lentiviral HSC transduction, but cytokines during a transduction period of 24 h increase the transduction rate and preserve the engraftment potential of transduced HSCs and inhibit apoptosis (235). 5. Prestimulation with early acting cytokines is also required for efficient gene transfer into human HSCs with oncoretroviral vectors. The cytokine combination of SCF, FLT3L, and TPO, compared with several cytokine combinations that contain IL-3 and/or IL-6 (146), induces better, more synchronous ex vivo CD133+ (143) or CD34+/Thy.1+ (146) cell expansion, yields higher transgene marking (147), improves survival (110–114), and maintains the multipotential engraftment ability of HSCs in NOD/SCID mice (115). Ex vivo stimulation with hematopoietic growth factors increases the expression of the envelope receptors on HSCs (116–119) and also maintains cell viability by inhibiting HSC apoptosis (149). Prolonged ex vivo manipulation of HSCs may result in either differentiation of HSCs


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and/or loss of their engraftment potential (114, 236). Human CD34+ cells enter active cycling after 36–48 h of cytokine stimulation (125, 126, 143, 148–154), while the expression of Pit-1 is concomitantly induced (131, 237). The length of prestimulation, the number of transduction cycles, and the time interval between the two transductions varies among various published retroviral gene transfer protocols (176, 238), so that total ex vivo cell manipulation time fluctuates between 60 h and 120 h. 6. Generation and selection of high-titer stable producer cell lines are beyond the scope of this chapter. Producer cell lines are usually derivatives of mouse embryonic fibroblasts, i.e., AM-12, PG13, GP+E86, PA317, or highly transfectable fetal human kidney cells, 293T, Phoenix, or human HT1080 fibrosarcoma cell line, i.e., FLYA13, TEFLYA. Cross-infection of the packaging cells with vector stocks originally obtained by transfection increases the number of high-titer packaging clones relative to direct transfection (181, 239). It is possible to generate stable high-titer clones by cross-infection with vector stocks derived from either Phoenix-Eco or VSV-G pseudotyped 293GPG cells (240). Stable bipartite retroviral packaging cell lines usually contain markers that allow selection against loss of the plasmid DNAs conferring the packaging functions. Using serum-free conditions for ex vivo transduction protocols increases the biosafety and decreases the immunogenicity of retroviral transduction (241). A wide variety of serum-free media have been developed and are shown to demonstrate better cell expansion then serum-containing media (125, 242). Retroviral packaging cells are serum-dependent for proliferation, but they could adapt to short-term culture in serum-free medium, allowing serum-free vector stock collection (13, 80, 100, 125, 178, 242–244); however, vector production usually requires a relatively prolonged incubation for vector harvest (245). Large-scale vector production either with stable producer cell lines, or by transient transfection requires a substantial amount of optimization, i.e., cell density, media content, media volume, incubation temperature, cell growth vessel, plasmid DNA concentration, harvest time, and cycles (125, 178, 242, 245). The data is usually specific to the producer cell line, vector design, and envelope protein. Reducing the vector harvest volume, i.e., 0.1 mL media/ cm2, increases the vector titers (10, 125, 178, 246), and repetitive harvests of vector stocks are usually feasible over several days (242, 247). The optimal incubation temperature for retroviral vector harvest is somewhat controversial. Some studies show greater retroviral vector inactivation and/or lower vector titers at 37°C when compared with 32°C (133). On the other hand, the viral particles produced at 37°C are less rigid than those produced at 32°C. They are therefore more stable

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and thus provide a higher transduction efficiency (125). We recommend comparing the titers of vector stocks harvested at 32°C and 37°C side by side for the selected high-titer packaging cell clones. VSV-G, RD114 pseudotyped vectors are usually concentrated with ultracentrifugation at 50,000×g for 90 min at 4°C, but the envelope proteins of amphotropic, ecotropic, or GALV vectors are not sheer-force resistant and cannot be concentrated with ultracentrifugation. The stress of centrifugation and filtration often cause the surface domain of those proteins to be shed. Sheer-force sensitive viral particles pseudotyped with Eco, Ampho, or GaLV envelopes, can only be concentrated up to 10-fold by either low-speed centrifugation (9,500 rpm in Beckman rotor JA-14 at 4°C for 12 h) (125), or centrifugation and filtering for 35 min at 3,000×g, 15°C, through centrifugal filter devices with a 100,000 molecular weight cut-off (178, 248). It is common research practice to filter vector stocks through a 0.45-μm filter to remove cellular debris. Do not use filters that contain detergents, because wetting agents will affect the integrity of the viral particles as well as the viability of the target cells. Frozen vector stocks collected in serum-free conditions and stored at −70°C are stable during a period of at least 12 months (125, 178). The half-life of frozen vector stocks is suggested to be biphasic. After 25–30% loss of vector titer upon the first freeze/thaw cycle, the decay of viral particles is slow, and the half-life of this stage varies from 18 to 41 months (249, 250). Interestingly, testing the frozen vector stocks only on target/indicator cell lines may be misleading, because transduction efficiency in primary cells may not show the same decrease (113, 251). This different outcome between cell types may be caused by differential cell surface receptor saturation of envelope receptors on indicator cell lines. Vector particle counts can be determined either by direct particle count using an electron microscope, or by indirect methods such as quantitative real-time PCR (125) or transduction of target/indicator cells. Vector stocks contain infectious viral particles as well as noninfectious particles. The vector titers defined by transduction of target cells depend in part on the level of expression of the envelope receptors (122, 247, 248, 252–254), which varies among target cells (132, 133). We recommend initially testing the transduction efficiency on both the type of primary cells that will be used for clinical application and also the various target/indicator cell lines, and for subsequent vector titrations, to select the cell line that gives transduction rates comparable to that obtained on primary cells. 7. Retroviral particle stability is susceptible to different factors, i.e., temperature, pH, ionic strength, shear stress. Retroviral


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particles move in a random Brownian motion, and have a relatively short half-life, 5–8 h, at 37°C (252) and only diffuse 0.5 mm during that half-life (89, 125). Viral particles do not diffuse more than 1 mm during a 12-h transduction period, but confining the transduction media height below 2 mm/ cm2 of surface area limits the nutrients for the cells (unpublished data). Additionally, the half-lives of VSV-G pseudotyped retroviral vectors drop dramatically below pH 6 and above pH 8 (124, 125), and ecotropic vectors also remain infectious in a narrow pH range from 5.5 to 8. Virus inactivation beyond these limits of pH is fast and irreversible. Limiting the transduction volume also influences the vector stability because of wider pH changes during the culture. Therefore, increasing the target cell uptake of the retroviral particles by forcing the particles to the vicinity of the target cells, i.e., polycations, spinoculation, or preloading, is a better option for enhancing the transduction efficiency. However, these parameters should be tested rigorously for each individual vector supernatant because nonpurified vector stocks contain soluble factors that might be toxic to the cells and reduce transduction efficiency, and those factors may also get concentrated during the process. Spinoculation of Phoenix-Eco-derived retroviral particles increased target cell death in our experimental systems but Ampho- or GALV-enveloped vectors did not alter target cell viability (unpublished data). We tested spinoculation on primary and different cancer cell lines at 500×g, 1,000×g, and 1,200×g for 1, 2, 4, and 6 h at 15°C. Cell viability decreased more than 50% after 4 and 6 h of spinoculation. We compared 2 h of spinoculation for transduction of mouse BM cells and human CD34+ cells. Spinoculation, at 1,000×g for 2 h at 15°C, increased transduction efficiency of the cells 20–35%, without substantial cell loss or vector copy number increase per cell (unpublished validation data). Adding polycations to the transduction media does not increase transduction efficiency for transductions held on RetroNectin®-coated plates (89, 115), and, furthermore, polycations may be toxic for target cells under serum-free conditions (125). For VSV-G-pseudotyped vectors, neither RetroNectin® nor polycations increase the transduction efficiency, but RetroNectin® provides growth supporting and anti-apoptotic effects on HSCs both in vitro and in vivo. The effect of RetroNectin® was of the same magnitude for all of the doses we tested (2–20 μg/cm2). 8. In this protocol, DNA is extracted from individual hematopoietic colonies. The colonies growing in semisolid media usually contain fewer than 200 cells, and any DNA isolation method needs at least a couple of thousand cells to start with. The amount of DNA extracted varies from picograms to a couple

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of nanograms. When the isolated material is used to detect the transgene of interest by PCR, a second control PCR using β-actin primers has to be run to prove the presence of DNA in the sample. The failure rate of isolating DNA from the samples depends on the size of the colony but is usually less than 15%. The number of colonies to pick depends on the expected transduction efficiency. There should be at least one colony among the picked colonies that will give a positive PCR product. Example: If the transduction efficiency in the hematopoietic progenitors is around 2%, you need to pick at least 58 colonies (1 out of 50 will be positive, and there can be a 15% failure in extracting DNA, so 50 + 15% × 50 = 58). 9. Molecular follow-up of the gene transfer studies require quantization of vector copy number and transgene expression. For that purpose, we use real-time PCR, which detects the fluorescence emitted during amplicon production of each PCR cycle. We prefer fluorescence labeled probes instead of SYBR Green, because they are not influenced by nonspecific amplification. Real-time PCR quantization offers a dynamic range of up to 107-fold, and consequently allows quantification of very low concentrations of target mRNA. For vector copy numbers determination with absolute quantization, we use 100 ng of DNA per reaction. cDNA plasmids are the preferred standards for absolute quantization. We generate standard curves with human serum albumin plasmid and our transgene plasmid dilutions in 100 ng of salmon sperm DNA (255). We design TaqMan probes longer than the primers (20–30 bases long with a melting temperature [Tm] value of 10°C higher than the Tm of the primers) that contain a fluorescent dye usually on the 5′ base, and a quenching dye typically on the 3′ base. Real-time PCR allows detection of multiple DNA sequences by designing each probe with a spectrally unique fluorophore and quencher pair. This multiplex real-time PCR design enables the target (transgene) and endogenous control (i.e., human serum albumin, 18S mRNA) to be amplified in single tube, however, the amplification efficiencies of the primer–probe sets used in multiplex reactions should be the same. The efficiency of the PCR should be 90–100%. Multiplex real-time PCR assays can be performed using multiple dyes with distinct emission wavelengths. We prefer making fourfold dilutions of plasmids in a series of eight dilutions, which allows us to detect 4 to 65,536 vector copy numbers per reactions. For transgene expression studies, we use the ΔΔCT relative quantitation method. Valid CT calculations require the efficiency of the target amplification and the efficiency of the reference amplification to be approximately equal. Relative transgene expression comparisons work best when the gene


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expression of the chosen endogenous control is more abundant and remains constant, in proportion to total RNA, among the samples. By using an invariant endogenous control as an active reference, quantitation of transgene mRNA can be normalized for differences in the amount of total RNA added to each reaction. We prefer either β-actin or 18S mRNA as the endogenous normalizer, because GAPDH expression may vary because of its upregulation in proliferating cells. The chosen endogenous mRNA should be proportional to the amount of input RNA. It is preferable to validate the chosen normalizer for the target cell or tissue. Primer probe design takes the center attention for transgene expression assays by using real-time reverse transcriptase PCR. For avoiding the false-positive results caused by amplification of contaminating genomic DNA, it is preferable to have primers spanning exon–exon junctions in the cDNA sequence. Reverse transcription of total RNA to cDNA should be done with random hexamers. We use 10–50 ng of target cDNA in our assays. References 1. Bertino JR. “Turning the tables”—making normal marrow resistant to chemotherapy. J Natl Cancer Inst 1990;82(15):1234–5. 2. Persons DA, Allay ER, Sawai N, et al. Successful treatment of murine beta-thalassemia using in vivo selection of genetically modified, drug-resistant hematopoietic stem cells. Blood 2003;102(2):506–13. 3. Havenga MJ, Werner AB, Valerio D, van Es HH. Methotrexate selectable retroviral vectors for Gaucher disease. Gene Ther 1998;5(10):1379–88. 4. Carella AM, Congiu AM, Gaozza E, et al. High-dose chemotherapy with autologous bone marrow transplantation in 50 advanced resistant Hodgkin’s disease patients: an Italian study group report. J Clin Oncol 1988;6(9):1411–6. 5. Kollmannsberger C, Mayer F, Kuczyk M, Kanz L, Bokemeyer C. Treatment of patients with metastatic germ cell tumors relapsing after high-dose chemotherapy. World J Urol 2001;19(2):120–5. 6. Vuento MH, Salmi TA, Remes KJ, Grenman SE. Hematological toxicity of salvage treatment after high-dose chemotherapy and conventional chemotherapy of ovarian cancer. Anticancer Res 2002;22(2B):1151–5. 7. Rapoport AP, Guo C, Badros A, et al. Autologous stem cell transplantation followed by consolidation chemotherapy for relapsed or refractory Hodgkin’s lymphoma. Bone Marrow Transplant 2004;34(10):883–90.

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14. Sugimoto Y, Sato S, Tsukahara S, et al. Coexpression of a multidrug resistance gene (MDR1) and herpes simplex virus thymidine kinase gene in a bicistronic retroviral vector Ha-MDR-IRES-TK allows selective killing of MDR1-transduced human tumors transplanted in nude mice. Cancer Gene Ther 1997;4(1):51–8. 15. Licht T, Goldenberg SK, Vieira WD, Gottesman MM, Pastan I. Drug selection of MDR1-transduced hematopoietic cells ex vivo increases transgene expression and chemoresistance in reconstituted bone marrow in mice. Gene Ther 2000;7(4):348–58. 16. Cowan KH, Moscow JA, Huang H, et al. Paclitaxel chemotherapy after autologous stem-cell transplantation and engraftment of hematopoietic cells transduced with a retrovirus containing the multidrug resistance complementary DNA (MDR1) in metastatic breast cancer patients. Clin Cancer Res 1999;5(7):1619–28. 17. Abonour R, Williams DA, Einhorn L, et al. Efficient retrovirus-mediated transfer of the multidrug resistance 1 gene into autologous human long-term repopulating hematopoietic stem cells. Nat Med 2000;6(6):652–8. 18. Capiaux GM, Budak-Alpdogan T, Takebe N, et al. Retroviral transduction of a mutant dihydrofolate reductase-thymidylate synthase fusion gene into murine marrow cells confers resistance to both methotrexate and 5-Fluorouracil. Hum Gene Ther 2003;14(5):435–46. 19. Sauerbrey A, McPherson JP, Zhao SC, Banerjee D, Bertino JR. Expression of a novel double-mutant dihydrofolate reductasecytidine deaminase fusion gene confers resistance to both methotrexate and cytosine arabinoside. Hum Gene Ther 1999; 10(15):2495–504. 20. Mineishi S, Nakahara S, Takebe N, Banerjee D, Zhao SC, Bertino JR. Co-expression of the herpes simplex virus thymidine kinase gene potentiates methotrexate resistance conferred by transfer of a mutated dihydrofolate reductase gene. Gene Ther 1997; 4(6):570–6. 21. Takebe N, Zhao SC, Adhikari D, et al. Generation of dual resistance to 4-hydroperoxycyclophosphamide and methotrexate by retroviral transfer of the human aldehyde dehydrogenase class 1 gene and a mutated dihydrofolate reductase gene. Mol Ther 2001;3(1):88–96. 22. Beausejour CM, Le NL, Letourneau S, Cournoyer D, Momparler RL. Coexpression of cytidine deaminase and mutant

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Chapter 35 Phase I Clinical Trial of Locoregional Administration of the Oncolytic Adenovirus ONYX-015 in Combination with Mitomycin-C, Doxorubicin, and Cisplatin Chemotherapy in Patients with Advanced Sarcomas Mateusz Opyrchal, Ileana Aderca, and Evanthia Galanis Summary Despite many advances in cancer therapy, metastatic disease continues to be incurable in the majority of cancer patients. There is an need for more efficient and less toxic treatments in this setting. Oncolytic virotherapy represents a novel promising direction in the treatment of cancer. Based on preclinical and clinical data, combination with standard chemotherapy has the potential to further increase the antitumor activity of oncolytic virotherapy in a synergistic manner. We present the design of a phase I clinical trial combining intratumoral injections of the oncolytic adenovirus ONYX-015 with systemic chemotherapy in patients with advanced sarcomas. Key words: Cisplatin chemotherapy, doxorubicin, mitomycin-C, oncolytic adenovirus, ONYX-015, sarcomas

1. Introduction ONYX-015 (dl1520) is chimeric human group C adenovirus that has been genetically engineered to incorporate deletions in the E1B-55k and E3B regions (1). The E1B protein in conjunction with E4ORF6 binds to the tumor suppressor protein p53 (2). This interaction causes p53 to be degraded and prevents it from causing cell cycle arrest, thus leading to a productive viral infection. Because approximately 50% of human cancers have p53 mutations, and these are often associated with chemotherapy resistance, this could represent an attractive approach in the treatment of cancer (3, 4). Subsequent work, however, has shown that ONYX-015 Wolfgang Walther and Ulrike S. Stein (eds.), Methods in Molecular Biology, Gene Therapy of Cancer, vol. 542 © Humana Press, a part of Springer Science + Business Media, LLC 2009 DOI: 10.1007/978-1-59745-561-9_35

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can grow efficiently in cancer cells with wild-type p53 (5, 6). This can at least in part be explained by p53 inactivation due to other mechanisms, such as mdm2 amplification (7, 8). In 1996, ONYX-015 was the first oncolytic virus to enter human clinical trials (9). The virus has demonstrated a remarkable safety profile, independent of the route of administration. The maximum tolerated dose has not been reached after intratumoral, hepatic artery, intraperitoneal, or systemic administration of the virus (8, 10–12). The initial trials tested the efficacy of the ONYX-015 as a single agent. Two phase II trials evaluated the virus in patients with recurrent head and neck cancer, after intratumoral injections. An approximately 15% response rate with no objective evidence of effect in uninjected tumors was observed (9, 13). No antitumor responses were seen, however, after intratumoral administration in patients with pancreatic cancer, intraperitoneal administration in patients with ovarian cancer, or intravenous administration in patients with colorectal cancer (12, 14, 15). These results emphasize the limited activity of ONYX-015 when used as single agent. Preclinical work both in vitro and in animal models has demonstrated synergistic activity between ONYX-015 and 5-fluorouracil (5FU) or cisplatin chemotherapy (7). In a subsequent phase II trial in recurrent head and neck cancer patients, approximately 65% of patients responded when treated with ONYX-015, cisplatin, and 5FU; this compares favorably with the historic response rate of 25–35% in this patient population (16). Injected tumors were significantly more likely to respond to the virus/chemotherapy combination as compared with noninjected tumors (p < 0.01). A phase III trial testing this approach in recurrent head and neck patients was aborted in the USA prior to its completion. In March of 2006, however, H101, a virus very similar to ONYX-015, was approved by the China’s State Food and Drug Administration for treatment of head and neck cancer by intratumoral injection in combination with chemotherapy. The H101 virus in addition to the E1B-55k gene deletion lacks all E3 proteins. This regulatory approval was based on a phase III clinical trial, which showed a 79% response rate in patients who received intratumoral administration of H101 in combination with platinum-based chemotherapy vs 40% in controls without virus treatment (p < 0.001). Shanghai Sunway Biotechnology, the company marketing H101, has also acquired US development rights to ONYX-015 (17). The protocol we describe here investigated an ONYX-015 chemotherapy combinatorial approach in patients with advanced sarcomas. Sarcomas represent a rational target for treatment with ONYX-015. They have a high frequency of p53 mutations ranging from 40% to 75%. In addition, MDM-2 gene amplification, which can result in functional p53 inactivation, occurs in another 10–30%

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of sarcoma cases (18, 19). Furthermore, in preclinical experiments, ONYX-015 has been shown to replicate effectively and cause significant cytotoxicity against the sarcoma line S4052 (18). In this trial, we combined ONYX-015 with antisarcoma agents that either are synergistic with ONYX-015, such as cisplatin, or for which ONYX-015 can reverse resistance, such as doxorubicin (7). The goals of study were: 1. To define the maximum tolerated dose (MTD) of ONYX-015 in combination with MAP chemotherapy (mitomycin C, doxorubicin, and cisplatin) in patients with metastatic sarcoma. 2. To assess ONYX-015 replication in sarcomas. 3. To assess the antitumor activity of the regimen (secondary endpoint). 4. To correlate, in a preliminary fashion, ONYX-015 replication with the presence of neutralizing antibodies. 5. To correlate any observed responses with the p53 and MDM-2 tumor status.

2. Materials 2.1. Laboratory Evaluation of Patient Samples 2.1.1. Immunohistochemistry for p53 and mdm2 Expression

1. Anti-p53 antibody M7146 (Dako, Glostrup, Denmark).

2.1.2. Plasma Neutralizing Antibody Titers

1. 12-Well plates (Fisher Scientific, Hampton, NH).

2. Anti-mdm2 antibody clone SMP14 (Dako). 3. HRP+/DAB+ chromogen substrate (Dako).

2. Agarose (Invitrogen Life Technologies, Carlsbad, CA). 3. HEK 293 cells (ATCC, Manassas, VA).

2.1.3. Detection of ONYX-015 Viral Genome in Plasma

1. QiaAmp Blood Kit (Qiagen, Almada, CA). 2. Carrier DNA (Dualsystems, San Mateo, CA). 3. Oligonucleotide primers. The following primers and probe where employed: forward primer 5′-GCTGGCGCAGAAGTATTCCA, reverse primer 5′-GTGCGGGTCTCATCGTACCT and the probe 5′FAM-ACCTTCCAGATCCGTCGACCTGCA-TAMRA. 4. DNA sequencer (Perkin-Elmer, Foster City, CA). 5. Standard polymerase chain reaction (PCR) reagents. 6. Standard reagents and apparatus for agarose gel electrophoresis.

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2.1.4. Detection of Adenovirus DNA in Tumor Biopsies

1. Biotinylated adenovirus DNA probe (Enzo Diagnostics, Farmingdale, New York). 2. Proteinase K (Invitrogen, Carlsbad, CA). 3. 4% paraformaldehyde. 4. Nitroblue tetrazolium/bromochloroindolyl phosphate (NBT /BCIP) (Roche Applied Science, Indianapolis, IN). 5. Nuclear fast red (Vector Laboratories, Burlingame, CA). 6. Streptavidin-alkaline phosphatase conjugate (Vector Laboratories).

3. Methods 3.1. Clinical Trial 3.1.1. Inclusion/ Exclusion Criteria Inclusion Criteria

1. Patients were required to have a histologic diagnosis of sarcoma beyond surgical cure with at least two measurable sites of metastatic disease. Injections of the viral agent were administered in one of the metastatic disease sites, while the remainder sites of metastatic disease served as control. 2. The injected lesion should have been between 1 and 10 cm in size. 3. 18 years of age or older. 4. Life expectancy ³3 months. 5. Adequate hematologic, renal, and hepatic function.

Exclusion Criteria

1. Poor performance score of Eastern Cooperative Oncology Group (ECOG) 3 or 4. 2. Uncontrolled infection. 3. Metastatic disease not amenable to intratumoral administration of the virus. 4. Other concurrent chemotherapy, immunotherapy, or radiation therapy. 5. More then two previous regimens for metastatic disease. 6. To avoid cardiotoxicity, patients with a previous maximum dose of doxorubicin ³250 mg/m2 were not eligible. The protocol was approved by the Mayo Clinic Institutional Review Board, and informed consent was obtained from all the patients.

3.1.2. Pretreatment Evaluation/Follow-Up Studies

1. Pretreatment evaluation included a history and physical examination, complete blood cell count, serum chemistries, and coagulation profile.

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2. Evaluation/measurement of injected and control tumors and assessment of disease status was performed by using appropriate imaging modalities, such as computed tomography (CT) scan, magnetic resonance imaging (MRI), or chest x-ray. 3. Follow-up studies included physical examination, complete blood cell counts, serum chemistries, and imaging performed every 4 weeks. 4. Neutralizing antibody titers were obtained at baseline and before cycle 2. 5. ONYX-015 genomes in the peripheral blood were quantified at baseline, on day 5 of cycle 1, and before cycle 2. In addition, one patient had extended sampling performed for viral kinetics, consisting of additional blood samples on days 8, 10, and 12 (3, 5, and 7 days after the day 5 dose) of cycle 1. 6. Tumor biopsy for confirmation of diagnosis and determination of p53 and mdm2 status was performed at baseline, and repeat biopsy to assess viral replication by in situ hybridization was obtained on day 5 of cycle 1 (Table 1 summarizes the test schedule). 3.1.3. Study Drug

The ONYX-015 virus was produced in human embryonic kidney 293 cells, as previously described, formulated in Tris buffer (10 μM Tris pH 7.4, 1 μM MgCl2, 150 μM NaCl, 10% glycerol) and supplied frozen (−20°C) in 0.5-mL vials (9). The viral particle (vp) to plaque-forming units (pfu) ratio was 20. Wild-type adenovirus was undetectable in all clinical lots of ONYX-015 at a limit of detection of 1 in 1011 particles. Thawed ONYX-015 was maintained at 2–8°C during dilution and handling and administered to patients within 2 h from viral preparation.

3.1.4. Treatment Schedule (see Note 1)

The study included following three viral dose levels per dose (Table 2): 1. Level 1: 109 pfu of virus per dose. 2. Level 2: 3 × 109 pfu per dose. 3. Level 3: 1010 pfu per dose. The virus was administered intratumorally in multiple sites throughout the tumor mass on days 1–5 of each treatment cycle for a total dose per cycle of: 1. 5 × 109 pfu in dose Level 1. 2. 1.5 × 1010 pfu in dose Level 2. 3. 5 × 1010 pfu in dose Level 3. The volume of viral administration was determined in conjunction with the injected tumor volume, the intent being to saturate as much of the tumor as possible with the viral solution. The injected tumor volume was calculated by multiplying the three dimensions of the lesion to be injected and dividing by 2. Subsequently

X X X

X X X X

Chemistry group AST, total Bili., Alk Phos, Creat, INR

Na, K, Mg

Chest X-ray

Evaluation/measurement of injected and control tumors and disease status (CXR, CT, MRI, etc.)2

Serum neutralizing antibodies

Tumor biopsy for viral replication

Tumor biopsy for confirmation of diagnosis and determination of p53, mdm-2 status

Viral titers (plasma)

X

Serum pregnancy test4

X

X3

Cycle 2 only

Cycles 2, 4, and 6 only

X

X

X

X

At 3 mo after treatment only; thereafter as clinically indicated

X

X

X

2

Day 8 biopsy was encouraged (if deemed safe) but was not mandatory. The same imaging modality should have been used for a given patient throughout the study. The study radiologist was to review all imaging studies. 3 As clinically indicated in association with MAP chemotherapy. 4 For women of childbearing potential only. Must be done

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