Oncolytic Viruses (Methods in Molecular Biology, v797)

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Author: David H. Kirn | Ta-Chiang Liu | Stephen H. Thorne

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METHODS

IN

MOLECULAR BIOLOGY™

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

For further volumes: http://www.springer.com/series/7651

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Oncolytic Viruses Methods and Protocols

Edited by

David H. Kirn Jennerex Inc., San Francisco, CA, USA and University of Oxford School of Medicine, Oxford, UK

Ta-Chiang Liu The Johns Hopkins University Medical Center, Baltimore, MD, USA and Washington University in St. Louis, St. Louis, MO, USA

Steve H. Thorne Department of Surgery and Immunology, University of Pittsburgh Cancer Institute, University of Pittsburgh, Pittsburgh, PA, USA

Editors David H. Kirn, MD Jennerex Inc. San Francisco, CA, USA and University of Oxford School of Medicine Oxford, UK [email protected]

Ta-Chiang Liu The Johns Hopkins University Medical Center Baltimore, MD, USA and Washington University in St. Louis St. Louis, MO, USA [email protected]

Steve H. Thorne, Ph.D. Department of Surgery and Immunology University of Pittsburgh Cancer Institute University of Pittsburgh Pittsburgh, PA, USA [email protected]

ISSN 1064-3745 e-ISSN 1940-6029 ISBN 978-1-61779-339-4 e-ISBN 978-1-61779-340-0 DOI 10.1007/978-1-61779-340-0 Springer New York Dordrecht Heidelberg London Library of Congress Control Number: 2011936969 © Springer Science+Business Media, LLC 2012 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. Cover illustration: Springer Images Printed on acid-free paper Humana Press is part of Springer Science+Business Media (www.springer.com)

Preface Since the ﬁrst report of an engineered oncolytic virus by Martuza et al. two decades ago, there has been a continuing and steady increase of interest in the ﬁeld. The keyword “oncolytic virus” is associated with nearly 300 publications from Pubmed in the year of 2009 alone. Herpes simplex virus (HSV) and adenovirus (Ad) were among the ﬁrst virus species to be engineered for oncolytic purposes, while the spectrum of virus species tested has since broadened to include vesicular stomatitis virus (VSV), reovirus, myxoma virus, vaccinia virus, measles virus, and Newcastle disease virus (NDV), among others. Although several of these virus species are inherently tumor-selective, others rely on attenuating or tumor-targeting modiﬁcations. During the early days of development, the majority of the assays utilized in oncolytic virus research were, not surprisingly, those commonly used in virology research. These include assays for bulk production, replication, cytopathic effects, and in vivo efﬁcacy. Over the last decade, the growth of knowledge in cancer biology, virology, and immunology, as well as advances in molecular biology, genetics, bioinformatics, and imaging technologies, have led to numerous and exciting novel discoveries, many of which have been incorporated into clinical trial designs. The long history of oncolytic adenovirus research has led to the development of several approaches designed to improve the selectivity and potency of the virus, some of which are also applicable to other virus species. In this volume, Giménez-Alejandre et al. describe the construction and puriﬁcation of capsid-modiﬁed adenoviruses, and Doronin and Shayakhmetov describe the construction of armed oncolytic adenoviruses. Protocols for many individual virus species are covered, including engineering and preparation of oncolytic HSV by Agarwalla and Aghi, propagation, puriﬁcation, and in vivo testing of oncolytic VSV by Diallo et al., and Msaouel et al. on retargeting oncolytic measles virus through ligand display. Properties of oncolytic reovirus and NDV are covered by Shmulevitz et al. and Fournier et al., respectively. Thorne describes the generation and testing of next generation of oncolytic vaccinia virus. As the host immune system plays a critical role in determining efﬁcacy of oncolytic viruses, we devote two chapters in the study of immune response. In vitro study of innate immune signaling and its impact on oncolytic virus is described by Heiber and Barber, whereas Dhar et al. describe the use of Syrian Hamster as an immunocompetent model for oncolytic adenovirus. Two imaging modalities are introduced in this volume: application of luciferase-expressing virus by Barry et al., and in vivo positron emission tomography (PET) imaging with the sodium iodide symporter as a reporter gene is described by Tran et al. Recent advances in stem cell research have led the ﬁeld in two distinct directions: the use of stem cells as carrier vehicles for oncolytic viruses, and the targeting of cancer stem cells. The former is described by Ahmed et al. while the later is covered by Alonso et al. Finally, the use of explant tissue samples from patients may provide useful information in predicting responses prior to clinical translation; this is covered in detail by Hallden.

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Ultimately, the success of this ﬁeld depends on breakthroughs in clinical studies. However, bench research remains vital for the translation of research in this ﬁeld. We hope the assays covered in this volume will assist you in your research. San Francisco, CA, USA St. Louis, MO, USA Pittsburgh, PA, USA

David H. Kirn Ta-Chiang Liu Steve H. Thorne

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

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1 Oncolytic Herpes Simplex Virus Engineering and Preparation . . . . . . . . . . . . . Pankaj K. Agarwalla and Manish K. Aghi 2 Construction of Capsid-Modified Adenoviruses by Recombination in Yeast and Purification by Iodixanol-Gradient . . . . . . . . . . . . . . . . . . . . . . . . Marta Giménez-Alejandre, Alena Gros, and Ramón Alemany 3 Construction of Targeted and Armed Oncolytic Adenoviruses. . . . . . . . . . . . . Konstantin Doronin and Dmitry M. Shayakhmetov 4 Syrian Hamster Tumor Model to Study Oncolytic Ad5-Based Vectors . . . . . . . Debanjan Dhar, Karoly Toth, and William S.M. Wold 5 Adenoviral Gene Expression and Replication in Human Tumor Explant Models. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Gunnel Halldén 6 Imaging Luciferase-Expressing Viruses . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Michael A. Barry, Shannon May, and Eric A. Weaver 7 In Vivo Positron Emission Tomography Imaging Using the Sodium Iodide Symporter as a Reporter Gene . . . . . . . . . . . . . . . . . . . . . . Lucile Tran, Nadège Dartial, Cecilia Hindorf, and Georges Vassaux 8 Maintaining and Loading Neural Stem Cells for Delivery of Oncolytic Adenovirus to Brain Tumors . . . . . . . . . . . . . . . . . . . . . . . . . . . . Atique U. Ahmed, Ilya V. Ulasov, Ronald W. Mercer, and Maciej S. Lesniak 9 Targeting Brain Tumor Stem Cells with Oncolytic Adenoviruses . . . . . . . . . . . Marta M. Alonso, Hong Jiang, Candelaria Gomez-Manzano, and Juan Fueyo 10 Propagation, Purification, and In Vivo Testing of Oncolytic Vesicular Stomatitis Virus Strains . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jean-Simon Diallo, Markus Vähä-Koskela, Fabrice Le Boeuf, and John Bell 11 Oncolytic Measles Virus Retargeting by Ligand Display. . . . . . . . . . . . . . . . . . Pavlos Msaouel, Ianko D. Iankov, Cory Allen, Stephen J. Russell, and Evanthia Galanis 12 Exploring Host Factors that Impact Reovirus Replication, Dissemination, and Reovirus-Induced Cell Death in Cancer Versus Normal Cells in Culture . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Maya Shmulevitz and Patrick W.K. Lee

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13 Analysis of Three Properties of Newcastle Disease Virus for Fighting Cancer: Tumor-Selective Replication, Antitumor Cytotoxicity, and Immunostimulation . . . . . . . . . . . . . . . . . . . . . . Philippe Fournier, Huijie Bian, József Szeberényi, and Volker Schirrmacher 14 Next-Generation Oncolytic Vaccinia Vectors . . . . . . . . . . . . . . . . . . . . . . . . . . Steve H. Thorne 15 Evaluation of Innate Immune Signaling Pathways in Transformed Cells. . . . . . Joshua F. Heiber and Glen N. Barber Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Contributors PANKAJ K. AGARWALLA s Harvard Medical School, Boston, MA, USA MANISH K. AGHI s Department of Neurosurgery, University of California at San Francisco, San Francisco, CA, USA ATIQUE U. AHMED s The University of Chicago, Chicago, IL, USA RAMÓN ALEMANY s Laboratori de Recerca Translacional, Institut Català d’Oncologia – IDIBELL, L´Hospitalet de Llobregat, Barcelona, Spain CORY ALLEN s Mayo Clinic, Rochester, MN, USA MARTA M. ALONSO s The University of Texas, M. D. Anderson Cancer Center, Houston, TX, USA GLEN N. BARBER s University of Miami School of Medicine, Miami, FL, USA MICHAEL A. BARRY s Department of Internal Medicine, Mayo Clinic, Rochester, MN, USA; Department of Immunology, Mayo Clinic, Rochester, MN, USA; Department of Molecular Medicine, Mayo Clinic, Rochester, MN, USA JOHN BELL s Ottawa Hospital Research Institute, Center for Innovative Cancer Research, University of Ottawa, Ottawa, ON, Canada HUIJIE BIAN s Fourth Military Medical University, Xian, China NADÈGE DARTIAL s INSERM U892, Nantes, France JEAN-SIMON DIALLO s Ottawa Hospital Research Institute and University of Ottawa, Ottawa, ON, Canada DEBANJAN DHAR s Saint Louis University, St. Louis, MO, USA KONSTANTIN DORONIN s Division of Medical Genetics, University of Washington, Seattle, WA, USA PHILIPPE FOURNIER s German Cancer Research Center(DKFZ), Heidelberg, Germany JUAN FUEYO s Department of Neuro-Oncology, The University of Texas, M. D. Anderson Cancer Center, Houston, TX, USA EVANTHIA GALANIS s Department of Molecular Medicine, Mayo Clinic, Rochester, MN, USA CANDELARIA GOMEZ-MANZANO s The University of Texas, M. D. Anderson Cancer Center, Houston, TX, USA MARTA GIMÉNEZ-ALEJANDRE s Institut Català d’Oncologia – IDIBELL, L´Hospitalet de Llobregat, Barcelona, Spain ALENA GROS s National Cancer Institute, US National Institutes of Health, Bethesda, MD, USA GUNNEL HALLDÉN s Centre for Molecular Oncology, Barts Cancer Institute, Queen Mary University of London, London, UK JOSHUA F. HEIBER s University of Miami School of Medicine, Miami, FL, USA CECILIA HINDORF s Imagerie Médicale, ONIRIS – Ecole Nationale Vétérinaire, Agroalimentaire et d’Alimentation Nantes Atlantique, Nantes, France IANKO D. IANKOV s Mayo Clinic, Rochester, MN, USA

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HONG JIANG s The University of Texas, M. D. Anderson Cancer Center, Houston, TX, USA DAVID H. KIRN s Jennerex Inc., San Francisco, CA, USA; University of Oxford School of Medicine, Oxford, UK FABRICE LE BOEUF s Ottawa Hospital Research Institute and University of Ottawa, Ottawa, ON, Canada PATRICK W.K. LEE s Department of Pathology and Microbiology & Immunology, Dalhousie University, Halifax, NS, Canada MACIEJ S. LESNIAK s The Brain Tumor Center, The University of Chicago, Chicago, IL, USA TA-CHIANG LIU s The Johns Hopkins University Medical Center, Baltimore, MD, USA; Washington University in St. Louis, St. Louis, MO, USA SHANNON MAY s Mayo Clinic, Rochester, MN, USA RONALD W. MERCER s The University of Chicago, Chicago, IL, USA PAVLOS MSAOUEL s Mayo Clinic, Rochester, MN, USA STEPHEN J. RUSSELL s Mayo Clinic, Rochester, MN, USA VOLKER SCHIRRMACHER s Tumor Immunology Program, German Cancer Research Center (DKFZ), Heidelberg, Germany DMITRY M. SHAYAKHMETOV s University of Washington, Seattle, WA, USA MAYA SHMULEVITZ s Department of Medical Microbiology and Immunology, University of Alberta, Edmonton, Alberta, Canada JÓZSEF SZEBERÉNYI s University of Pécs Medical School, Pécs, Hungary KAROLY TOTH s Saint Louis University, St. Louis, MO, USA STEVE H. THORNE s Department of Surgery and Immunology, University of Pittsburgh Cancer Institute, University of Pittsburgh, Pittsburgh, PA, USA LUCILE TRAN s INSERM U948 and Institut des Maladies de l’Appareil Digestif, CHU Hôtel Dieu, Nantes, France ILYA V. ULASOV s The University of Chicago, Chicago, IL, USA MARKUS VÄHÄ-KOSKELA s Ottawa Hospital Research Institute and University of Ottawa, Ottawa, ON, Canada GEORGES VASSAUX s INSERM U948 and Institut des Maladies de l’Appareil Digestif, CHU Hôtel Dieu, Nantes, France ERIC A. WEAVER s Mayo Clinic, Rochester, MN, USA WILLIAM S.M. WOLD s Department of Molecular Microbiology and Immunology, Saint Louis University, St. Louis, MO, USA

Chapter 1 Oncolytic Herpes Simplex Virus Engineering and Preparation Pankaj K. Agarwalla and Manish K. Aghi Abstract Herpes simplex virus-1 (HSV-1) is an enveloped, double-stranded DNA virus that has been used with modiﬁcation as an oncolytic virus against a number of tumor types. Modiﬁcations that make HSV-1 replication-conditional, i.e., selectively divide in replicating cells make it fulﬁll a prerequisite criteria for oncolytic viruses. Other appealing features of HSV-1 as an oncolytic virus include its large, modiﬁable genome; its sensitivity to antiviral agents, such as ganciclovir; and its lack of host cell integration. Here, we review the methods of HSV-1 engineering, through traditional recombination techniques as well as through bacterial artiﬁcial chromosome (BAC) technology. We then describe protocols for titering, ampliﬁcation, and puriﬁcation of engineered HSV-1-derived oncolytic viruses. Key words: Oncolytic virus, Virus titer, Virus puriﬁcation, HSV-BAC, G47$

1. Introduction Herpes simplex virus-1 (HSV-1) is an enveloped, double-stranded DNA virus with several advantages for use as an oncolytic virus: a large genome suitable for insertion of foreign genes; tropism for neural cells; a safety mechanism in its sensitivity to agents, such as ganciclovir; high titers can be generated; and it does not integrate into the host genome, so it is unlikely to be oncogenic (1, 2). These advantages have led to the use of HSV-1s genetically engineered to be replication-conditional, i.e., selectively dividing in replicating cells, as oncolytic viruses in the treatment of numerous cancer types (2–4). Here, we describe the basic structure of wildtype HSV-1 and the protocols for modiﬁcation, puriﬁcation, ampliﬁcation, and titration of HSV-1 for use as an oncolytic virus. We also describe several speciﬁc examples of oncolytic HSVs modiﬁed with the methods presented here.

David H. Kirn et al. (eds.), Oncolytic Viruses: Methods and Protocols, Methods in Molecular Biology, vol. 797, DOI 10.1007/978-1-61779-340-0_1, © Springer Science+Business Media, LLC 2012

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HSV-1 has four main components: a core that contains double-stranded DNA (dsDNA), an icosadeltahedral capsid, an amorphous tegument, and an outer lipid bilayer envelope with glycoprotein spikes (3, 4). The DNA is arranged as two unique sequences – Unique Long (UL) and Unique Short (US) – that are each ﬂanked by repeat sequences. Homologous recombination at these repeat sequences results in four possible permutations of linear and inverted UL and US sequences. During infection and after cell entry mediated by surface glycoproteins, the viral genome circularizes and begins transcription of immediate early (IE) genes, which are involved with transcription regulation of the host cell and virus. The IE genes also contribute to transcription regulation of early (E) and late (L) genes (2–4). The ﬁrst engineered HSV-1 oncolytic virus had a mutation in the viral thymidine kinase (TK) gene, and showed killing of glioma cells in vitro and in models of glioma in vivo (5, 6). This mutant is replication-conditional because the mutant virus can only replicate in dividing cells because only dividing cells like tumor cells express sufﬁcient amounts of mammalian TK to complement the lack of viral TK. Unfortunately, viral expression of TK is essential for the antiviral efﬁcacy of nucleoside analogues like ganciclovir or acyclovir. Viral thymidine kinase monophosphorylates these nucleoside analogues (viral TK is much more efﬁcient than human nucleoside kinases at monophosphorylating antiviral nucleoside analogues), which are then further phosphorylated by cellular kinases, ultimately producing ganciclovir or acyclovir triphosphate, which are incorporated into elongating DNA chains, after which they interrupt DNA synthesis. Because of the inability to use nucleoside analogues as a safety mechanism for this ﬁrst-generation, viral-TK mutated vector, signiﬁcant safety concerns were raised and, in fact, neurotoxicity was seen at high doses (6). In an alternative HSV-1 vector, a mutation in the viral G134.5 gene was introduced. The G134.5 gene and its product, ICP34.5, allow normal HSV to subvert the host’s “shut-off” response against infection. Once infected with HSV, a normal cell will activate protein kinase R (PKR) which in turn phosphorylates and inactivates eukaryotic initiation factor-2A (eIF-2A), thereby shutting down protein synthesis in the normal host cell. ICP34.5 restores protein synthesis by activating protein phosphatase-1A which dephosphorylates and restores eIF-2A function (2, 6). Mutations in this gene, G134.5, result in an HSV that cannot replicate in normal cells, which abrogate protein synthesis machinery. In malignant cells, however, the activation of PKR is less pronounced, likely due to other mutations, and thus, the ICP34.5-mutant HSV is replication-conditional. Another viral gene that can be mutated to render HSV-1 replicationconditional is UL39, which encodes ICP6, the large subunit of viral ribonucleotide reductase. HSVs with ICP6 mutations can only replicate in dividing cells like tumor cells because only tumor cells

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express sufﬁcient levels of mammalian ribonucleotide reductase to complement the viral mutation. However, while ICP34.5 and ICP6 single mutant HSVs are sensitive to antiviral nucleoside analogues and lack that speciﬁc safety concern seen with the original TK-deﬁcient HSVs, ICP34.5 and ICP6 single mutant HSVs have undergone few preclinical studies because of the concern that single mutant HSVs would pose a safety risk if they were to undergo in vivo recombination and restoration of the wild-type HSV phenotype (2). Double-mutant viruses are theoretically safer as the chances of recombination and restoration of the wild-type HSV phenotype are decreased (2, 6, 7). Therefore, a second-generation virus, termed G207, was created which harbors an insertion of the Escherichia coli lacZ gene into the UL39 gene, which encodes ICP6, and has deletion of both copies of ICP34.5 (7). MGH-1, which has the same modiﬁcations as G207, was created by a different group using a slightly different transfection method (8). Finally, a third generation HSV-1 mutant, G47$, was created by modifying G207 by using bacterial artiﬁcial chromosome (BAC) technology to delete the nonessential viral gene A47 (9, 10). The deletion created in G47$ places the late US11 gene under control of the immediate-early A47 promoter, which enhances the growth of G47$ relative to G207 because earlier expression of US11 prevents the premature termination of protein synthesis that slows the growth of G34.5-mutated HSVs like G207. In addition, deletion of the A47 gene prevents the downregulation of MHC class I expression seen in HSVs expressing A47, which could enhance the antitumor immune response (11). Importantly, G47$ was produced using BAC technology, an advance which allows for more straightforward HSV-1 engineering, including the “arming” of oncolytic HSVs to express other gene products that might improve oncolytic virotherapy. Here, we describe the initial preparation and storage of wild type and recombinant HSV-1, as can be obtained either commercially or through another laboratory. Next, we review the recombination methods used in engineering oncolytic HSVs. We also review protocols for oncolytic HSV production and titration.

2. Materials 2.1. Materials Needed for HSV Work of Any Kind

1. Biosafety level 2 (BL2) and a biocontainment hood: HSV-1 is a human pathogen and requires these precautions. 2. A virucidal agent such as LpHse (Steris): 7.7% o-phenylphenol, 7.6% p-tertiary amylphenol, 84.7% inert ingredients. Used at a 1:256 dilution (1/2 oz/gal distilled water).

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2.2. Materials for Titration, Ampliﬁcation, and Puriﬁcation of HSV (Subheadings 3.1–3.4)

1. African green monkey kidney cells [Vero cells, American Type Culture Collection (ATCC)]. 2. DMEM tissue culture medium (Mediatech, Inc.). 3. Fetal bovine serum (FBS) and calf serum (CS) (Mediatech, Inc.). 4. Heat-inactivated FBS (iFBS): Made by heating FBS to 56°C for 30 min and then stored at −20°C. 5. 1% Penicillin/streptomycin solution (Invitrogen). 6. Sterile Dulbecco’s phosphate-buffered saline (DPBS) supplemented with 50 mg/L streptomycin sulfate, 100 mg/L kanamycin monosulfate, 1,000 mg/L glucose, 36 mg/L sodium pyruvate, 0.133 g/L calcium chloride (CaCl22H2O), and 0.1 g/L magnesium chloride (MgCl26H2O) (Sigma-Aldrich). 7. Sterile PBS (Mediatech, Inc., Manassas, VA) enriched with 1,000 mg/L glucose. 8. BD Falcon 6-well tissue culture plates and T150 tissue culture ﬂasks (BD Biosciences). 9. Human anti-HSV IgG (Gamunex, Talecris Biotherapeutics). 10. Ethanol–dry ice bath (Sigma-Aldrich). 11. Sonicator bath at 4°C, standard tabletop tissue culture centrifuge, standard tabletop rocking shaker, standard tissue culture microscope, and high-speed ﬂoor centrifuge (Fisher Scientiﬁc, Inc.). 12. Virus buffer: 150 mM NaCl and 20 mM Tris–HCl at pH = 7.5 (Sigma-Aldrich). 13. Virus stock (e.g., Strain F obtained commercially or virus stock obtained from previous ampliﬁcations/puriﬁcations as outlined below). 14. X-Gal concentrated stock solution: 50 mg/mL X-Gal substrate (Sigma-Aldrich) dissolved in DMSO. Solution should be stored at 4°C in the dark (aluminum foil wrap around a 50-mL conical tube). 15. Diluted X-Gal solution that can be used once the day of titration: 0.5 mL of X-Gal stock (50 mg/mL), 0.05 mL of 1 M MgCl2, 1 mL of 125 mM potassium ferricyanide (in PBS), and 1 mL of 125 mM potassium ferrocyanide (in PBS) mixed in PBS for a total volume of 25 mL (PBS in the ﬁnal solution should come out to 1×). Filter X-Gal solution after mixing to prevent crystal formation. 16. Neutral red solution: 7 mL of ddH2O, 1 mL of methanol, and 40 ML of neutral red stock solution (Sigma-Aldrich). 17. Glutaraldehyde (0.2%)/paraformaldehyde (2%) solution (SigmaAldrich).

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18. 10 mL Oak Ridge High-Speed Polypropylene Copolymer centrifuge tubes (Fisher Scientiﬁc, Inc.). 19. Millex syringe ﬁlters: 5 Mm Millex SV (PVDF), 0.45 Mm Millex-HV (durapore/PVDF), and 0.22 Mm Millex (PVDF) ﬁlters (Millipore). 20. PBS with 30% sucrose (Sigma-Aldrich). 21. PBS with 10% glycerol (Sigma-Aldrich). 22. Nunc-brand cryotubes (Fisher Scientiﬁc, Inc.). 2.3. Materials for Engineering Oncolytic HSV (Subheadings 3.5–3.12)

1. HSV-G47$ BAC DNA: Obtained as described by Fukuhara et al. (10). HSV-G47$ BAC expresses GFP, chloramphenicol resistance, an FRT locus, and a loxP locus within the ICP6 gene. 2. pVec9 shuttle vector: Obtained as described by Fukuhara et al. (10), contains a loxP site, a lacZ gene, a transgene multiple cloning site with a pCMV promoter, an FRT site, and a kanamycin resistance gene. 3. 10× Cre buffer: 500 mM Tris–HCl, pH 7.5, 330 mM NaCl, 100 mM MgCl2. 4. TE buffer: 10 mM Tris–HCl and 1 mM EDTA (pH = 8.0) (Sigma-Aldrich). 5. 50× TAE buffer, pH 7.6–7.8: 242 g Tris–HCl, 100 mL 0.5 M EDTA, pH 8.0, and 57.1-mL glacial acetic acid; then, add enough volume of Millipore H2O to dissolve solids, then add HCl to get ﬁnal pH to 7.6–7.8, then use Millipore H2O to get ﬁnal volume to 1,000 mL. 6. 3 M Sodium acetate, glucose, ethanol, glycerol, isopropanol, agarose, TAE buffer (Sigma-Aldrich). 7. LB plates with kanamycin (10 Mg/mL) and chloramphenicol (12.5 Mg/mL) (Sigma-Aldrich). 8. Shaking incubator at 37°C (Fisher Scientiﬁc, Inc.). 9. SOC medium (per protocol by QIAGEN). 10. Electroporation cuvettes with 0.1-cm gap (Gene Pulser, Bio-Rad, Inc.). 11. E. coli electroporation competent cells DH10B (Invitrogen). 12. Screwtop microcentrifuge tubes (Fisher Scientiﬁc, Inc.). 13. Gene Pulser Xcell electroporator (Bio-Rad, Inc.). 14. Solution 1 (25 mM Tris–HCl, 10 mM EDTA, 50 mM glucose): Add 250 ML of 1 M Tris–HCl (pH 8.0), 200 ML of 0.5 M EDTA, and 90 mg of glucose into a total of 10 mL of ddH2O (Sigma-Aldrich). 15. Lysozyme 25 mg/mL: Add 250 mg of lysozyme (SigmaAldrich) to 10 mL of 10 mM Tris–HCl at pH 8.0.

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16. QIAGEN plasmid mini and maxi kits, including buffers QBT, QC, and QF (QIAGEN). 17. QIAGEN-tip 500 columns (QIAGEN, Dusseldorf). 18. DNA spectrophotometer (Fisher Scientiﬁc, Inc.). 19. Low- and high-molecular weight DNA ladders (Invitrogen). 20. Standard DNA electrophoresis setup, including loading dye (Bio-Rad, Inc.). 21. Sarstedt centrifuge tubes (Nümbrecht). 22. pFLPe plasmid (available from laboratory of Prof. Rudolf Jaenisch, MIT, Cambridge, MA). 23. Opti-MEM solution without any supplements (Invitrogen). 24. Lipofectamine 2000 (Invitrogen). 25. RSB buffer: 10 mM Tris–HCl at pH 8.0, 10 mM KCl, and 1.5 mM MgCl2 (Sigma-Aldrich). 26. Phenol, chloroform, and isoamyl (puriﬁcation grade) (SigmaAldrich).

3. Methods Subheadings 3.1–3.4 below deal primarily with the titration, ampliﬁcation, and puriﬁcation of HSV-1, protocols which can be applied not only to wild-type HSV-1, but also to modiﬁed virus. In short, when any HSV-1 virus is received from an outside source, such as a vendor or another laboratory, it needs to be quantiﬁed in order to know the correct amounts for ampliﬁcation. Once ampliﬁed in a cell line, the virus should be puriﬁed and requantiﬁed to serve as a stable virus stock. Subheadings 3.5–3.12 are protocols to make an “armed” oncolytic virus from the G47$ backbone (10). These protocols can also be applied in general to manipulation and puriﬁcation of oncolytic viruses. For example, the transfection procedure to transfect plasmid DNA into a cell line can be used for simple recombination to modify standard backbones. A procedure similar to the one presented here for G47$ manipulation was used for the creation of MGH-1, which was created from R3616, a virus from the strain F backbone that has both copies of ICP34.5 deleted (12) (see Note 1). 3.1. Virus Titration

1. Establish Vero cell culture in 6-well tissue culture plates day before titration at approximately 3 × 105 cells/2 mL of DMEM supplemented with 10% CS and 1% penicillin/streptomycin. 2. Grow cells overnight. 3. On the day of titration, ensure that cells are approximately 60–70% conﬂuent and appear healthy.

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4. Wash twice with virus wash buffer (DPBS with supplements and 1% iFBS) and ensure that cells are not dry during steps. 5. While cells are in last wash, prepare virus stock by performing three freeze–thaw cycles followed by sonication at 30% output for 1 min. 6. Use a serial dilution technique to create 10−2 and 10−4 dilutions of virus stock in virus wash buffer (e.g., 10 ML into 1 mL of virus wash buffer for 10−2. Then, take 10 ML of new solution and place into 1 mL of virus wash buffer for 10−4 dilution of original virus). 7. Add 0.7 mL of each virus dilution per well (three wells per dilution). 8. Rock 6-well plate at room temperature for 5 min. 9. Place 6-well plate at 37°C for 1–2 h. 10. Remove virus and add 2 mL/well with DMEM supplemented with 1% iFBS and 0.1% human IgG at 37°C. 11. Incubate infected cells appropriately for approximately 2–3 days until plaques develop. Plates should be level in incubator. 12. If titrating virus with lacZ gene insertion, proceed to protocol 1.1B, otherwise, remove media and ﬁx with methanol (2 mL/well) for 5 min. 13. Remove methanol and let cells dry. 14. To stain and count cells, incubate each well with 1 mL of Giemsa stain that is prediluted 1:20 in ddH2O. Ensure that stain is evenly distributed. 15. Incubate in stain for 5–10 min carefully checking intensity of stain. 16. Once appropriately stained, remove Giemsa stain and rinse with tap water gently. 17. Dry plates and count plaques using stereomicroscope to obtain virus titer, namely plaque-forming units (pfu) per amount of total virus stock used. 18. Repeat with various dilutions as necessary to obtain plates that can easily be counted for pfu. This will allow for the most accurate titer to be obtained. 3.2. X-Gal-Based Staining of Virus with lacZ Expression

1. Follow protocol in Subheading 3.1 until cells are ready for staining. 2. Wash cells with sterile PBS without calcium/magnesium. Note: if cells do not adhere well to the plate, do not wash before ﬁxation. 3. Remove wash carefully.

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Fig. 1. Plaques of HSV-G47Delta/Vec9/empty. (a) Photograph of one well from 24-well plate after X-Gal staining during virus puriﬁcation of HSV-G47Delta/Vec9/empty. Viral plaques (shown here in black and white) typically stained blue with a white center which is area of cytopathic effect for each plaque. Background would typically be stained with neutral red. This example demonstrates pure virus since all plaques express lacZ. If this were not a pure population of lacZ-expressing virus, then one would see white plaques without the ring of lacZ-expressing infected cells. (b) 10× photomicrograph demonstrating a few viral plaques after X-Gal/neutral red staining during virus titration of pure HSV-G47Delta/Vec9/empty virus. The center of each plaque reﬂects cytopathic effect with a ring of virus-infected cells with expression of lacZ that would typically appear blue but is merely dark in this black and white photo.

4. Warm X-Gal staining solution to 37°C fully and mix well before use. 5. Fix cells with cold glutaraldehyde/paraformaldehyde ﬁxative (see Subheading 2) at 2 mL/well and let sit for 5–10 min at room temperature. 6. Wash cells three times with sterile PBS without calcium/ magnesium. 7. Incubate with X-Gal staining solution (see Subheading 2) at 0.6 mL/well at 37°C for 3 h in nonsterile incubator. 8. Aspirate remaining X-Gal stain and wash cells with tap water at room temperature. 9. Counterstain with neutral red solution (see Subheading 2) for 2–10 min at room temperature while noting color change. When color has changed appropriately, quench with tap water and count plaques as described above (Fig. 1). 10. Plates can be stored in the open at room temperature. 3.3. Virus Ampliﬁcation in Vero Cells

1. Grow healthy low-passage Vero cells in two large T150 tissue culture flasks on the day before amplification at 8 × 106 cells/flask. 2. The next day, cells should be approximately ten million cells per T150 or 70–80% conﬂuent. Ensure that they appear healthy. 3. Aspirate media and wash cells twice with PBS and 1% iFBS at 37°C.

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4. Dilute virus stock in PBS and 1% iFBS to obtain an MOI of 0.01 just prior to infection. MOI is obtained from initial titration above. 5. Place approximately 8 mL of diluted virus into each T150 ﬂask of Vero cells at 37°C. 6. Rock ﬂasks slowly for 10 min at room temperature. 7. Incubate at 37°C and 5% CO2 for 90 min. Ensure that ﬂask remains level horizontally during infection. 8. Aspirate viral inoculum and add 25 mL of DMEM with 3% iFBS per 150-mL ﬂask and incubate at 34.5°C and 5% CO2. 9. Wait until complete cytopathic effect has been obtained when cells are rounded and refractile, but have not yet detached from the plate. This is the optimal time for harvesting of virus with the highest titers. 10. When total CPE has been obtained, dislodge cells by ﬁrmly banging ﬂasks on hard surface and by scraping cells if necessary. 11. Pipette cells gently up and down on ﬂask surface to mix cells and assist in removing cells from ﬂask surface. 12. Centrifuge cells in tabletop tissue culture centrifuge at approximately 2,500 × g for 5 min. 13. Remove all but approximately 1 mL of supernatant. 14. Add 1 mL of virus buffer (for a total volume of 2 mL) and resuspend pellet in cryotubes. 15. Rapidly freeze in EtOH–dry ice bath and store at −80°C. 16. Repeat procedure until a total of ten ﬂasks are prepared and stored to provide a total 20 mL of ampliﬁed, but unpuriﬁed virus stock in a 1:1 mix of media and virus buffer. 3.4. Virus Puriﬁcation (see Note 2)

1. Combine 20 mL of ampliﬁed virus stock from Subheading 3.1 into a 50-mL BD Falcon centrifuge tube. 2. Freeze/thaw 3× in EtOH–dry ice bath. 3. Sonicate at 30% output for 1 min at 4°C. 4. Spin tubes at 500 × g for 10 min in standard tissue culture tabletop centrifuge. 5. Remove supernatant and respin supernatant at 2,500 × g for 10 min. 6. Save last supernatant and ﬁlter using 5 Mm Millex SV (PVDF) ﬁlter followed by 0.45-Mm Millex-HV (durapore/PVDF) ﬁlter. Use approximately 10 mL per ﬁlter. Since virus tends to stick to tubes, use a little virus buffer to rinse tubes and ﬁlter. 7. Filter 2 mL of sterile 30% sucrose in PBS with 0.22-Mm Millex PVDF ﬁlter and add to bottom of presterilized 10-mL Oak Ridge centrifuge tubes.

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8. Layer 5 mL of double-ﬁltered virus supernatant from step 5 very carefully over sucrose–PBS cushion in Oak Ridge tubes. This is done by adding the supernatant very slowly over the side of the tube, so two layers are clearly visible. 9. Centrifuge at 22,000 × g for 90 min at 4°C in ﬂoor-top highspeed centrifuge. 10. Remove supernatant and resuspend pellet carefully in 2 mL of PBS with 10% glycerol. PBS with 10% glycerol should be preﬁltered with 0.22 Mm PVDF ﬁlter. 11. Let pellet stand in PBS + 10% glycerol at 4°C for 2 h on shaker. 12. After 2 h, sonicate on high for 30 s, vortex, and pipette up and down to create homogenous solution of virus. 13. Aliquot small portion for titering of virus and place remainder in cryotubes for storage at −80°C. 3.5. Site-Speciﬁc Recombination of HSV-G47D BAC and pVec9 Shuttle Vector (see Note 3)

The goal of this section is to produce a large DNA known as BACG47$/Vec9/transgene, which has both chloramphenicol and kanamycin resistance as well as GFP and lacZ expression. 1. If needed, speciﬁc transgenes can be inserted into the pVec9 shuttle vector through standard recombination protocols, generating pVec9/transgene. 2. Combine 1 Mg of HSV-G47$ BAC DNA, 1 ML 10× Cre buffer, 1 ML Cre recombinase, and approximately 50 ng of pVec9 shuttle vector into a total volume of 10 ML of RNAse/DNAsefree sterile H2O. 3. Incubate mixture for 30 min at 37°C. 4. Inactivate recombinase by incubating for 10 min at 70°C. 5. Incubate at room temperature for 10 min. 6. To begin ethanol precipitation, add 40 ML of TE buffer. 7. Add 5 ML of 3 M sodium acetate and mix gently by pipetting. 8. Add 125 ML of ethanol and invert very gently. Since the DNA is large, aggressive mixing will damage it. 9. Incubate at −80°C for 30 min. 10. Centrifuge in tabletop microcentrifuge for 15 min at 4°C at 16,500 × g. 11. Discard supernatant and wash pellet with 500 ML of 70% ethanol. 12. Centrifuge in tabletop microcentrifuge for 5 min at 4°C at 16,500 × g. 13. Discard the supernatant and dry pellet to air, but do not let it dry out completely. 14. Dissolve in 5 ML of sterile H2O.

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After the initial recombination in Subheading 3.5 has occurred, BAC-G47$/Vec9/transgene is formed. In this section, we electroporate BAC-G47$/Vec9/transgene into bacteria, which can then be used to select for appropriate recombination events through double-antibiotic resistance genes. 1. Chill four electroporation cuvettes on ice. 2. Warm two antibiotic-selection plates at 37°C. 3. Thaw on ice one frozen vial of DH10B E. coli cells. 4. Transfer 20 ML of E. coli cells each to four new screwtop microcentrifuge tubes cooled on ice. 5. Immediately before each transformation add 1 ML of DNA mixture obtained from Subheading 2.1. Mix completely by tapping gently. 6. Transfer cell/DNA mixture to chilled electroporation cuvette on ice and tap it gently so that the mixture sits evenly. There should be no air bubbles and any condensation on the outside of the cuvette should be wiped carefully. 7. Use the following electroporator settings: 1.5 kV, 200 7, 25 MF. 8. Wipe off any condensation on the side of cuvette and place it in the chamber. 9. Remove the cuvette immediately from the chamber and add 500 ML of SOC medium at room temperature. 10. Transfer the E. coli diluted in SOC medium to sterile microcentrifuge tube. 11. Incubate in shaking incubator at 37°C for 45 min. 12. Plate 100–400 ML of the cells into two antibiotic-selective plates and incubate overnight at 37°C. Exact volume of cells used should be titrated to efﬁciency of transduction.

3.7. Bacterial Puriﬁcation and “Mini-Prep” Ampliﬁcation

1. From the two antibiotic-selective plates from Subheading 2.2, pick individual colonies and inoculate each clone into a separately labeled microcentrifuge tube with 10 ML of sterile ddH2O. 2. Create a replica plate by taking 1 ML of the picked colonies from step 1 and plate it on another antibiotic plate that is demarcated in grid fashion. Grow overnight and then store plate at 4°C. 3. Take remaining 9 ML of bacteria in ddH2O and amplify in 10 mL of LB plus antibiotics. Grow overnight in standard shaking conditions. Create stock by making ﬁve labeled cryotubes and adding 850 ML of bacteria in LB and 150 ML of sterile glycerol per tube. Snap-freeze in EtOH–dry ice bath and store at −80°C. This should leave approximately 5 mL of ampliﬁed bacteria in LB for DNA isolation.

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4. Harvest bacterial cells by centrifugation at 9,000 × g in standard tabletop centrifuge for 10 min at 4°C. Discard supernatant. (After centrifugation, it is possible to store cells at −20°C for later work). 5. Resuspend cell pellet in 200 ML of Solution 1 without lysozyme. 6. Transfer to new microcentrifuge tube. 7. Add 25 ML lysozyme (see Subheading 2) and mix gently. 8. Add 400 ML of P2 from QIAGEN kit and mix well gently. Leave on ice for 2 min. 9. Add 400 ML of P3 from QIAGEN kit and mix well gently. Keep on ice for 5 min. 10. Centrifuge for 10 min at 13,000 rpm in standard tabletop microcentrifuge at 4°C. 11. Transfer supernatant to a new 2-mL microcentrifuge tube. The total volume should be approximately 1 mL at this stage. 12. Add 1 ML of RNAse from QIAGEN kit at 100 mg/mL to make a ﬁnal concentration of 0.1 mg/mL. 13. Incubate at room temperature for 15 min. 14. Precipitate DNA by adding 700 ML of room temperature isopropanol. 15. Centrifuge immediately for 10 min at 16,500 × g (microcentrifuge) at room temperature. 16. Discard supernatant and wash pellet with 500 ML of 70% ethanol at room temperature. 17. Centrifuge for 5 min at 16,500 × g (microcentrifuge) at room temperature. 18. Remove the supernatant and allow pellet to air-dry for 1 min at 37°C. 19. Dissolve pellet with 50 ML of TE buffer. 20. Measure DNA concentration using spectrophotometer at optical density of 260 nm per standard procedures. 3.8. Hind III Restriction Analysis and DNA Gel Electrophoresis

1. Combine 20 ML of 10× NEBuffer 2 and 2 ML of Hind III enzyme together with 78 ML of ddH2O for a total of 100 ML of digestion mix. 2. Take ﬁve clones of BAC-G47$/Vec9/transgene from Subheading 3.5 and combine 10 ML of each sample clone with 10 ML of digestion mix. Incubate at 37°C for 2 h. Ideally, there should be at least 1 Mg of DNA for the digestion mix to work properly. 3. BAC-G47$ and BAC-G47$/Vec9/empty should be used as controls. For each control sample, combine 1 ML of DNA,

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9 ML of ddH2O, and 10 ML of digestion mix. Incubate at 37°C for 2 h. 4. For gel electrophoresis, create a 0.5% agarose/TAE 300-mL gel using standard procedures. 5. Load 20 ML of each sample and 2 ML of loading dye into each lane. 6. Two DNA ladders should be used – a low-molecular weight and a high-molecular weight ladder. For low-molecular weight ladder, follow manufacturer’s instructions. To make high-molecular weight ladder, combine 4 ML of high-molecular weight DNA ladder with 14 ML of TE. Incubate in water bath at 65°C for 5 min. Add 2 ML of loading dye and load total 20 ML into lane. 7. Run gel at 50 V for 16–24 h. 3.9. “Maxi-Prep” Ampliﬁcation

1. Once the correct clones are identiﬁed, amplify these clones and appropriate controls by inoculating clones from the replica plate into 2 mL of LB/chloramphenicol (12.5 Mg/mL) and incubating in shaking incubator at 37°C for 3 h. 2. Add this 2 mL preculture to a ﬂask of 200 mL LB/chloramphenicol (12.5 Mg/mL) and grow overnight with good aeration in shaking incubator at 37°C. 3. Centrifuge 200 mL of bacterial culture at 2,500 × g for 10 min at 4°C. Remove supernatant by decantation. 4. Resuspend cell pellet in 16 mL of Solution 1 (without lysozyme). 5. Add 1.6 mL of lysozyme (see Subheading 2) and mix gently. 6. Add 32 mL of P2 and mix well gently. Leave at room temperature for 5 min. 7. Add 32 mL of P3 and mix well gently. Keep on ice for 10 min. 8. Centrifuge for 30 min at 5,000 × g in standard ﬂoor centrifuge at 4°C. 9. Transfer supernatant to a new centrifuge tube. 10. Centrifuge again for 30 min at 5,000 × g in standard ﬂoor centrifuge at 4°C. 11. Transfer the supernatant to a clean Erlenmeyer ﬂask through a layer of sterile gauze. 12. Add 80 ML of RNAse at 100 mg/mL to make a ﬁnal concentration of 0.1 mg/mL. 13. Incubate at room temperature for 15–30 min. 14. Using a QIAGEN-tip 500 column, preequilibrate columns with 10 mL of QBT buffer.

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15. Apply DNA solution to the column (approximately 2.5× of 30-mL volume). 16. Warm 15 mL of QF buffer to 65°C. 17. Wash columns with 30 mL of QC three times. 18. Elute with 15 mL of QF buffer at 65°C into a Sarstedt 30-mL tube. 19. Precipitate DNA by adding 10.5 mL of room temperature isopropanol to each tube and mix gently. 20. Centrifuge for 60 min at 8,000 × g in ﬂoor-top centrifuge at 4°C. 21. Resuspend the DNA pellet with 15 mL of 80% ethanol. 22. Centrifuge for 15 min at 8,000 × g in ﬂoor-top centrifuge at 4°C. 23. Remove the supernatant and resuspend the pellet in 1 mL of 70% ethanol. Transfer to new screwcap microcentrifuge tube. 24. Centrifuge at 16,500 × g in tabletop microcentrifuge for 10 min at 4°C. 25. Remove supernatant. 26. Pulse centrifuge to remove any residual ethanol. 27. Dry the pellet for 2 min at room temperature without letting it overdry. 28. Dissolve the pellet with 100 ML of TE buffer. 29. Dilute 4 ML of DNA into a total volume of 200 ML of MQ. Measure optical density at 260 nm and calculate concentration of DNA. 30. Repeat Hind III digestion and gel electrophoresis after “maxiprep” ampliﬁcation to ensure that the appropriate DNA has been obtained for transfection. This DNA is referred to as BAC-G47$/Vec9/transgene. 3.10. Transfection of G47D /Vec9/ Transgene and pFLPe into Vero Cells

Cotransfecting the BAC-G47$/Vec9/transgene plasmid along with pFLPe, a plasmid that expresses FLPe, into Vero cells ensures that, within the Vero cell, expression of FLPe will create another recombination event, generating G47$-transgene that expresses lacZ but not GFP. Transcription of this DNA produces viral particles for further puriﬁcation and ampliﬁcation. Of note, the large BAC-G47$/Vec9/transgene does not produce viral particles without the FLPe recombination due to the presence of a lambda stuffer sequence. 1. Establish Vero cell culture in 6-well tissue culture plates day before titration at approximately 3 × 105 cells/well with 2 mL of DMEM supplemented with 10% CS. 2. Grow cells overnight.

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3. On the day of transfection, ensure that cells are approximately 70–80% conﬂuent and appear healthy. 4. For each well, prepare one tube with 1.5 Mg of pHSV-BACshuttle, 0.5 Mg of pFLPe, and 250 ML of Opti-MEM I without any supplements. Gently mix. 5. For each well, prepare another tube with 20 ML of lipofectamine 2,000 and 250 ML of DMEM. Gently mix and incubate for 5 min at room temperature. 6. Combine tubes from steps 5 and 6. Gently mix and incubate for 20 min at room temperature. 7. While incubating the combined mixture from both tubes, remove media from 6-well plate and wash with DMEM once. 8. Add 2 mL of DMEM + 1% iFBS to each well after the wash. 9. Add combined DNA-lipofectamine mix to each well and rock gently for 5 min at room temperature. 10. Incubate at 37°C for 4 h. 11. Remove media and replace with DMEM + 1% iFBS. Ensure that there are GFP-positive cells when changing the media. 12. When 50–80% CPE is observed, harvest viral particles and cells by scraping. 13. Centrifuge cells in tabletop tissue culture centrifuge at approximately 700 × g for 5 min. 14. Remove all but approximately 1 mL of supernatant. 15. Add 1 mL of virus buffer (for a total volume of 2 mL) and resuspend pellet in cryotubes. 16. Rapidly freeze in EtOH–dry ice bath and store at −80°C. 17. In order to plaque-purify by limiting dilution in the next step, perform titering of virus stock as described in Subheading 3.1. Since this new virus has the recombination with the LacZ insertion, the X-Gal staining protocol in Subheading 3.2 should be followed. 3.11. PlaquePuriﬁcation of Recombinant Virus from pHSV-BACShuttle

Plaque-puriﬁcation by limiting dilution will isolate a single viral clone that can be further tested to ensure accuracy of the recombination. 1. On the day before the titration, plate Vero cells in 96-well plates at 1 × 104 cells/well with 150 ML of DMEM supplemented with 10% CS. Make enough plates to be able to detect positive recombinant viruses at <0.5 pfu/well. 2. Remove media using a multichannel pipetter and wash cells with DPBS supplemented with glucose and 1% iFBS. 3. Dilute recombinant virus in DPBS supplemented with glucose and 1% iFBS at a concentration that will be sufﬁcient to infect cells at 0.3 pfu/well.

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4. Remove media from cells and infect cells at 0.3 pfu/well in 50 ML. Rock for 10 min at room temperature and incubate at 37°C for 2 h on a rocker. 5. After 2 h, remove virus inoculum. Ensure that inoculum is treated with bleach and disposed properly. 6. Add 150 ML DMEM and 1% iFBS (without human IgG) and incubate at 37°C. 7. On day 2, check the wells for plaques under the microscope and mark wells that contain only one plaque. 8. When plaques are large enough (usually on day 4), scrape cells and transfer both cells and media to a new labeled 96-well plate using a multichannel pipetter. Seal the plate with paraﬁlm and store at −35°C in a box clearly labeled as biohazard. 9. Pick a few clones out of the marked wells and titer virus using X-gal staining as outlined in Subheading 3.2. 3.12. HSV-G47D Viral DNA Ampliﬁcation, Puriﬁcation, and Analysis

Once titer is determined, the viral DNA needs to be ampliﬁed and puriﬁed to conﬁrm that the correct recombination has taken place. 1. On day 0, plate Vero cells in 1T150 ﬂask per clone using DMEM supplemented with 10% CS. Cells should be approximately 90–95% conﬂuent on day 1. It is important to have control clones as well. 2. Remove media and wash cells with DPBS supplemented with glucose and 1% iFBS. 3. Dilute virus at MOI 1.5 in 2 mL of PBS supplemented with glucose and 1% iFBS. Add to ﬂask. 4. Rock ﬂask for 10 min and incubate at 34.5°C for 2 h with rocking every 15 min. 5. Add 13 mL of DMEM supplemented with 1% iFBS and incubate at 34.5°C until complete cytopathic effect (approximately 24 h). 6. On day 2, remove media (If there are many cells in the media, spin cells at 700 × g for 5 min and resuspend in 1 mL of PBS. This can be added to cells in the next step). Dislodge cells by gentle tapping of ﬂask. Scrape off attached cells that remain. Transfer into 15-mL BD Falcon centrifuge tube. 7. Wash any remaining cells in ﬂask with 2 mL of PBS and add to centrifuge tube with other cells. At the end of this, there should be one centrifuge tube with as many cells as possible from one ﬂask. 8. Spin cells at 500 × g for 5 min in standard tabletop centrifuge. Discard supernatant and wash cells with 10 mL RSB. Spin again at same settings and resuspend with 1 mL RSB.

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9. Let cells sit on ice for 20 min to allow swelling. For the following steps, ensure that cells and supernatants are kept at 4°C until the proteinase K is added. 10. Add 50 ML of 10% NP40 to give ﬁnal concentration of 0.5% in 1 mL of RSB and cells. Incubate at room temperature for 10 min. 11. Add 10 ML of RNAse A (10 mg/mL in ddH2O) and mix gently. 12. Spin tube at 700 × g for 5 min in standard tabletop centrifuge. Remove the cytoplasmic supernatant and save at 4°C. 13. Resuspend the pellet, which contains the nuclei, in 0.4 mL RSB. Add 10 ML of 10% Triton X-100. Invert gently and spin down at 2,000 rpm for 5 min in standard tabletop microcentrifuge. 14. Combine both cytoplasmic and nuclei supernatants. 15. Add 70 ML of 0.2 M EDTA and 60 ML of 10% SDS to the combined supernatants. Mix gently to avoid air bubbles. 16. Add 25 ML of proteinase K (10 mg/mL in 2 mM CaCl2 and 50 mM Tris–HCl at pH = 8.0). 17. Incubate overnight at 37°C. 18. On day 3, note total volume of sample. Extract DNA by adding a half volume of phenol and another half volume of chloroform– isoamyl alcohol extraction mixture (phenol–chloroform–isoamyl should be at the following ratio – 25:24:1). 19. Rotate gently and centrifuge at room temperature for 5 min at 3,000 × g (microcentrifuge). After phases are separated, remove and save aqueous phase into an Oak Ridge centrifuge tube using a wide bore or cut pipette tip. 20. Repeat extraction on nonaqueous phase using only a half volume of chloroform–isoamyl alcohol (24:1). Save aqueous phase in previous tube. 21. Repeat extraction on nonaqueous phase using only half volume of chloroform (without isoamyl alcohol) and let it evaporate under chemical hood for 15 min. 22. Combine all aqueous phases and note new volume. 23. Begin precipitation of DNA by adding sodium acetate (3 M stock) so that ﬁnal concentration is 0.3 M. Mix gently. 24. Next, add 2.5 times the volume of ethanol. 25. Allow mixture to precipitate for 2 h at −20°C. 26. Centrifuge in high-speed ﬂoor-top centrifuge at 4°C for 30 min at 7,000 × g. 27. Add 1 times the volume of ethanol without disrupting the pellet. Respin at 100 × g for 5 min at 4°C.

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28. Let the pellet dry to air brieﬂy and resuspend in 50 ML of TE buffer. Transfer to microcentrifuge tube and wash original tube with another 25–50 ML of TE buffer and add to previous DNA. 29. Measure DNA using spectrophotometer and perform restriction enzyme digestion followed by gel electrophoresis to conﬁrm proper virus. Once conﬁrmed, this virus may be ampliﬁed and puriﬁed by the methods outlined in Subheading 1.

4. Notes 1. All institutional safety guidelines should be followed for the safe handling and decontamination/disposal of materials that come into contact with the virus, which usually includes disinfecting with virucidal agent then autoclaving cell culture materials that have come into contact with HSV-1 particles, including but not limited to pipettes, ﬂasks, culture media, and tubes. In tissue culture, HSV-1 infected plates need not be separated from uninfected plates because virus transmission only occurs through ﬂuids during soniﬁcation by the generation of an aerosol. 2. In Subheading 3.4, if the homogenous solution of virus remains turbid, repeat steps 1–13. 3. Throughout Subheadings 3.5–3.12, appropriate controls should be made and processed as well, including BAC-G47$ and BAC-G47$/Vec9/empty. BAC-G47$ is the HSV BAC DNA without the Cre-mediated recombination and BACG47$/Vec9/empty is the BAC DNA after recombination with an empty shuttle vector that contains no transgene. 4. In Subheading 3.6, once a few colonies have started to grow overnight, it is important to isolate clones and check for accuracy of recombination before creation of the virus.

Acknowledgments The authors wish to thank Drs. Samuel Rabkin, Riyuchi Kanai, and Slawomir Antoszczyk for the ﬁgures and assistance. PKA is supported by a continuing clinical research grant from the Doris Duke Charitable Foundation. MKA was supported by a research grant from the American Brain Tumor Association (ABTA) at the time studies cited in this chapter were completed.

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References 1. Chiocca, E. A. (2002). Oncolytic viruses. Nat Rev Cancer 2, 938–50. 2. Aghi, M., and Chiocca, E. A. (2003). Genetically engineered herpes simplex viral vectors in the treatment of brain tumors: a review. Cancer Invest 21, 278–92. 3. Shen, Y., and Nemunaitis, J. (2006). Herpes simplex virus 1 (HSV-1) for cancer treatment. Cancer Gene Ther 13, 975–92. 4. Aghi, M., and Martuza, R. (2005). Oncolytic viral therapies - the clinical experience. Oncogene 24, 7802–16. 5. Martuza, R. L., Malick, A., Markert, J. M., Ruffner, K. L., and Coen, D. M. (1991). Experimental therapy of human glioma by means of a genetically engineered virus mutant. Science 252, 854–6. 6. Markert, J. M., Parker, J. N., Buchsbaum, D. J., Grizzle, W. E., Gillespie, G. Y., and Whitley, R. J. (2006). Oncolytic HSV-1 for the treatment of brain tumours. Herpes : the journal of the IHMF 13, 66–71. 7. Mineta, T., Rabkin, S. D., Yazaki, T., Hunter, W. D., and Martuza, R. L. (1995). Attenuated multi-mutated herpes simplex virus-1 for the treatment of malignant gliomas. Nat Med 1, 938–43.

8. Kramm, C. M., Chase, M., Herrlinger, U., Jacobs, A., Pechan, P. A., Rainov, N. G., SenaEsteves, M., Aghi, M., Barnett, F. H., Chiocca, E. A., and Breakeﬁeld, X. O. (1997). Therapeutic efﬁciency and safety of a second-generation replication-conditional HSV1 vector for brain tumor gene therapy. Hum Gene Ther 8, 2057–68. 9. Ino, Y., Saeki, Y., Fukuhara, H., and Todo, T. (2006). Triple combination of oncolytic herpes simplex virus-1 vectors armed with interleukin-12, interleukin-18, or soluble B7-1 results in enhanced antitumor efﬁcacy. Clin Cancer Res 12, 643–52. 10. Fukuhara, H., Ino, Y., Kuroda, T., Martuza, R., and Todo, T. (2005). Triple gene-deleted oncolytic herpes simplex virus vector double-armed with interleukin 18 and soluble B7-1 constructed by bacterial artiﬁcial chromosome-mediated system. Cancer Research 65, 10663–8. 11. Todo, T., Martuza, R. L., Rabkin, S. D., and Johnson, P. A. (2001). Oncolytic herpes simplex virus vector with enhanced MHC class I presentation and tumor cell killing. Proc Natl Acad Sci USA 98, 6396–401. 12. Chou, J., Kern, E. R., Whitley, R. J., and Roizman, B. (1990). Mapping of herpes simplex virus-1 neurovirulence to gamma 134.5, a gene nonessential for growth in culture. Science 250, 1262–6.

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Chapter 2 Construction of Capsid-Modiﬁed Adenoviruses by Recombination in Yeast and Puriﬁcation by Iodixanol-Gradient Marta Giménez-Alejandre, Alena Gros, and Ramón Alemany Abstract Adenovirus represents a valuable tool for the treatment of cancer, but tumor targeting remains a pending issue. Most common procedures to modify adenovirus genome are time-consuming due to the requirement of multiple cloning steps, and the low efﬁcacy of the recombination process. Here, we present a new method for homologous recombination in yeast to fast construct recombinant adenoviruses. Also, an alternative procedure to purify viral stocks, based on iodixanol gradient is described. Compared to classical methods, iodixanol is nontoxic to cells, which avoids desalting to use in vitro and in vivo. Moreover, viral stocks are more viable and it can be used for large-scale puriﬁcations. Finally, a protocol for analyzing blood persistence of modiﬁed vector in in vivo biodistribution is presented. Key words: Adenovirus, Detargeting, Homologous recombination, Iodixanol, Biodistribution

1. Introduction Many efforts in virotherapy of cancer with adenoviruses are focused on the delivery of viral particles to tumor cells. However, the hepatotropism of Ad5 following systemic administration limits its efﬁcacy and results in toxicity. In order to retarget adenovirus to desired cells, it is necessary to abrogate liver transduction (liver detargeting) and to expose heterologous peptides, which are speciﬁc for tumor cells (tumor targeting), on the external surface of the virus. The design of optimized vectors with modiﬁed tropism for clinical use requires knowledge and understanding of adenovirus biology and in vivo biodistribution.

David H. Kirn et al. (eds.), Oncolytic Viruses: Methods and Protocols, Methods in Molecular Biology, vol. 797, DOI 10.1007/978-1-61779-340-0_2, © Springer Science+Business Media, LLC 2012

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Based on in vitro data, it was postulated that when virus is administered intravenously, hepatocyte entry was mediated by a direct interaction of Ad capsid with its cellular receptors CAR and integrins. However, the double ablation of CAR and integrinbinding sites failed to reduce hepatocyte transduction in vivo, suggesting the involvement of other interactions for Ad liver entry (1–3). Thereafter, it was suggested that HSPGs played a role in liver transduction through a direct interaction with KKTK motif in the ﬁber shaft (4), as mutation of this motif resulted in a signiﬁcant reduction in liver transduction. However, subsequent studies reported that this low transduction was attributed to a defect in postinternalization steps, suggesting a blockade at the trafﬁcking level, i.e., receptor-mediated endocytosis, capsid disassembly, endosomal escape, or nuclear translocation (5). In 2005, Shayakhmetov et al. reported a novel pathway for liver transduction, showing that blood factors play a major role in targeting Ad to hepatocytes (6). Human coagulation factor IX and C4-binding protein was proposed to bridge the ﬁber knob to HSPGs and low-density lipoprotein receptor-related protein on the hepatocyte surface. A triple mutation within the ﬁber knob was shown to inhibit these interactions, reducing virus load in the liver by up to 50-fold, although these modiﬁcations did not completely abrogate liver transduction. In recent years, two independent studies have described a direct interaction between vitamin K-dependent coagulation factors, predominantly factor X, and the trimeric hexon of the Ad capsid, in directing hepatocyte transduction (7, 8). Recently, Alba and collaborators identiﬁed factor X binding sites on the adenovirus hexon, and demonstrated that a single point mutation in HVR7 was enough to completely block FX-mediated transduction in vitro and in vivo (9). In humans, adenovirus interactions with blood cells, mainly erythrocytes and platelets (10, 11), may compromise virus access to tumor cells. Therefore, capsid modiﬁcations to avoid these interactions could also increase virus bioavailability. As long as liver detargeting is achieved, the opportunity for tumor targeting improves. In these terms, numerous strategies have been developed in an attempt to confer a novel tropism for tumor cells. Coating of targeted-polymers to adenovirus surface or the use of bi-adaptor molecules, such as diabodies and fusion proteins, have been successfully incorporated, nongenetically, by chemical conjugation (12–14). However, genetic modiﬁcations of the adenovirus genome are required in order to maintain such modiﬁcations in the virion progeny. In this sense, genetic incorporation of small peptides within various capsid locations, the use of knobless ﬁbers fused to super-stable antibodies, afﬁbodies or the use of pseudotyped vectors which incorporate ﬁbers derived from other Ad serotypes, have been shown to increase infectivity of target cells (15–17).

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This chapter deals with methods to modify capsid proteins, to purify recombinant adenovirus and determine biodistribution in tumor-bearing mice in order to construct truly targeted adenoviruses for cancer treatment. Commonly, Ad is constructed using homologous recombination in bacteria (18). As more versatile strategy, we present the use of yeast to construct the recombinant virus genome. Recombination only requires 40-bp homology tails at the extremes of the insert and a positive-negative selection can be applied. Iodixanol is an alternative method to CsCl to purify Ad. Compared to the commonly used CsCl (not described here), iodixanol is faster and does not require desalting the obtained virus for in vitro and in vivo applications.

2. Materials 2.1. Adenovirus Construction by Homologous Recombination in Yeast

Reagents (when not speciﬁed can be purchased at Sigma): 1. Polyethylene glycol 3350 (50%, w/v): Dissolve 50 g of PEG 3350 in 30 ml of H2O in a 150-ml beaker on a stirring hot plate. Cool down the solution to room temperature; ﬁll volume up to 100 ml, mix thoroughly by inversion and autoclave. Store, securely capped, at RT. Evaporation of water from the solution will increase the concentration of PEG and severely reduce the yield of transformants. 2. 10× AA Solution: 5.7 g Brent supplement mixture-His–Leu– Try–Ura + 0.5 g Leucine (Do not add for LEU-) + 0.2 g tryptophan + 0.1 g histidine + 0.1 g uracil (Do not add for URA-). Add ddH2O up to 500 ml and autoclave 15 min. Store at 4°C. 3. 1.0 M lithium acetate: Dissolve 5.1 g of lithium acetate dihydrate in 50 ml of H2O, sterilize by autoclaving, and store at RT. 4. Transformation mix: 240 Ml of PEG (50%, w/v) + 36 Ml 1.0 M lithium acetate + 10 Ml Boiled SS-Carrier DNA (10 mg/ ml) + 74 Ml [(vector) + (insert)] DNA plus H2O (Milli-Q, autoclaved). 5. YPDA++ (Yeast extract/peptone/dextrose/adenine rich medium) : 5 g yeast extract + 10 g bacto-peptone. Add ddH2O up to 450 ml and autoclave 20 min. Then, add 50 ml 20% glucose; 20 ml 0.5% adenine (previously ﬁltered through 22 Mm ﬁlter). For YPDA++ plates, add 15 g bacto agar before autoclave. 6. SC (basic medium): 3.35 g yeast nitrogen base [YNB w/o AA, with ammonium sulfate (Difco)]. Add ddH2O up to 400 ml and autoclave. Then, add 50 ml of 20% glucose and 50 ml of 10× AA solution (URA- or LEU-, or URA/LEU-). For SC plates, add 15 g of bacto agar before autoclave.

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7. SC plates with 5-ﬂuoroorotic acid (5-FOA): (a) In a beaker, mix: 0.63 g BSM–His–Leu–Try–Ura + 0.04 g uracil (plasmids that grow in FOA plates have lost the URA gene) + 0.02 g Tryp + 0.01 g His + 0.05 g leucine (do not add if the plasmid has CAL) + 0.5 g FOA + 3.5 g [YNB w/o AA, with ammonium sulfate (Difco)] + 10 g glucose/ dextrose. Add ddH2O up to 250 ml. Stir and heat on a stir plate to dissolve. Try to keep temperature below 45°C. It may take a while to dissolve 5-FOA. Filter-sterilize when dissolved and still warm. (b) Mix: 10 g bacto agar + 250 ml ddH2O. Autoclave and cool down to 45°C. Add ﬁltered mixture (a) and mix thoroughly in an independent ﬂask. Pour in plates (protect from light). 8. Yeast miniprep mix: 2% Triton-X100, 1% SDS, 0.1 M NaCl, 10 mM Tris–HCl, pH 8.0, and 1 mM EDTA. 9. 425–600 Mm glass beads. 2.2. Puriﬁcation of Viral Prestocks by Banding on Iodixanol

1. Dulbecco’s modiﬁed Eagle’s medium (DMEM) (Gibco/BRL) supplemented with 10 or 5% fetal bovine serum (FBS; Hyclone). 2. HEK-293 cells or other adenovirus packaging cell lines. 3. Tetrachloroethylene (Fisher Scientiﬁc). 4. Ultracentrifuge: Beckman Coulter Optima L90K o L100XP and rotor SW40Ti (Beckman Coulter). Polyallomer centrifuge tubes for SW40Ti rotor (Beckman Coulter). 5. Stock solutions: L

Sol A: 60% iodixanol (if it is not sterile we ﬁlter sterilize).

L

Sol B: 10× PBS containing 10 mM MgCl2 and 25 mM KCl (10× PBS-MK).

L

Sol C: PBS containing 1 mM MgCl2 and 2.5 mM KCl (PBS-MK).

L

Sol D: 2 M NaCl in PBS-MK.

6. Solutions for iodixanol gradient (prepare from stock solutions): L

Sol 1 (54% iodixanol in PBS-MK): 13.5 ml Sol A + 1.5 ml sol B.

L

Sol 2 (40% iodixanol in PBS-MK): 4 ml Sol 1 + 1.4 ml Sol C.

L

Sol 3 (25% iodixanol in PBS-MK). 2.5 ml Sol 1 + 2.9 ml Sol C.

L

Sol 4 (15% iodixanol in 1 M NaCl PBS-MK): 1.5 ml Sol 1 + 2.7 ml Sol D + 1.2 ml Sol C.

7. PBS 1× Ca++/Mg++ (Gibco), Glycerol anhydrous (Fluka).

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3. Methods 3.1. Adenovirus Construction by Homologous Recombination in Yeast

Homologous recombination in yeast confers several advantages compared to homologous recombination in bacteria, i.e., the high efﬁciency of the process and the requirement of shorter regions of homology (40 bp). The adenovirus plasmids should contain three elements for the correct selection and ampliﬁcation in yeast: a centromere “CEN,” an autonomously replicating sequence “ARS” and a yeast selection gene (URA or LEU that allows the yeast to grow in media without uracil or leucine, respectively). The fragments containing the centromere, ARS, and uracil or leucine, are referred as CAU or CAL, respectively. Such fragment can be ampliﬁed by PCR from the plasmid pRS416 for the URA selection and from pRS425 for the LEU selection (Stratagene). The primers to amplify this fragment should contain 15 nt at their 3c ends corresponding to the beginning and end (complementary sequence) of the CAU or CAL fragment, and 40 nt tails at their 5c ends that are homologous to the site targeted in the adenovirus plasmid (see Note 1). Once the CAU/CAL fragment is inserted on the Ad plasmid (CAL is always recommended as it allows positive–negative selection in the future), two different strategies are followed depending on the presence of restriction sites: cut-repair or URA-positive– negative selection (Fig. 1). If one or two restriction sites are available in the pAdCAU or pAdCAL plasmids (vector), it can be linearized and recombined with a DNA sequence (insert) that shares at least 40 nt of homology at both sides of the cut. With the recombination, the plasmid is circularized again and yeast can grow in URA- or LEU-deﬁcient plates. If no restriction sites are available, URA fragment can be inserted at position to be modiﬁed without the need to cut it (URA fragment can be ampliﬁed from pRS416 using primers with tails of 40 nt homologous to ﬂanking sequences of position to be modiﬁed). A positive selection (incorporation of URA) using uracil and leucine-deﬁcient plates will select recombinant clones. Then, the URA gene is replaced for the desired fragment using a negative selection (loss of URA) in FOA and leucine-deﬁcient plates (see Note 2). Only those plasmids which have lost the URA gene will grow (5-FOA gives a toxic compound in the presence of URA which will not let these yeast grow). Once yeast colonies are obtained, the DNA from yeast plasmid (low copy) is isolated and transformed to bacteria (high copy) in order to analyze it. A similar system has been published by Hokanson et al. (19).

3.1.1. Preparation of the Insert

There are different types of inserts: the CAU or CAL to adapt a regular bacterial plasmid to grow in yeast, the URA insert to use positive–negative selection in a plasmid containing LEU in its

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#!,

&IBER PCR CAL

P23 Select in Leu- plates #!, )NSERT

52!

8

8

8

8

)NSERT 8

&IBER

52!

a) No unique restriction site Select in Leu- Ura - plates #!,

#!, Select in FOA Leu - plates

-ODIFIED &IBER

8 &IBER

b) Unique restriction site

#!, Select in Leuplates

#!,

Fig. 1. Adenovirus construction by homologous recombination in yeast. Main steps involved in the generation of recombinant adenoviruses using homologous recombination in yeast. Centromere-autonomously replicating sequences – Leucine gene (CAL) renders a bacterial plasmid competent for yeast growth. The adenovirus genomic plasmid with CAL can be used to insert a DNA of interest either via positive–negative selection with the URA gene (a) or via cut-repair when a unique restriction site is available at the targeted site (b).

backbone, and a regular DNA insert obtained by restriction or PCR to repair (recircularize) a linearized yeast plasmid. The common requirement is that the 5c and 3c ends (a minimum of 40 bp) of the insert are homologous to a region in the receptor vector that will be replaced. 1. Primer design: Oligos to amplify CAL (CAU uses the same oligos, but the template for PCR is pRS416 instead of pRS425): Forward: 40 bp homology upstream of targeted site + ACCT GGGTCCTTTTCATCAC. Reverse: 40 bp homology downstream of targeted site (reverse orientation) + CATCTGTGCG GTATTTCACA. Oligos to amplify URA (from pRS416): Forward: 40 bp homology upstream of targeted site + TCAATTCATCATTTTT. Reverse: 40 bp homology downstream of targeted site (reverse orientation) + GTAATAACTGATATAA. Examples to introduce URA fragment into the ﬁber, penton base, or hexon in order to introduce detargeting punctual mutations are given

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Table 1 Primers used for positive URA-selection to construct tropism-modiﬁed adenoviruses Protein modiﬁed

Phenotype (ablation)

Primer orientation Sequence

Fiber (knob)

CAR

Forward Reverse

Fiber (shaft)

Heparan sulfates

Forward Reverse

Penton base

Integrin

Forward Reverse

Hexon

Coagulation factor X

Forward Reverse

GAAAATGGAGTGCTACTAAACAATTCCTTCCT GGACCCTGAA TCAATTCATCATTTTT AGGCTGTGCCTTCAGTAAGATCTCCATTTCTA AAGTTCCA GTAATAACTGATATAA CCTTACCTCCCAAAATGTAACCACTGTGAGCC CACCTCTC TCAATTCATCATTTTT TAACTGTGAGGGGTGCAGATATTTCCAGGTT TATGTTTGA GTAATAACTGATATAA GGCAATGCAGCCGGTGGAGGACATGAACGAT CATGCCATT TCAATTCATCATTTTT CGGCCTCAGCGCGCTTCTCCTCAGCCCGTGT GGCAAAGGTGTAATAACTGATATAA TGGATGGGAAAAAGATGCTACAGAATTTTCAG ATAAAAATTCAATTCATCATTTTT TTAGATTGATTTCCATGGCAAAATTATTTCCAA CTCTTATGTAATAACTGATATAA

Example of primers to insert URA at different sites of capsid proteins to ablate the interaction with the Ad receptors. These primers can be used to amplify URA gene by PCR from the template pRS416. Each primer contains a fragment of homology with the extremes of URA gene (15 b-underlined) and a fragment of homology with each site of insertion in the ﬁber, penton base, or hexon protein (40 b-tail). All sequences are given from 5c to 3c. After recombination, recombinant colonies that have incorporate URA fragment will be selected in URA- and LEU-deﬁcient plates

in Table 1. Once URA fragment is inserted, it needs to be replaced by the insert (ampliﬁed by PCR or restriction) containing such modiﬁcations (Table 2). In addition to these punctual mutations, larger modiﬁcations, such as pseudotyping, afﬁbodies, or knobless ﬁbers, can also be constructed with targeting aims using this strategy. 2. PCR mix: 0.5 Ml template DNA (20 ng) (e.g., pRS416); 25 Ml Ex-Taq 2×; 1 Ml 20 MM oligo F; 1 Ml 20 MM oligo R; 22.5 Ml Milli-Q-autoclaved H2O. 3. PCR program (PCR fragment 1 kb): 1 min at 95°C , 30× (30 s at 95°C – 30 s at 55°C – 2 min 30 s at 72°C), 5 min at 72°C. 4. After the PCR, digest the template plasmid adding 10 U of DpnI (only cuts the methylated DNA template, not the PCR product) to the PCR and incubate 2 h at 37°C. Then, use phenol–chloroform extraction and clean with gel puriﬁcation, ethanol precipitation, and resuspend in ddH2O.

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Table 2 Inserts used to construct tropism-modiﬁed adenoviruses Modiﬁcation Protein modiﬁed

Phenotype (ablation)

Sequence

Y477A

Fiber

CAR

AAATGGAGTGCTACTAAACAATTCC TTCCTGGACCCTGAAGCTTGGAA CTTTAGAAATGGAGATCTTACTGA AGGCACAGCCTA

GATK

Fiber

Heparan sulfates

CCTTACCTCCCAAAATGTAACCACT GTGAGCCCACCTCTCGGCGCCAC CAAGTCAAACATAAACCTGGAAAT ATCTGCACCCCTCACAGTTA

RAE

Penton base

Integrins

GGCAATGCAGCCGGTGGAGGACATG AACGATCATGCCATTCGCGCTGA GACCTTTGCCACACGGGCTGAGGA GAAGCGCGCTGAGGCCG

E451Q

Hexon

Coagulation Factor X

TGGATGGGAAAAAGATGCTACAGAA TTTTCAGATAAAAATCAGATAAGAG TTGGAAATAATTTTGCCATGGAAA TCAATCTAA

Once URA is inserted at ﬁber, penton base, and hexon sites to be modiﬁed, it should be replaced with these inserts (obtained by PCR or restriction cut) containing the corresponding mutations using a negative selection. After recombination, recombinant colonies that have lost URA gene will be selected in FOA- and LEU-deﬁcient plates

3.1.2. Homologous Recombination in Yeast: Preparing Competent Yeast and Cotransformation with Vector and Insert

Protocols for yeast transformation have been adapted from Gietz and Woods (20, 21). Day 1 1. In a 50-ml Falcon, inoculate 5 ml of YPDA++ (yeast extract, peptone, dextrose, adenine rich medium) with one colony of yeast or 10 Ml of yeast glycerol stock (strain YPH857). 2. Incubate O/N in a rotary shaker at 30°C. Day 2 1. Dilute O/N growth 1/10 in YPDA++ to measure OD600 (use YPDA++ as blank, 1 OD600 = 1.5 × 107 cells/ml). Calculate dilution to prepare 50 ml of prewarmed YPDA++ at OD600 = 0.15 (0.15 OD600 = 2.25 × 106 cells/ml). 2. Incubate the ﬂask on a rotary shaker at 30°C and until exponential growth is achieved (OD600 = 0.4–0.9, approximately 5 h). 3. Transfer the 50 ml to Falcon and spin down at 3,000 × g for 5 min, room temperature. 4. Decant supernatant and wash pellet with 25 ml ddH2O. Spin again (3,000 × g, 5 min).

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5. Decant supernatant and resuspend the cells in 1 ml of ddH2O. 6. Boil salmon sperm or Herring DNA (10 mg/ml) in a boiling water bath (use 10 Ml for each transformation) for 5 min and then keep on ice while harvesting the cells. 7. Transfer the yeast suspension to an Eppendorf, centrifuge for 30 s at 6,500 × g in a microcentrifuge, and discard the supernatant. 8. Add ddH2O to a ﬁnal volume of 1 ml and vortex-mix vigorously to resuspend the cells. 9. Pipette 100 Ml samples (108 cells) into 1.5-ml microcentrifuge tubes, one for each transformation, centrifuge at 6,500 × g at RT for 30 s, and remove the supernatant. 10. Make up the transformation mix: 240 Ml PEG (50%, w/v) + 36 Ml 1.0 M lithium acetate + 10 Ml boiled SS-Carrier DNA (10 mg/ml) + 74 Ml plasmid DNA and insert plus H2O (Milli-Q, autoclaved). 11. Add 360 Ml of transformation mix to each transformation tube and resuspend the cells by mixing vigorously (vortex). 12. Incubate the tubes in a 42°C water bath for 40 min. 13. Microcentrifuge at 6,500 × g at room temperature for 30 s and remove the supernatant with the micropipette. 14. Pipette 1.0 ml of H2O (Milli-Q, autoclaved) into each tube, stir the pellet with a micropipette tip and vortex vigorously. 15. Plate appropriate dilutions of the cell suspension onto SC-URA or SC-LEU plates. Spread gently (few movements). Use plates without URA or LEU or both according to the presence of URA, LEU, or both genes, respectively, in the vector or the insert. 16. Incubate at 30°C for 2–3 days until yeast colonies appear. There should be more colonies in the plates with (vector) + (insert), than in the (vector) and (insert)-alone controls. 3.1.3. Yeast Plasmid Extraction after Yeast Transformation

1. Seed 2 ml of liquid SC-URA (or SC-LEU) O/N at 30°C with the desired colony. 2. Transfer 1.5 ml to Eppendorf tube. Centrifuge at maximum speed 5 s in a microcentrifuge. Discard supernatant and resuspend the yeast pellet in residual liquid. 3. Add 400 Ml of Yeast Miniprep mix (2% Triton X-100, 1% SDS, 0.1 M NaCl, 10 mM Tris–HCl, pH 8.0, 1 mM EDTA) and mix. 4. Add 400 Ml phenol:chloroform:isoamyl alcohol (25:24:1). 5. Add 0.3 g glass beads. Close Eppendorf with paraﬁlm. 6. Vortex for 2 min at 4°C. 7. Centrifuge at maximum speed 5 min.

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8. Take 300 Ml of supernatant and add 600 Ml of EtOH + 2% NaAc. Invert and leave at −80 or −20°C to increase DNA precipitation. 9. Centrifuge at maximum speed 20 min at room temperature. 10. Resuspend in 25 Ml of H2O or TE (H2O is better when planning to transform this DNA by electroporation). 11. Use 2 Ml for electrocompetent bacteria (e.g., DH5A strain) transformation. Next day, pick up colonies and purify the plasmid DNA by classical bacterial Miniprep. 12. Identify positive recombinants and check their genomic structure by a battery of informative restriction enzymes. 3.2. Puriﬁcation of Viral Prestocks by Banding on Iodixanol

An E1-deletion renders adenovirus incapable of replicating by itself, so a permissive cell line like HEK-293, that provide the E1 genes, should be used to generate and amplify recombinant viruses reaching high titers up to 1012 vp/ml. When a complete cytopathic effect is observed between 48–72 h after the infection on HEK-293 monolayers, up to 90% of adenoviral vectors remain intracellularly. The classical procedure to release the virus from the cells to the supernatant is based on repeated freezing and thawing cycles (up to three times). However, this procedure may destroy the virions if the sample is frosted and thawed too many times, and it would reduce the titer of the stock. Here, we describe an alternative method as efﬁcacious as freezing and thawing in releasing the virus from the cell, but less timeconsuming. Virus puriﬁcation is essential in order to remove defective particles, remove cell debris and media components and concentrate the virus to a level suitable for injection. Classical methods to purify adenovirus are based on density gradient centrifugation with cesium chloride. However, the concentrations used to band the virus are hyperosmotic and the exchange of buffer of virus preparation is required. A new method based on iodixanol overcomes this problem, as is nontoxic to cells and can be administered in vivo (22). Furthermore, other advantages as an increase of functional to physical particles ratio and the possibility of a large-scale puriﬁcation pose this method as the election for virus puriﬁcation.

3.2.1. Virus Ampliﬁcation

1. Amplify the cells. Typically, ten 15-cm-diameter plates of 293 cells yield approximately 4 × 1012 vp in total (30,000 vp/cell). 2. Infect cells with either 300 vp/cell or do a 1/8–1/10 dilution from a cell extract (usually, cell extracts contain 1 × 109 TU/ml and a cell extract from a 15-cm-diameter plate can be used to infect 8–10 plates of 15 cm diameter).

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3. Leave the cells in the incubator for 48–72 h. Check the cytopathic effect 48 h postinfection. Make sure to harvest the cells when these are rounded but not completely detached. If you wait too long, most of the virus will be in the supernatant, and this virus is lost during the puriﬁcation process. 4. Harvest the cells and the medium by pipetting the cells and centrifuge the 50-ml Falcon tubes at 3,000 × g during 5 min. 5. Keep 40 ml of the supernatant and discard the rest by aspiration. Resuspend the cells from each Falcon tube carefully and join the cell suspensions into one Falcon tube. Leave approximately 5 ml of supernatant and transfer the cell suspension to a new 15-ml Falcon tube. 3.2.2. Preparing the Virus Cell Extract for the Iodixanol Gradient

1. Add an equal volume of tetrachloroethylene. 2. Shake vigorously until a single phase is formed. This lyses the cells, releasing the virus. 3. Centrifuge for 20 min at 3,000 × g to separate organic and aqueous phases. Lower layer corresponds to tetrachloroethylene, cell debris remains in the interphase and upper layer corresponds to medium containing the virus (see Note 3). 4. Pipette off upper layer and transfer to a new tube. Store at 4°C.

3.2.3. Preparing the Iodixanol Gradient

1. Lay the following amounts of each solution carefully into each Beckman ultracentrifuge tubes to form the step gradient: 0.5 ml Sol 1 + 1.8 ml Sol 2 + 1.8 ml Sol 3 + 1.8 ml Sol 4. 2. Add 6.5 ml of the cell extract onto each tube. 3. Ultracentrifuge at 155,000 × g at 10°C for 1 h. Adenovirus will appear as a bluish white band. 4. Remove and discard the upper portion by suction. 5. Carefully remove the band with a pipette and collect into a 50-ml Falcon tube and place it on ice. Try to take the virus (bluish band) in volume as small as possible to reduce virus dilution (1.5–2.5 ml). 6. Aliquot the virus and store at −80°C (see Note 4).

3.3. In Vivo Delivery of Adenovirus Vectors to Tumors

After capsid modiﬁcation, the behavior of adenoviruses injected systemically can be altered. To analyze these changes, it is important to distinguish between two terms that are sometimes confusing: biodistribution and transduction. The former, refers to the virus fate once it is administered and depends on anatomical features like fenestration size, and the interaction with blood cells and proteins. On the other hand, transduction includes two additional steps, gene transfer (dependent on the capsid) and expression (dependent on gene promoter).

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The decrease in liver transduction due to detargeting mutations sometimes is reﬂected in a higher viremia. Methods to evaluate virus biodistribution (e.g., RT-PCR) and transduction (e.g., luciferase expression) are extensively explained elsewhere, so here we focus on a method to determine viremia and blood persistence after systemic administration of adenovirus. 3.3.1. Systemic Administration of Adenovirus

1. Prepare the virus at the desired concentration in 150 Ml of PBS at room temperature. For most vectors and mouse strains, the lethal dose 50 is around 5 × 1010–1011 vp/mouse. 2. Load the virus in a 0.5- to 1-ml syringe, insert a 28-G needle, and remove the air from the syringe. 3. Immobilize the animal in a small chamber. For 25–35 g animals, a 50-ml Falcon tube with V-shaped cut next to the cap as a tail exit and a hole at the bottom to allow breathing can be used. 4. Localize the tail veins at the sides of the tail and clean the tail with ethanol. The warmth of a lamp will dilate the vein and would help to better visualize it. 5. Inject the needle with the syringe as horizontal as possible. A site at three-quarters the length of the tail toward the tip is a good point to start because, in case of misinjection, it is possible to use the same vein closer to the animal body. The needle should enter 2–3 mm into the vein. 6. Start the injection slowly. If a resistance is noted or the injection site becomes white, it indicates that the solution is not entering the vein. 7. Upon retraction of the needle, the injection site should bleed if the injection is correct. Apply a pressure for 30 s to stop bleeding.

3.3.2. Determination of Viremia and Blood Persistence

1. At different times, postinjection (1c, 5c, 10c, 15c, 20c, 30c, 60c, 4 h, and 24 h), bleed the animal from the tail, cutting it 0.5 cm from the tip. Control the bleeding by applying light pressure on the tail and collect one drop of blood into a heparinized tube. 2. Centrifuge 5 s to bring the blood to the bottom of the microcentrifuge tube. Keep the tube on ice until all time points have been collected. 3. Spin down the cells brieﬂy and take an aliquot of plasma to assay the virus concentration. 4. Viral particles can be measured by RT-PCR and functional particles by antihexon staining can also be analyzed depending on vector modiﬁcation.

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4. Notes 1. As the only plasmid that can grow in yeast is the recombination product, it is not necessary to open or linearize the adenovirus genomic plasmid for the recombination. However, if a site with a unique enzyme is available, then such a site should be targeted. That is, the 40-bp ﬂanking regions of the CAU or CAL fragments should fall upstream and downstream of this site. Then, linearization of the adenovirus genomic plasmid with this unique enzyme increases the rate of homologous recombination. 2. Alternatively to the negative selection step the URA gene can be ﬂanked by unique restriction sites and cut it after the positive selection to proceed as in a cut-repair protocol. 3. Usually, 20 min of centrifugation is sufﬁcient. Upper layer sometimes remains slightly turbid (centrifugation time can be increased if upper layer is still very turbid). 4. Virus eluted in iodixanol can be frozen at −80°C and preserves the virus well, but iodixanol does not allow quantiﬁcation of physical titer by OD260. In order to solve this either, the virus can be maintained in iodixanol and calculate the physical titer with another method (quantitative PCR, protein quantiﬁcation, etc.) or the virus can be passed through sephadex G100 column and eluted in PBS+/+5% glycerol.

Acknowledgments We thank to Manel Cascalló and Lynda Coughlan for critical reading of the manuscript. References 1. Alemany, R., and Curiel, D. T. (2001) CARbinding ablation does not change biodistribution and toxicity of adenoviral vectors, Gene therapy 8, 1347–1353. 2. Bayo-Puxan, N., Cascallo, M., Gros, A., Huch, M., Fillat, C., and Alemany, R. (2006) Role of the putative heparan sulfate glycosaminoglycan-binding site of the adenovirus type 5 ﬁber shaft on liver detargeting and knob-mediated retargeting, The Journal of general virology 87, 2487–2495. 3. Einfeld, D. A., Schroeder, R., Roelvink, P. W., Lizonova, A., King, C. R., Kovesdi, I., and Wickham, T. J. (2001) Reducing the native tropism of adenovirus vectors requires removal

of both CAR and integrin interactions, Journal of virology 75, 11284–11291. 4. Smith, T. A., Idamakanti, N., Rollence, M. L., Marshall-Neff, J., Kim, J., Mulgrew, K., Nemerow, G. R., Kaleko, M., and Stevenson, S. C. (2003) Adenovirus serotype 5 ﬁber shaft inﬂuences in vivo gene transfer in mice, Human gene therapy 14, 777–787. 5. Kritz, A. B., Nicol, C. G., Dishart, K. L., Nelson, R., Holbeck, S., Von Seggern, D. J., Work, L. M., McVey, J. H., Nicklin, S. A., and Baker, A. H. (2007) Adenovirus 5 ﬁbers mutated at the putative HSPG-binding site show restricted retargeting with targeting peptides in the HI loop, Mol Ther 15, 741–749.

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6. Shayakhmetov, D. M., Gaggar, A., Ni, S., Li, Z. Y., and Lieber, A. (2005) Adenovirus binding to blood factors results in liver cell infection and hepatotoxicity, Journal of virology 79, 7478–7491. 7. Kalyuzhniy, O., Di Paolo, N. C., Silvestry, M., Hofherr, S. E., Barry, M. A., Stewart, P. L., and Shayakhmetov, D. M. (2008) Adenovirus serotype 5 hexon is critical for virus infection of hepatocytes in vivo, Proceedings of the National Academy of Sciences of the United States of America 105, 5483–5488. 8. Waddington SN, M. J., Bhella D, Parker AL, Barker K, Atoda H, Pink R, Buckley SM, Greig JA, Denby L, Custers J, Morita T, Francischetti IM, Monteiro RQ, Barouch DH, van Rooijen N, Napoli C, Havenga MJ, Nicklin SA, Baker AH. (2008) Adenovirus serotype 5 hexon mediates liver gene transfer., Cell 132, 397–409. 9. Alba, R., Bradshaw, A. C., Parker, A. L., Bhella, D., Waddington, S. N., Nicklin, S. A., van Rooijen, N., Custers, J., Goudsmit, J., Barouch, D. H., McVey, J. H., and Baker, A. H. (2009) Identiﬁcation of coagulation factor (F)X binding sites on the adenovirus serotype 5 hexon: effect of mutagenesis on FX interactions and gene transfer, Blood 114, 965–971. 10. Carlisle, R. C., Di, Y., Cerny, A. M., Sonnen, A. F., Sim, R. B., Green, N. K., Subr, V., Ulbrich, K., Gilbert, R. J., Fisher, K. D., Finberg, R. W., and Seymour, L. W. (2009) Human erythrocytes bind and inactivate type 5 adenovirus by presenting Coxsackie virus-adenovirus receptor and complement receptor 1, Blood 113, 1909–1918. 11. Stone, D., Liu, Y., Shayakhmetov, D., Li, Z. Y., Ni, S., and Lieber, A. (2007) Adenovirusplatelet interaction in blood causes virus sequestration to the reticuloendothelial system of the liver, Journal of virology 81, 4866–4871. 12. Morrison, J., Briggs, S. S., Green, N., Fisher, K., Subr, V., Ulbrich, K., Kehoe, S., and Seymour, L. W. (2008) Virotherapy of ovarian cancer with polymer-cloaked adenovirus retargeted to the epidermal growth factor receptor, Mol Ther 16, 244–251. 13. Korn, T., Nettelbeck, D. M., Volkel, T., Muller, R., and Kontermann, R. E. (2004) Recombinant bispeciﬁc antibodies for the targeting of adenoviruses to CEA-expressing tumour cells: a comparative analysis of bacterially expressed single-chain diabody and tandem scFv, The journal of gene medicine 6, 642–651.

14. Dmitriev, I., Kashentseva, E., Rogers, B. E., Krasnykh, V., and Curiel, D. T. (2000) Ectodomain of coxsackievirus and adenovirus receptor genetically fused to epidermal growth factor mediates adenovirus targeting to epidermal growth factor receptor-positive cells, Journal of virology 74, 6875–6884. 15. Bayo-Puxan, N., Gimenez-Alejandre, M., Lavilla-Alonso, S., Gros, A., Cascallo, M., Hemminki, A., and Alemany, R. (2009) Replacement of adenovirus type 5 ﬁber shaft heparan sulfate proteoglycan-binding domain with RGD for improved tumor infectivity and targeting, Human gene therapy 20, 1214–1221. 16. Magnusson, M. K., Henning, P., Myhre, S., Wikman, M., Uil, T. G., Friedman, M., Andersson, K. M., Hong, S. S., Hoeben, R. C., Habib, N. A., Stahl, S., Boulanger, P., and Lindholm, L. (2007) Adenovirus 5 vector genetically re-targeted by an Afﬁbody molecule with speciﬁcity for tumor antigen HER2/neu, Cancer gene therapy 14, 468–479. 17. Coughlan, L., Vallath, S., Saha, A., Flak, M., McNeish, I. A., Vassaux, G., Marshall, J. F., Hart, I. R., and Thomas, G. J. (2009) In vivo retargeting of adenovirus type 5 to alphavbeta6 integrin results in reduced hepatotoxicity and improved tumor uptake following systemic delivery, Journal of virology 83, 6416–6428. 18. He, T. C., Zhou, S., da Costa, L. T., Yu, J., Kinzler, K. W., and Vogelstein, B. (1998) A simpliﬁed system for generating recombinant adenoviruses, Proceedings of the National Academy of Sciences of the United States of America 95, 2509–2514. 19. Hokanson, C. A., Dora, E., Donahue, B. A., Rivkin, M., Finer, M., and Mendez, M. J. (2003) Hybrid yeast-bacteria cloning system used to capture and modify adenoviral and nonviral genomes, Hum Gene Ther 14, 329–339. 20. Gietz, R. D., and Woods, R. A. (2002) Transformation of yeast by lithium acetate/ single-stranded carrier DNA/polyethylene glycol method, Methods in enzymology 350, 87–96. 21. Gietz, R. D., and Woods, R. A. (2006) Yeast transformation by the LiAc/SS Carrier DNA/ PEG method, Methods in molecular biology (Clifton, N.J) 313, 107–120. 22. Peng, H. H., Wu, S., Davis, J. J., Wang, L., Roth, J. A., Marini, F. C., 3rd, and Fang, B. (2006) A rapid and efﬁcient method for puriﬁcation of recombinant adenovirus with arginine-glycine-aspartic acid-modiﬁed ﬁbers, Analytical biochemistry 354, 140–147.

Chapter 3 Construction of Targeted and Armed Oncolytic Adenoviruses Konstantin Doronin and Dmitry M. Shayakhmetov Abstract Oncolytic (replication-competent) adenoviruses (Ads) represent the most advanced platform for cancer gene therapy. These viral vectors ablate tumors by killing tumor cells in the process of virus replication. As progeny virions are released, they infect remaining cancer cells, generating a bystander effect. Ads engineered for increased cancer speciﬁcity produce less damage to normal tissues. First-generation oncolytic Ads have demonstrated acceptable levels of safety while the efﬁcacy was observed only in combination with chemotherapy and/or radiation. Second-generation oncolytic Ads are armed with therapeutic transgenes to increase release, spread, and bystander effect for enhancing the efﬁcacy. Third-generation oncolytic Ads are armed vectors with capsid modiﬁcations for transductional detargeting from normal tissues and targeting to cancer cells. Chemical modiﬁcation of the capsid additionally improves therapeutic window. Here, we describe methods for generation and characterization of advanced-generation oncolytic Ads. Key words: Adenovirus, Oncolytic virotherapy, Genetic vectors, Animal models, Experimental neoplasms

1. Introduction Adenoviruses (Ads) belong to the family Adenoviridae. Ads have icosahedral protein capsids without a lipid envelope; their genome is a double-stranded DNA of approximately 30–40 kilobase pairs (kbp). There are 51 serotypes of Ads divided into groups A–F. Ad type 5 (Ad5) from group C has been used for the past three decades as a vector for gene transfer. Historically, replication-competent Ad5 vectors were initially described. Later, with the development of complementing cell lines expressing some of Ad5 proteins in trans, it became possible to construct replication-deﬁcient vectors (1). Replication-deﬁcient Ad5 vectors have been used in numerous gene therapy studies aimed to replace defective genes with correct

David H. Kirn et al. (eds.), Oncolytic Viruses: Methods and Protocols, Methods in Molecular Biology, vol. 797, DOI 10.1007/978-1-61779-340-0_3, © Springer Science+Business Media, LLC 2012

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ones (replacement gene therapy). It was found, however, that the transient nature of expression from these vectors is a major limitation. In cancer gene therapy, however, transient expression does not represent an obstacle since the goal in this case is to kill infected cancer cells. As replication-deﬁcient vectors have higher cloning capacities and are generally considered a safer alternative, these vectors were used in gene therapy studies initially for delivery of therapeutic genes to cancer cells in tumors. It was soon realized that replication-competent Ad vectors might have advantages over replication-deﬁcient vectors in the context of cancer gene therapy because these vectors are self-amplifying within tumors and they can reinfect tumor cells that were not infected upon initial transduction. Repeating cycles of reinfection result in elimination of tumors. This envisaged scenario led to expectations that oncolytic (replication-competent) Ad can be used for systemic (intravascular) treatment rather than the intratumoral injection route that has been used in the majority of clinical trials. Wild-type Ads of various serotypes were used in clinical trials as early as 1956 and were found to lack signiﬁcant anticancer activity (2). A ﬁrst-generation cancerspeciﬁc vector ONYX-015 with a deletion of E1B-55K was believed at that time to confer selectivity of replication in p53 negative (cancer) versus p53 positive (normal) cells. This vector was evaluated in clinical trials in the 1990s and was found to lack activity as a single agent although some activity was found in combination with chemotherapy (3). More recently, a vector similar to ONYX-015 but lacking the E3 region (Oncorine, H101) was approved for clinical use in China against head and neck cancers using intratumoral injection in combination with chemotherapy. The approval of Oncorine was an important milestone for the ﬁeld of cancer gene therapy and oncolytic viruses; however, more efﬁcacious oncolytic Ads need to be developed to win regulatory approval in other countries. In addition, for commercial success, oncolytic Ads will have to be more efﬁcacious than existing therapeutic options to provide a cost-efﬁcient alternative to surgery, chemotherapy, and radiation (4). Here, we describe techniques for the construction of advancedgeneration oncolytic (replication-competent) Ad vectors with enhanced cancer selectivity and anticancer efﬁcacy. We have previously described vectors in which replication was rendered cancerselective by the modiﬁcation of Ad transactivator E1A protein, and tissue speciﬁcity of replication was conferred by substitution of lung-speciﬁc SP-B promoter for native E4 promoter (5, 6). Anticancer efﬁcacy of oncolytic viruses can be increased by expression of therapeutic genes (7). Here, we describe methods for construction of Ad5-based vectors expressing therapeutic transgenes in the context of oncolytic Ad or from a binary expression system. Additionally, tropism of Ad5-based vectors is determined by the presence of coxsackie-adenovirus receptor (CAR) on the target

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tumor cells. Changing Ad5 vector ﬁbers to ﬁbers targeted to a different receptor, CD46, allows circumvention of the problem of low expression of CAR by some cancer types, thus increasing the efﬁcacy of tumor transduction (8). Finally, chemical modiﬁcation of Ad5 capsid with high molecular weight polyethylene glycol (PEG) changes virus biodistribution, increases efﬁcacy, and can be readily applied to various Ad vectors without a necessity to change the virus genome (9).

2. Materials 1. Escherichia coli strains DH5A and BJ5183. 2. Cell lines: 293, 293N3S (Microbix), A549 (human lung carcinoma), LNCaP (human prostate carcinoma), Hep3B (human hepatocellular carcinoma), American Type Culture Collection (ATCC). KB and HeLaS3 spinner cells were kindly provided by William S.M. Wold (St. Louis University, St. Louis, MO). VK10-9 cells (293 expressing E4 and pIX) were kindly provided by Valeri Krougliak (Mount Sinai School of Medicine, New York). 3. Dulbecco’s modiﬁed Eagle’s medium (DMEM), 2× DMEM, and Joklik-modiﬁed MEM. 4. Fetal bovine serum (FBS). 5. Sodium bicarbonate, glutamine, penicillin/streptomycin, trypsin–ethylenediaminetetraacetic acid (EDTA), phosphatebuffered saline (PBS). 6. Neutral red, sterile stock solution 0.3 g/l in water. 7. 5% sodium deoxycholate. 8. CsCl. 9. 1 M Tris–HCl, pH 8.0. 10. 0.5 M EDTA. 11. TE buffer: Make with 10 mM Tris–HCl, pH 8.0, 1 mM EDTA. 12. Pronase E. 13. 2.5 M CaCl2. 14. BBS transfection buffer: Prepare 2× stock with 50 mM BES, 280 mM NaCl, 1.5 mM Na2HPO4, pH 6.95. 15. Cell factories (Corning, Sigma-Aldrich). 16. Spinner ﬂasks (Bellco). 17. Phenol:chloroform:isoamyl alcohol (25:24:1, v/v). 18. 10% sodium dodecyl sulfate (SDS).

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19. 10% Triton X-100. 20. Benzonase (Novagen, Merck KGaA). 21. Slyde-A-Lyzer dialysis cassettes 10,000 MWCO (Pierce). 22. Econo-Pac 10DG desalting columns (Bio-Rad). 23. 20-kDa succinimide-activated PEG (m-SCM-20K PEG; JenKem Technology). 24. Sephadex G-100 (GE Healthcare). 25. Matrigel (BD Biosciences). 26. Nude (nu/nu) mice, 4- to 6-week-old (Harlan Sprague Dawley). 27. C57BL/6 mice, 4- to 6-week-old (Jackson Laboratory). 28. ABI 7900HT Real-Time PCR System (Applied Biosystems). 29. Kodak In vivo F imaging system (Carestream). 30. DNeasy blood and tissue kit (Qiagen). 31. D-luciferin (Molecular Imaging Products). 32. Alanine aminotransferase kit (BioTron Diagnostics).

3. Methods 3.1. Transcriptional Targeting and Arming of Oncolytic Adenoviruses

Several approaches were described for targeting Ad replication to cancer cells as opposed to normal cells and for the arming of oncolytic Ads with auxillary therapeutic genes. We have described viruses with mutations in E1A transactivator abolishing binding of the mutant proteins to pRB and p300/CBP, thus restricting virus replication to actively dividing (cancer) cells. The vector could be additionally restricted in replication to lung cancers by placing the E4 region under the control of the tissue-speciﬁc SP-B promoter, which is active in lung tissue. To increase their antitumor efﬁcacies, the vectors were engineered to overexpress the adenovirus death protein (ADP, E3-11.6K), resulting in a more efﬁcacious release of Ad from infected cells and consequently in the increased spread of Ad within the tumors (5, 6). To further increase the efﬁcacy of oncolytic Ads, we have suggested the expression of additional therapeutic genes, either from the same oncolytic virus backbone (10) or in the context of a binary Ad vector system. These consist of a replication-defective ﬁrstgeneration Ad vector expressing the therapeutic transgene under the control of regulatable Tet-On system and a replicationcompetent Ad, which provides both oncolytic functions and helper functions for replication-deﬁcient Ad vector (7).

3 3.1.1. Construction of Oncolytic Vectors Combining Mutations in E1A, E3, and E4 Regions by Homologous Recombination Between Viral Genomic DNA and Shuttle Plasmid

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Mutations in E1A were shown to confer cancer selectivity to oncolytic Ads. Double mutants of E1A that prevent binding to pRB and p300/CBP were shown to provide more efﬁcient selectivity as compared to a single mutation preventing binding only to pRB (11). Homologous recombination in 293 cells between genomic DNA of Ad dl1101/1107 [kindly provided by Stanley Bayley (McMaster University, Toronto, ON)] and a shuttle plasmid pLKH encompassing BamHI (60 m.u.) to the right terminus (100 m.u.) [kindly provided by Linda Hawkins (St. Louis University, St. Louis, MO)] was implemented to construct virus vectors KD3, KD3-IFN (5, 10) (Fig. 1a). To construct KD1-SPB with E4-promoter substitution, we transfected E4-expressing 293 cells VK10-9 cells (6, 12). This method can be applied for combining various mutations between existing Ad vectors and new mutations introduced into shuttle plasmids. The shuttle plasmid pFG23 analogous to pLKH is also commercially available (Microbix). 1. Prepare deproteinized genomic Ad DNA. Desalt CsCl-banded virus by dialysis against 100 volumes of 10 mM Tris–HCl, pH 8.0. Add 1/10 volume of 10% SDS to puriﬁed desalted Ad and add Pronase E (Sigma) to a ﬁnal concentration of 1 mg/ml in 10 mM Tris–HCl, pH 8.0, 1 mM EDTA. After incubation at 37°C for 3 h, purify viral DNA by one extraction with phenolchloroform and one extraction with chloroform (see Note 1). 2. Precipitate DNA by addition of 1/10 volume of 3 M NaOAc (pH = 5.2) and 2 volumes of EtOH. After incubation for 1 h at −20°C, pellet DNA by centrifugation at 12,000 × g for 15 min; wash once with 70% EtOH. Repellet by centrifugation at 12,000 × g for 5 min and dissolve in TE buffer (10 mM Tris– HCl, pH 8.0, 1 mM EDTA). One OD260 unit of Ad results in 40 Mg of genomic DNA (see Note 2). Digest viral DNA with SpeI and EcoRI to reduce the background of wild-type virus after transfection. SrfI can also be used in combination with SpeI and EcoRI to further reduce the background of the wildtype virus. 3. Preparation of shuttle plasmid DNA. Introduce mutations into E3 and/or E4 regions of shuttle plasmid pLKH using PCRbased protocol with pfu polymerase. Deletions in E3B result in a modiﬁcation of splicing pattern resulting in increased expression of ADP (E3-11.6 K) (5, 10) (Fig. 1b). Transgenes replacing E3B are expressed strictly late in the infectious cycle, such that expression can be completely blocked by Ara-C, an inhibitor of DNA replication. Deletion of the entire E3 region and insertion of transgenes downstream of ADP gene results in both overexpression of ADP and strictly late expression of transgenes (10) (Fig. 1b).

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Fig. 1. Construction of oncolytic Ad vectors by homologous recombination between viral genomic DNA and shuttle plasmid. (a) Modiﬁcations are introduced to E3 and E4 region in shuttle plasmid. Viral DNA digested with SpeI and EcoRI is cotransfected with shuttle plasmid into 293 or VK10-9 cells. (b) Rescued vectors carry E1A mutations for cancer speciﬁcity, overexpress ADP (E3-11.6K) and express IFN-alpha for increased anticancer activity. E4 promoter replacement by a tissuespeciﬁc promoter results in increased selectivity.

4. Prepare plasmid DNA for transfections by double banding in isopycnic CsCl gradient (see Note 3). 5. Cotransfection of genomic DNA with shuttle plasmid. We routinely use modiﬁed calcium-phosphate precipitation technique for cotransfection (13). Plate low (<50) passage 293 cells in 60 mm dishes 1 day prior to transfection in 5 ml of

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DMEM/10%FBS. Medium with high concentrations of phosphate, such as RPMI1640, cannot be used with this technique. Cells should be 70–90% conﬂuent at the time of transfection (see Note 4). 6. Mix 5 Mg of viral DNA and 15 Mg of shuttle plasmid DNA in 450 Ml of water. Genomic DNA digested with restriction enzymes is used directly without additional puriﬁcation steps. Add 500 Ml of 2× BBS buffer (50 mM BES; 280 mM NaCl; 1.5 mM Na2HPO4; pH 6.95). Add 50 Ml of 2.5 M CaCl2, mix. Add precipitate to 293 cells (or to VK10-9 cells for E4 promoter-modiﬁed vectors) at 1/10 volume of the medium (0.5 ml per 5 ml of medium in 60-mm dish). 7. After overnight incubation with precipitate, replace the medium with DMEM/5% FBS/0.8% agarose overlay. The overlay medium is made by mixing equal volumes of 2× DMEM/10% FBS equilibrated at 37°C and 1.6% agarose equilibrated at 56°C. Neutral red (30 Mg/ml) may be added to overlays to facilitate visualization of plaques. 8. For wild-type E3 or ADP-overexpressing viruses, plaques start to appear on approximately day 7 after transfection. Pick the plaques, resuspended in 100 Ml of DMEM, and freeze at −70°C. 3.1.2. Construction of Vectors Expressing Transgenes from E1 Region and E3 Region by Homologous Recombination Between Two Plasmids

We have described vectors expressing transgenes from expression cassettes inserted either in wild-type E1 region of Ad or substituting for the deleted E1 region (ﬁrst generation Ad vectors) (7, 9) (Fig. 2). Left-hand shuttle plasmids were pXC-1 (Microbix) containing the wild-type E1 region of Ad5 or p722 (14), which is a derivate of pdlE1SP1A (Microbix). p722 was constructed by subcloning SspI-Bst1107I fragment of pdlE1SP1A between SspI and PvuII sites in pBluescript SK(+) (Invitrogen) backbone resulting in a high-copy vector (14). Expression cassettes were cloned into unique HpaI site in pXC-1 located downstream of E1A ORFs or in unique sites of p722 polylinker (see Note 5). Resulting plasmids were cotransfected into 293 cells along with pBHGE3KD3, which has the same version of E3 as vector KD3 for achieving ADP overexpression, while maintaining the cloning capacity characteristic for E3-deleted Ads (14). Replication-competent viruses derived from cotransfection of pXC-1-based plasmids and pBHGKD3E3 are fully replication-competent and were able to carry a large expression cassette for CMV-EGFPLuc (2.8 kbp) (9). Theoretically, the cloning capacity of this vector is 3.3 kbp, including the capacity of Ad5 capsid to encapsidate genomes exceeding wild type by 5% (1,800 bp) and an additional cloning capacity resulting from E3 deletion (1,500 bp). Replication-deﬁcient viruses derived from cotransfection of p722-based plasmids and pBHGKD3E3 were able to carry large expression cassettes for pharmacologically

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Fig. 2. Construction of oncolytic Ad vectors by homologous recombination between two plasmids. (a) Transgenes can be inserted into intact E1 region in pXC-1 or substitute for deleted E1 region in pdlE1SP1A. pBHGKD3E3 carries ADPoverexpressing version of E3 from KD3 vector. After cotransfection into 293 cells, pBHGKD3E3, which lacks encapsidation signal, replicates in cells but can only be rescued into virus after homologous recombination with a shuttle plasmid. (b) Rescued viruses carry either CMV-EGFP expression cassette for bioluminescence imaging or Tet-On-TRAIL cassette for increased anticancer efﬁcacy.

(Tet-On) regulated expression of transgenes (7). The cloning capacity of vectors derived from p722-pBHGKD3E3 combination is 6.4 kbp. pBHG10 and pBHG11 carry larger deletions in E3 and allow for insertion of transgenes up to 8 kbp (15), however, pBHGKD3E3 allows for overexpression of ADP resulting in earlier appearance of plaques (days 7–10 after transfection) and 1.8 kbp of additional cloning capacity as compared with pBHGE3, which carries wild-type Ad5 E3 (15). 1. Cotransfection of shuttle plasmid with pBHGKD3E3 for homologous recombination in 293 cells. Insert the expression cassette into the polylinker of left-hand shuttle plasmid or into HpaI site in pXC-1 (see Note 5).

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2. Prepare plasmid DNA by double banding in CsCl gradients (see Note 3). Mix 5 Mg of left-hand shuttle plasmid with 15 Mg of pBHGKD3E3 and make the calcium-phosphate precipitate as described in Subheading 3.1.1. Add precipitate to a monolayer of 293 cells (1/10 of the medium volume) and incubate overnight. After incubation, remove medium and add agarose overlay. 3. Alternatively, regular liquid medium can be used instead of overlay. pBHGKD3E3 is derived from pBHG10 which carries a deletion of Ad encapsidation signal resulting in the absence of wild-type virus background after transfection (see Note 6). Plaques of recombinant vectors appear on days 7–10 after transfection. 3.2. Transductional Detargeting and Targeting of Oncolytic Adenoviruses

Ad5 uses CAR as a primary receptor. CAR is ubiquitously expressed in the human body and this imposes limitations on systemic administration of Ad vectors in human patients. Substituting Ad5 ﬁber with cancer-speciﬁc targeting moieties would result in oncolytic Ads with increased cancer selectivity at the level of transduction. We have described Ad vectors with Ad5 ﬁbers replaced with Ad35 ﬁbers, which use CD46 as a receptor (16). Oncolytic Ads based on this ﬁber replacement strategy were constructed and demonstrated to have changed transduction tropism (8, 17) (Fig. 3). To construct

Fig. 3. Construction of chimeric Ad5GFP/F35 with Ad5 capsid and Ad35 ﬁber. (a) Modiﬁed ﬁber gene consisting of Ad5 tail and Ad35 shaft and knob was inserted into Ad5 L5 region using PCR-based protocol and transferred by recombination in E. coli into shuttle plasmid pAdHM4. Ad genome was released from pAdHM4GFPF35 by digestion with PacI and transfected into 293 cells resulting in virus rescue. (b) Structure of the chimeric Ad5/Ad35 ﬁber. Italicized amino acid residues are conserved in ﬁbers of most Ad serotypes.

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the chimeric vector, the Ad5 ﬁber gene in pAdGFP was substituted by an Ad5/35 chimeric ﬁber gene generated by the two-step PCR protocol using pfu polymerase in a shuttle vector carrying CMVEGFP cassette in E3. Chimeric Ad5/35 ﬁber gene consists of (1) sequences corresponding to the Ad5 ﬁber 5c nontranslated region and the ﬁrst 132 bp of the ﬁber tail domain, (2) the Ad35 shaft and knob domains (nt 132–991), and (3) the Ad5 E4 region, including the Ad5 ﬁber polyadenylation signal domains. The resulting plasmid was named pAdGFP/F35. To generate full-length E1/ E3-deleted vector genomes, modiﬁed E3 and ﬁber regions were transferred by homologous recombination in RecA+ E. coli strain BJ5183 to pAdHM4 containing Ad5 genome with E1 and E3 regions deleted and Ad5 sequences ﬂanked by PacI sites (18). Hexon of Ad5 was recently shown to bind blood factors with high afﬁnity (19) resulting in transduction of liver hepatocytes in vivo. Modiﬁcation of hexon in HVR5 or HVR7 results in reduced transduction of hepatocytes (19, 20). We have shown that insertion of BAP peptide into HVR5 of oncolytic Ad hexon results in an improved therapeutic window (21). We have constructed hexon-modiﬁed vector by homologous recombination between shuttle pAdHexBAP plasmid (22) and genomic virus DNA. 1. Substitute Ad5 ﬁbers with chimeric ﬁbers in plasmids. Introduce mutation into Ad5 ﬁber gene in an appropriate shuttle plasmid containing the ﬁber region using the two-step PCR protocol. 2. Sequence the resulting plasmid clones to ensure that no additional mutation occurred. 3. Excise modiﬁed ﬁber gene with restriction enzymes ﬂanking the ﬁber gene. Recombine modiﬁed ﬁber gene with pAdHM4 (18) linearized by SrfI. Analyze resulting recombinants by PCR. Amplify correct plasmid clones in E. coli DH5A and purify by double-CsCl banding. 4. Verify the correct structure of the chimeric ﬁber gene by restriction endonuclease digestion and sequencing. 5. Rescue recombinant virus with modiﬁed ﬁbers from plasmids. To produce recombinant viruses, digest full-genome plasmids with PacI to release the viral genomes and transfect into 293 cells as described in Subheading 3.1.1. 6. Collect plaques and store at −70°C. 7. To modify hexon, construct hexon-modiﬁed shuttle plasmids. Introduce mutations into hexon gene in a shuttle plasmid (e.g., pAdHex) by using PCR-based protocol with pfu polymerase. 8. Rescue of recombinant virus with modiﬁed hexon from plasmids. Digest genomic Ad DNA with PmeI and cotransfect with the shuttle plasmid into 293 cells as described in Subheading 3.1.1.

3

3.3. PlaquePuriﬁcation, Propagation, Genome Analysis 3.3.1. Plaque Assay

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Plaque assay is useful for the puriﬁcation of recombinant virus and for determining virus titers (see Note 7). 293 cells are routinely used for propagation of E1-deleted Ad vectors. A549 cells rather than 293 should preferably be used for plaque-puriﬁcation and propagation of Ads with modiﬁcations in E1 region because 293 cells carry E1 region and wild-type version of E1 can be restored in recombinant virus due to homologous recombination. 1. Plate cells on 60-mm dishes 1 day prior to assay. Cells should be conﬂuent at the time of use. 2. Prepare serial dilutions of virus in serum-free DMEM. Pipette tips should be changed after each dilution and barrier tips should be used. 3. Remove medium and add 0.5 ml virus per dish. Adsorb the virus on the monolayer for 1 h with periodical rocking of the dish for even distribution of virus over the monolayer. 4. Add 10 ml of overlay medium containing 0.8% agarose and 33 Mg/ml of neutral red (see Note 8). 5. Depending on studied Ad, plaques will appear between days 3 and 30 postinfection. For wild-type Ad5 ﬁnal titers may be counted on day 15 (very few new plaques are going to appear later). For vectors with small plaques appearing late (e.g., E3-11.6K (ADP)-negative vectors), it is preferable to count plaques on day 30 after infection. 6. For plaque-puriﬁcation, pick plaques with agarose plugs using barrier pipette tips or Pasteur pipettes, resuspend in 100 Ml of DMEM and store at −70°C.

3.3.2. Infection of Cells in Monolayer and Extraction of Virus DNA from Infected Cells

1. Grow 293 or A549 cells in 48- or 24-well plates to conﬂuency. Add 10 Ml of plaque resuspended in DMEM. Wait for complete cytopathic effect (CPE), i.e., all cells are rounded up and detached from the plastic surface. Depending on the vector and the cells, this will take 2–14 days. Collect cells and the medium, freeze-thaw three times to release the virus from cells, freeze at −70°C. This will result in a stock with a titer of approximately 108 plaque forming units (PFU)/ml. 2. Grow 293 or A549 cells in 60-mm dishes to semiconﬂuency. Remove medium and add 50 Ml of virus diluted in 0.5 ml of DMEM (approximately 5 PFU/cell). Adsorb for 1 h with periodical rocking of the dish to spread the virus uniformly. Add 5 ml of DMEM/10% FBS to each dish and incubate for 2 days. Usually, CPE is complete on days 2–3 (all cells are rounded up). Harvest infected cells in medium by pipetting. Pellet cells by low speed centrifugation (300 × g, 10 min). Cell pellets can be frozen at this point.

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3. To make a concentrated stock, resuspend the pellet in 1 ml of DMEM, freeze-thaw three times to release the virus, add glycerol to 10%, freeze at −70°C. 4. To extract virus DNA, add 300 Ml of extraction buffer (150 mM NaCl, 10 mM Tris–HCl, pH 8.0, 0.5% Triton X-100) to cell pellet from step 2, mix, incubate on ice for 30 min with periodical shaking. Spin down in a microcentrifuge at 12,000 × g for 10 min. Collect the supernate, add pronase E (1 mg/ml) and SDS to 10%. Incubate at 37°C for 1 h. Extract once with phenol-chloroform, separate phases by centrifugation at 12,000 × g for 5 min, collect supernatant, extract once with chloroform, centrifuge for 5 min at 12,000 × g, and collect supernate. Add 50 Ml of 3 M NaOAc, pH 5.2 and 1 ml of EtOH. Incubate at −20°C for 1 h, pellet DNA in microcentrifuge at 12,000 × g for 10 min. Wash DNA pellet with 1 ml of 70% EtOH, centrifuge at 12,000 × g for 10 min. Remove and discard the supernate, dissolve DNA pellet in 50 Ml of TE or water. This DNA can be analyzed by restriction digestion or PCR. RNA can be removed from the samples by adding RNase A (1 Ml of 10 mg/ml stock solution) and incubation at 37°C for 1 h. 3.3.3. Large-Scale Vector Preparation and Puriﬁcation

Ads can be grown on a large scale in cell monolayers in 15-cm dishes, in cell factories, in roller bottles, or in spinner culture. 293 cells and spinner-adapted 293N3S cells (23) express E1 proteins and are used for growing E1-deleted vectors. For growing E1Amodiﬁed oncolytic vectors, we routinely use A549 cells grown as a monolayer in cell factories and spinner KB or HeLaS3 cells. 1. Grow 6 l of spinner KB cells in Joklik MEM/5% HS or one cell factory of A549 cells in DMEM/5% FBS. This should result in approximately 1010 cells. Infect cells with 5–20 PFU/cell of Ad. For spinner culture, reduce the infection volume to 1/10 of original volume, infect with spinning for 6 h, and then add fresh medium to the original culture volume (see Note 9). For cell factory, adsorb virus on cells for 3 h in 200 ml of medium with periodical rocking for even virus distribution on the surface. After adsorption, add fresh medium to the original volume. Incubate for 48–72 h. 2. Collect cells in the medium from spinners or cell factory. Pellet cells by centrifugation at 300 × g for 15 min in 500-ml Corning conical-bottom centrifuge bottles. Turn off the brake. Cell pellets can be frozen at this point. 3. Resuspend the cell pellet (10 ml) in 10 ml of 50 mM Tris– HCl, pH 8.0, 1 mM MgCl2. Add 1/10 volume (2 ml) of 5% sodium deoxycholate, mix by rotating for 10 min (see Note 10). Complete lysis of cells results in a very viscous solution.

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Add 250 U of benzonase, incubate at 37°C for 1 h. Centrifuge for 20 min at 1,400 × g and collect the supernatant. Add 0.51 g of CsCl to each one ml of virus-containing supernatant. 4. Transfer the solution to Beckman OptiSeal ultracentrifuge tubes. Centrifuge in 70Ti rotor at 93,000 × g in Beckman ultracentrifuge at 4°C for 16 h. 5. Stop the centrifuge without the brake. The virus band will be in the middle of the tube. Collect the virus band with a syringe by puncturing the side of the tube. Dilute the virus band in rebanding solution (0.51 g of CsCl per ml in 50 mM Tris–HCl, pH 8.0). Reband the virus by ultracentrifugation for 16 h. 6. Collect the virus band in minimal volume (approximately 1 ml) and dialyze against two 1 l changes of 10 mM Tris–HCl, pH 8.0 at 4°C for 2 h. 7. Dilute the virus as needed in 10 mM Tris–HCl, pH 8.0, add sterile glycerol to 10%, aliquot, and store frozen at −70°C. 8. Virus concentrations can be determined by measuring OD260. One unit corresponds to 1.1 × 1012 viral particles (vp) of Ad5. For determining the biological activity, virus should be titered by plaque assay or limiting dilution assay (see Note 11). Virus particle to plaque-forming unit ratio for good preparations of the vectors is usually around 10. 3.4. Chemical Modiﬁcation with Polyethylene Glycol

Puriﬁed Ad capsids can be chemically modiﬁed to modulate in vivo biodistribution and anticancer properties. PEG-modiﬁed (PEGylated) proteins are currently in clinical use. PEGylation leads to improved pharmacokinetics (extended half-life) of protein in vivo. PEGylation was shown to reduce innate and adaptive immune responses to Ad and reduce viral binding to platelets, red blood cells, and Kupfer cells. We have recently shown that PEGylation of oncolytic Ad with high molecular weight (20 kDa) PEG efﬁciently detargets Ad from hepatocytes and increases anticancer efﬁcacy in animal models of cancer (9). 1. PEGylation of adenovirus with 20-kDa PEG. Desalt Ad using Econo-Pac 10DG desalting columns (Bio-Rad, Hercules, CA), replacing the CsCl (or Tris) buffer with KPBS (136 mM NaCl, 2.6 mM KCl, 1.7 mM KH2PO4, 10 mM K2HPO4). Do not use Tris-based buffers, as this will interfere with the PEGylation reaction. 2. Add 20-kDa succinimide-activated PEG to the virus sample at a concentration of 10 mg per 1 × 1012 viral particles per ml of reaction mixture. Do not increase the concentration of virus or PEG in the reaction mixture as this will result in virus precipitation (24). Rotate the reaction mixture at room temperature for 1 h.

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3. After incubation, remove the unbound PEG by gel-ﬁltration on a Sephadex G-100 size-exclusion column. Process the mock-treated Ad in the same way but in the absence of PEG. 4. Assess properties of PEGylated Ad preparations. The degree of PEGylation can be estimated by Western blotting with antibodies against speciﬁc Ad proteins, e.g., hexon (24). Titering PEGylated Ad is not feasible, as PEGylation results in an almost complete loss of infectivity of Ad in vitro (9, 24). Vector concentration can be measured by quantitative real-time polymerase chain reaction (qPCR) and expressed as viral genomes per ml (see Note 12). 3.5. Characterization of Oncolytic Ads In Vivo 3.5.1. Anticancer Activity in Animal Models of Cancer

The goal of studies on oncolytic Ads is to produce efﬁcient and safe anticancer agents. Testing anticancer activity in animal models provides valuable information on efﬁcacy. Preliminary data on efﬁcacy is usually obtained in xenograft models of cancer, with human cancer cells grown subcutaneously or orthotopically in immunodeﬁcient nude mice. Alternatively, immunocompetent animal tumor models (Cotton rats, Syrian hamsters) can be used. However, as rodent cells are less permissive to Ad replication than human cells, decreased anticancer activity of oncolytic viruses might be expected as compared to human xenografts. 1. Study oncolytic vectors in subcutaneous xenograft models. Grow LNCaP or Hep3B cells as monolayers. Collect cells with trypsin–EDTA. Neutralize trypsin promptly by adding DMEM/10% FBS (see Note 13). 2. To establish subcutaneous tumors, inject 107 tumor cells in 100 Ml of DMEM/50% Matrigel subcutaneously in the hind ﬂank of nude mice. Addition of Matrigel greatly improves the percentage of mice developing tumors and reduces variation in tumor size and time of appearance. To avoid shearing cells during injection, draw the cells in DMEM/Matrigel into the syringe without the needle attached. Use 23-gauge needles for injection. 3. LNCaP form established tumors at 30 days after injection of cells. Hep3B form established tumors by day 21 (see Note 14). Inject established tumors intratumorally with 109–1011 vp of Ad diluted in 100 Ml of PBS. Alternatively, inject Ad vector in 100 Ml of PBS into tail vein using tuberculin syringe. 4. Depletion of liver macrophages (Kupfer cells) and anticoagulation treatment were shown to increase the anticancer efﬁcacy of oncolytic Ad delivered systemically (25). To deplete macrophages, inject 3 × 1010 vp of ﬁrst-generation replication deﬁcient Ad in 100 Ml of PBS 4 h before oncolytic Ad injection. To deplete coagulation blood factors, 1 day before Ad injection inject mice subcutaneously with 133 Mg of warfarin in 100 Ml of peanut oil (see Note 15).

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5. Measure two perpendicular diameters of tumors twice a week. Euthanize mice when tumor volume exceeds 2 ml. Calculate tumor volume as width2 × length × 1/2. 6. Study oncolytic Ad vectors in orthotopic lung metastasis xenograft model. Inject 3 × 106 A549 cells into the tail vein of nude mice. 7. After 10 days, inject these mice intravenously with 5 × 1010 vp of oncolytic Ad. 8. At day 7 after virus injection, euthanize mice, collect the lungs, and perfuse lungs with India ink solution. Count metastases per lung lobe. 3.5.2. Biodistribution and Hepatotoxicity

1. Use real-time PCR for studying the biodistribution in xenograft models. Inject nude mice bearing xenograft tumors intratumorally or intravenously. At various time points postinjection (days 0, 1, and 2), euthanize animals and collect tissues (blood, tumor, liver, spleen, lungs). 2. Extract DNA from tissue samples with DNeasy blood and tissue kit (Qiagen). 3. Perform qPCR with QuantiTect SYBR green PCR master mix and primers speciﬁc for Ad or transgene sequences. Generate standard curves with plasmid or Ad DNA diluted in puriﬁed DNA from mock-injected animals. Perform qPCR on an ABI 7900HT Real-Time PCR System in absolute quantiﬁcation mode. Analyze results with ABI analysis software. 4. Use bioluminescence imaging for studying transgene expression and vector spread. Inject tumor-bearing mice with ﬁreﬂy luciferase-expressing oncolytic Ad. 5. At 24 h after injection and at later time points, anesthetize animals with ketamine and xylazine (intraperitoneally) and also intraperitoneally inject with 100 Ml of D-luciferin (20 mg/ml). Five minutes after injection, place animals into a cassette and using a Kodak In-Vivo Imaging System F, image for 3 min (see Note 16). Process and analyze images using Kodak imaging software (Carestream Health). 6. Study hepatotoxicity of Ad vectors in mice. Wild-type Ad5 displays abortive replication in the mouse liver after intravenous injection of doses close to 3 × 109 PFU (3 × 1010 vp) per mouse accompanied by high levels of liver toxicity (26). C57BL/6 mice can be used to assess hepatotoxicity of Ad vectors due to transduction of and replication in hepatocytes. Inject C57BL/6 mice intravenously via the tail vein with 1010 to 1011 vp of Ad. On day 3 after injection, collect serum samples and measure alanine aminotransferase (ALT) levels with an ALT kit (BioTron Diagnostics, Hemet, CA) as described by the manufacturer (see Note 17).

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4. Notes 1. It is important to use fresh phenol. Use of oxidized phenol will result in viral DNA damage and reduction of recombinant virus rescue efﬁcacy. 2. It is important at this stage to avoid complete drying of the pellet as this can result in poor solubility of viral DNA. 3. Qiagen maxi-prep kit can also be used with the technique modiﬁed for large plasmids; however, plasmids prepared by double banding in CsCl give higher numbers of plaques upon transfection. 4. Lipofectamine 2000 (Invitrogen) can also be used for transfection. 5. Expression cassette inserted into HpaI site in pXC-1 does not need to carry a polyadenylation signal because there is endogenous E1A polyadenylation signal downstream of the insert. 6. Use of agarose overlay may still be advantageous as two-plasmid cotransfection system does produce deletion/rearrangement mutants with low frequency. 7. Vectors with various modiﬁcations of genome and/or capsids vary in their ability to form plaques; therefore, in many cases it is preferable to determine vector titer as OD260 units (virus particles). One OD260 unit equals 1.1 × 1012 virus particles. 8. Serum should not be heat-inactivated if A549 cells are used in plaque assay. Heat inactivation of serum results in reduced viability of A549 cells under overlay. 9. Adsorption of virus in reduced volume of medium is crucial for uniform infection and obtaining high yields. 10. Alternatively, virus can be extracted with 0.1% deoxycholate, by freezing-thawing cells four times, or by ultrasound. 11. Plaque-forming unit titers determined by limiting dilution assay are 2–3 times higher than titers determined by plaque assay. 12. OD260 assay can also be used for determining concentration of viral particles PEGylated with 20-kDa PEG; however, with 5-kDa PEG, OD260 assay results are affected by PEGylation and should not be used (9). 13. Prolonged incubation in trypsin results in reduced cell viability and increases variation in tumor size and time of appearance. 14. Tumors are established when blood vessels are formed providing abundant blood supply. At that time, Hep3B tumors become very aggressive, doubling in size every 3–7 days. If left untreated, they cross the threshold of 2,000 mm3 in 7–14 days.

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15. Make sure that warfarin in peanut oil stays at the site of injection and does not leak out. This will result in variation in efﬁcacy of warfarinization. 16. Bioluminescence signal weakens with time. To make comparisons between groups imaged separately and on different days, it is important to keep consistent time between injection of luciferin and start of imaging. 17. Depending on cancer-speciﬁc features of the vector and the dose, mice could be found dead or moribund on day 3. High levels of AST and ALT can be observed on day 2 after injection in such high toxicity/dose combinations while all animals are still alive.

Acknowledgment This work was supported by R01CA141439 to D.M. Shayakhmetov. References 1. Graham, F. L., Smiley, J., Russell, W. C., and Nairn, R. (1977) Characteristics of a human cell line transformed by DNA from human adenovirus type 5. J Gen Virol. 36, 59–74. 2. Huebner, R. J., Rowe, W. P., Schatten, W. E., Smith, R. R., and Thomas, L. B. (1956) Studies on the use of viruses in the treatment of carcinoma of the cervix. Cancer. 9, 1211–1218. 3. Liu, T. C., Galanis, E., and Kirn, D. (2007) Clinical trial results with oncolytic virotherapy: a century of promise, a decade of progress. Nat Clin Pract Oncol. 4, 101–117. 4. Yamamoto, M. and Curiel, D. T. (2010) Current issues and future directions of oncolytic adenoviruses. Mol Ther. 18, 243–250. 5. Doronin, K., Toth, K., Kuppuswamy, M., Ward, P., Tollefson, A. E., and Wold, W. S. (2000) Tumor-speciﬁc, replication-competent adenovirus vectors overexpressing the adenovirus death protein. J Virol. 74, 6147–6155. 6. Doronin, K., Kuppuswamy, M., Toth, K., Tollefson, A. E., Krajcsi, P., Krougliak, V., et al. (2001) Tissue-speciﬁc, tumor-selective, replication-competent adenovirus vector for cancer gene therapy. J Virol. 75, 3314–3324. 7. Shashkova, E. V., Kuppuswamy, M. N., Wold, W. S., and Doronin, K. (2008) Anticancer activity of oncolytic adenovirus vector armed with IFN-alpha and ADP is enhanced by pharmacologically controlled expression of TRAIL. Cancer Gene Ther. 15, 61–72.

8. Shayakhmetov, D. M., Papayannopoulou, T., Stamatoyannopoulos, G., and Lieber, A. (2000) Efﬁcient gene transfer into human CD34(+) cells by a retargeted adenovirus vector. J Virol. 74, 2567–2583. 9. Doronin, K., Shashkova, E. V., May, S. M., Hofherr, S. E., and Barry, M. A. (2009) Chemical modiﬁcation with high molecular weight polyethylene glycol reduces transduction of hepatocytes and increases efﬁcacy of intravenously delivered oncolytic adenovirus. Hum Gene Ther. 20, 975–988. 10. Shashkova, E. V., Spencer, J. F., Wold, W. S., and Doronin, K. (2007) Targeting interferonalpha increases antitumor efﬁcacy and reduces hepatotoxicity of E1A-mutated spreadenhanced oncolytic adenovirus. Mol Ther. 15, 598–607. 11. Sauthoff, H., Pipiya, T., Heitner, S., Chen, S., Bleck, B., Reibman, J., et al. (2004) Impact of E1a modiﬁcations on tumor-selective adenoviral replication and toxicity. Mol Ther. 10, 749–757. 12. Krougliak, V. and Graham, F. L. (1995) Development of cell lines capable of complementing E1, E4, and protein IX defective adenovirus type 5 mutants. Hum Gene Ther. 6, 1575–1586. 13. Chen, C. and Okayama, H. (1987) Highefﬁciency transformation of mammalian cells by plasmid DNA. Mol Cell Biol. 7, 2745–2752.

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14. Kuppuswamy, M., Spencer, J. F., Doronin, K., Tollefson, A. E., Wold, W. S., and Toth, K. (2005) Oncolytic adenovirus that overproduces ADP and replicates selectively in tumors due to hTERT promoter-regulated E4 gene expression. Gene Ther. 12, 1608–1617. 15. Bett, A. J., Haddara, W., Prevec, L., and Graham, F. L. (1994) An efﬁcient and ﬂexible system for construction of adenovirus vectors with insertions or deletions in early regions 1 and 3. Proc Natl Acad Sci USA. 91, 8802–8806. 16. Gaggar, A., Shayakhmetov, D. M., and Lieber, A. (2003) CD46 is a cellular receptor for group B adenoviruses. Nat Med. 9, 1408–1412. 17. Sova, P., Ren, X. W., Ni, S., Bernt, K. M., Mi, J., Kiviat, N., et al. (2004) A tumor-targeted and conditionally replicating oncolytic adenovirus vector expressing TRAIL for treatment of liver metastases. Mol Ther. 9, 496–509. 18. Mizuguchi, H. and Kay, M. A. (1998) Efﬁcient construction of a recombinant adenovirus vector by an improved in vitro ligation method. Hum Gene Ther. 9, 2577–2583. 19. Kalyuzhniy, O., Di Paolo, N. C., Silvestry, M., Hofherr, S. E., Barry, M. A., Stewart, P. L., et al. (2008) Adenovirus serotype 5 hexon is critical for virus infection of hepatocytes in vivo. Proc Natl Acad Sci USA. 105, 5483–5488. 20. Alba, R., Bradshaw, A. C., Parker, A. L., Bhella, D., Waddington, S. N., Nicklin, S. A.,

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et al. (2009) Identiﬁcation of coagulation factor (F)X binding sites on the adenovirus serotype 5 hexon: effect of mutagenesis on FX interactions and gene transfer. Blood. 114, 965–971. Shashkova, E. V., May, S. M., Doronin, K., and Barry, M. A. (2009) Expanded anticancer therapeutic window of hexon-modiﬁed oncolytic adenovirus. Mol Ther. 17, 2121–2130. Campos, S. K. and Barry, M. A. (2004) Rapid construction of capsid-modiﬁed adenoviral vectors through bacteriophage lambda Red recombination. Hum Gene Ther. 15, 1125–1130. Graham, F. L. (1987) Growth of 293 cells in suspension culture. J Gen Virol. 68 (Pt 3), 937–940. O’Riordan, C. R., Lachapelle, A., Delgado, C., Parkes, V., Wadsworth, S. C., Smith, A. E., et al. (1999) PEGylation of adenovirus with retention of infectivity and protection from neutralizing antibody in vitro and in vivo. Hum Gene Ther. 10, 1349–1358. Shashkova, E. V., Doronin, K., Senac, J. S., and Barry, M. A. (2008) Macrophage depletion combined with anticoagulant therapy increases therapeutic window of systemic treatment with oncolytic adenovirus. Cancer Res. 68, 5896–5904. Duncan, S. J., Gordon, F. C., Gregory, D. W., McPhie, J. L., Postlethwaite, R., White, R., et al. (1978) Infection of mouse liver by human adenovirus type 5. J Gen Virol. 40, 45–61.

Chapter 4 Syrian Hamster Tumor Model to Study Oncolytic Ad5-Based Vectors Debanjan Dhar, Karoly Toth, and William S.M. Wold Abstract Oncolytic (replicating) adenovirus (Ad) vectors are emerging as a promising form of a cancer therapy agent. There has been a need for an appropriate animal model to study oncolytic Ad since human Ad replication is usually species speciﬁc. We have shown that Syrian (golden) hamsters are an appropriate animal model to study human Ad5-based vectors. Syrian hamsters are immunocompetent, and they allow human Ad5 replication in normal tissues as well as in Syrian hamster cancer cells. The development of the Syrian hamster as a model to study oncolytic Ad vectors has opened avenues to explore the role of host immune response and preexisting immunity in Ad vector efﬁcacy and toxicity/biodistribution following Ad vector administration. Since most of the normal tissues in the Syrian hamster are permissive for human Ad5 replication, Ad vectors can be studied in the context of orthotopic cancer model developed in Syrian hamsters. Key words: Oncolytic adenovirus vector, Syrian hamster model, Immunosuppression, Neutralization, Cancer gene therapy, Intraperitoneal tumors

1. Introduction Oncolytic adenovirus (Ad) vectors are under study as cancer therapy agents. A major obstacle in the oncolytic Ad ﬁeld is the lack of a proper animal model to study these vectors. Because of the species speciﬁcity of Ads, such vectors are commonly evaluated in immunodeﬁcient mice bearing human tumor xenografts. However, since these mice are immunodeﬁcient and nonpermissive for human Ads, this model cannot adequately address the effect of the host immune system on the vector-infected tumor or the toxicity and biodistribution of the vector in normal tissues. We have developed a novel Syrian hamster model for the study of oncolytic Ad serotype 5 (Ad5)-based vectors because the hamsters are both immunocompetent and permissive David H. Kirn et al. (eds.), Oncolytic Viruses: Methods and Protocols, Methods in Molecular Biology, vol. 797, DOI 10.1007/978-1-61779-340-0_4, © Springer Science+Business Media, LLC 2012

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for human Ad5 (1). We have further used this model to study the effects of preexisting anti-Ad5 immunity (which is prevalent in the human population) on intratumoral vector efﬁcacy (2) and the role of preexisting immunity and passive immunity in vector-associated toxicity (3). Additionally, we compared the hamster and the mouse models to determine the toxicity and biodistribution of Ad vectors following an intravenous (iv) injection of the vector (4, 5). We have also used Syrian hamsters to develop a pancreatic cancer model to study oncolytic Ads (6) and to study the effect of immunosuppression on oncolytic Ad vector antitumor efﬁcacy (7). Since Syrian hamsters are permissive for Ad5 replication, they provided us with a very promising model to study anti-Ad drugs (8). Apart from our group, the Syrian hamster is also being used by other groups to study various Ad vectors (9–12). This chapter describes how to immunosuppress the hamster, generate preexisting immunity against Ad, and how to determine the anti-Ad5 neutralizing antibody (NAb) titers from various tissues. Additionally, we describe methods for establishing cell lines from Syrian hamster tumors, passaging transplantable tumors, ways to generate disseminated pancreatic tumors, and how to oral gavage a hamster. The handling, maintenance, and some basic techniques with Syrian hamsters have been described previously (13). We have used the Syrian hamster model to work primarily with Ad5-based vectors. We presume that the model would be useful for other Ad serotypes in Species C (Ad1, 2, 5, 6) because these serotypes are highly related to Ad5. In our experience, the Syrian hamster is not permissive, or is minimally permissive for human Ads that are not in Species C.

2. Materials 1. Syrian (golden) hamsters (Mesocricetus auratus), 4- to 5-weekold females (Harlan Sprague Dawley, Indianapolis, IN). 2. Dulbecco’s modiﬁed Eagle’s medium (DMEM). 3. Fetal bovine serum (FBS). 4. Trypsin–ethylenediaminetetraacetic acid (EDTA). 5. Virus: Puriﬁed and titered Ad5. 6. Human lung carcinoma cell line A549 [American Type Culture Collection (ATCC)]. 7. Cyclophosphamide (CP) (Sigma-Aldrich).

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8. Phosphate-buffered saline (PBS). 9. 0.9% sodium chloride (NaCl) solution. 10. Virus dilution buffer: 10 mM Tris–HCl, pH 8.2 and10% glycerol. 11. Water bath, 56°C. 12. Antibiotic Baytril (Bayer HealthCare). Make 100 mg/l stock in sterile water. 13. TEK-fresh rodent bedding (Harlan Sprague Dawley). 14. Teklad 2018/2918 rodent food (Harlan Sprague Dawley). 15. Sterile microtubes with screw caps, 1.5 ml (Sarstedt). 16. Blood collection tube (BD Microtainer with EDTA; Becton Dickinson). 17. Serum separator tube (BD Microtainer with serum separator; Becton Dickinson). 18. TissueLyser homogenizer (Qiagen). 19. Tungsten carbide beads, 3 mm (Qiagen). 20. 25-G and 28-G needles (Becton Dickinson). 21. 1-ml and 3-ml tuberculin syringe (Becton Dickinson). 22. 70% ethanol. 23. Multichannel pipettes (8 and 12 channels) (Gilson). 24. Sterile ﬂat bottom and round bottom 96-well tissue culture plates (TPP). 25. N95 mask. 26. Tumor implant needle 13-G × 3¼ (Cadence Science). 27. Animal feeding needle 16-G × 3 w/3-mm ball (Cadence Science). 28. Surgical glue (SkinStitch Co.). 29. 0.05%trypsin–0.53 mM EDTA (Invitrogen). 30. Panomics Cancer Cell Isolation Kit (Panomics Kit) (Panomics Inc.). This kit contains Tumor Cell Digestion Solution, Tumor Cell Suspension Solution, Tumor Cell Puriﬁcation Solution, and 100-Mm mesh-size strainers. 31. 5,000 U/ml penicillin–5,000 Mg/ml streptomycin (Invitrogen). 32. 10 nM nonessential amino acids (Invitrogen). 33. 100 mM sodium pyruvate (Invitrogen). 34. Syrian hamster pancreatic cancer cells (SHPC6 cells, generated in our laboratory and available upon request).

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3. Methods 3.1. Immunosuppression of Syrian Hamsters and Recovery from Immunosuppression

1. Syrian hamsters are immunosuppressed by administering CP. CP is dissolved in 0.9% NaCl to a ﬁnal concentration of 20 mg/ ml and then ﬁlter sterilized (see Notes 1 and 2). 2. CP is administered twice every week by intraperitoneal injection for the duration of the study using 25-G needles (see Note 3). 3. Hamsters are weighed every week to determine the CP dose. 4. The initial dose of CP is 140 mg/kg body weight, and the subsequent doses are 100 mg/kg. 5. This protocol results in signiﬁcant immunosuppression by 7–10 days post ﬁrst dose of CP (2, 7, 8). 6. Immunosuppressed hamsters are housed in sterile caging with cellulose/paper bedding (Harlan TEK-Fresh) (to prevent irritation and infection of feet) and fed irradiated chow. 7. The antibiotic Baytril is added to the drinking water at a ﬁnal concentration of 100 mg/l. 8. The level of immunosuppression is monitored by periodic analysis of complete blood cell counts. Blood is collected in anticoagulant tubes (BD microtainer with EDTA) from anesthetized animals either via the jugular vein or via retro-orbital bleeds. 9. The health of the hamsters should be monitored closely. Expect general complications from anemia, such as paleness of mucous membranes, lethargy, weight loss, decreased food, and water consumption. Hemoglobin levels should be periodically checked to prevent severe anemia. The normal level of hemoglobin in a Syrian hamster is 15.6–18.9 g/dl. In our experiments, immunosuppressed hamsters with hemoglobin levels of ~8–10 g/dl survived well and were able to recover from immunosuppression. 10. CP dosing is stopped to allow the hamsters to recover from immunosuppression. Depending on the length and extent of immunosuppression, it takes around 14–20 days for the white blood cells (WBC) and red blood cells (RBC) to recover to normal levels.

3.2. Intramuscular Immunization of Syrian Hamsters with Adenovirus

1. Syrian hamsters are immunized with a single dose of Ad5 [1 × 1010 plaque forming units (pfu)/hamster or 2 × 1011 virus particles (vp)/hamster] (14, 15). 2. Puriﬁed and titered Ad5 stocks are diluted appropriately in buffer containing 10 mM Tris–HCl, pH 8.2 and 10% glycerol. 3. With a 28-G needle, the virus is injected into the muscle of the hind thigh. Swab the area with 70% ethanol before injecting the virus.

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4. The amount of NAb generated by the immunization is measured by a neutralization assay (Subheading 3.4) of the serum collected from the hamsters. 5. A strong immune response is detected within 2 weeks after immunization (2). 6. For boosting the immune response, the same dose of Ad5 is injected intramuscularly at 14 or 15 days postimmunization. 3.3. Passive Immunization of Syrian Hamsters Against Ad5

1. Hamsters in a small group are immunized and boosted with Ad5 as discussed in Subheading 3.2. The number of hamsters that need to be immunized depends on the number of hamsters that need to be passively immunized and the NAb titer of the immune serum (see Note 4). 2. Serum is collected 14 days after the boost and pooled. The NAb titer of the pooled serum is determined as discussed in Subheading 3.4. This serum is used for passive immunization. 3. The hamsters that need to be passively immunized are immunosuppressed at least 7–10 days prior to passive transfer of immune serum. 4. The immunosuppressed hamsters are passively immunized by injecting the immune serum through an intraperitoneal injection. 5. The amount of serum transferred depends on the NAb titer of the pooled serum and the amount of NAb that needs to be achieved in the recipient hamster. In our experience, 0.4 ml of pooled serum with a NAb titer of 1:10240 when transferred into an immunosuppressed hamster weighing ~100 g generates an NAb titer of ~1:320 at 1 day after injection of the immune serum. The NAb titer is determined using the assay described in Subheading 3.4. 6. Passive immunization may be repeated to maintain or increase the NAb level in the immunosuppressed hamsters.

3.4. Determining NAb Titers in Serum, Ascites, and Tumors

1. To obtain serum, blood is collected in serum separator tubes (BD Microtainer with serum separator). The blood samples are allowed to stay at room temperature for ~30 min, and then centrifuged for 10 min in a swing bucket centrifuge or 15 min in a ﬁxed angle centrifuge at 1,000–1,300 RCF. The serum is separated in the top layer and is collected in a fresh tube. The serum can be stored at −20°C. 2. Ascites samples are centrifuged for 10–15 min at 1,000– 1,300 RCF to pellet the cell debris. If the ascites sample contains blood, then a serum separator tube may be used as mentioned above. 3. Tumors are collected from the hamsters at sacriﬁce and chopped into small pieces (as small as possible) using a razor blade.

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The tumor pieces are collected in 1.5-ml sterile screw cap tubes. Multiple tubes are used if the tumor is large. Usually for one tube, 1–2 g tumor is enough leaving some space for adding buffer. The tumors are weighed and 300-Ml sterile PBS is added into each tube. The tumors are homogenized with a single tungsten carbide bead, using a Qiagen TissueLyser instrument. Tumor homogenates are freeze-thawed three times, sonicated for 7 min, and centrifuged. The supernatant is collected and used for determining NAb titers. 4. One day before performing the assay, A549 cells are plated in 96-well plates at 8 × 105 cells per plate in a volume of 100 Ml per well. 5. Serum, ascites, and tumor samples are incubated at 56°C for 30 min to inactivate complement. 6. Two samples are assayed per plate. Using four replicate wells, each sample is diluted two fold across a round-bottomed 96-well plate in medium containing 20% FBS to normalize the total serum concentration across the plate. The ﬁrst row contains 90-Ml media and 10-Ml test sample (making it 1:10 diluted). From the ﬁrst row, 50-Ml sample is added into the next row containing 50-Ml media (making the second row 1:20 diluted). Similarly, the samples are diluted two fold across the plate except the last row which contains no sample in order to observe the effect of virus only. 7. Dilutions of test samples are incubated with 100 pfu/well of Ad5 (50-Ml volume) for 1 h at 37°C. This makes the test sample dilution in the ﬁrst row to be 1:40. After 1 h of incubation, the sample-virus mixtures (100-Ml total volume) are transferred to the 96-well plate containing A549 cells. 8. The cells are incubated for 1 h at 37°C. After incubation, the medium is removed and replaced with fresh DMEM containing 5% FBS. 9. Wells are individually scored (positive or negative) for cytopathic effect (CPE) at 7–10 days postinfection. 10. NAb titers are determined by the highest dilution of serum that results in at least 50% inhibition of CPE (two of four wells positive for CPE). 3.5. Passaging Transplantable Tumors

1. Surgically remove the tumor under aseptic conditions. Alternatively, if the experimental design or the condition of the animal warrants it, the animal can be sacriﬁced. The tumor mass should be removed from the dead animal as soon as possible. 2. In a biosafety cabinet, place the tumor tissue into a tissue culture dish containing DME medium (without serum). Remove nontumor tissue (connective tissue, fat, etc.) and necrotic tumor tissue using sterile scalpels and forceps.

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3. Cut the viable tumor mass into approximately 1 mm × 3 mm pieces and wash them in DME medium. 4. The tumor fragments that are not needed for immediate implantation can be stored frozen in DME, 10% DMSO at 80°C. Use the same methods and equipment for freezing the tumor fragments as for freezing cell lines. 5. Implant the tumors subcutaneously into recipient hamsters. Clip the hair of the animals at the site of implantation (usually, the hind ﬂanks or the interscapular area) and disinfect it with 70% ethanol. Remove the stylet of the implant needle and load a single tumor fragment into the needle. Insert the needle subcutaneously, then place the stylet into the bore of the needle and push it all the way in. Remove the needle together with the stylet, and close the wound with surgical glue. 3.6. Establishing Cell Lines from Syrian Hamster Tumors

1. Remove the tumors under aseptic conditions when they reach approximately 2-ml size (~16-mm diameter). Prepare tumor fragments as described in Subheading 3.5, steps 2 and 3. 2. Resuspend the fragments in 20 ml of DME (use a 50-ml conical centrifuge tube) and centrifuge at 300 × g for 10 min. Discard the supernatant. 3. Prepare a single-cell suspension of the tumor tissue as follows. Resuspend the pellet from step 2 in 10-ml Tumor Cell Digestion Solution [from the Panomics Cancer Cell Isolation Kit (Panomics kit)] and incubate at 37°C for 3 h with gentle agitation. 4. Add 10 ml of Tumor Cell Suspension Solution (from the Panomics kit) to the cells, and pass it through a 100-Mm mesh strainer (provided in the Panomics kit). Collect the ﬂowthrough in a new 50-ml conical centrifuge tube. 5. Centrifuge the cell suspension from step 4 at 300 × g for 10 min. Discard the supernatant. Resuspend the pellet in 20 ml of Tumor Cell Suspension Solution. 6. Add 20-ml Tumor Cell Puriﬁcation Solution (from the Panomics kit) to a 50-ml conical centrifuge tube and centrifuge at 300 × g for 2 min. This creates a gradient in the Cell Puriﬁcation Solution. Gently overlay the cell suspension from step 5 on top of the Tumor Cell Puriﬁcation Solution. Pipette the cell suspension on the wall of the tube to avoid disturbing the interphase. 7. Place the tube in a stand and do not disturb it for 6 min. Tumor cell aggregates will settle to the bottom of the tube while debris and RBC remain in suspension. 8. After 6 min, gently insert a 10-ml pipette all the way down to the bottom of the tube with the cell suspension, and remove 6 ml of liquid. Pipette this 6 ml into a fresh 15-ml conical centrifuge tube.

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9. Wait 6 min and repeat step 8. This second 6 ml of medium will have some contaminating debris, but it still contains a substantial amount of tumor cells. This will serve as a backup culture in case something happens to the cells collected in step 8. 10. Centrifuge the two tubes from steps 8 and 9 at 300 × g for 10 min. Resuspend the pellets in 5 ml DME with 10% FBS, sodium pyruvate, nonessential amino acids, 50 U/ml penicillin, and 50 Mg/ml streptomycin. Seed the cells from both tubes into two 60-mm dishes. 11. Next day, change the medium, using the same composition as in step 10. After this, expand the cells. Certain slow-growing cells will require the use of “conditioned medium.” For these cells, save ca. half of the used medium at each passage, and add it back to the freshly split cells. 12. At this point, the cell pool might be heterogeneous. Make multiple freezings of this cell pool. 13. Isolate individual clones by the limiting-dilution method. Seed ~30 cells (0.3 cells/well) of a single-cell suspension of the pool into a 96-well plate. Monolayers growing in any well should be arisen from a single cell. 3.7. Induction of Intraperitoneal Disseminated Pancreatic Tumors

1. Culture SHPC6 hamster pancreatic cancer cells in DME with 10% FBS, 1 mM sodium pyruvate, nonessential amino acids, 50 U/ml penicillin, and 50 Mg/ml streptomycin (see Note 5). 2. Harvest the cells by washing them off the plastic surface with the medium in the tissue culture dish [SHPC6 cells adhere to the surface very loosely (see Note 6)]. Alternatively, collect the medium in the tissue culture dish and add 2 ml of 0.05% trypsin, 0.53 mM EDTA to the cells that adhere to the surface. After 5 min at 37°C, add back the medium that was in the dish and resuspend the cells by trituration. Centrifuge the cell suspension at 300 × g for 10 min. Resuspend the pellet in serumfree DME, and centrifuge again at 300 × g for 10 min. Resuspend the pellet in serum-free DME at 2 × 107 cells/ml. 3. Disinfect the skin of the hamster at the lower right quadrant of the abdomen by wiping it with 70% ethanol, and inject 500 Ml of cell suspension (1 × 107 cells) into the peritoneal cavity using a 25-G needle attached to a 3-cc syringe. Tumor nodules of ~1-mm diameter will form in the peritoneal cavity by 4 days after injection, and the animals will develop serosanguinous ascites (indicated by a distended abdomen) by 8 days after injection.

3.8. Oral Gavage with Syrian Hamsters

1. Bend the feeding needle at its halfway point at about 20° angle. This will help lead the needle into the esophagus. 2. Load the substance to be injected into the syringe (see Note 7). Wipe off any excess drug from the outside of the needle; hamsters

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have a sense of taste and will struggle excessively if the substrate is foul tasting. 3. Scruff the animal with your left hand (if you are right-handed) and hold it so that your thumb is at the corner of the hamster’s mouth and your index ﬁnger is against the neck/head of the hamster. Keep the animal’s nose pointing upward. This step is critical; once the animal is held correctly, the insertion of the needle is very simple. 4. Hold the syringe so that the needle is parallel with the hamster’s body, and that it curves away from the animal when pointing downward. 5. Insert the needle into the right corner of the hamster’s mouth and gently slide it downward. Avoid getting it trapped under the tongue. When you reach the pharynx (approximately 2 cm from the mouth), the animal will swallow it. The needle should slide down to about three-quarters of its length without any resistance. If any resistance is encountered, the needle should be withdrawn, the animal should be put down, and the procedure should be repeated with the holding of the hamster adjusted. 6. Inject the drug by ﬁrmly holding the syringe between your thumb and middle ﬁngers and pushing on the plunger with your index ﬁnger. Be sure that you are not pressing the needle against the stomach wall of the hamster. Pull out the needle only after you have injected all the liquid. 7. Observe the animal that it does not have dispnea (a sign of injection into the trachea) and that it does not vomit back the inoculum. Hamsters are generally easy to gavage; in our extensive experience, we never had an animal that was misinjected.

4. Notes 1. CP is a carcinogen and should be handled very carefully. Gloves, gowns, and N95 masks are recommended. 2. CP solutions are stored at 4°C in the dark. CP loses its potency if stored for more than 2 weeks at 4°C. 3. Before injecting CP, the abdomen should be wiped with 70% ethanol to prevent any infection. 4. A maximum of about 2–3 ml serum can be collected from a hamster that weighs ~120 g at sacriﬁce. 5. For induction of intraperitoneal disseminated pancreatic tumors, SHPC6 cells from one 90% conﬂuent 150-mm dish are needed for every two animals.

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6. When passaging SHPC6 cells, take care to collect all cells from the dish; otherwise, in the long run you will select for the more adherent or the more anchorage-independent cell population. 7. The needle hub will hold approximately 200 Ml; allow for this when preparing.

Acknowledgments We thank Jacqueline Spencer, Maria Thomas, and other members of our laboratory for outstanding research and discussions. This research was supported by grant R01 CA118022 from the National Institutes of Health. References 1. Thomas, M. A., Spencer, J. F., La Regina, M. C., Dhar, D., Tollefson, A. E., Toth, K., et al. (2006) Syrian hamster as a permissive immunocompetent animal model for the study of oncolytic adenovirus vectors. Cancer Res. 66, 1270–6 2. Dhar, D., Spencer, J. F., Toth, K., and Wold, W. S. (2009) Effect of preexisting immunity on oncolytic adenovirus vector INGN 007 antitumor efﬁcacy in immunocompetent and immunosuppressed Syrian hamsters. J. Virol. 83, 2130–9 3. Dhar, D., Spencer, J. F., Toth, K., and Wold, W. S. (2009) Pre-existing Immunity and Passive Immunity to Adenovirus 5 Prevents Toxicity Caused by an Oncolytic Adenovirus Vector in the Syrian Hamster Model. Mol. Ther. 17, 1724–32 4. Lichtenstein, D. L., Spencer, J. F., Doronin, K., Patra, D., Meyer, J. M., Shashkova, E. V., et al. (2009) An acute toxicology study with INGN 007, an oncolytic adenovirus vector, in mice and permissive Syrian hamsters; comparisons with wild-type Ad5 and a replication-defective adenovirus vector. Cancer Gene Ther. 16, 644–54 5. Ying, B., Toth, K., Spencer, J. F., Meyer, J., Tollefson, A. E., Patra, D., et al. (2009) INGN 007, an oncolytic adenovirus vector, replicates in Syrian hamsters but not mice: comparison of biodistribution studies. Cancer Gene Ther. 16, 625–37 6. Spencer, J. F., Sagartz, J. E., Wold, W. S. M., and Toth, K. (2009) New pancreatic carcinoma model for studying oncolytic adenoviruses in the permissive Syrian hamster. Cancer Gene Ther. 16, 912–22

7. Thomas, M. A., Spencer, J. F., Toth, K., Sagartz, J. E., Phillips, N. J., and Wold, W. S. (2008) Immunosuppression enhances oncolytic adenovirus replication and antitumor efﬁcacy in the Syrian hamster model. Mol. Ther. 16, 1665–73 8. Toth, K., Spencer, J. F., Dhar, D., Sagartz, J. E., Buller, R. M., Painter, G. R., et al. (2008) Hexadecyloxypropyl-cidofovir, CMX001, prevents adenovirus-induced mortality in a permissive, immunosuppressed animal model. Proc. Natl. Acad. Sci.USA 105, 7293–7 9. Bortolanza, S., Alzuguren, P., Bunuales, M., Qian, C., Prieto, J., and Hernandez-Alcoceba, R. (2007) Human adenovirus replicates in immunocompetent models of pancreatic cancer in Syrian hamsters. Hum. Gene Ther. 18, 681–90 10. Bortolanza, S., Bunuales, M., Otano, I., GonzalezAseguinolaza, G., Ortiz-de-Solorzano, C., Perez, D., et al. (2009) Treatment of pancreatic cancer with an oncolytic adenovirus expressing interleukin-12 in Syrian hamsters. Mol. Ther. 17, 614–22 11. Bortolanza, S., Bunuales, M., Alzuguren, P., Lamas, O., Aldabe, R., Prieto, J., et al. (2009) Deletion of the E3-6.7K/gp19K region reduces the persistence of wild-type adenovirus in a permissive tumor model in Syrian hamsters. Cancer Gene Ther. 16, 703–12 12. Matthews, K., Noker, P. E., Tian, B., Grimes, S. D., Fulton, R., Schweikart, K., et al. (2009) Identifying the safety proﬁle of Ad5.SSTR/ TK.RGD, a novel infectivity-enhanced bicistronic adenovirus, in anticipation of a phase I clinical trial in patients with recurrent ovarian cancer. Clin. Cancer Res. 15, 4131–7

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13. Thomas, M. A., Spencer, J. F., and Wold, W. S. (2007) Use of the Syrian hamster as an animal model for oncolytic adenovirus vectors. Methods Mol. Med. 130, 169–83 14. Tollefson, A. E., Kuppuswamy, M., Shashkova, E. V., Doronin, K., and Wold, W. S. (2007)

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Preparation and titration of CsCl-banded adenovirus stocks. Methods Mol. Med. 130, 223–35 15. Tollefson, A. E., Hermiston, T. W., and Wold, W. S. M. (1998) Preparation and titration of CsCl-banded adenovirus stocks. 21, 1–9

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Chapter 5 Adenoviral Gene Expression and Replication in Human Tumor Explant Models Gunnel Halldén Abstract Promising results have been reported from numerous studies with replication-selective oncolytic adenoviral mutants as novel treatments for a variety of cancers. Most of these studies were performed in cancer cell lines, dissociated tumor tissue, or animal models, and the predictive utility for efﬁcacy and safety in the clinical setting is unclear. Indeed, the outcome of many clinical trials with viral mutants that demonstrated high efﬁcacy preclinically has so far been disappointing, necessitating better test models. To this end, we developed a methodology using primary human cancer specimens for evaluation of cytotoxicity ex vivo including colorectal liver metastasis, ovarian, breast, colon, and prostate carcinomas. Under optimized culture conditions, primary human tumor tissue remained viable for up to 48 h, enabling evaluation of viral mutants in tissue with intact morphology. This assay may have great utility to investigate novel viral mutants and to identify treatment sensitive cancers by assessing speciﬁc oncolytic mutants in individual cases. Key words: Ex vivo, Replication-selective, Preclinical model, Viral localization, Human tumor specimens, Immunohistochemistry

1. Introduction Since the ﬁrst clinical evaluation of the oncolytic adenoviral mutant dl1520 (Onyx-015), several novel mutants have been developed for clinical testing by targeting other cellular pathways involved in tumor progression (1, 2). Currently trials with '24 mutants are underway targeting recurrent ovarian cancer and malignant gliomas (http://www.clinicaltrials.gov). Numerous mutants are also being developed preclinically to improve efﬁcacy (3–6).

David H. Kirn et al. (eds.), Oncolytic Viruses: Methods and Protocols, Methods in Molecular Biology, vol. 797, DOI 10.1007/978-1-61779-340-0_5, © Springer Science+Business Media, LLC 2012

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Even though many novel adenoviral mutants have shown promise as future oncolytic agents in preclinical studies clinical results have so far been disappointing. One reason for the differences in observed and anticipated efﬁcacy is that cancer cell lines in culture might not reﬂect the biology of tumors in vivo. Nor does dissociated epithelial tumor tissue reﬂect the in situ situation in regard to viral infection and replication due to loss of cell polarity, changes in protein expression patterns, and loss of tissue structure. Other reasons are the heterogeneity of human tumors and successful treatment in one case might not be efﬁcacious in other patients. Additional difﬁculties are the limited availability of human adenovirus-permissive immunocompetent animal models because of the strict species speciﬁcity for adenoviral replication. Normal murine cells do not support human adenoviral replication and the only semipermissive species are the Syrian hamster and the Cotton rat (7–10). To date, the majority of in vivo studies are primarily performed in nude mice with human tumor xenografts derived from cancer cell lines making the applicability of preclinical results to the clinical situation unclear. We have previously developed murine tumor models to evaluate the contribution of the host immune response to antitumor efﬁcacy and cytotoxicity in the presence of immune factors (11). However, these models are also limited in regard to predicting toxicity and viral clearance in patients. This highlights the need to identify the optimal target cancers for various oncolytic agents and the limitations with currently available preclinical models. Several groups have developed alternative test models for evaluation of oncolytic viruses in human tumor specimens prior to clinical evaluation. In one study, a perfusion system for liver wedges was described evaluating replication of a mutant Herpes simplex virus (HSV) ex vivo (12). Although, replication was demonstrated after 48 h rather large tumor specimens were required. Another study used smaller liver tumor samples that were sectioned into 250-Pm slices, demonstrating good tissue viability and adenoviral gene expression up to 36 h ex vivo (13). Tissue from patients with Barrett’s esophagus was also viable and infectable when kept in culture ex vivo for up to 24 h (14). We have recently optimized a culture system using very small samples of intact human adenocarcinoma tissue for evaluation of viral infectivity and replication in individual patient cases (15). Dependent on the speciﬁc tissue, samples were viable up to 48 h, early and late viral gene expression was detectable from 17 to 48 h (Figs. 1–3) and expression of cellular receptors such as Coxsackievirus adenovirus receptor (CAR) and DvE3- and DvE5-integrin was demonstrated. This ex vivo method could serve as relevant transitional model from cell lines and murine studies prior to clinical applications for novel oncolytic mutants.

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Fig. 1. Viability of human ovarian serous adenocarcinoma tissue cultured over time ex vivo. Tissue specimens were cut into blocks (<1 mm3) and incubated under the conditions described in text. After 20 and 40 h, samples were ﬁxed in PFA, parafﬁn-embedded, and either stained with H and E (upper panels) to monitor tissue structure or with the anti-PCNA antibody (lower panels) for determination of proliferating cells. Average ratio of PCNA positive to total epithelial cell population in this specimen was 12–23% with lower intensity staining in the 40-h samples original magniﬁcation r 400.

2. Materials 2.1. Tissue Collection

1. Fresh tumor samples from surgery or biopsy sampling (see Note 1). 2. Iscove’s modiﬁed Dulbecco’s medium (IMDM) and fetal calf serum (FCS) (Gibco/Invitrogen). 3. ViaSpan tissue preservation ﬂuid (University of Wisconsin (UW) solution; Bristol-Myers Squibb). 4. Penicillin–streptomycin (10,000 U/ml; Sigma-Aldrich).

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Fig. 2. Human lung carcinoma cells A549 were inoculated subcutaneously in C57Bl6 nu/nu mice and grown as untreated xenografts for 25 days. The tumors were surgically removed and immediately processed for ex vivo culture by cutting thin sections <1 mm. Tissue sections were incubated in 5% FCS-IMDM and infected with the replication-selective Ad'E1ACR2 at 1 × 107 vp/2–4 sections in 200-Pl media. Infection media was removed after 2 h and 10%FCS in IMDM added. The tissue was ﬁxed in 4% PFA and processed for parafﬁn embedding 36 h later. Antibody staining and detection for E1A (a, b) and hexon (c) as described in Subheading 3.5. Original magniﬁcation ×100.

Fig. 3. Human tumor specimens processed and cultured ex vivo followed by infection with adenovirus mutants. (a–c) Ovarian serous adenocarcinoma samples were infected with adenovirus wild type virus (Ad5) (a), replication-defective dl 312 (b) and replication-selective Ad'E1ACR2 (c) at 2 × 109 vp/3 tissue blocks in 200 Pl for 4 h. Media was replaced with 10% FCS in IMDM, ﬁxed in PFA and processed for parafﬁn embedding 40 h later. Antibody staining and detection of hexon was as described in Subheading 3.5. Original magniﬁcation ×100 (a, b) and ×200 (c) inserts ×600. (d–f) Antibody detection of E1A in human tumor specimens infected with the Ad'E1ACR2 mutant at 2 × 109 vp/2–4 tissue blocks in 200 Pl for 4 h and processed after 25 h as described in text. (d) Mammary epithelial carcinoma tissue, (e) rectal carcinoma tissue, and (f) ovarian carcinoma tissue, original magniﬁcation ×400. (a–f) All tissue samples were counterstained by H/E.

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1. Sterile ﬁltered phosphate-buffered saline (PBS). 2. Ethanol 70%. 3. Sterile cell culture plastic plates, scalpels, tweezers, and scissors. 4. Tissue slicer such as the Brendel/Vitron Tissue Slicer (Vitron) with an O2/CO2 (95%/5%) gas cylinder.

2.3. Tissue Incubation

1. Sterile ﬁltered media supplements and growth factors: Transferrin, insulin, hydrocortisone, epidermal growth factor (EGF), transforming growth factors (TGF; D and/or E), estradiol, dihydrotestosterone (DHT), insulin-like growth factor (IGF), hepatocyte growth factor (HGF) (Sigma-Aldrich), and mibolerone (PerkinElmer). 2. IMDM supplemented with 5 and 10% FCS, 100 U/ml penicillin– streptomycin, 4 Pg/ml transferrin and the growth factors relevant for the speciﬁc tissue such as 10 PM insulin, 1 PM IGF, 1 PM hydrocortisone, 10 nM EGF, 10 nM TGF, 10 nM HGF, 1 PM estradiol (ovarian and breast), 100 nM DHT, or 1 nM mibolerone (prostate) (PerkinElmer).

2.4. Virus Infection

1. High-titer viruses at a stock concentration of at least 1 × 1010 viral particles/ml (vp/ml) or higher if possible, including replicating viral mutants and replication-defective control mutants (see Note 2). 2. Incubator at 37°C with 5% CO2/air for incubation up to 48 h.

2.5. Fixation of Tissue and Immunohistochemistry

1. Sterile 4% paraformaldehyde (PFA) in PBS. 2. Parafﬁn and tissue cassettes (Leica, UK). 3. Optimum cutting temperature (O.C.T.) compound and TissueTek Cryomold (Sakura Finetek). 4. Hematoxylin and Eosin (H/E) for immunohistochemistry (IHC). 5. Antibodies for detection of viral and cellular antigens for example; mouse monoclonal anti-E1A (M73; Gene Tex Inc.), rabbit polyclonal anti-E1A (Sauta Biotechnology), goat polyclonal anti-hexon (Chemicon), mouse monoclonal anti-proliferating cellular nuclear antigen (PCNA) (Dako), mouse monoclonal human speciﬁc anti-coxsackie and adenovirus receptor (CAR) (ATCC), anti-human Dv-integrin (Chemicon) for detection of DvE3 and DvE5 integrins. 6. Other IHC reagents; Dako Target Retrieval solution (Dako) for PCNA staining, relevant secondary antibodies and detection reagents, e.g., 3,3c-diaminobenzidine (DAB) (Dako).

2.6. Analysis of Viral Replication and Genome Ampliﬁcation

Viral replication by tissue culture infective dose (TCID50) (see Note 3): 1. HEK293 or A549 cells (ATCC).

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2. 96-well plates and 10% FCS in Dulbecco’s modiﬁed Eagle’s media (DMEM) (Gibco/Invitrogen). Viral genome ampliﬁcation by qPCR (see Note 4). 3. QIAmp DNA Blood Mini Kit (Qiagen). 4. Relevant primers and PCR detection reagents.

3. Methods Human tissue specimens for research purposes can only be obtained after review of each protocol by national and local committees that monitor adherence to ethical regulations. In addition to approval from the local research ethics committees including all institutions where the work is undertaken, informed consent must be given by each patient similar to the procedures for participation in clinical trials (see Note 1). Recording of the amount of tissue collected, used experimentally, disposed, or stored is also required. The specimens should be processed as soon as possible after surgical removal to minimize hypoxic damage and tissue necrosis. For most specimens, the ideal time is less than 2 h after surgery with even shorter time for larger specimens, benign and normal adherent tissue (see Note 5). The tissue should be kept at 4°C at all times during the preparation. Equipment must be sterile and all solutions prepared and precooled such as media, growth factors, and virus stocks to be ready for use often requiring a team effort. Precautions have to be taken to prevent cross-contamination of the regular cell stocks in the culture facilities since the tissue specimens are not sterile, cannot be screened for contaminants and should be handled as if containing unknown viruses, bacteria, and mycoplasma (see Note 6). Since the viability ex vivo of any human tissue is limited it is almost always necessary to perform the study without knowing the exact tumor type or the quality of the tissue. We have obtained valuable information from a variety of surgical and biopsy specimens (Figs. 1 and 3), but occasionally a sample will not be viable ex vivo, limiting the study (see Notes 7 and 8). 3.1. Collection and Preparation of Tissue Specimens

1. Immediately after surgical removal the tissue specimens (usually <1 cm3) are placed on ice in precooled 10%FCS in IMDM supplemented with 100 U/ml penicillin–streptomycin and is transported in a covered and sealed icebox to the research laboratory. 2. The specimens are transferred to a large (10–15 cm) cell culturetreated dish with 10% FCS in IMDM placed on a layer of ice. Using sterile scalpels, scissors, and tweezers gently dissect the specimen to remove and discard necrotic or connective tissue areas. Care should be taken not to compress the tissue at any time (see Note 9).

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3. Whenever possible, slice the tissue into thin slices (<1 mm); however, most often this is not possible due to tissue structure and dicing of specimens into crossed blocks or cubes of <1 mm3 is more feasible. Alternatively, a mechanical tissue slicer can be used to generate thin slices (Subheading 3.2 below). Inspect specimen carefully to identify eventual adherent normal tissue that is a valuable source of control tissue (see Note 10). 4. Rinse the processed tissue in cold PBS three times, collect homogeneous tissue sections or blocks, and transfer to 24-well plates with 0.8 ml/well of 10% FCS in IMDM with transferrin and the relevant growth factors. Ideally, one tissue block is placed in each 24-well; however, if plenty of tissue is available it is more practical to place 2–3 blocks in each well. 5. Set aside 3–4 representative tissue blocks, wash in PBS, and place in ﬁxative (Subheading 3.4). This starting material is used as references in the IHC to determine quality of tissue at the beginning of the study. Dependent on the variation in tissue morphology within the specimen, it might be necessary to use representative sections for each area (see Note 11). 6. The plates are incubated at 37°C in 5% CO2/air for 10–30 min while discarding unused tissue (biological waste) and preparing virus dilutions (Subheading 3.3). 3.2. Tissue Preparation Using a Mechanical Tissue Slicer

1. For larger tissue specimens (>2 cm3), a mechanical tissue slicer can be used to cut thin (>250 Pm) sections that can improve viability (see Note 12). 2. All parts in contact with tissue must be sterilized (autoclave or 70% ethanol). Flush the sectioning compartment and collecting tubing with the ViaSpan solution after thorough washing with sterile PBS. Keep the ViaSpan solution and collecting vials on ice at all times. Adjust the O2/CO2 (95%/5%) ﬂow rate to the maximum setting that does not create bubbles in the solution. 3. Tissue cores are cut by using the accompanying coring devices (4–8 mm diameter) on the dissected specimen on ice. 4. Set the desired slice thickness dependent on tissue and turn on power for the sectioning blade. Cut by moving the metal core evenly back and forth and collect sections in the ViaSpan solution on ice. In our studies, good-quality sections of t300 Pm could be generated for liver tumor specimens. 5. Carefully rinse and place the sections in 24-well plates (1–5 sections/well) in 10% FCS in IMDM with transferrin and relevant growth factors as described above (Subheading 3.1, steps 5–7). 6. The process is repeated to cut cores from other areas of the specimen.

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3.3. Viral Infection of Tissue Ex Vivo

1. Aspirate media from each 24-well and add the minimal volume that covers the tissue, 200–400 Pl containing the desired concentration of virus in 5% FCS in IMDM supplemented with transferrin and relevant growth factors. In our experience, when determining tissue sensitivity to various mutants a dose of 2 × 109 vp/well resulted in reproducible data (see Notes 2 and 13). 2. After 2 h the virus solution is removed and fresh 10% FCS in IMDM (0.5–0.8 ml) is added to the tissue. 3. Alternatively, if the tissue is poorly infectable or higher levels of viral uptake is required, the infection media can remain in the wells for an additional 2 h, followed by aspiration and addition of the 10% FCS-IMDM (0.5–0.8 ml). The virus solution can also be left in the wells for the entire incubation time if an equal or greater volume of 10% FCS media is added to make up a total volume of 0.5–0.8 ml (see Notes 13 and 14). 4. The infected tissue samples are incubated at 37°C in 5% CO2 and at selected time points, e.g., 6, 12, 18, 24, and 48 h, samples are removed for analysis (the following sections). 5. Change media every 24 h and inspect tissue for microorganism contamination by light microscopy.

3.4. Fixation of Tissue and IHC

3.5. General Histopathology and IHC

Wash tissue slices/blocks in prewarmed (37°C) PBS 3 × 1 ml and place in 4% PFA (PBS-buffered) in Eppendorf tubes at 4°C for 48 h (see Note 15). 1. Wash ﬁxed tissue sections in PBS and place in tissue cassettes for parafﬁn embedding or in cryomolds with the O.C.T. compound (see Note 16). 2. Section and process embedded tissue according to standard histology protocols. 3. Tissue quality and survival is evaluated by examining cytopathic effect (CPE) and general tissue and cell morphology by staining with Hematoxylin and Eosin (H/E) (see Note 17). 4. The fraction of proliferating cells in each section is used as a more quantitative indicator of tissue viability and can be estimated by staining for PCNA according to the following protocol. Parafﬁn sections are dewaxed in xylene, rehydrated in ethanol of decreasing concentrations, unmasked with Dako Target Retrieval solution, followed by blocking of nonspeciﬁc binding with normal rabbit serum for 30 min. A mouse monoclonal primary antibody against human PCNA is applied at 1:200 dilution in PBS and incubated for 1 h at 18–24, wash in PBS for 3 × 5 min, incubate with a biotin-conjugated rabbit anti-mouse antibody (Dako) for 30 min and wash 2 × 5 min in PBS. Treat sections with 0.03% hydrogen peroxide in methanol

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for 20 min, wash in PBS for 2 × 3 min, and incubate with streptavidin–HRP complex (Dako) for 30 min. Develop using the substrate 3,3c-diaminobenzidine (DAB) (Dako) as a chromogen. Monitor development of staining by microscopy and stop after 5–15 min by immersion in distilled water. Counterstain with Hematoxylin, dehydrate in increasing concentrations of ethanol, histoclear with xylene, and mount for light microscopy analysis. 5. To detect other cellular and viral proteins the above staining protocol can be followed with adjustments to optimize for each speciﬁc antigen. We use a goat polyclonal hexon-speciﬁc anti-Ad5 antibody at a dilution of 1:200 to determine viral uptake and assembly and a rabbit polyclonal anti-E1A antiserum at a dilution of 1:50 to determine viral replication and early gene expression. The mouse monoclonal human speciﬁc coxsackie and adenovirus receptor (CAR) and Dv-integrin antibodies, can be used at 1:1,000 for detection of cellular CAR and for detection of DvE3 and DvE5 integrins respectively, both in frozen sections. 6. The respective secondary antibodies are added and detection can be with DAB as described above. 3.6. Quantiﬁcation of CPE, PCNA, Hexon, and E1A Staining by IHC

1. Scoring of staining for each treatment group (each 24-well) can be performed as follows using 3–6 sections/treatment group: in an area including 1,000 cells on each slide count the number of positive cells and compare to the total number of live epithelial cells in each section determined by H/E staining. 2. To quantify virus-induced CPE- and PCNA-positive cells, the proportions of cells demonstrating CPE or staining can be expressed as percentages after averaging all slides in each treatment group. 3. For hexon and E1A staining the following criteria can be used to quantify the ratios of positively stained cells: −, no staining, +, <20% of cells positive, ++, 20–40% of cells positive, and +++, >40% positive. Statistical analysis is by t-test, two-tailed analysis of variance (Anova).

3.7. Functional Assay to Determine Viral Replication by the Limiting Dilution Method (TCID50 )

1. After the initial 2–4 h infection of the tissue samples, wash the tissue and add fresh media (d0.5 ml) to each culture well. 2. At speciﬁc time points, 12–48 h postinfection, collect tissue and media together, divide sample into two, one half for replication and the other for qPCR analysis (Subheading 3.8); freeze both aliquots at −80°C. 3. For the replication assay, freeze–thaw samples three times by rapidly immersing in 37°C water bath and in N2(l) (see Note 18).

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4. Plate 10,000 HEK293 (or A549) cells per well in 96-well plates in a total 200 Pl 10% FCS in DMEM, one to three plates/sample. 5. Add 10–20 Pl of undiluted or diluted samples (dependent on tissue type and experimental conditions) to the 12 wells in the top row using a multichannel pipette. Mix by carefully pipetting up and down and transfer to the next row and so on, leaving the last row of wells as a control without virus. 6. Incubate the infected cells at 37°C and determine CPE 7–10 days postinfection. If possible, each sample is determined in duplicates or triplicates in the TCID50 assay, and is expressed as infectious units (IU)/ml and averaged (see Note 18). Results can be presented as the ratio of each replication-competent virus to a replication defective mutant in the absence of viral replication and gene expression. An internal standard with known activity is always included to enable comparison between assays. 7. Statistical analysis is by t-test, two-tailed analysis of variance (Anova). 3.8. Determination of Viral DNA Ampliﬁcation

Total DNA is prepared from the harvested and frozen aliquots described above (Subheading 3.7, step 2) using the QIAmp DNA Blood Mini Kit according to the manufacturer’s instructions. Quantify DNA and add 20 ng of total DNA in 5 Pl to each PCR. Primers targeting any adenoviral region will work as long as they speciﬁcally amplify the respective gene. Viral DNA is analyzed at speciﬁc time points and normalized to total cellular DNA or a cellular housekeeping gene such as glyceraldehyde 3-phosphate dehydrogenase (GAPDH) (see Note 4).

4. Notes 1. Specimens must only be obtained after appropriate ethics approvals and informed consent according to speciﬁc national and local regulations. In the UK, applications need to be approved by the research institution and also by the local NHSethics committee at each hospital where the clinical procedures are performed. Annual reports to the hospital committees ensure that the application remains active. 2. It is essential to include nonreplicating viral mutants as a control for particle toxicity in absence of viral replication with or without viral gene expression. We frequently use dl312 (E1A and E3B deleted) and AdGFP (E1 replaced by CMVGFP). If undiluted stock viruses are added to the tissue, it is important to limit the volume of addition dependent on the stock-buffer composition. For example, more than 0.5% CsCl2

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or 3% glycerol is usually toxic to the tissue that is often more sensitive than cultured cell lines. 3. In our experience, the TCID50 method is preferred over the plaque assay due to its higher sensitivity and good reproducibility. 4. For detection of viral genomes, we routinely use hexon primers and SYBR Green according to standard protocols; hexonforward: 5c-GGACAGGCCTACCCTGCTAAC-3c and reverse: 5c-TGCTGTCAACTGCGGTCTTG-3c. However, any viral gene can be used for detection. Viral genomes are normalized to cellular DNA or GAPDH ampliﬁed in the same sample. If possible, the increase in viral particles over time can also be expressed as the ratio of particles at each time point relative to amount of virus taken up by cells after 3–6 h. 5. Close collaboration with the surgical team is a necessity with dedicated personnel for immediate collection of tissue in media on ice and transport to the research laboratory. Local regulations for transport of biological material must also be observed; for example, in the UK public transport is, in most cases, not allowed. 6. At our center, we use separate cell culture hoods and incubators for surgical specimens to minimize the risk of cross contamination. An important aspect of working with human material is also the health and safety of the personnel in direct contact with the tissue. Extra care to avoid direct contact, cuts by needles and sharps should be taken because of the risk of infectious diseases spreading, mainly HIV and HBV. 7. We have had good results with several tumor types except prostate cancers. While tissue viability was good transduction determined by IHC was rather poor. One possible reason is that epithelial cells that are sensitive to adenovirus infection are located in typical small gland-like structures in the prostate tumors that might require longer exposure times for viral penetration. Obtaining larger specimens to enable cutting of thin Pm-slices could potentially improve viral transduction in this tissue. 8. Since adenovirus has a relatively long life cycle, with signiﬁcant gene expression ﬁrst after 12 h, the described ex vivo model system might be more suitable for faster replicating viruses such as poxviruses and HSV. CPE, viral gene expression, and replication might be more easily detected at earlier time points with these viruses when tissue is still in good condition. 9. Avoid compression of tissue, since this will affect viability; lift rather than pinch tissue and change scalpel often to prevent pressure damage on surrounding tissue. 10. Frequently, normal adherent tissue is removed with the tumor tissue and can be found around the edges. Normal tissue has

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greatly reduced viability ex vivo but can generate valuable information. However, we have only been able to keep normal (and benign) samples in good condition for a maximum of 24 h that enables detection of viral gene expression, but only limited replication. 11. Tumor specimens are often heterogeneous and similar sections or blocks should be used for comparison. It is also important to identify and exclude connective or necrotic tissue. Owing to the limited viability ex vivo, it is usually not possible to histologically deﬁne the tumor tissue prior to the experimental set up and can only be determined retrospectively. Despite careful sampling and selection of homologous tumors, we have observed a high degree of cell heterogeneity, both in tumors derived from different patients as well as in metastatic tumors from the same patient. Consequently, the sensitivity to viral infection and the ability to support viral replication were highly variable. In some cases, the majority of epithelial cells demonstrated intracellular hexon expression while in other cases only a few viral particles could be detected intracellularly. Other tissue specimens showed an accumulation of viral particles along the cell surface with few particles entering the cells. Therefore, multiple tissue sections or blocks from various areas of the tumor will result in more accurate information. 12. To use a mechanical device such as the Brendel/Vitron slicer requires larger tissue specimens than for manual sectioning. In our laboratory, we were only able to obtain large enough tumor specimens of colorectal metastasis from liver resections. In our experience, specimens need to be t2 cm3 to enable the use of both the coring tool and successful sectioning. Although thinner sections improve viability, the slicer requires large volumes (1–2 l) of the ViaSpan solution that is relatively expensive and has a short shelf life. Very thin sections are more difﬁcult to collect and process for IHC while tissue viability should be improved. 13. For general screening purposes of various viral mutants, a higher dose of 2 × 109 vp/well enables detection of virus even after short incubation times. It is possible to perform dose– response analysis by diluting virus over a range of concentrations (e.g., 500–2 × 109 vp/well), taking into account that the lower doses might not be detectable if viability over time or infectability is poor. 14. When tissue had a high mucous content, we achieved better viral transduction if the tissue sections/blocks were pretreated with hyaluronidase 1.5 mg/ml in IMDM for 30 min prior to infection. 15. At this point, it is important to not compress the tissue; transfer from well to tube with PFA by lifting tissue rather than pinching.

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16. For the antibodies described in this protocol, parafﬁn-embedded tissue worked well for detection of PCNA, E1A, and hexon, while detection of CAR and integrins were only possible in frozen sections. In our experience, tissue morphology is superior in parafﬁn-embedded sections and should be used whenever possible. 17. If tissue viability appears to be poor (<50% of cells are intact), no further staining with speciﬁc antibodies yields meaningful results and the study should be terminated. However, it is often still possible to determine type of cancer for future reference. 18. There are many methods for calculating the infectivity using the TCID50 assay; we have set up an Excel template based on the Reed and Muench method (15,16). The following formulas are used in the template: TCID50 = 10L − d(s − 0.5), TCID50/ml = TCID50 × V where L = lowest dilution with CPE, d = log difference between dilutions, s = sum of proportions of CPE-containing wells/row, and V = infection volume in ml. Convert to infectious units by the following formula: (IU)/ ml = log TCID50 × 0.69 (16).

Acknowledgments This work was supported by grants from the Barts & The London Charity and Cancer Research UK (C633-A6253/A6251). The author would like to thank the patients and surgical teams at the Hammersmith Hospital, London and the Veterans Administration Hospital, Palo Alto for their cooperation, and Steve Thorne, Yaohe Wang, Vipul Bakta, Celina Costas, and Suresh Radhakrishnan for their contributions in developing this method. References 1. Parato, K. A., Senger, D., Forsyth, P. A., and Bell, J. C. (2005) Recent progress in the battle between oncolytic viruses and tumours, Nat Rev Cancer 5, 965–976. 2. Liu, T. C., Galanis, E., and Kirn, D. (2007) Clinical trial results with oncolytic virotherapy: a century of promise, a decade of progress, Nat Clin Pract Oncol 4, 101–117. 3. de Vrij, J., Willemsen, R. A., Lindholm, L., and Hoeben, R. C., GIANT consortium (2010) Adenovirus-Derived Vectors for Prostate Cancer Gene Therapy, Hum Gene Ther 21(7), 795–805.

4. Matthews, K., Noker, P. E., Tian, B., Grimes, S. D., Fulton, R., Schweikart, K., Harris, R., Aurigemma, R., Wang, M., Barnes, M. N., Siegal, G. P., Hemminki, A., Zinn, K., Curiel, D. T., and Alvarez, R. D. (2009) Identifying the safety proﬁle of Ad5.SSTR/TK.RGD, a novel infectivity-enhanced bicistronic adenovirus, in anticipation of a phase I clinical trial in patients with recurrent ovarian cancer, Clin Cancer Res 15, 4131–4137. 5. Freytag, S. O., Stricker, H., Peabody, J., Pegg, J., Paielli, D., Movsas, B., Barton, K. N.,

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Brown, S. L., Lu, M., and Kim, J. H. (2007) Five-year follow-up of trial of replication-competent adenovirus-mediated suicide gene therapy for treatment of prostate cancer, Mol Ther 15, 636–642. 6. Öberg, D., Yanover, E., Sweeney, K., Adam, V., Costas, C., Lemoine, N.R., and Halldén, G. (2010) Improved potency and selectivity of an oncolytic E1ACR2 and E1B19K deleted adenoviral mutant (Adśś) in prostate and pancreatic cancers. Clin Canc Res 16, 541–53. 7. Russell, W. C. (2009) Adenoviruses: update on structure and function, J Gen Virol 90, 1–20. 8. Thomas, M. A., Spencer, J. F., Toth, K., Sagartz, J. E., Phillips, N. J., and Wold, W. S. (2008) Immunosuppression enhances oncolytic adenovirus replication and antitumor efﬁcacy in the Syrian hamster model, Mol Ther 16, 1665–1673. 9. Thomas, M. A., Spencer, J. F., and Wold, W. S. (2007) Use of the Syrian hamster as an animal model for oncolytic adenovirus vectors, Methods Mol Med 130, 169–183. 10. Steel, J. C., Morrison, B. J., Mannan, P., AbuAsab, M. S., Wildner, O., Miles, B. K., Yim, K. C., Ramanan, V., Prince, G. A., and Morris, J. C. (2007) Immunocompetent syngeneic cotton rat tumor models for the assessment of replication-competent oncolytic adenovirus, Virology 369, 131–142. 11. Wang, Y., Hallden, G., Hill, R., Anand, A., Liu, T. C., Francis, J., Brooks, G., Lemoine, N., and Kirn, D. (2003) E3 gene manipulations affect oncolytic adenovirus activity in immuno-

competent tumor models, Nat Biotechnol 21, 1328–1335. 12. Kasuya, H., Pawlik, T. M., Mullen, J. T., Donahue, J. M., Nakamura, H., Chandrasekhar, S., Kawasaki, H., Choi, E., and Tanabe, K. K. (2004) Selectivity of an oncolytic herpes simplex virus for cells expressing the DF3/MUC1 antigen, Cancer Res 64, 2561–2567. 13. Kirby, T. O., Rivera, A., Rein, D., Wang, M., Ulasov, I., Breidenbach, M., Kataram, M., Contreras, J. L., Krumdieck, C., Yamamoto, M., Rots, M. G., Haisma, H. J., Alvarez, R. D., Mahasreshti, P. J., and Curiel, D. T. (2004) A novel ex vivo model system for evaluation of conditionally replicative adenoviruses therapeutic efﬁcacy and toxicity, Clin Cancer Res 10, 8697–8703. 14. Marsman, W. A., Buskens, C. J., Wesseling, J. G., Offerhaus, G. J., Bergman, J. J., Tytgat, G. N., van Lanschot, J. J., and Bosma, P. J. (2004) Gene therapy for esophageal carcinoma: the use of an explant model to test adenoviral vectors ex vivo, Cancer Gene Ther 11, 289–296. 15. Wang, Y., Thorne, S., Hannock, J., Francis, J., Au, T., Reid, T., Lemoine, N., Kirn, D., and Hallden, G. (2005) A novel assay to assess primary human cancer infectibility by replicationselective oncolytic adenoviruses, Clin Cancer Res 11, 351–360. 16. O’Reilly, D., Muller, L., and Luckow, V. (1994) Virus Methods, in Bculovirus expression vectors: A laboratory manual, pp 132–134, Oxford University Press, Oxford.

Chapter 6 Imaging Luciferase-Expressing Viruses Michael A. Barry, Shannon May, and Eric A. Weaver Abstract Optical imaging of luciferage gene expression has become a powerful tool to track cells and viruses in vivo in small animal models. Luciferase imaging has been used to study the location of infection by replicationdefective and replication-competent viruses and to track changes in the distribution of viruses in mouse models. This approach has also been used in oncolytic studies as a noninvasive means to monitor the growth and killing of tumor cells modiﬁed with luciferase genes. In this chapter, we describe the techniques used for luciferase imaging as have been applied to track replication-defective and replicationcompetent adenoviruses in mouse and hamster models of oncolysis and virus pharmacology. Although these methods are simple, the process of obtaining accurate luciferase imaging data has many caveats that are discussed. Key words: Luciferase, In vivo imaging, Luminescence, Luciferin, Adenovirus

1. Introduction The technologies available to the basic scientist to track and localize viruses and tumor cells have historically been quite primitive. In most cases, virus and cell trafﬁcking has been assessed by the use of terminal assays in which the animal must be sacriﬁced and the cells or viruses are tracked after the animal is “taken apart” either at the organ level or in tissue sections. These “grind and ﬁnd” assays are quite laborious requiring that one actually sections the whole animal to be certain of the tissue localization of the virus to ensure that all sites are observed and unexpected localization sites are not missed. Furthermore, these terminal assays obviate the ability to perform kinetic studies in one animal over many time points. Given these difﬁculties, noninvasive and nonterminal virus and cancer cell tracking was needed. One approach that partially satisﬁes

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this need is to “arm” viruses or cancer cells with reporter genes that can be detected by imaging. Reporter genes encode proteins that are easily detected in cells and in intact animals with sensitive imaging systems. The most used reporter genes include betagalactosidase, luciferases, green ﬂuorescent protein (GFP) and its varied color derivatives, and alkaline phosphatase. Of these, luciferase and ﬂuorescent proteins can be used to varying degrees for optical imaging at visible wavelengths of light in small animals. One can also use reporter genes, such as thymidine kinase or the sodium iodide symporter, for high-energy PET and SPECT imaging in small animals, large animals, and in humans. For most researchers, optical imaging is simpler and more easily obtainable in the laboratory setting than radioactive imaging for PET or SPECT. Since PET and SPECT imaging are the subject of another chapter, they are not discussed further here. One can in some cases directly image reporters, like GFP and other ﬂuorescent proteins in living animals. In practice, high background ﬂuorescence and scatter in the green, red, and far red wavelengths make the “noise” of imaging too high to easily detect most current ﬂuorescent proteins in vivo (1, 2). Newer, far red ﬂuorescent proteins to date are still difﬁcult to image in mice (M.A. Barry et al., unpublished observations), but future near-infrared ﬂuorophores may circumvent this difﬁculty. Given these issues, luciferase imaging is arguably the best choice for noninvasive, inexpensive, and nonradioactive imaging in small animals. Given this, we have “armed” replication-defective and replication-competent adenovirus serotype 5 (Ad5) viruses with luciferase and GFP–luciferase reporter genes to track (1) sites of infection, (2) persistence of infection, (3) spread of virus, and (4) elimination of virus due to immune responses (1, 3–11). In addition, one can monitor immune responses against these proteins in immunocompetent mice (i.e., H-2d-restricted T cells in BALB/c mice vs. GFP antibodies against luciferase (7, 8)). These virus persistence–immune response studies of course cannot be performed in immunodeﬁcient models using human tumor xenografts. With these applications in mind, below we provide the simple protocol for imaging codon-optimized ﬁreﬂy luciferase with its substrate luciferin after Ad5 infection. Similar approaches can be applied for other luciferases (e.g., Gaussia luciferase, Renilla luciferase). These other luciferases use coelenterazine rather than luciferin as a substrate, so injections of substrate and timing of imaging are different. In our hands, the coelenterazine substrate is less soluble and does not distribute as well as luciferin, so can be more difﬁcult to use.

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2. Materials 1. Dulbecco’s phosphate buffered saline (DPBS) (Gibco/BRL). 2. Luciferin (Molecular Imaging Products Company) diluted to 20 mg/ml in DPBS, ﬁlter sterilize using a 0.22-Mm ﬁlter. Aliquot and store at −80°C. Protect from light exposure and avoid repeated freeze–thaws. 3. Ketamine (Fort Dodge), 100 mg/ml. 4. Xylazine (Vet Tek) diluted to 20 mg/ml using sterile ddH2O. 5. Isoﬂurane (Cardinal Health).

3. Methods 3.1. Animal Preparation for Luciferase Imaging

Appropriate animal protocol and biosafety approvals must be obtained before performing these experiments. Investigators should apply appropriate biosafety containment in instruments and rooms whenever imaging is performed. The following is an example of luciferase imaging after intramuscular (i.m.) injection of a replication-defective Ad5 vector (Ad-Luc). 1. Dilute Ad-Luc in DPBS to 2e11 virus particles (vp)/ml. 2. Using a 0.3-ml 29-G ½p syringe, inject 0.025 ml of virus into each quadriceps of the mouse. 3. After 24 h, place the mouse in an isoﬂurane induction chamber (3–5%) (see Note 2). 4. Once the animal is under sedation, inject 0.2 ml of luciferin (20 mg/ml in sterile DPBS) intraperitoneally (i.p.). Inject the luciferin substrate 5–10 min prior to image capture. 5. Working quickly, transfer the mouse to the imaging system and maintain sedation with 1–3% isoﬂurane. We have found that mice can be maintained for extended lengths of time without risk to the animals at these isoﬂurane levels. 6. Image the mouse dorsally using a 10-min exposure with 4 × 4 binning or 1 × 1 binning depending on signal intensity.

3.2. Image Capture Using Lumazone Imager

Below is provided the steps for a Lumazone (Mag Biosystems Lumazone, in vivo imaging system; software version 2.0) (see Note 3). 1. Make sure that cap is on the white light source in the imaging cabinet. 2. Open Lumazone software. The “Lumazone Analyzer” box should display.

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3. Select “Conﬁgure”: (a) Choose “Luciferin” from Experiment Type drop down. (This only affects how the image is titled after multichannel acquire is chosen and imaging is complete). (b) Select “Chemiluminescence” box. (c) Select “Brightﬁeld” box. (d) Select “OK.” 4. Select focus/exposure: (a) Select on “Chemi” button (for chemiluminescence): –

Make sure that “Adjust Exp for Binning” box is checked.

–

Choose binning setting.

–

Set “Exp Pvw” time (should show in MM:ss:mmm).

–

If “Adjust Exp for Binning” box is not checked, you also need to set the “Exp Acq” time.

(b) Select “BF” (for brightﬁeld). (c) Select “Live” to check and/or modify the focus of the image. (d) Select “Close.” (e) Select “Multichannel Acquire” in the “Lumazone Analyzer” box to begin taking the image. (f) Once the image has been captured, select on the “Best Fit Display Range or Contrast Equalization” button to bring the picture into focus – complete for each image. (g) Two images display. One image is the BF/white light image. One image is the chemiluminescence image. (h) Save the images. 3.3. Image Display

The bright light and chemiluminescent images are captured as grayscale images (Fig. 1). In order to create an image that is more esthetically pleasing and easier to interpret by eye, the images can be pseudocolored and overlaid onto a white light image of the animals. 1. Load the experiment. This opens up an image of the light emitted in vivo and a white light image of the animals (Fig. 1a). 2. Select the Calibrated LUT function on the analyze toolbar and select the image you want to calibrate. When prompted, select “No” in order to create a new image for calibration. This brings up an entirely new image window and does not alter the original image (Fig. 1b).

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Fig. 1. Operational screens for bright light and chemiluminescent images.

3. Two windows open when the calibrated image is created. Select Best Fit in the Display Range Window and use the LUT range to adjust the image and remove low-level background signal (Fig. 1c). 4. Select OK on the Calibrated LUT window and Yes to placing a calibration bar on the image. 5. Modify the calibration bar as needed and place in an area of the image that does not show signal (Fig. 1d). 6. Make changes to the font and overall appearance of the calibration bar and select OK. This creates a pseudocolored image with a calibration bar. 7. In order to show the position of the signal relative to the animal, an overlaid image can be created.

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8. Select Overlay Images and Yes and the image is calibrated. Select the calibrated image and then the white light image of the animals and an overlaid image is created (Fig. 1e). 3.4. Image Data Analysis

After capturing an image, data analysis need to be performed. There are several ways to capture and analyze the data. The primary issue of concern is light contamination, either from outside the imaging chambers from other animals or from background signal inherent to all electronics. The following is an example of image analysis performed using the Lumazone Imaging System. 1. Capture an image as previously described. 2. Load the experiment to be analyzed. 3. The ﬁrst thing that needs to be done is to subtract the background signal. This can be done using gray values without intensity calibration or using photons as determined by intensity calibration. 4. Deﬁne an area of interest (AOI) in the image outside of the area, where the mice are imaged Edit > New AOI (Fig. 2a). 5. In order to deﬁne background levels, the AOI background signal levels need to be determined. Open up count/size: Measure > Count/Size. Select the range of intensity to measure. For this procedure, the range should be the full scope of the range. For the Lumazone, this range is 0 to 65,535. Select the measurements to be determined using the Count/Size function: Measure > Count/Size > Measure > Select Measurements. In this case, select the Den./Inten. (mean). Select count on the Count/ Size function. To view the value, either click on the AOI or select Measure > Count/Size > Measure > View > Object Attributes. In this example, the Den./Inten. (mean) = 811.51459. 6. Once the background signal levels have been determined, they can be subtracted from the image. Open up image operations: Process > Operations. Check the following: Operation = Subtract, second operand = Number, Put result in = New Image. Enter the background signal (811.51459), deselect the AOI, and select apply (Fig. 2b). Select the Best Fit icon to view the best display range and contrast equalization. 7. Create an AOI over the area of the image to be quantitated. In the Count/Size function, select the measurements to acquire. In general, the Den./Inten. (sum) values are the most representative of the signal being measured. However, in some cases, the Den./Inten. (mean) may be more informative. 8. Depending on the values desired, gray values or converted photons/second can be reported. Select (none) in the intensity calibration if gray values are desired or select the calibration curve that was established when the unit was installed. In this

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Fig. 2. Image data analysis.

case, “Lumazone Bottom” gives values reported as photons/ second by converting gray values to photons using the calibration curve. Select Apply. 9. In the Count/Size function, select Count and Measure. Double click on the AOI to view the object attributes. In this example, there are 9.01972e9 photons/second emitted from the AOI (Fig. 2b) (see Note 1).

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4. Notes 1. Although the basic procedure of image capture during luciferase imaging is relatively simple, luciferase imaging is a dynamic procedure. As with all catalytic reactions, the signal is constantly changing. Luciferin, for example, is being degraded by the enzyme luciferase. Therefore, the procedures should be performed in a consistent and timely manner. Also, another issue of concern is how long of an exposure is needed. This changes from experiment to experiment and is dependent upon many factors, including how much virus was administered, the route, and even the position of the animal. Overexposure as shown in Fig. 2c can lead to a loss of valuable data. In this example, the exposure time should be reduced. Another issue is that light can be reﬂected off other objects, but still represents signal from the animal (Fig. 2c). A chamber that has dividers to separate the animals during luciferase imaging helps eliminate this problem. 2. Alternative anesthesia (Subheading 3.1): If isoﬂurane is not available, the mice can be anesthetized with ketamine and xylazine. Dilute the ketamine (27.77 mg/ml) and xylazine (1.11 mg/ml) in sterile dH2O and inject 0.1 ml i.p. into a 20to 25-g mouse using a 1.0-ml 26-G 5/8p syringe. The mouse is sedated for approximately 30 min. 3. Subheading 3.2: Each imaging system requires different steps in order to obtain both the chemiluminescent luciferase image as well as the bright-ﬁeld white image of the animal itself.

Acknowledgments We would like to thank all past and present Barry laboratory members for their experiences in imaging that have been distilled here. This work has been supported by the Muscular Dystrophy Association, the Ralph C. Wilson Foundation, the Propionic Acidemia Foundation, R01 AI06709, R01 AI065304, R01 CA136945-01A2, and by the NIH P50 CA91956 Prostate Cancer SPORE grant at the Mayo Clinic. References 1. Blum, J. S., J. S. Temenoff, H. Park, J. A. Jansen, A. G. Mikos, and M. A. Barry. 2004. Development and characterization of enhanced green ﬂuorescent protein and luciferase expressing cell line for non-destructive evaluation of

tissue engineering constructs. Biomaterials 25:5809–5819. 2. Adams, K. E., S. Ke, S. Kwon, F. Liang, Z. Fan, Y. Lu, K. Hirschi, M. E. Mawad, M. A. Barry, and E. Sevick-Muraca. 2007. Comparison of

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visible and near-infrared wavelength excitable ﬂuorescent dyes for molecular imaging of cancer. J Biomed Optics 12:024017-024011024019. Mok, H., D. J. Palmer, P. Ng, and M. A. Barry. 2005. Evaluation of polyethylene glycol modiﬁcation of ﬁrst-generation and helper-dependent adenoviral vectors to reduce innate immune responses. Mol Ther 11:66–79. Hofherr, S. E., S. Mok, F. C. Gushiken, J. A. Lopez, and M. A. Barry. 2007. Polyethylene Glycol Modiﬁcation of Adenovirus Reduces Platelet Activation, Endothelial Cell Activation, and Thrombocytopenia. Human Gene Therapy 18:837–848. Shashkova, E. V., K. Doronin, J. S. Senac, and M. A. Barry. 2008. Macrophage depletion combined with anticoagulant therapy increases therapeutic window of systemic treatment with oncolytic adenovirus. Cancer research 68:5896–5904. Hofherr, S. E., E. V. Shashkova, E. A. Weaver, R. Khare, and M. A. Barry. 2008. Modiﬁcation of adenoviral vectors with polyethylene glycol modulates in vivo tissue tropism and gene expression. Mol Ther 16:1276–1282.

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7. Weaver, E. A., and M. A. Barry. 2008. Effects of Shielding Adenoviral Vectors with Polyethylene Glycol (PEG) on Vector-speciﬁc and Vaccine-mediated Immune Responses. Hum Gene Ther 19:1369–1382. 8. Weaver, E. A., P. N. Nehete, S. S. Buchl, J. S. Senac, D. Palmer, P. Ng, K. J. Sastry, and M. A. Barry. 2009. Comparison of replicationcompetent, ﬁrst generation, and helper-dependent adenoviral vaccines. PLoS ONE 4:e5059. 9. Shashkova, E. V., S. M. May, K. Doronin, and M. A. Barry. 2009. Expanded Anticancer Therapeutic Window of Hexon-modiﬁed Oncolytic Adenovirus. Mol Ther Online Sept 15. 10. Doronin, K., E. V. Shashkova, S. M. May, S. E. Hofherr, and M. A. Barry. 2009. Chemical modiﬁcation with high molecular weight polyethylene glycol reduces transduction of hepatocytes and increases efﬁcacy of intravenously delivered oncolytic adenovirus. Hum Gene Ther 20:975–988. 11. Chen, C. Y., S. May, and M. A. Barry. 2010. Targeting Adenoviruses with Factor X-Single Chain Antibody Fusion Proteins. Hum Gene Ther 21(6):739–749.

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Chapter 7 In Vivo Positron Emission Tomography Imaging Using the Sodium Iodide Symporter as a Reporter Gene Lucile Tran, Nadège Dartial, Cecilia Hindorf, and Georges Vassaux Abstract Information regarding the biodistribution and kinetics of spread of oncolytic viruses is crucial for safety considerations in the design of future, more efﬁcient reagents. Although optical imaging can be used to gain this information in rodent models, imaging with radioactive isotopes presents the advantage of being directly applicable to humans, and preclinical imaging data can provide a basis for the design of clinical protocols. In this chapter, we describe the use of the Na/I symporter as a reporter gene, associated with 124 I-NaI as a radiotracer, to monitor the anatomical localization, as well as the propagation, of oncolytic viruses using dedicated, preclinical positron emission tomography scanners. Key words: Gene therapy, Oncolytic virus, Reporter gene, Sodium iodide symporter, Positron emission tomography, Molecular imaging

1. Introduction Molecular imaging of gene expression has important applications in gene therapy. This technology can provide unique information on the anatomic site, level, and duration of gene transfer. This information has safety implications and can be used to improve the design of new vectors. Optical (bioluminescence, ﬂuorescence) imaging and imaging with radioactive isotopes [scintillation camera and positron emission tomography (PET)] are the most commonly used imaging modalities by the gene therapy community. The former is highly relevant for preclinical studies in rodents (see Chapter 10), but the attenuation of light originating from deeper tissues limits its applications in large animals and humans. Imaging using

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radiotracers is, therefore, the method of choice to monitor gene transfer in humans. Molecular imaging of gene expression requires a reporter gene as well as a tracer (a radiotracer in the case of imaging using radioisotopes). The tracer accumulates in tissues in which the reporter gene is expressed and a camera is able to detect the accumulation of the radioactivity in these tissues. The two main modalities of detection of gene transfer are single photon emission-computed tomography (SPECT) and PET. SPECT imaging is based on the detection of individual photons emitted by the isotope (1). Detection by a PET camera requires tracers that emit positrons, the so-called beta plus emitters. The positron travels through the medium until it has lost its kinetic energy and then forms a pair with an electron. The electron–positron pair is immediately annihilated and two 511 keV photons are emitted in opposite directions (180° apart). The detector in the PET camera registers all events on a ring of detectors around the radioactivity source, but processes only events that occur simultaneously (within a very short time window) in two detectors situated 180° apart (2). Because of its potential in terms of quantiﬁcation, as well as its high sensitivity, PET is the technique of choice to collect information on the anatomic location, magnitude, and kinetics of gene expression. Different reporter systems have been described in the literature: the herpes simplex thymidine kinase (HSV1-tk) gene associated with its radiolabeled substrates (3, 4) and the dopamine D2 (5), somatostatin type 2 (6), and the norepinephrine (7) receptors associated with their radioligands have all been used to monitor gene transfer by either SPECT or PET. In this chapter, we focus on another reporter system which uses the sodium/iodide symporter (NIS) as a reporter gene. NIS can be associated with different radiohalogen for either SPECT (123I or 99mTc-pertechnetate) or PET (124I) imaging (8). NIS is a transmembrane protein using the sodium gradient to concentrate iodide in cells. It is mainly expressed in the thyroid and the stomach (8, 9). The use of NIS does not require extensive radiochemistry to generate probes, and relevant radioisotopes are commercially available for both gamma camera imaging (planar and SPECT) and PET imaging. The approach relies on more than 50 years of experience of imaging NIS in thyroid-related disorders (9). Many studies have successfully used the NIS gene to monitor gene transfer in preclinical studies (for review, see refs. 8, 9). As far as oncolytic viruses are concerned, preclinical imaging of adenoviruses (10–12), measles (13–15), and vesicular stomatitis viruses (16) has been reported. A phase I clinical study has recently been conducted, validating the imaging in humans using an oncolytic adenovirus encoding NIS (17). This chapter details a protocol for imaging NIS in living, anesthetized mice using a PET camera.

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2. Materials 1. Oncolytic viruses encoding NIS comprise adenoviruses (10–12, 17), measles (13–15), and vesicular stomatitis viruses (16). 2. Mice are usually used in these studies. The strain and the sex depend on the pathology targeted by the gene therapy protocol associated with the imaging project (see Note 1). 3. Levo-thyroxine (Levothyrox, Merck): Prepare a solution at 5 mg/mL in water (see Note 2). 4. Isoﬂurane (Forene, Abbott) or any other approved anesthetic. 5. Phosphate-buffered saline (PBS): Prepare a 10X stock containing 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. All these chemicals are available from Sigma. Prepare in sterile conditions a 1× solution by diluting the 10X stock in sterile water. 6. Na[124I] can be obtained from IBA (http://group.iba-worldwide.com) (see Note 3). 7. Different dedicated small animal PET systems are commercially available from different manufacturers: Siemens Medical Solutions (http://www.smed.com), GE Healthcare (http:// www.gehealthcare.com), Bioscan (http://www.bioscan.com) (see Note 4). 8. Anesthesia system, chamber, and isoﬂurane pump (Summit medical equipment company, Bend, OR).

3. Methods 3.1. General Considerations

3.2. Preparation of the Study Animals

Radioisotopes should be used only by persons trained in radiation protection. Time, distance, appropriate shielding, and knowledge of the radioactive source are the four keywords for practical radiation protection which always must be applied in order to minimize the radiation dose received. Radioactive waste disposal must be handled in accordance with national regulations. Radioisotopes must be handled in a designated area. It is important to image the animals according to very strict protocols, for example by performing the imaging at the same time point post radiotracer injection, to obtain reliable and comparable images. 1. Mice are handled in a germ-free environment and given food and water ad libitum. 2. Mice are acclimatized for 1 week before the imaging experiments.

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3. Control animals not injected with the viral vector expressing the NIS gene need to be included. In these animals, iodide accumulation is expected in the thyroid (unless levo-thyroxine is administered) and stomach as a result of endogenous NIS expression and in the bladder as a result of excretion in the urine. 4. Mice can be fed with levo-thyroxine (5 mg/mL) in their drinking water to suppress thyroidal NIS expression. Ideally, this treatment should start 1 week before the imaging procedure. 3.3. Injection of the Viral Vector Expressing the NIS Imaging Gene

3.4. Preparation of 124 I-NaI for Injection into Mice

1. Inject the appropriate dose of the viral vector. The dose and schedule of injection depend on the particular experiment (see Note 5). 2. The vector can be injected intraperitoneally, subcutaneously, and intravenously via the tail vein or intratumorally. The volume of injection varies depending on the titer of the virus, but we recommend that this volume does not exceed 1 mL for intraperitoneal administration, 0.2 mL for subcutaneous injection, 0.2 mL for intravenous injection, and 0.1 mL for intratumoral injection. Repeated injections can be envisaged. 1. Dilute the 124I with PBS or NaCl (0.9%) to a suitable activity concentration (18.5 MBq/ml). 2. Fill the syringe. 3. Measure the activity in the syringe with a dose calibrator. It should correspond approximately to 1.85 MBq. The volume of the injected solution must be greater than 50 ML (typically, 100 ML). 4. Record the exact measurement (activity and time of measurement). 1. Inject the entire amount of Na[124I] contained in the syringe (typically, 1.85 MBq).

3.5. Injection of the Radioactive Solution into Mice

2. Record the injection time.

3.5.1. Intraperitoneal Injection

3. Measure the remaining activity in the syringe and record it together with the time of measurement. 4. The mice are kept awake during 5 min before the anesthesia to allow the dissemination of the radioactive iodide solution throughout the whole body. 5. Place the mouse in the anesthesia chamber in an atmosphere of 2% isoﬂurane (see Note 6). 6. Record the time point at which the mouse is anesthetized. 7. The anesthetized mouse is placed onto a sealed, prewarmed (37°C) imaging chamber connected to the anesthetic gas system (2% isoﬂurane) (see Note 7).

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1. Place the mouse into the anesthesia chamber in an atmosphere of 2% isoﬂurane. 2. Place the nose of the anesthetized mouse in the cone receiving a constant ﬂux of 2% isoﬂurane. Implant a tail vein cannula (see Note 9). 3. Place the anesthetized mouse in a sealed, prewarmed (37°C) imaging chamber connected to the anesthetic gas system (2% isoﬂurane) (see Notes 6–8). 4. Insert the syringe in the tubing of the cannula. Inject the entire amount of Na[124I] contained in the syringe. Remove the syringe, ﬁll it with 0.05 mL PBS, and ﬂush out the radioactivity remaining in the tubing. 5. Record the injection time.

3.6. PET Imaging Session

1. Once the mouse is well-positioned, the bed is moving inside the camera (see Note 9). 2. Data acquisition can start as soon as possible after the time of injection for the intravenous procedure or at around 10 min post injection for the intraperitoneal procedure. Tests in our laboratory have shown that both procedures provide equivalent data, as long as the whole data acquisition is completed within 1 h after radiotracer injection. 3. A manual describing the image acquisition should be provided by the microPET manufacturer (this stage is apparatus-speciﬁc). 4. Appropriate energy window settings and timing window settings must be optimized as the decay scheme of 124I is relatively complicated (this stage is apparatus-speciﬁc). 5. The duration of the PET acquisition is usually between 20 and 60 min.

3.7. Image Reconstruction and Analysis

1. Use the software provided by the manufacturer to reconstruct the image. 2. The analysis of the images can be performed via various software programs, for example via the A Medical Imaging Data Examiner (AMIDE) program (18) or by ImageJ (http:// rsbweb.nih.gov/ij/), both available free on line. 3. Images can be visualized coronally, sagittally, or transversally (a coronal image is presented in Fig. 1). 4. The activity in certain regions of the images can be quantiﬁed by drawing regions of interest (ROIs) in the images, if the microPET is calibrated for quantitative imaging.

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Fig. 1. Visualization of NIS expression in anesthetized mice upon tail vein injection of 5 × 108 PFU of a wild-type adenovirus encoding NIS (AdIP1 in ref. 10) or saline buffer (control). The images presented show coronal views and the chosen slices lie at the level of the thyroid region (T). 124I tracer is seen in the stomach (S) and the bladder (B) of the control animal. Tracer uptake in the AdIP1-treated animal is in the upper peritoneal cavity/abdomen consistent with adenoviral homing to the liver (L) when administered systemically.

4. Notes 1. Animal experiments must be conducted in accordance with the animal care policy approved by your institution and by national regulations. 2. To dissolve levo-thyroxine in water, crush the tablets (5 mg/ mL) mechanically with a mortar and warm the solution under agitation to homogenize. 3. [124I]Iodide can also be produced using the 124Te(p,n)124I reaction using irradiation of a [124Te]tellurium(IV) oxide target with 12.5 MeV protons followed by dry distillation in a quartz apparatus (see protocol and relevant references in (19)). 4. The instructions regarding how to set up the microPET camera are provided by the manufacturer. This process is apparatusspeciﬁc. In addition, an optimization procedure is required to use 124I. 5. If the virus is expected to infect an area in the proximity of the thyroid, it is advised to feed mice with levo-thyroxine 1 week before the scan. This suppresses iodide uptake by the thyroid. 6. To reduce iodide uptake by the bladder, pressure can be exerted on the animal’s bladder region to eliminate the urine. However,

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note that the urine is highly radioactive. To avoid contamination, absorb the urine with a thick layer of paper and dispose of it in a lead-protected waste bin. 7. Wrap the mouse in cling ﬁlm if the imaging chamber is not heated. 8. Tail vein injection requires training. To increase success, the tail can be dilated by heat (incubate in water at 37°C). 9. Perform preliminary tests to ﬁnd the optimal position of the mouse during the microPET acquisition.

Acknowledgments This work is supported by grants from INSERM, la Ligue Nationale Contre le Cancer, ARC, by a grant from the “Paris scientiﬁques régionaux” scheme from the Région Pays de la Loire and by grant 0607 – 3D1615 – 66/AO INSERM from the French National Cancer Institute (INCa). References 1. Buck, A. K., Nekolla, S., Ziegler, S., Beer, A., Krause, B. J., Herrmann, K., Scheidhauer, K., Wester, H. J., Rummeny, E. J., Schwaiger, M., and Drzezga, A. (2008) Spect/Ct. J Nucl Med 49, 1305–19. 2. Cherry, S. R. (2006) The 2006 Henry N. Wagner Lecture: Of mice and men (and positrons)--advances in PET imaging technology. J Nucl Med 47, 1735–45. 3. Tjuvajev, J. G., Avril, N., Oku, T., Sasajima, T., Miyagawa, T., Joshi, R., Safer, M., Beattie, B., DiResta, G., Daghighian, F., Augensen, F., Koutcher, J., Zweit, J., Humm, J., Larson, S. M., Finn, R., and Blasberg, R. (1998) Imaging herpes virus thymidine kinase gene transfer and expression by positron emission tomography. Cancer Res 58, 4333–41. 4. Mullerad, M., Eisenberg, D. P., Akhurst, T. J., Adusumilli, P. S., Riedl, C. C., Bhargava, A., Gonen, M., Finn, R., Scardino, P. T., and Fong, Y. (2006) Use of positron emission tomography to target prostate cancer gene therapy by oncolytic herpes simplex virus. Mol Imaging Biol 8, 30–5. 5. Liang, Q., Satyamurthy, N., Barrio, J. R., Toyokuni, T., Phelps, M. P., Gambhir, S. S., and Herschman, H. R. (2001) Noninvasive, quantitative imaging in living animals of a mutant dopamine D2 receptor reporter gene

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in which ligand binding is uncoupled from signal transduction. Gene Ther 8, 1490–8. McCart, J. A., Mehta, N., Scollard, D., Reilly, R. M., Carrasquillo, J. A., Tang, N., Deng, H., Miller, M., Xu, H., Libutti, S. K., Alexander, H. R., and Bartlett, D. L. (2004) Oncolytic vaccinia virus expressing the human somatostatin receptor SSTR2: molecular imaging after systemic delivery using 111In-pentetreotide. Mol Ther 10, 553–61. Brader, P., Kelly, K. J., Chen, N., Yu, Y. A., Zhang, Q., Zanzonico, P., Burnazi, E. M., Ghani, R. E., Serganova, I., Hricak, H., Szalay, A. A., Fong, Y., and Blasberg, R. G. (2009) Imaging a Genetically Engineered Oncolytic Vaccinia Virus (GLV-1h99) Using a Human Norepinephrine Transporter Reporter Gene. Clin Cancer Res 15, 3791–801. Baril, P., Martin-Duque, P., and Vassaux, G. (2009) Visualization of gene expression in the live subject using the Na/I symporter as a reporter gene: applications in biotherapy. Br J Pharmacol 159(4), 761–71. Hingorani, M., Spitzweg, C., Vassaux, G., Newbold, K., Melcher, A., Pandha, H., Vile, R., and Harrington, K. The Biology of the Sodium Iodide Symporter and its Potential for Targeted Gene Delivery. Curr Cancer Drug Targets 2010 Mar;10(2), 242–67.

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10. Merron, A., Peerlinck, I., Martin-Duque, P., Burnet, J., Quintanilla, M., Mather, S., Hingorani, M., Harrington, K., Iggo, R., and Vassaux, G. (2007) SPECT/CT imaging of oncolytic adenovirus propagation in tumours in vivo using the Na/I symporter as a reporter gene. Gene Ther 14, 1731–8. 11. Peerlinck, I., Merron, A., Baril, P., Conchon, S., Martin-Duque, P., Hindorf, C., Burnet, J., Quintanilla, M., Hingorani, M., Iggo, R., Lemoine, N. R., Harrington, K., and Vassaux, G. (2009) Targeted radionuclide therapy using a Wnt-targeted replicating adenovirus encoding the Na/I symporter. Clin Cancer Res 15, 6595–601. 12. Merron, A., Baril, P., Martin-Duque, P., de la Vieja, A., Tran, L., Briat, A., Harrington, K. J., McNeish, I. A., and Vassaux, G. Assessment of the Na/I symporter as a reporter gene to visualize oncolytic adenovirus propagation in peritoneal tumours. Eur J Nucl Med Mol Imaging 2010 Jul;37(7), 1377–85. 13. Dingli, D., Peng, K. W., Harvey, M. E., Greipp, P. R., O’Connor, M. K., Cattaneo, R., Morris, J. C., and Russell, S. J. (2004) Image-guided radiovirotherapy for multiple myeloma using a recombinant measles virus expressing the thyroidal sodium iodide symporter. Blood 103, 1641–6. 14. Hasegawa, K., Pham, L., O’Connor, M. K., Federspiel, M. J., Russell, S. J., and Peng, K. W. (2006) Dual therapy of ovarian cancer using measles viruses expressing carcinoem-

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bryonic antigen and sodium iodide symporter. Clin Cancer Res 12, 1868–75. Msaouel, P., Iankov, I. D., Allen, C., Aderca, I., Federspiel, M. J., Tindall, D. J., Morris, J. C., Koutsilieris, M., Russell, S. J., and Galanis, E. (2009) Noninvasive imaging and radiovirotherapy of prostate cancer using an oncolytic measles virus expressing the sodium iodide symporter. Mol Ther 17, 2041–8. Goel, A., Carlson, S. K., Classic, K. L., Greiner, S., Naik, S., Power, A. T., Bell, J. C., and Russell, S. J. (2007) Radioiodide imaging and radiovirotherapy of multiple myeloma using VSV(Delta51)-NIS, an attenuated vesicular stomatitis virus encoding the sodium iodide symporter gene. Blood 110, 2342–50. Barton, K. N., Stricker, H., Brown, S. L., Elshaikh, M., Aref, I., Lu, M., Pegg, J., Zhang, Y., Karvelis, K. C., Siddiqui, F., Kim, J. H., Freytag, S. O., and Movsas, B. (2008) Phase I study of noninvasive imaging of adenovirus-mediated gene expression in the human prostate. Mol Ther 16, 1761–9. Loening, A. M., and Gambhir, S. S. (2003) AMIDE: a free software tool for multimodality medical image analysis. Mol Imaging 2, 131–7. Groot-Wassink, T., Aboagye, E. O., Glaser, M., Lemoine, N. R., and Vassaux, G. (2002) Adenovirus biodistribution and noninvasive imaging of gene expression in vivo by positron emission tomography using human sodium/ iodide symporter as reporter gene. Hum Gene Ther 13, 1723–35.

Chapter 8 Maintaining and Loading Neural Stem Cells for Delivery of Oncolytic Adenovirus to Brain Tumors Atique U. Ahmed, Ilya V. Ulasov, Ronald W. Mercer, and Maciej S. Lesniak Abstract Despite recent advancements in the treatment of cancer, the prognosis for patients with malignant brain tumors remains poor. The success of currently available therapies has been limited in part because of the disseminated nature of these tumors. Furthermore, most of these tumors, when in a high-grade form, are resistant to chemo- and radiotherapy. Taking the above considerations into account, effective treatment of these cancers not only requires the development of new means to target tumor burdens that have dispersed signiﬁcantly from their site of origin, but also therapeutic approaches which can appropriately discriminate between tumor cell and normal brain. In the past two decades, novel approaches involving the use of oncolytic adenoviruses to target malignant brain tumors have undergone extensive investigation and proven to be an effective mode of antiglioma therapy. While the use of various oncolytic adenoviruses has been proven to be safe for local delivery in preclinical and clinical trials, the successful application of this approach in the clinic has been hampered by the host immune response against the viral vector. The discovery of the inherent tumor-tropic properties of neural stem cells (NSCs) provides a unique opportunity that employs NSCs as a cellular vehicle to track tumor cells and deliver therapeutic oncolytic virus. This presents a novel platform for targeted delivery of oncolytic adenovirus to disseminated tumors selectively while hiding the therapeutic virus from the host immune system. NSC loaded with an oncolytic adenovirus offer a more selective and effective method of targeting satellite tumor burdens and this chapter reviews the methodology associated with this unique approach. Key words: Cell carrier, Neural stem cell, Oncolytic adenovirus, Glioblastoma, Virus loading

1. Introduction 1.1. Oncolytic Virus

Cancer is a multistage genetic disease that involves alterations in multiple molecular pathways related to growth control and cell death (1). There are many genes that have been identiﬁed in recent years, which could be potential targets for novel cancer therapy.

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The new knowledge of the molecular mechanisms underlying oncogenesis and the development of the viral vector as a vehicle for gene delivery have permitted the formulation of the concept of cancer gene therapy. Strategies for cancer gene therapy adopt ideas and technologies ranging from generating the immune response against tumor antigens to directly attacking tumor cells. However, the therapeutic efﬁcacy of most of the cancer gene therapy approaches is signiﬁcantly compromised by the inability of the current viral vectors to deliver genes in vivo and target systemic metastasis. To overcome this problem, researchers have used the viruses’ ability to spread from their site of infection to the neighboring cells. Infected cells are killed, as they become the factories for producing the new infectious viral particles (VPs). The process of infection is particularly attractive to cancer gene therapy because it not only ampliﬁes therapeutic genes in a tumor-selective manner, but also has the potential to lyse and kill the infected tumor cells. The use of replicating viruses against cancer, also referred to as virotherapy, has a long history that began in the early 1900s with the anecdotal reports of temporary cancer remissions in patients who received a viral immunization or had a natural viral infection (2). The success of this approach depends on our ability to identify and engineer viruses that replicate speciﬁcally in tumor cells, but not in the normal cells. These viruses, termed oncolytic viruses, are essentially tumor-speciﬁc, self-replicating, lysis-inducing cancer killers. Many oncolytic viruses that belong to several viral families have been identiﬁed or engineered. They include herpes simplex viruses, adenovirus, retroviruses paramyxoviruses, and poxviruses (3). These viruses can be categorized into four major groups on the basis of their oncolytic restriction: (1) mutation/deletionderived viruses, (2) transcriptionally targeted oncolytic viruses, (3) transductionally targeted oncolytic viruses, and (4) “naturally smart” viruses. Oncolytic viruses for cancer exploit the difference of the molecular makeup between the tumor cells and their normal counterparts; they also utilize recombinant DNA technology to engineer viral vectors to selectively replicate in the tumor cells and destroy them. 1.2. Oncolytic Viral Therapy for Malignant Glioma

Despite recent advancement in surgical techniques and adjuvant radio- and chemotherapy, the median survival for patients with GBM is approximately 12–15 months and it remains one of the deadliest classes of human cancer (4). Current methods of cancer treatment include surgical excision of the tumor, chemotherapy, tumor radiation, and various combinations of these therapies. The two main problems that exist with these current therapies are lack of treatment speciﬁcity and extensive peripheral toxicity. These issues have initiated the development of novel approaches to cancer therapy in order to improve the patient lifestyles and increase their survival (5). Cancer gene therapy allows us to utilize our knowledge

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of the molecular mechanisms underlying oncogenesis as well as our ability to develop novel viral vectors for targeted delivery of therapeutic genes (6). During the past two decades, a number of promising oncolytic viruses have demonstrated antiglioma activity in both preclinical and clinical settings (7). However, the host’s immune system imposes a new, greater challenge to the development of an effective oncolytic viral (OV) therapy against human cancers. Our immune system, which evolved to control viral infections, does not have the sophistication to distinguish between malicious pathogens and therapeutic viruses. So, most of the therapeutic viruses delivered at the tumor site do not exist as free-ﬂoating particles for longer than few minutes at most (8), which inhibits their ability to infect satellite tumor burdens and limits antitumor effects. 1.3. Neural Stem Cell

Neural stem cells (NSCs) of the CNS have recently received a great deal of attention because of their therapeutic potential for neurological disorders. NSCs are deﬁned as CNS progenitor cells that have the capacity for self-renewal and multipotent potential to differentiate into three major cells in CNS: neuron, astrocytes, and oligodendrocytes (9). In the last decade, many in vitro and in vivo studies demonstrated that NSCs have the unique, inherent properties to migrate throughout the brain and target invasive solid tumors, including gliomas (10, 11). This provides a novel platform for targeted delivery of anticancer agents to disseminated tumors selectively while leaving healthy surrounding neurons intact. Recent studies have shown that NSCs can be engineered to deliver a variety of therapeutic genes to selectively destroy gliomas in animal models (12, 13).

1.4. NSC as a Cell Carrier: Guided Trojan Horse for Systemic Oncolytic Virus Delivery

Early in vivo experiments with oncolytic viruses revealed that infected virus-producing cells could also mediate antitumor activity when administered in the place of naked virus (14). This led to the hypothesis that producer cells can be used to hide the therapeutic virus from the host immune system and delivered systemically in order to travel precisely to the disseminated tumor burden. Our lab, along with others, has shown that NSC can be used as a carrier for stealth delivery of an oncolytic adenovirus in vivo for antiglioma therapy (12). At least in theory, oncolytic viruses can be ideal anticancer agents to be loaded into NSC for the following reasons: ﬁrst, oncolytic viruses can replicate selectively in the tumor cells and thus should be able to amplify the therapeutic gene efﬁcacy at the tumor sites; secondly, once the oncolytic virus is released from the loaded NSC at the delivery sites, it can also distinguish tumors from normal tissues and induce tumor cell-speciﬁc cytolysis. In order to achieve the maximum delivery and therapeutic efﬁcacy in vivo, NSCs need to be loaded/infected with the maximum dose of oncolytic virus without affecting the carrier cell survival and their tumor homing property. The kinetics with which the virus

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interacts with the carrier cell must be compatible with the in vivo trafﬁcking of the carrier cell to the tumor. The timing of the oncolytic virus life cycle is a critical determinant factor for this phase. To act as Trojan horse vehicles and successfully hide the oncolytic virus from the host immune system, the cell carriers should ideally reach the tumor site before the viral antigen is displayed on their surface. Moreover, the ex vivo virus loading capacity must be synchronized with the in vivo delivery route and the migration rate of the carrier cell in order to maximize the delivery of the therapeutic oncolytic virus at the tumor sites. Each virus has very different kinetics of entry, replication, and progeny release. Basically, careful ex vivo examination of the interaction between the cell carrier (CC) and the individual virus needs to be done in order to maximize therapeutic virus delivery in vivo at the tumor site. Here, we present a detailed overview of oncolytic adenovirus delivery techniques using NSCs as cell carriers. The protocol outlined in this chapter describes techniques necessary to load an oncolytic adenoviral vector into NSC for a stealth in vivo therapeutic virus delivery approach for antiglioma therapy.

2. Materials 2.1. Cell Culture

1. Normal tissue culture equipment, i.e., incubators with CO2 laminar, biohazard ﬂow hood. 2. Heat inactivated fetal bovine serum (Life Technologies). 3. Dulbecco’s modiﬁed Eagle’s media (DMEM; Life Technologies) containing 50 U/ml of penicillin–streptomycin (Life Technologies) for A549 cell culture. 4. 0.25% Trypsin (Invitrogen). 5. 15-ml conical tubes. 6. 0.22- and 0.45-Mm ﬁlters. 7. Sterile phosphate buffer saline (PBS), pH 7.4 (Invitrogen). 8. Hemocytometer. 9. Trypan blue (Sigma).

2.2. Maintenance of Neural Stem Cells

1. ReNcell NSC Maintenance Medium (CHEMICON) contains DMEM/F12 w/o HEPES, w/L-glutamine human serum albumin, human transferring, putrescine dihydrochloride, human recombinant insulin, L-thyroxine, triiodothyronine, progesterone, sodium selenite, heparin, and corticosterone (see Note 1). 2. ReNcell NSC Freezing Medium (CHEMICON). 3. Basic ﬁbroblast growth factor (bFGF; FGF-2; speciﬁc activity >2 × 106 U/mg CHEMICON).

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4. Epidermal growth factor (EGF; speciﬁc activity >1 × 107 U/mg; CHEMICON). 5. Laminin (Sigma). 6. DMEM/F12 w/o HEPES, w/L-Glutamine (CHEMICON). 7. Accutase (CHEMICON). 2.3. Large-Scale Production of Oncolytic Adenovirus Vectors

1. A549 cell. 2. 175-mm ﬂask. 3. DMEM medium. 4. Virus dialysis buffer: 100 mM Tris–HCl (pH 7.4), 10 mM MgCl2, and 10% (v/v) glycerol.

2.4. Puriﬁcation of Adenovirus by CsCl Banding

1. 10 and 100 mM Tris–HCl (pH 8.0), autoclave sterilized. 2. 5% sodium deoxycholate, ﬁlter sterilized. 3. Glycerol, autoclave sterilize. 4. Beckman SW 28 rotor and ultra-clear tubes. 5. Slide-A-Lyser dialysis cassettes (Pierce, Rockford, IL, USA). 6. Dialysis buffer: 10 mM Tris–HCl (pH 8.0).

2.5. Viral Particle Number Determination

1. Virus lysis buffer: 0.1% sodium dodecyl sulfate (SDS), 0.02 M Tris–HCl (pH 7.4), 1 mM EDTA. 2. Spectrophotometer capable of reading optical density at 260 nm. 3. Measuring cuvette. 4. 1.5-ml Eppendorf tubes. 5. Heating blocks.

3. Methods 3.1. Large-Scale Adenovirus Production

Because the purity and functional integrity of recombinant viral vectors’ preparation are extremely critical for ex vivo loading into carrier cells as well as in vivo therapeutic efﬁcacy, this part of the methods section brieﬂy summarizes methods used for the propagation and analysis of oncolytic adenoviral stocks to be loaded into stem cells for in vivo delivery. 1. The propagation of oncolytic adenoviral vectors is performed in the A549 cell line. Because most of the viruses remain associated with the infected cells until the cells are lysed at a very late phase of infection, high-titer stocks can be prepared by concentrating infected A549 cells. Infect A549 cell monolayer with the “low-passage” adenovirus stock in the 75-mm dish

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and gradually scaling up to a 150-mm dish. For each large-scale preparation, use at least twenty to thirty 150-mm dishes. 2. When a complete cytopathic effect (CPE) is reached (~48 h post infection, no more than 96 h), collect the cells in the media (20–25 ml for each 150-mm dish). Spin down the cells at 129 × g (800 rpm) for 10 min in the Eppendorf 5810R benchtop centrifuge. Discard the supernatant and resuspend the cell pellets in 1.5–3.0 ml of 0.1 M Tris–HCl buffer (pH 8.0) per 150 mm pellet (see Note 2). 3. Release the virus by snap freezing the cell plate in liquid nitrogen and thawing in the 37°C water bath. Repeat this process three times. Transfer the freeze/thaw cell lysate to Beckman (25 × 89 mm) polyallomer centrifuge tubes and spin out the cell debris at 4,000 × g (7,697 × g) for 10 min in the Beckman CS-15R benchtop centrifuge using prechilled (4°C) F0630 rotor. Alternatively, centrifuge the samples in 50-ml Falcon tubes in a Sorvall RC5C-Plus using the SLA-600TC rotor at 4,890 × g for 10 min. Remove the supernatants and transfer them to clean 50-ml Falcon tubes. 3.2. Vector Puriﬁcation by CsCl Ultracentrifugation

Puriﬁcation of an oncolytic Ad vector is a three-step process: 1. A discontinuous CsCl gradient that removes the majority of the cellular debris and unpackaged viral particles 2. A continuous CsCl gradient to completely separate infectious and defective viral particles 3. The removal of CsCl from concentrated viral stock by desalting All the steps in this protocol require the use of 30-ml centrifuge tubes for Beckman SW28 rotors or equivalent. Other tube size can be used as long the solution volumes are adapted (see Note 3).

3.2.1. Discontinuous Gradient

1. Prepare the following CsCl solutions and ﬁlter sterilize: (a) TD = 8 g NaCl + 0.38 g KCl + 0.1 g Na2HPO4, 3 g Tris base per liter H2O, pH to 7.5 with HCl (b) 1.25 g/ml = 36.16 g CsCl + 100 ml TD (c) 1.35 g/ml = 51.2 g CsCl + 100 ml TD (d) 1.40 g/ml = 62 g CsCl + 100 ml TD 2. Prechill a swing bucket rotor (Beckman SW28 or equivalent) to 4°C. 3. Prepare CsCl gradients by carefully layering 7.6 ml of 1.4 g/ ml of CsCl beneath 11.4 ml of 1.25 g/ml CsCl solution in Beckman (25 × 89 mm) ultra-clear centrifuge tubes. Finally, very carefully overlay the gradients with the cell-free media containing viral particles from above (~19 ml per gradient).

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If the viral stock is less than 19 ml, use 0.1 mM Tris–HCl (pH 7.9) to complete the volume. 4. Make sure that the tubes are well-balanced, and centrifuge at 100,000 × g in SW 28 rotor for 2 h at 15°C. Sorvall Discovery 100S ultracentrifuge with Superspin rotors can be used alternatively. 5. In a laminar ﬂow hood, very carefully remove the tubes from the rotor, and then secure one tube with a three-pronged clamp attached to a stand. Generally, two milky layers are observed; the upper band of which consist of low-density, empty, assembled adenoviral particles while the lower band represents mature encapsidated viral particles. Collect the lower band by puncturing either side of the tube using a 5-ml syringe and 19-gauge needle (see Note 4). 6. Collect all the virus bands together. 3.2.2. Continuous Gradient

1. Load the collected virus from the previous step into Beckman 13 × 15-mm ultra-clear centrifuge tubes. Fill the tubes to within 2–3 mm to the top with 1.35 g/ml CsCl solution. 2. Centrifuge at 150,000 × g at 15°C for 16–20 h using the Beckman SW 55 swing out rotor. Sorvall Discovery 100S ultracentrifuge with TH-660 rotor can be used alternatively. 3. After the centrifuge, the continuous gradient looks like the step gradients one, except there is only one band. Collect the virus bands, which should be located centrally to the tube. Keep the CsCl-puriﬁed viral fraction on ice at all time.

3.2.3. Virus Desalting

1. Transfer the viral vector into Pierce slide-A-Lyzer dialysis cassette. 2. Dialysis in 500-ml dialysis buffer for 30 min, twice. 3. Repeat the dialysis in 1,000-ml dialysis buffer for 1 h for three times. 4. Remove the vector suspension from the dialysis cassette and aliquot in sterile Eppendorf tubes in such volume that repeated freeze–thaw can be avoided in order to prevent loss of activity. Keep vector aliquots in a −80°C freezer.

3.3. Viral Particle Determination

This method determines the titer of a viral stock suspension using viral particles as units by establishing the correlation between the number of VPs and the DNA contained. This can easily be measured by absorbance at 260 nm (OD260) using a spectrophotometer. Each OD260 unit represents approximately 1.1 × 1012 adenovirus particles. The OD260 reading between 0.1 and 1.0 should only be used to do the ﬁnal calculation because only at this range can the OD260 reading accurately reﬂect the amount of DNA for most spectrophotometers.

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1. Prepare virus lysis buffer. 2. Prepare the following dilution of puriﬁed viral stock with the virus lysis buffer: 1:3 dilution = 33.3 Ml viruses + 66.7 Ml of VLB 1:5 dilution = 20 Ml viruses + 80 Ml of VLB 1:10 dilution = 10 Ml viruses + 90 Ml of VLB 3. Mix brieﬂy by vortex and incubate at 56°C for at least 10 min. 4. Centrifuge the tubes and allow the samples to cool down. 5. Switch on the spectrophotometer, use the 260/280 – program, and let the UV lamp warm up for at least 10 min before reading samples. 6. Measure OD260 in a spectrophotometer for each sample. Use 100 Ml of VLB as blank. 7. To calculate the number of viral particles per ml in the stock, use the following formulas: OD260 × dilution factor × 1.1 × 1012 = VP/ml. Disregard the OD260 reading out of 0.1–1.0 range. 3.4. Viral Titration by Rapid Titer Assay

This assay takes advantage of the fact that every cell infected with adenovirus expresses viral proteins. We can measure the percent of cells that are infected in a speciﬁc stock of virus by monitoring the number of cells expressing viral protein (15). The viral titers that can be obtained by using this assay are much quicker (within 48 h) than any other conventional assay. These results are very comparable to other titer protocols (15). The complete kit for such an assay can be purchased from Clontech (Adeno-X™ Rapid Titer Kit). This kit measures the production of viral hexon proteins in infected cells by immunohistochemical analysis. 1. Seed 1 × 104 healthy HEK 293 cells in each well of a 96-well plate for 24 h at 37°C. 2. Using media as a diluent, prepare tenfold serial dilution of viral stock from 10−2 to 10−8 ml. 3. Add viral dilution drop wise to well. Each dilution of virus should be assayed in triplicates to ensure accuracy. 4. Incubate cells at 37°C in 5% CO2 for 48 h. 5. After 48 h, aspirate the media and allow cells to dry in hood for 10 min. 6. Fix the cells by very gently adding ice-cold 100% methanol to each well. 7. Incubate the plate at −20°C for 10 min. 8. Aspirate methanol. Gently wash wells three times with PBS. At this point, cells can be stored at 4°C for 48 h in PBS before moving on to the next steps.

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9. Dilute the mouse anti-hexon antibody 1:1,500 in PBS containing 1% BSA. 10. Aspirate the PBS from ﬁnal wash. Then, add anti-hexon antibody dilution to each well. Incubate for 1 h at 37°C. 11. Remove the anti-hexon antibody. Then, gently rinse the wells three times with PBS. 12. Dilute the HRP-conjugated rat anti-mouse antibody 1:850 to each well. Incubate for 1 h at 37°C. 13. Before removing the rat anti-mouse HRP-conjugated antibody, prepare the DAB working solution by diluting 10× DAB substrate 1:10 with a 1× stable peroxidase buffer. Allow the DAB working solution to come to room temperature (see Note 5). 14. Aspirate the HRP-conjugated antibody. Gently rinse each well three times with PBS + 1% BSA. 15. Add the DAB working solution to each well. Incubate at room temperature for 10 min. 16. Observe under the microscope. All the adenovirus-infected cells should become brown. If one is having difﬁculties identifying the positive cells from brown staining, the cells can be incubated longer in the DAB working solution. 17. Count a minimum of three ﬁelds of brown and black positive cells using a microscope with a 20× objective, and calculate the mean number of positive cells in each well. 18. Calculate infectious units (IUs)/ml for each well as follows: ¤ Infected cells ³ ¤ Fields ³ ¥¦ ´µ r ¥¦ ´ Field Well µ . Volume virus (ml) r (Dilution factor) For 96-well plate (area = 0.32 cm2) and 20× objective (ﬁeld area = 0.64 mm2), ﬁelds/well = 50. For the 10× objective (ﬁeld area = 2.54 mm2), ﬁelds/well = 12.6. 3.5. Culture and Maintenance of NSC

3.5.1. Thawing Neural Stem Cell Lines

Human NSCs (ReNCell) were obtained from Millipore and maintained according to the manufacturer’s protocol. Brieﬂy, these NSCs were isolated from the cortical region of 14-week-old fetal tissue and immortalized by retroviral transduction and insertion of the c-myc gene. Cells were characterized according to their expression of nestin, SOX-2, CD133, and CD44 stem cell markers. Subcultures of human NSCs for experimentation were conducted as follows. 1. Coat the tissue culture plastic with laminin (Sigma) at a concentration of 20 Mg/ml in serum-free DMEM in 37°C and 5% CO2 atmospheric conditions 4 h before NSC plating. 2. Thaw a frozen vial as quickly as possible by placing the vial in 37°C water bath for 1–2 min.

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3. Immediately remove the vial from the water bath and decontaminate by wiping the vial with 70% ethanol. 4. Transfer the contents of the vile into 15-ml Falcon tube with 4 ml of ReNCell NSC Maintenance Medium, supplemented with 20 ng/ml bFGF and 20 ng/ml EGF (complete ReNCell medium). 5. Centrifuge at 246 r g for 5 min. Aspirate the medium from the tube and resuspend the cell pellet with 8 ml of complete ReNCell medium. 6. Pipet the medium containing NSCs into one laminin-coated 10-cm dish. Gently swirl dish to evenly distribute cells. 7. Place dishes into 37°C and 5% CO2 incubator. 3.5.2. Maintaining the Neural Stem Cell Lines

1. Precoat the tissue culture plastic dishes as described above with laminin (Sigma) 4 h before NSC plating. 2. Discard the culture medium by aspiration. Rinse the cell monolayer once with 1× PBS. 3. Detach the NSCs by adding 3 ml/10-cm dish of Accutase solution and incubate at 37°C for 2–3 min. Observe the cells under an inverted microscope until all the cells are detached from the plate (see Note 6). 4. Add 3 ml of complete medium and transfer the cells by gently pipetting into 15-ml Falcon tube. 5. Centrifuge the cell solution at 246 u g, resuspend the pellet in the 3-ml complete ReNCell medium, and transfer 1 ml into laminin-precoated 10-cm dish (see Notes 7 and 8).

3.6. Ex Vivo Optimization of the Therapeutic Virus Loading into Carrier Cells

The goal of the ex vivo loading phase is not only to infect as many carrier cells as possible, but also to keep them alive post in vivo delivery. This is so the cells can home in on the tumor and produce a high amount of therapeutic virus. This part of the chapter brieﬂy describes the optimization experiments of these important parameters in the in vitro setting.

3.6.1. Cell Viability Analysis of the Cell Carrier

In order to determine the maximum ex vivo virus loading dose that has minimal impact on the survival of the carrier cell: 1. Plate 1 × 104 cells/well NSCs in a 96-well plate. 2. On the following day, infect the cell monolayer with 0.01– 1,000 IU/cells (tenfold dilution) of the oncolytic virus. Infect each IU in triplicate wells. 3. After 2 h of incubation, remove the virus-containing media, wash the cell two times with PBS, and add fresh complete ReNCell medium. 4. 72 h and 96 h post infection, count the viable cells in each infected condition by using Trypan blue exclusion method.

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1. Transfer medium from the infected well to 1.5-ml Eppendorf tubes. Detach the cell monolayer by adding 50 Ml Accutase solution. 2. Transfer the detached cells from each well to the corresponding 1.5-ml Eppendorf tube. 3. Mix 10 Ml of cell suspension with 10 Ml of trypan blue solution. 4. Transfer immediately to the counting chamber of the hemocytometer. 5. Viable cells exclude trypan blue while dead cells stain blue due to trypan blue uptake. Count the viable cells in three squares in the counting areas of the hemocytometer. Average the three counts. 6. Calculate the total number of viable cells using the following formula: Viable cells (cells/ml) = Average count × dilution factor (2) × 104

3.6.3. Dose–Response Analysis of Viral Replication and Progeny Release

1. Plate carrier cells at a density of 2.5 × 104 cells/well in 24-well laminin-precoat tissue culture plate. 2. On the following day, infect the cell monolayer with oncolytic adenovirus as described above (Subheading 3.6.1, step 2). 3. 72 h post infection, collect the supernatant from the infected monolayer. Detach the infected cells by Accutase treatment, neutralize with the complete medium, and transfer the cell suspension in a 1.5-ml Eppendorf tube. Centrifuge the cell solution at 1,000 × g and resuspend the pellet in the 50 Ml PBS. 4. Release the virus by snap freezing the cell plate in liquid nitrogen and thaw in the 37°C water bath. Repeat this process three times. Transfer the freeze/thaw cells into tabletop centrifuge tubes and spin out the cell debris at 6,172 r g for 10 min. 5. Measure the virus titer in the supernatants (progeny released by the carrier cell) and in the cell plate (viral replication in the carrier cell) by using the protocol for rapid titer assay described above.

3.6.4. Preparing Neural Stem Cells for Ex Vivo Loading

1. Remove the culture medium by aspiration. Rinse the cell monolayer once with 1× PBS. 2. Detach the NSCs by adding 3 ml Accutase solution/10-cm to the dish of Accutase solution and incubate at 37°C for 2–3 min. 3. Add an equal volume of ReNCell complete medium and transfer the cell suspension into 15-ml Falcon tube. 4. Measure the cell concentration in the cell suspension by hemocytometer. 5. Centrifuge the cell solution at 1,000 × g. Remove the medium from the tube by aspiration and wash the pellet with PBS two times. 6. After the second wash, resuspend the NSCs in PBS at the appropriate cell concentration (see Note 9).

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7. Add the proper dose of oncolytic virus in the NSC suspension and gently triturate the cells. Incubate at room temperature for 2 h. 8. Centrifuge the cell solution at 1,000 × g and wash the pellet three times with 1× PBS. 9. After the last spin, remove the supernatant by aspiration and resuspend the pellet in PBS at the appropriate cell concentration. 10. Place virus-loaded NSCs on ice until ready to use (see Note 10).

4. Notes 1. Maintenance medium should be stored at −20°C until ready to use. Upon thawing, the maintenance medium should be stored at 2–8°C and given a 1-month expiration date. 2. All 150-mm dishes do not have to be prepared concurrently. It is often convenient to prepare ten dishes at time. 3. The contaminants, such as unpackaged virus, may have a similar density to that of the mature virion. This can cause the distance between the two bands to be very small after the CsCl gradient separation. In the large-diameter tube, such as 30-ml tube, bands of similar density of virus appear thinner and further separated. Thus, the recovery of viral bands is easier in the 30-ml tube compared to small-volume tube. 4. The area between the defective (upper band) and infectious viral particles (lower band) may appear turbid. Avoid removing this turbid area. 5. Do not allow the 10× DAB substrate to warm to room temperature. 6. Do not keep the NSCs in the Accutase solution for more than 5 min. The dish can be gently tapped to help the cells detach. 7. Initially, the NSCs grow rapidly, doubling 8–10 h in the presence of FGF and EGF. Never let cells reach conﬂuence as it causes the cells to differentiate. If the cells split 1:2 or 1:3 ratio from 70 to 80% conﬂuent dish, cells need to split 2–3 times a week. After 25–30 passages, the NSCs start to slow down. 8. The most common problem for passaging NSC is the fact that cells appear healthy when plated, but die overnight or fail to grow due to the residual Accutase solution. Care needs to be taken to avoid this. 9. In order to achieve optimum ex vivo loading, the carrier cells need to incubate with the oncolytic virus in a minimum volume (5 × 106 carrier cells in 100-Ml volume). 10. Prior to in vivo injection, cells can be maintained on ice for 1–2 h.

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References 1. Hanahan, D., and Weinberg, R. A. (2000) The hallmarks of cancer, Cell 100, 57–70. 2. Kelly, E., and Russell, S. J. (2007) History of oncolytic viruses: genesis to genetic engineering, Mol Ther 15, 651–659. 3. Guo, Z. S., Thorne, S. H., and Bartlett, D. L. (2008) Oncolytic virotherapy: molecular targets in tumor-selective replication and carrier cell-mediated delivery of oncolytic viruses, Biochimica et biophysica acta 1785, 217–231. 4. Lesniak, M. S. (2007) Advances in neurooncology: novel therapies and clinical trials, Expert review of anticancer therapy 7, S1. 5. Lesniak, M. S. (2005) Brain tumors: controversies and challenges in management, Expert review of neurotherapeutics 5, 1–2. 6. Lesniak, M. S. (2006) Gene therapy for malignant glioma, Expert review of neurotherapeutics 6, 479–488. 7. Parker, J. N., Bauer, D. F., Cody, J. J., and Markert, J. M. (2009) Oncolytic viral therapy of malignant glioma, Neurotherapeutics 6, 558–569. 8. Fisher, K. (2006) Striking out at disseminated metastases: the systemic delivery of oncolytic viruses, Current opinion in molecular therapeutics 8, 301–313. 9. Brustle, O., Spiro, A. C., Karram, K., Choudhary, K., Okabe, S., and McKay, R. D. (1997) In vitro-generated neural precursors participate in mammalian brain development, Proceedings of the National Academy of Sciences of the United States of America 94, 14809–14814.

10. Aboody, K. S., Brown, A., Rainov, N. G., Bower, K. A., Liu, S., Yang, W., Small, J. E., Herrlinger, U., Ourednik, V., Black, P. M., Breakeﬁeld, X. O., and Snyder, E. Y. (2000) Neural stem cells display extensive tropism for pathology in adult brain: evidence from intracranial gliomas, Proceedings of the National Academy of Sciences of the United States of America 97, 12846–12851. 11. Benedetti, S., Pirola, B., Pollo, B., Magrassi, L., Bruzzone, M. G., Rigamonti, D., Galli, R., Selleri, S., Di Meco, F., De Fraja, C., Vescovi, A., Cattaneo, E., and Finocchiaro, G. (2000) Gene therapy of experimental brain tumors using neural progenitor cells, Nature medicine 6, 447–450. 12. Tyler, M. A., Ulasov, I. V., Sonabend, A. M., Nandi, S., Han, Y., Marler, S., Roth, J., and Lesniak, M. S. (2009) Neural stem cells target intracranial glioma to deliver an oncolytic adenovirus in vivo, Gene therapy 16, 262–278. 13. Aboody, K. S., Najbauer, J., and Danks, M. K. (2008) Stem and progenitor cell-mediated tumor selective gene therapy, Gene therapy 15, 739–752. 14. Coukos, G., Makrigiannakis, A., Kang, E. H., Caparelli, D., Benjamin, I., Kaiser, L. R., Rubin, S. C., Albelda, S. M., and Molnar-Kimber, K. L. (1999) Use of carrier cells to deliver a replication-selective herpes simplex virus-1 mutant for the intraperitoneal therapy of epithelial ovarian cancer, Clin Cancer Res 5, 1523–1537. 15. Bewig, B., and Schmidt, W. E. (2000) Accelerated titering of adenoviruses, BioTechniques 28, 870–873.

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Chapter 9 Targeting Brain Tumor Stem Cells with Oncolytic Adenoviruses Marta M. Alonso, Hong Jiang, Candelaria Gomez-Manzano, and Juan Fueyo Abstract In 2004, brain tumor stem cells (BTSCs) were isolated from surgical human malignant gliomas. This cancer cell population has been identiﬁed as the root for tumor initiation and resistance to therapies. Thus, it is imperative to develop new therapies that can eradicate this subpopulation to improve the prognosis of patients with brain tumors. Our group previously reported the antiglioma effect of the tumor-selective oncolytic adenovirus Delta-24-RGD that is now being tested in a phase I clinical trial for patients with malignant gliomas. We also showed that Delta-24-RGD infects, replicates in, and induces cell death in BTSCs. Interestingly, we observed that adenoviral-infected cells undergo autophagy and that autophagyrelated cytoplasmic vacuolization might be part of the lysis process. Here, we summarize the materials and methods used in our study as follows: establishment of neurosphere cultures from surgical samples of human glioblastoma multiformes; assessment of stem cell markers; examination of adenoviral receptors in BTSCs; evaluation of the cytotoxicity induced by oncolytic adenoviruses; and assessment of autophagy in oncolytic adenovirus-infected BTSCs in vitro, and ﬁnally we describe a method to detect upregulation of the autophagy-related protein Atg5 in tumors treated with Delta-24-RGD. Key words: Brain tumors, Brain tumor stem cells, Glioma, Oncolytic adenovirus, Delta-24-RGD, Autophagy

1. Introduction The most common primary brain tumor in adults is glioblastoma multiforme (GBM). GBMs remain universally fatal and, despite the use of maximal therapy, the median survival duration is slightly more than 1 year (1). These dismal survival statistics underscore the need for new therapeutic approaches for GBM. The ﬁnding of brain tumor stem cells (BTSCs), the putative subpopulation causing the

David H. Kirn et al. (eds.), Oncolytic Viruses: Methods and Protocols, Methods in Molecular Biology, vol. 797, DOI 10.1007/978-1-61779-340-0_9, © Springer Science+Business Media, LLC 2012

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resistance of these tumors to conventional therapies, was made by applying principles of cell culture and analysis of normal neural stem cells (NSCs) and identifying cell surface markers that allow for isolation of distinct tumor cell populations that can then be studied in vitro and in vivo (2, 3). BTSCs can be enriched from a population of brain tumor cells by sorting dissociated suspensions of tumor cells for those expressing the NSC marker CD133 (4). CD133+ cells, which also express the NSC marker nestin but not differentiated neural lineage markers, represent a minority of the entire brain tumor cell population, exclusively generate clonal tumor spheres in suspension culture, and exhibit increased self-renewal capacity (3). In addition, implantation of human BTSCs into the brain of immunocompromised animal model results in the formation of tumors that phenotypically resemble the original primary tumor (3). The identiﬁcation of BTSCs provides a powerful tool for investigating the tumorigenic process in the central nervous system and for developing therapies targeted speciﬁcally to this compartment. Due to the tumor-initiating properties of BTSCs, therapies that destroy, induce differentiation, and/or prevent the invasion/metastasis of cancer stem cells are potentially curative. An approach utilized to design novel, rational, effective treatments of GBM is based in the use of oncolytic adenoviruses. We previously reported the antiglioma effect of the tumor-selective adenovirus Delta-24-RGD, the genome which encompasses a 24-bp deletion in the E1A gene region encoding the amino acids responsible for binding Rb protein (5) and displays enhanced infectivity through the insertion of an RGD-4C motif to the ﬁber HI loop (6). Delta-24-RGD has proven to be highly effective in inducing the suppression of brain tumors in mice. Of importance, our group also showed that Delta-24-RGD is an efﬁcacious therapeutic agent against BTSCs. Interestingly, Delta-24-RGD elicited autophagic cell death in this cancer subpopulation. Autophagy has been proposed as the basis for the mechanism of cell death during adenoviral infection (7). In agreement with this hypothesis, we showed that adenoviral infection is followed by upregulation of the levels of a critical protein in the autophagy pathway, Atg5, in host cells (8, 9). In addition, in Delta24-RGD-treated BTSC-derived xenografts, we observed Atg5 overexpression and colocalization with adenoviral proteins (9) suggesting that increased levels of Atg5 could be used as a surrogate marker of adenovirus activity in human gliomas treated with oncolytic adenoviruses in future clinical trials. In this review, we summarize the material and methods used in our studies. It includes the establishment of neurosphere cultures from surgical samples of human GBMs; assessment of stem cell markers through polymerase chain reaction (PCR); examination of adenoviral receptors in BTSCs via ﬂow cytometry; evaluation of the cytotoxicity induced by oncolytic adenoviruses; and assessment of autophagy in oncolytic adenovirus-infected BTSCs in vitro and in vivo.

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2. Materials 2.1. Establishment of Neurosphere Cultures from Surgical Samples of Human GBMs

1. Artiﬁcial cerebrospinal ﬂuid (CSF): 125 mM NaCl, 5 mM KCL, 1.3 mM MgCl2, 2 mM CaCl2, 26 mM NaHCO3, 10 mM D-glucose. It should be stored at 4°C. 2. Low Ca2+ CSF: 125 mM NaCl, 5 mM KCl, 3.2 mM MgCl2, 0.1 mM CaCl2, 26 mM NaHCO3, 10 mM D-glucose. It should be stored at 4°C. 3. Collagenase (Sigma) is dissolved at 1% in phosphate-buffered saline (PBS) and stored at −20°C. 4. Hyaluronidase (Sigma) is dissolved at 2% in PBS and stored at −20°C. 5. Trypsin/ethylenediaminetetraacetic acid (EDTA) solution (GIBCO Invitrogen): 1×. It should be stored at 4°C. 6. 70-Mm cell strainer (BD Biosciences). 7. Solution of 0.9 M sucrose in 0.5× Hanks’ balanced salt solution. It should be stored at 4°C. 8. Red blood cell (RBC) lysis buffer: Dissolve 8.26 g of NH4Cl, 1 g of KHCO3, and 0.037 g of EDTA in 1 L of deionized distilled water (ddH2O). Mix well and autoclave. Store up to 6 months at room temperature or 4°C. RBC lysis buffer should always be used at room temperature. 9. Tris–EDTA (TE) buffer: 10 mM Tris–Cl, pH 7.5, 1 mM EDTA. 10. Trypsin inhibitor solution (Sigma): 10 mg/mL in sterile water with 0.9% NaCl and 0.9% benzyl alcohol, pH 5.7–6.2. It should be stored at 4°C. 11. Dulbecco’s modiﬁed Eagle’s medium (DMEM) supplemented with F-12 medium (GIBCO Invitrogen). 12. Neuro-stem cell medium: DMEM/F-12 medium, B27 supplement. Add fresh epidermal growth factor (EGF; 20 ng/ mL) and basic ﬁbroblast growth factor (bFGF; 20 ng/mL) at every preparation. EGF is dissolved in 10% bovine serum albumin (BSA; 50 ng/mL), and bFGF is dissolved in 5 mM Tris–Cl, pH 7.0 (50 ng/mL), and stored at −80°C.

2.2. Assessment of Stem Cell Markers Using PCR

1. RNeasy total RNA isolation kit (Qiagen). 2. Ultraviolet spectrophotometer for RNA quantiﬁcation. 3. Materials required for cDNA synthesis: (a) 200 ng RNA. (b) Thermocycler. (c) 10 mM dNTP mix: 10 mM each dATP, dCTP, dGTP, and dTTP.

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(d) 50 MM Oligo(dt) primer (Invitrogen). (e) SuperScript III First-Strand Synthesis System for reverse transcriptase (RT)-PCR (Invitrogen). (f) Water: PCR grade. (g) 0.2-mL thin-walled PCR tubes. 4. Materials required for PCR: (a) Input cDNA from the previous synthesis. (b) Speciﬁc primers for the genes of interest (see Subheading 3.2 for the primer sequences). (c) 10 mM dNTP mix: 10 mM each dATP, dCTP, dGTP, and dTTP. (d) Taq DNA polymerase (Roche Applied Science). (e) Water: PCR grade. (f) 0.2-mL thin-walled PCR tubes. (g) Sterile reaction tubes for preparing master mixes and dilutions. (h) Thermocycler. 5. Agarose, Type I: Low EEO (Sigma). 6. Tris–acetate–EDTA (TAE) running buffer (1×): Dissolve the following in 1 L of ddH2O: 48.4 g of Tris base, 10.9 g of glacial acetic acid, and 2.92 g of EDTA. Store at room temperature. 7. 100 bp molecular weight marker (Invitrogen). 8. Mini agarose gel electrophoresis unit. 2.3. Examination of Adenoviral Receptors in BTSCs Using Flow Cytometry

1. BTSCs. 2. Neuro-stem cell medium. 3. 400–600 U/mL Accutase solution (Sigma–Aldrich). 4. PBS. 5. Antibody binding buffer: Prepare a solution containing PBS, pH 7.2, 0.5% BSA, and 2 mM EDTA. Keep buffer cold (4–8°C). 6. Human FcR blocking reagent (Miltenyi Biotec) to prevent FcR-mediated antibody labeling. 7. 5 Mg/mL propidium iodide for the exclusion of dead cells (Sigma) without cell ﬁxation. 8. Mouse monoclonal anti-integrin-AvB3 antibody (Chemicon International): 1:200 dilution. 9. Mouse monoclonal anti-integrin-AvB5 antibody (Chemicon International): 1:400 dilution. 10. Mouse monoclonal anti-coxsakie adenoviral receptor (CAR) antibody (Upstate Biotechnology): 1:2,000 dilution.

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11. Fluorescein isothiocyanate-labeled goat anti-mouse IgG secondary antibody (Santa Cruz Biotechnology): 1:100 dilution. 12. FACSCalibur (BD Biosciences). 2.4. Evaluation of the Cytotoxicity Induced by Oncolytic Adenoviruses

1. Oncolytic adenovirus: Store at −80°C. 2. BTSCs. 3. Neuro-Stem cell medium. 4. Trypsin, 0.5% (10×) with EDTA 4Na, liquid (Invitrogen). 5. 10 mg/ml soybean trypsin inhibitor (10×, Sigma). 6. Flat-bottom, 96-well plates. 7. CellTiter 96® AQueous One Solution Cell Proliferation Assay (MTS) (Promega Life Science). 8. Plate reader.

2.5. Assessment of Autophagy in Oncolytic AdenovirusInfected BTSCs In Vivo and In Vitro 2.5.1. Induction of Acidic Vesicular Organelles

1. Oncolytic adenovirus: Store at −80°C. 2. BTSCs. 3. Neuro-Stem cell medium. 4. 400–600 U/mL Accutase solution (Sigma–Aldrich). 5. Six-well plates. 6. 10 mg/mL acridine orange solution (Sigma) (in ddH2O). 7. PBS. 8. FACSCalibur (BD Biosciences).

2.5.2. Electron Microscopy

1. Oncolytic adenovirus: Store at −80°C. 2. BTSCs. 3. Neuro-Stem cell medium. 4. Glass coverslips. 5. Fixing solution containing 3% glutaraldehyde plus 2% paraformaldehyde in 0.1 M cacodylate buffer, pH 7.3. 6. OsO4: 1% in 0.1 M cacodylate buffer, pH 7.3. 7. 1% Millipore-ﬁltered uranyl acetate (UA). 8. 100, 95, and 70% ethanol. 9. Spurr’s low-viscosity medium. 10. Lead citrate. 11. Ultracut microtome (Leica) stained with uranyl acetate and lead citrate using a Leica EM stainer. 12. Electron microscope (JEM 1010 transmission electron microscope; JEOL).

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2.5.3. Immunoﬂuorescent Analysis of Adenoviral Fiber and Atg5 Protein Expression in the Brain of Mice Bearing BTSC Xenografts

1. Parafﬁn slides containing BTSCs previously infected with an oncolytic adenoviral vector. 2. Mouse monoclonal antibodies speciﬁc for adenoviral ﬁber protein (4D2; Lab Vision): 1:500 dilution. 3. Rabbit polyclonal anti-Atg5 antibody (Cosmo Bio Co., Ltd.): 1:200 dilution. 4. Texas Red (ﬁber)- or Alexa Fluor 488 (Atg5)-conjugated secondary antibodies (Molecular Probes): 1:500 dilution. 5. PBS. 6. Xylene. 7. 95 and 70% alcohol. 8. Distilled water. 9. 10% goat serum in PBS. 10. 0.0 l M citrate buffer, pH 6.0. 11. “Auto buffer”: 0.01 M Tris–Cl, pH 7.4, 1 mM CaCl2, 1 mM MgCl2, 0.1% Triton X-100. 12. 1% BSA in auto buffer. 13. Steamer (used for antigen retrieval). 14. PAP Pen (Kiyota International). 15. ProLong Antifade Kit (#P-7481; Molecular Probes).

3. Methods 3.1. Establishment of Neurosphere Cultures from Surgical Samples of Human GBMs

1. The samples are processed right after surgery. Cut tumors into small pieces using a scalpel (see Note 1). 2. Incubate tissue pieces in 5 mL of CSF for 10 min. 3. Spin down tissue samples at 239 r g (microcentrifuge) for 5 min and aspirate the supernatant. 4. Wash the tissue once with low Ca2+ CSF (add CSF at a volume sufﬁcient to cover the tissue). 5. Spin down tissue samples at 1,500 rpm (microcentrifuge) for 5 min. Discard supernatant. 6. Add 4.25 mL of low Ca2+ CSF plus 500 ML of 1% collagenase and 250 ML of 2% hyaluronidase and incubate tissue for 1 h at 37°C (see Note 2). 7. Spin down samples at 1,500 rpm (microcentrifuge) for 5 min and discard the supernatant. 8. Add 2–5 mL of TE buffer to the pellet and incubate for 10 min at room temperature.

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9. Add an equal volume of trypsin inhibitor to TE buffer-treated tissue samples. 10. Immediately add DMEM/F-12 medium to enzyme-treated tissue and dissociate cells by pipetting up and down. 11. Place the strainer in a 50-mL tube and then pass cells through the strainer to achieve a single cell suspension. 12. Spin down these cells at 1,500 rpm (microcentrifuge) for 5 min and aspirate the supernatant. 13. Resuspend cells in 5–10 mL of a 0.9 M sucrose solution. 14. Centrifuge cells at 664 r g (microcentrifuge) for 20 min (at this point, if a great deal of white debris remains, repeat steps 13 and 14). 15. Wash cells with 5–10 mL of DMEM/F-12 medium twice and add RBC lysis buffer to the cells. 16. Incubate the cells at room temperature for 5 min to allow the RBC lysis buffer to work and spin down cells at 1,500 rpm (microcentrifuge) for 5 min. Discard the supernatant. 17. Wash cells three times with DMEM/F-12 medium. Discard the supernatant. 18. Add 10 mL of Neuro-Stem cell medium to the cells and place them in a 100-mm cell culture dish. Allow at least 10 days for the cells to grow (see Note 3). The cells grow as neurosphere in neuro-stem cell medium (Fig. 1).

Fig. 1. Neurosphere cultures were established from acute cell dissociation of human glioblastoma multiforme surgical specimens and maintained in neuro-stem cell medium. Note that the cells grow as spheroids. This ﬁgure has been modiﬁed from a previous publication (9).

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3.2. Assessment of Stem Cell Markers Using PCR

1. Once the BTSC cultures are established, extraction of the RNA can proceed and markers of stemness can be examined. 2. To obtain a sufﬁcient yield of RNA, at least 2–3 × 105 cells should be collected. 3. Collect cells and spin them down at 1,000 rpm (microcentrifuge) for 1 min. Discard the supernatant. 4. Wash the cells twice with PBS. It is important to aspirate as much medium as possible (without disturbing the cell pellet; see Note 4). 5. Proceed to extracting the RNA using the RNeasy kit following the manufacturer’s instructions. 6. Measure the RNA at 260/280 nm using the spectrophotometer. Once the RNA concentration is calculated, cDNA synthesis can proceed. 7. For each cell line, prepare a 0.2-mL tube containing 200 ng of the sample RNA. Higher concentrations of RNA can be used and diluted accordingly with ddH2O at the end of the synthesis. 8. Add 1 ML of the 50-MM oligo(dt) primer and 1 ML of 10 mM dNTPs and bring the mixture to a ﬁnal volume of 13 ML. 9. Incubate the mixture at 65°C for 5 min and then place on ice for at least 1 min. 10. Add 4 ML of 5× RT buffer and 2 ML of 0.1 M dithiothreitol to the mixture. 11. Incubate the mixture at 42°C for 2 min and then place it on ice and collect sample using brief centrifugation. 12. Add 1 ML of the SuperScript II RT enzyme to the mixture and mix gently. 13. Incubate the mixture at 42°C for 50 min. 14. Terminate the reactions by incubating the samples at 85°C for 5 min. Chill on ice. We recommend using 10% of the volume of the ﬁrst-strand reaction (2 ML) as template for following PCR (see Note 5). 15. Prepare a master mix for each gene of interest, including one for the housekeeping gene. The volume of each master mix typically should be 110% of the volume required for all of the samples. The extra volume allows for loss of sample volume during pipetting. The master mix includes the following (25-ML ﬁnal volume per sample): (a) Reverse primer (10 MM): 0.5 ML (b) Forward primer (10 MM): 0.5 ML (c) dNTPS (10 mM): 0.25 ML (d) PCR buffer (10×): 2.5 ML

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(e) Taq polymerase enzyme: 0.25 ML (f ) Water: 19 ML 16. Dispense 23 ML of the master mix into each cDNA sample and place it in the thermocycler. 17. After cycling, store the samples at −20°C for later use if needed. 18. Check the PCR product in a 2% agarose gel for size and speciﬁcity. For instance, a 50-mL gel requires 50 mL of TAE buffer and 1 g of agarose. Dissolve the agarose in the TAE buffer, bringing it to a boil in a microwave. Let it cool down and add 0.5 Mg/mL ethidium bromide to the mixture before pouring the gel into the casting device. 19. Once the gel has solidiﬁed, place it in the electrophoresis tank. 20. Load the samples in the gel. Use an appropriately sized marker; in this case, a 100-bp marker is appropriate because the fragments being ampliﬁed range from 100 to 400 bp. 21. Run the gel at 5 V/cm for approximately 1 h. Samples should be checked to optimize time and voltage. 22. Visualize the PCR product using a transilluminator (see Note 6). Primer Sequences

3.3. Examination of Adenoviral Receptors in BTSCs

CD133-F

TGGATGCAGAACTTGACAACGT

CD133-R

ATACCTGCTACGACAGTCGTGGT

Nestin-F

GGCAGCGTTGGAACAGAGGT

Nestin-R

CATCTTGAGGTGCGCCAGCT

Bmi1-F

AGCAGAAATGCATCGAACAA

Bmi1-R

CCTAACCAGATGAAGTTGCTGA

Sox2-F

CCCCCGGCGGCAATAGCA

Sox2-R

TCGGCGCCGGGGAGATACAT

GAPDH-F

AGCTGAACGGGAAGCTCACT

GAPDH-R

TGCTGTAGCCAAATTCGTTG

1. Collect BTSCs by spinning them down at 1,000 rpm (microcentrifuge) for 1 min. 2. Discard the supernatant and add 500 ML of Accutase to the pellet. Pipette the pellet up and down and place cells in the incubator at 37°C for 5 min. 3. Add 4 mL of Neuro-Stem cell medium to the pellet and, after pipetting the cells up and down at least ﬁve times, pass them through a strainer. This step ensures single cell suspension (see Note 7).

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4. Resuspend up to 106 nucleated cells in 80 ML of antibody-binding buffer. 5. Add 20 ML of FcR blocking reagent to the cells. 6. Add antihuman integrin mouse AvB5 antibody (1:400 dilution), antihuman integrin mouse AvB3 antibody (1:200 dilution), or anti-CAR antibody (1:2,000 dilution) to the cells at 4°C for 1 h. 7. Wash the cells once with 1 mL of PBS. 8. Incubate cells with the ﬂuorescein isothiocyanate-labeled goat anti-mouse IgG secondary antibody (1:100 dilution) at room temperature for 1 h (see Note 8). 9. Wash the cells twice with PBS and stain them with propidium iodide at 25 Mg/mL for 15 min at room temperature. 10. Resuspend the samples in 0.5–1.0 mL of PBS (depending on the number of cells, fewer than one million cells for a 0.5-mL volume is recommended). 11. Analyze cell samples using a FACScan cytometer (Becton Dickinson) with a 488-nm argon laser for excitation. Fluorescence is detected using a 520-nm band-pass ﬁlter, and all cytometric data are analyzed using CellQuest software (Becton Dickinson). 3.4. Evaluation of the Cytotoxicity Induced by Oncolytic Adenoviruses

1. Dissociate BTSCs and plate them in 96-well plates at 2 × 104 cells/50 ML/well in triplicate. 2. Cells should be immediately infected with oncolytic adenoviruses at a range of desired doses in a volume of 50 ML. The dose range allows establishment of indexes, such as the 10, 50, and 100% inhibitory concentrations (IC10, IC50, IC100). The ﬁnal volume after infection should be 100 ML. 3. Incubate the cells for 6 days. 4. On the sixth day, add 10 ML/well 10× trypsin with EDTA to the cells and incubate at 37°C for 15 min. 5. Add 10 ML/well 10× trypsin inhibitor and pipette the cells ten times (see Note 9). 6. Add 22 ML/well MTS reagent to the cells and incubate at 37°C for 4 h. 7. Read the plate with a plate reader. 8. Data can be shown as the percentage of viable cells in treated samples relative to that of mock-treated cells (equal to 100%).

3.5. Assessment of Autophagy in Oncolytic AdenovirusInfected BTSCs In Vivo and In Vitro

As mentioned above, several complementary techniques can be used to monitor adenovirally induced autophagy:

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Autophagy is characterized by the development of acidic vesicular organelles (10, 11). The cytoplasm and nucleoli of acridine orangestained cells ﬂuoresce bright green and dim red, respectively, whereas acidic compartments ﬂuoresce bright red (11). Therefore, autophagy can be assessed by quantifying acidic vesicular organelles in the cells using supravital cell staining with acridine orange as described below. 1. Dissociate BTSC spheroids using Accutase as described previously and immediately infect the cells with adenoviruses. Dose depends on adenoviral infectivity of used cell line. 2. Seventy-two hours later, dissociate the cells using Accutase. 3. Add acridine orange (1 Mg/mL; Sigma–Aldrich) to the medium for 15 min (see Note 10). 4. Collect the cells using centrifugation at 1,000 rpm (microcentrifuge) for 1 min. 5. Wash the cells with PBS. 6. Analyze the cells with a ﬂow cytometer. Green (510–530 nm, FL1-H channel) and red (>650 nm, FL3-H channel) ﬂuorescence emissions from 104 cells illuminated with blue (488 nm) excitation light are measured with a FACSCalibur from BD Biosciences using CellQuest software (San Jose, CA, USA). Control (untreated) cells are used to set the parameters for analysis of this cell sample. Place a horizontal cursor across the sample to achieve an arbitrary value above the cursor of ±5% of AVOs in the control cells. Anything above this cursor in the test sample is considered percentage of AVOs and thus indicative of the existence of an autophagic process (see Note 11).

3.5.2. Electron Microscopy

To date, electron microscopy has been the only reliable method of monitoring autophagy. The autophagy phenomenon is identiﬁed by the existence of autophagosomes and autolysosomes. The autophagosome is a double-membrane structure containing undigested cytoplasmic material, including organelles, whereas the autolysosome is a single-membrane structure containing cytoplasmic components at various stages of degradation. Because clear differentiation between autophagosomes and autolysosomes is sometimes difﬁcult, these structures are often generalized as “autophagic vacuoles.” Importantly, double-membrane structures containing cytoplasmic components are not always autophagosomes. For example, when a portion of a cell protrudes into the next cell as with a hernia and the protrusion is cut perpendicular to the axis, it can appear to be a double-membrane structure. Thus, conventional electron microscopic images of autophagic samples must be assessed carefully. 1. Allow neurospheres to grow on glass coverslips and infect them with Ad-DeltaE1 (a replication-incompetent adenovirus that

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serves as a negative control) or Delta-24-RGD at 10 pfu/cell for 72 h (dose depends on the cell line used). 2. Fix the cells in a solution containing 3% glutaraldehyde plus 2% paraformaldehyde in 0.1 M cacodylate buffer (pH 7.3) for 1 h (see Note 12). 3. Postﬁx the samples in 1% OsO4 in the same buffer for 1 h. 4. Stain the en bloc with 1% Millipore-ﬁltered uranyl acetate (see Note 13). 5. Dehydrate samples in ethanol at increasing concentrations and then inﬁltrate and embed them in Spurr’s low-viscosity medium. 6. At this point, the samples are polymerized in a 70°C oven for 2 days. 7. Cut ultrathin sections of the samples using a Leica Ultracut microtome and stain them with uranyl acetate and lead citrate using a Leica EM stainer (see Note 14). 8. Examine the sections using a JEM 1010 transmission electron microscope (JEOL, USA, Inc.) at an accelerating voltage of 80 kV. Digital images of the sections are obtained using the AMT Imaging System (Advanced Microscopy Techniques) (Fig. 2).

Fig. 2. Electron microscopy photographs showing (a) mock-infected and (b) Delta-24-RGDinfected BTSCs. Please note that cytoplasmic vacuoles are present in the virus-infected cells but not in the mock-infected cells. Close-up images of the Delta-24-RGD-infected cells in (b) show (c) a cluster of Delta-24-RGD virions (white arrow) in the nucleus and (d ) complex autophagic vacuolar bodies in the cytoplasm. This ﬁgure has been modiﬁed from a previous publication (9).

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All the steps in this procedure are performed at room temperature in a light-protected humidity box unless otherwise noted (see Note 15). 1. Deparafﬁnize the slides containing the BTSC xenografts (untreated and treated with the adenovirus) by immersing them in xylene twice for 5 min with occasional agitation. Rehydrate the slides by passing them through a graded series of ethanol for 5 min each in the following order: 95, 70, 50, and 25%. Wash the slides for 2 × 5 min in PBS. Keeping the sections from drying out during the immunohistochemical procedure is very important, as a high background of autoﬂuorescence may result. 2. Place the slides in a slide rack in a Coplin jar and cover the slides with 0.01 M citrate buffer. Place the slides in the steamer for 5 min. Once antigen retrieval is complete, remove the slides from the steamer and allow them to cool for 20 min at room temperature. Wash the slides with distilled water for 5 min. 3. Remove each slide one at a time and label and circle the sections using a PAP pen to minimize the number of reagents needed for the procedure. Immediately after circling the sections, place the slides in the humidity box and cover the sections with PBS. 4. Prepare 10% normal goat serum by diluting normal goat serum into 1% BSA in auto buffer. Incubate the sections in 10% goat serum for 30 min (see Note 16). 5. Prepare a 1:500 dilution of monoclonal anti-adenoviral ﬁber (4D2, Lab Vision, Fremont, CA) and a 1:200 dilution of the rabbit polyclonal ATG5 antibody (Cosmo Bio Co., LTD), diluting them into 1% BSA in auto buffer. Gently tap the normal goat serum from each section and replace it with the primary antibody. Apply two drops of mouse and rabbit isotype control antibodies to sections designated as negative controls. Incubate the slides overnight at 4°C. 6. Wash the slides in PBS three times for 2 min. Prepare 10% goat serum by diluting normal goat serum into 1% BSA in auto buffer. Incubate the samples for 30 min. This step appears to reduce background ﬂuorescence. 7. Prepare the secondary antibody by diluting Texas Red (ﬁber)- or Alexa Fluor 488 (Atg5)-conjugated secondary antibodies into 10% normal goat serum prepared with 1% BSA/auto buffer at 1:500. Incubate the slides for 30 min with a secondary antibody and then wash them three times for 2 min in PBS. 8. Cover the slides with ProLong antifade mounting medium according to the manufacturer’s directions, taking care to remove any bubbles formed beneath the coverslips. Although the slides can be viewed immediately, the ProLong data sheet states that antifade protection is highest after the mounting medium is allowed to dry overnight (see Note 17).

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4. Notes 1. Smaller fragments of tissue dissociate better and yield a higher number of neurospheres. 2. Check frequently the status of the tissue dissociation. The time required for tissue dissociation depends on the size of the tumor fragments. In addition, cell viability is reduced with prolonged digestion. 3. Plating the cells at a higher density facilitates a faster generation of neurospheres. The interval between subcultures is often determined by the growth rate of the neurospheres. It is important not to allow the neurospheres achieve a big size because cell viability may be impacted in successive subcultures. 4. Successful RNA isolation requires fast processing and careful handling of the tissue or cells prior to isolation. Be extremely cautious with the washings to avoid excessive losing of cells. 5. At this moment, the samples can be stored at 4 or −20°C for a later use. 6. This technique is very useful to detect signiﬁcant differences in expression levels. However, if you want to quantify or detect subtle differences in expression, we recommend the use of quantitative RT-PCR. 7. It is crucial to obtain a single cell suspension in order to achieve a homogeneous labeling of the cells. In addition, it ensures that the cells ﬂow properly through the cytometer. 8. We highly recommend the use of primary antibodies already labeled with your ﬂuorochrome of choice. It saves you an extra step, which means saving time and the possibility to introduce more variations and errors. 9. It is necessary to dissociate the neurospheres at this time because MTS is not able to penetrate into the spheres efﬁciently, resulting in underestimation of the cell viability. 10. Although some degree of variability in the labeling might be observed, a 15-min loading is sufﬁcient to attain a homogeneous staining. 11. The intensity of the red ﬂuorescence is proportional to the degree of acidity and/or the volume of the cellular acidic compartment. Therefore, we could measure a change in the degree of acidity and/or the fractional volume of their cellular acidic compartment. 12. Most of the chemicals used for processing specimens for electron microscopy are extremely hazardous, especially glutaraldehyde, formaldehyde, osmium tetroxide, embedding medium in liquid form, lead citrate, and uranyl acetate. Extreme care should be taken when handling these chemicals. All steps must be performed

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in a fume hood and gloves should be worn all the time. Osmium tetroxide, glutaraldehyde, and propylene oxide/resin waste should be collected in bottles for safe disposal. 13. It is important to carry out this part of the technique in dark conditions to prevent uranyl acetate from being precipitated, as UA is photoreductive. 14. Before staining the ultrathin section, it is important to air dry for 2 h or overnight. 15. Brief tissue ﬁxation shortly after interruption of vital functions preserves morphology and antigenicity. 16. It is preferable to use BSA with immunohistochemical grade. 17. If the ﬂuorescence signal is too weak or the background is too high, check the composition of buffers and the working concentrations for the primary and secondary antibodies.

Acknowledgments We thank Donald Norwood (Department of Scientiﬁc Publication, The University of Texas M. D. Anderson Cancer Center, Houston, TX) for editorial assistance. This work was supported by Alex Lemonade Stand Foundation, Brain Tumor Society, Marcus Foundation, and NIH P50CA127001. References 1. Furnari, F.B., Fenton, T., Bachoo, R.M., et al. (2007) Malignant astrocytic glioma: genetics, biology, and paths to treatment. Genes Dev 21, 2683–710. 2. Galli, R., Binda, E., Orfanelli, U., et al. (2004) Isolation and characterization of tumorigenic, stem-like neural precursors from human glioblastoma. Cancer Res 64, 7011–21. 3. Singh, S.K., Clarke, I.D., Terasaki, M., et al. (2003) Identiﬁcation of a cancer stem cell in human brain tumors. Cancer Res 63, 5821–8. 4. Reynolds, B.A., and Weiss, S. (1992) Generation of neurons and astrocytes from isolated cells of the adult mammalian central nervous system. Science 255, 1707–10. 5. Fueyo, J., Gomez-Manzano, C., Alemany, R., et al. (2000) A mutant oncolytic adenovirus targeting the Rb pathway produces anti-glioma effect in vivo. Oncogene 19, 2–12. 6. Fueyo, J., Alemany, R., Gomez-Manzano, C., et al. (2003) Preclinical characterization of the antiglioma activity of a tropism-enhanced adenovirus targeted to the retinoblastoma pathway. J Natl Cancer Inst 95, 652–60.

7. 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. 8. Alonso, M.M., Jiang, H., Yokoyama, T., et al. (2008) Delta-24-RGD in combination with RAD001 induces enhanced anti-glioma effect via autophagic cell death. Mol Ther 16, 487–93. 9. 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. 10. Daido, S., Kanzawa, T., Yamamoto, A., Takeuchi, H., Kondo, Y., and Kondo, S. (2004) Pivotal role of the cell death factor BNIP3 in ceramide-induced autophagic cell death in malignant glioma cells. Cancer Res 64, 4286–93. 11. Paglin, S., Hollister, T., Delohery, T., et al. (2001) A novel response of cancer cells to radiation involves autmophagy and formation of acidic vesicles. Cancer Res 61, 439–44.

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Chapter 10 Propagation, Puriﬁcation, and In Vivo Testing of Oncolytic Vesicular Stomatitis Virus Strains Jean-Simon Diallo, Markus Vähä-Koskela, Fabrice Le Boeuf, and John Bell Abstract Oncolytic viruses are self-amplifying therapeutics that speciﬁcally replicate in and kill cancer cells. We have previously shown that vesicular stomatitis virus (VSV) can be used as an oncolytic virus. A strain of VSV harboring a mutation in the M protein (VSV$51) was found to exhibit enhanced tumor selectivity over its wild-type counterpart due to its inability to overcome antiviral programs in normal cells and due to the frequent defects in antiviral signaling pathways observed in the majority of tumors. VSV$51 can harbor transgenes, is easily propagated and puriﬁed to high titers, and shows potent oncolytic activity in several mouse models, including syngeneic CT26-lacZ subcutaneous colon carcinoma models. However, VSVneutralizing antibodies targeting mainly the VSV-G surface glycoprotein arise within 3–5 days following the initial dose. This should be considered for strategies aiming at increasing the effectiveness of VSV through delivery of additional doses of virus or aiming to prolong VSV replication in vivo. Key words: Vesicular stomatits virus, Oncolytic, Puriﬁcation, Syngeneic mouse model, Neutralizing antibodies

1. Introduction Vesicular stomatitis virus (VSV) is a small negative-strand RNA virus belonging to the Rhabdovirus family. The VSV genome (approximately 11 kbp) encodes ﬁve distinct genes referred to as the nucleocapsid (N), polymerase proteins (P and L), the matrix protein (M), and the surface glycoprotein (G). The VSV virion is bullet-shaped and consists of viral RNA wrapped tightly within the ribonucleocapsid (an array of N, P, L proteins) to which binds M protein. The virion becomes enveloped with G protein trimers

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upon budding from the host cell, which is the main target of neutralizing antibodies generated against the virus (1). VSV has been explored as an oncolytic virus due in part to its rapid life cycle and its potent cytolytic activity. Being an RNA virus that replicates in the cytoplasm, VSV is unlikely to recombine with cellular DNA and cannot integrate the genome. Combined with low human exposure to the virus and the absence of preexisting immunity in the population, this makes VSV an attractive candidate for human cancer therapy (2). Interferon (IFN) signaling is thought to be defective in approximately 70–75% of all tumors (3). In light of this, we and others have explored VSV strains that exploit this tumor-speciﬁc defect to enhance tumor targeting (4, 5). One of these mutant strains is VSV$51 which harbors a mutation in the M protein at methionine 51. Whereas wild-type M protein can block the nuclear export of cellular antiviral genes and their subsequent expression, the M$51 protein cannot and so VSV$51 replication is quickly subdued in normal cells that have intact antiviral signaling pathways (4).

2. Materials 2.1. Cell Culture and Virus Propagation

1. HyQ High glucose Dulbecco’s modiﬁed Eagle medium (DMEM) supplemented with 10% fetal bovine serum (FBS) (HyClone, Waltham, MA). 2. Solution of trypsin (0.25%) and ethylenediamine tetraacetic acid (EDTA) (1 mM) from Gibco/BRL. 3. ViCell (Beckman/Coulter) automated cell counter (alternately any standard hemocytometer can be used). 4. Phosphate buffer saline (PBS, HyClone). 5. 0.20 Mm Bottle top ﬁltration unit (Millipore).

2.2. Virus Puriﬁcation

1. OptiPrep 60% (w/v) (Sigma). 2. Tris stock solution (100 ml): Dissolve 12.1 g Tris base into 100 ml ddH2O. Filter and store at 4°C. 3. EDTA stock solution (100 ml): Dissolve 3.72 g EDTA Na2·2H2O into 100 ml ddH2O. Filter and store at 4°C. 4. NaCl stock solution (100 ml): Dissolve 5.84 g NaCl into 100 ml ddH2O. Filter and store at 4°C. 5. Solution B (100 ml): To 50 ml water, add 30 ml Tris stock solution and 3.0 ml EDTA stock solution. Adjust pH to 7.4 with 5 M HCl, and adjust volume to 100 ml with ddH2O. Filter and store at 4°C.

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6. Solution C (100 ml): To 50 ml water, add 10 ml NaCl stock solution, 5.0 ml Tris stock solution, and 0.5 ml EDTA stock solution. Adjust pH to 7.4 with 1 M HCl, and adjust volume to 100 ml with ddH2O. Filter and store at 4°C. 7. Solution D: Add 5 volumes of OptiPrep (Sigma) to 1 volume solution B. 8. Gradient maker (e.g., Auto Densi-Flow, LabConco). 9. Ultra-clear thinwall Beckman ultra-centrifuge tubes (11-ml maximum volume) for use with Beckman SW41Ti rotor. 2.3. Virus Titering

1. 2× concentrated DMEM prepared from powder (Gibco/BRL) supplemented with 20% FBS. 2. 1% agarose dissolved in deionized water. 3. 3:1 methanol: Glacial acetic acid ﬁxative solution. 4. Coomassie brilliant blue solution: 0.5 g Coomassie Brillant Blue R (Sigma) is dissolved in 250 ml 40% methanol and subsequently ﬁltered on Whatman paper. 250 ml 20% acetic acid is then added to the ﬁltered solution.

2.4. Mouse Tumor Implantation, Treatment with Virus, and Imaging

1. Syringes. 2. IVIS200 Series Imaging System (Xenogen). 3. 3% Isoﬂurane (Baxter). 4. D-Luciferin, Products).

10

mg/ml

in

PBS

(Molecular

Imaging

5. Electronic caliper. 2.5. Collection of Mouse Serum and Neutralizing Antibody Assay

1. Restraint tube (perforate 50-ml Falcon tube). 2. Vaseline. 3. Beveled syringe needle. 4. Heparinized collection tubes.

3. Methods A key consideration in using VSV or VSV$51 for both in vitro and in vivo experimentation is starting from a pure high-titer stock of virus containing as little defective uninfectious particles as possible. To this end, virus is propagated by infecting sensitive cells (e.g., Vero cells) at a low multiplicity of infection and then collected from supernatants prior to extensive cell lysis. The virus is subsequently puriﬁed on gradient to eliminate cellular contaminants, such as DNA, protein, and cellular exosomes (6, 7), which can induce antiviral programs and impact the permissiveness of cells

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in vitro or induce immune responses in vivo. It is possible to quantify how many infectious VSV particles have been obtained using viral plaque assays. Once viral titers have been assessed, virus can then be used for in vitro or in vivo testing alone or in combination with other agents (8, 9). In vitro, virus spread can be followed easily when VSV is engineered to express ﬂuorescent proteins using ﬂuorescence microscopy. For example, we use VSV variants in which have been inserted green ﬂuorescent protein (GFP) or ﬁreﬂy luciferase (FLuc) between the G and L genes driven by an intergenic VSV promoter. FLuc-expressing variants are useful as they can be imaged in vivo in real time using injectable luciferin and in vivo imaging systems (IVISs). The oncolytic properties of VSV can be observed in xenografts implanted in immunocompromised animals or in immunocompetent mice harboring syngeneic tumors. The latter is preferred since immune responses can both positively and negatively impact oncolytic virus efﬁcacy (10, 11). Subcutaneous models are convenient since tumors can be injected directly and volumes post infection can be followed using a caliper (Fig. 3). Viral replication can be followed by IVIS as well. Infection of mice with VSV leads to production of neutralizing antibodies from B cells that typically recognize the G-protein. These antibodies effectively inactivate the virus and generally arise 4–5 days post infection when virus is given intravenously (Fig. 4). VSV-neutralizing antibody titers can be easily measured from sera extracted from mouse blood obtained by saphenous vein bleeds at various times post infection. 3.1. Propagation of VSV in Cell Culture

1. Prior to infection with VSV, Vero African green monkey kidney cells are seeded at 1 × 104 cells per cm2 in FBS supplemented with DMEM in 150-mm cell culture petri dishes (25 ml per plate). 2. When cultures have reached roughly 95% conﬂuency (~3–4 days), they can be infected with a preexisting virus stock at 2 × 105 plaque forming units (PFUs) per 150-mm petri (an MOI of ~0.01, see Note 1). 3. For infection of ten plates (see Note 2), 2 × 106 PFUs of stock is diluted in 50 ml of serum-free DMEM and vortexed thoroughly (see Note 3). 4. Supernatant is removed from each petri dish and 5 ml of virus mixed in serum-free DMEM is added to each plate. Petris are then placed in a 37°C 5% CO2 humidiﬁed incubator for 45 min (see Note 4). 5. After incubation, the serum-free media is removed and 25 ml of DMEM containing 10% FBS is added to each petri. Infected cultures are then placed in a 37°C 5% CO2 humidiﬁed incubator.

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6. When approximately 50% cytopathic effect (CPE) (cell rounding) is observed (usually, after approximately 24 h), the supernatants are collected in 50-ml Falcon tubes (see Note 5). 7. The supernatants are centrifuged at 780 × g for 5 min to pellet heavy cellular debris. 8. Cleared supernatants are ﬁltered through 0.2 Mm membranes (Millipore) and then either used directly or puriﬁed further (see Note 6). 3.2. Puriﬁcation of VSV

Puriﬁed VSV crude obtained in Subheading 3.1 should be further concentrated by ultracentrifugation prior to gradient puriﬁcation. 1. Virus can be pelleted using a Beckman Coulter Avanti J25I Ultracentrifuge at 14,000 rpm (28,000 × g) for 1 h 30 min at 4°C in polypropylene centrifuge bottles (JA-14 rotor). The virus pellet should be visible at this stage and supernatant should be carefully discarded, removing as much media as possible without disturbing the pellet (see Note 7). 2. To purify the virus using OptiPrep® (see Note 8), the virus pellet isolated following ultracentrifugation is resuspended in maximum 2 ml of solution C (see Note 9). 3. For a single 12-ml gradient (accommodating up to 2 ml of resuspended virus), prepare 5 ml of 15% OptiPrep® solution by diluting 1.5 ml of solution D (which is 50% OptiPrep®) with 3.5 ml of solution C. Also prepare 5 ml of 35% OptiPrep® solution by diluting 3.5 ml of solution D with 1.5 ml of solution C. 4. Transfer 5 ml of the 35% OptiPrep® solution into the ﬁrst chamber of a gradient maker (the chamber closest to the exit valve on a LabConco Auto Densi-Flow model gradient maker) and 4.5 ml of the 15% OptiPrep® solution into the second chamber (furthest away from the exit valve). 5. Let ~0.5 ml of the 35% solution leave the exit valve before opening the valve connecting the two gradient chambers. This generates 9.5 ml 15–35% continuous OptiPrep® gradient which can be collected in clear SW-41 Ti thin-walled ultracentrifuge tubes (Beckman # 344059). 6. When the gradient has been obtained, carefully layer up to 2 ml of virus suspension onto the gradient. 7. Ultracentrifuge virus and gradient at 36,000 rpm (160,000 × g) for 1.5 h at 4°C. 8. If the pellet has been properly dispersed, a single band should be visible roughly mid-gradient (see Fig. 1). Collect this band by ﬁrst removing the OptiPrep® above it and then aspirating the band carefully using a P 1000 pipet (should take at least 600 Ml).

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Fig. 1. VSV puriﬁcation on OptiPrep® Gradient. VSV band in a 5–50% OptiPrep Gradient obtained starting from 10 × 150-mm petris of Vero cells following a 24-h incubation. Note that a single band is obtained approximately mid-gradient. Multiple bands, some with macroscopic ﬂakes, may be observed if pellet is insufﬁciently dispersed in solution C prior to gradient puriﬁcation.

9. To avoid repeated freeze-thawing, make many small-volume aliquots (e.g., 10 Ml) and store at least at −80°C. This virus should remain stable for several months. 3.3. Quantiﬁcation of Infectious VSV Particles by Viral Plaque Assay

VSV obtained from various sources, including infected cell cultures, tissues, and puriﬁed virus stocks, can be quantiﬁed by viral plaque assays. 1. Vero cells are plated in six-well plates at densities such that they have reached approximately 95% conﬂuence the following day. In six-well plates, this equates to approximately 1 × 106 cells per well (see Note 10). 2. Serum-free media is then used to do serial dilutions of virus (see Note 11).

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Fig. 2. Viral plaque assay for quantifying VSV titers. Vero cells were plated and inoculated with serial dilutions of a supernatant obtained from VSV$51-infected cells then coated with an agarose/DMEM overlay. Twenty-four hours later, wells were ﬁxed with methanol–acetic acid ﬁxative, agarose overlay was removed, and ﬁxed cells were stained with Coomassie brilliant blue. The arrow points to a typical plaque. The middle well contains 25 plaques, an appropriate number to calculate viral titers which in this case comes to (25/10−5) × 2 = 5 × 106.

3. The media covering the plated Vero cells is then removed and 500 Ml of diluted virus stock (for each dilution) is used to infect Vero cells which are then incubated for 45 min at 37°C in a 5% CO2 humidiﬁed incubator. 4. During this time, a 1% agarose solution prepared in steriledeionized H2O should be brought to a boil and put in a 42°C water bath. 2× concentrated DMEM containing 20% FBS should be warmed in a 37°C water bath as well. 5. After the 45-min incubation, the media covering infected Vero cells should be removed. 1:1 volumes of 1% agarose: 2× DMEM, 20% FBS at 42°C should then be mixed together and 1–2 ml of this mixture should be used to cover each well of Vero cells infected with virus dilutions (see Note 12). 6. Cells should then be put back in a 37°C 5% CO2 humidiﬁed incubator for another 24 h (see Note 13). 7. After incubation, 1 ml methanol–acetic acid ﬁxative (3:1 ratio) is added on top of the agarose layer in each well and allowed to incubate at room temperature for 1 h. 8. Subsequently, the agarose can be washed or lifted off the wells using a strong water current (e.g., under a tap). 9. Fixed Vero cells are stained using 1 ml of a Coomassie blue solution and incubated for 30 min at room temperature on a plate shaker (at low speed). 10. Resulting viral plaques can be easily visualized (see Fig. 2) and plates can be stored indeﬁnitely. 11. To calculate viral titers, plaques are counted ideally at a dilution step, where between 10 and 100 plaques are visible.

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12. The number of plaques is then divided by the appropriate dilution and the resulting number is multiplied by 2 to give a titer in PFU/ml (since 500 Ml were used to infect cells). For example, if 25 plaques were counted in the well where a 1/106 dilution was used, the titer of the initial undiluted sample is (25/(10−6)) × 2 = 5 × 107 PFU/ml. 3.4. In Vivo Testing of Oncolytic VSVD51

VSV$51 has been shown to be an effective oncolytic in several tumor models. Because it is well-recognized that immune responses play a major role in oncolytic virus efﬁcacy, it is best to study this oncolytic strain in immunocompetent models. One model we frequently use is the CT26 colon carcinoma model, which is syngeneic in Balb/C mice (see Note 14). This model is convenient because tumors grow homogenously with relatively good reproducibility without ulcerating or metastasizing too quickly. As such, they are particularly suitable for measuring tumor volumes. 1. Balb/C mice can be injected subcutaneously in the ﬂanks with 3 × 105 cells (>95% viable) resuspended in 100 Ml serum-free media or PBS. 2. Ten days later, tumors reach approximately 200 mm3 as measured using a caliper and the formula Volume = (Length × (Width)2)/2. 3. VSV$51 can be injected either intratumorally or intravenously at doses of up to 1 × 109 PFU (see Note 15). 4. Tumor volumes may be measured at regular intervals. Delayed tumor progression can be observed even after a single injection of VSV$51 delivered intratumorally (Fig. 3). 5. Virus replication can be followed in vivo using IVIS if VSV$51expressing Fluc is used. To do this, mice are injected with 200 Ml of luciferin substrate (10 mg/ml) intraperitoneally. Five minutes later, mice are anesthetized using isoﬂurane and then put in the IVIS for photon measurement (see Note 16). Exposure times vary generally between 30 s and 3 min. 6. If so desired, tumors or other organs (for toxicity studies, for example) can be collected by dissection and used as a starting material for viral plaque assays to conﬁrm the presence (or absence) of virus. 7. To do this, tissues (whole or partial) are dissected, weighed, and homogenized in 500 Ml PBS. It is important to drench mice in ethanol 70% prior to dissection to minimize the risk of contamination (see Note 17). 8. A fraction of this homogenate can be used for viral plaque assays (Subheading 3.3, see Note 18).

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Fig. 3. Injection of VSV$51 intratumorally delays progression of syngeneic CT26 tumors. Mice were implanted with 3 × 105 VSV$51-sensitive CT26-lacZ cells (see Note 14). Tumors were injected with 5 × 107 PFU of VSV$51 (in 50 Ml PBS) 11 days following tumor implantation. Tumor volumes were measured every 2–3 days using an electronic caliper. Volumes are expressed in mm3 and were calculated according to the formula in Subheading 3.4, step 2. Five and six mice were included in PBS (full lines) and VSV$51 (dashed lines) groups, respectively.

3.5. Following the Production of VSVNeutralizing Antibodies In Vivo

VSV is quite sensitive to neutralizing antibodies present in the serum of immunized individuals; as such, additional doses of VSV may not reach tumors following the rise of neutralizing antibody titers. It may be useful for investigators to monitor neutralizing antibody titers, for example in response to cotreatment with drugs that can impact innate and elicited immune responses, such as cyclophosphamide (12). To measure VSV-neutralizing antibodies, fresh whole blood from infected animals or individuals is required. In mice, whole blood can be collected repeatedly by saphenous bleed. 1. Mice are put facing downward in a restraint tube (e.g., perforated 50-ml Falcon tube). 2. The mouse tail is wrapped around the index ﬁnger and held by the middle ﬁnger. 3. Light pressure is applied by the index ﬁnger and the middle ﬁnger on the lower abdomen and legs toward the wall of the restraint tube, securing the mouse in place. At this stage, the mouse should have its head in the restraint tube and its hind legs sticking out of the tube. 4. Using the thumb, light pressure is applied just above the hip, extending the leg. 5. Vaseline is then applied to the extended leg so that the saphenous vein can be easily visualized.

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6. A small puncture at a 45° angle is done directly on the saphenous vein and blood is drawn using heparinized collection tubes (see Note 19). 7. Blood contained within the capped heparinized collection tubes is spun down at 3,400 × g in an Eppendorf minifuge for 5 min to separate serum from blood cells. 8. Serum (fraction on top) should be transferred gently to a fresh tube using a micropipet (see Note 20). 9. Collected serum can be tested for its ability to neutralize VSV infection (see Note 21). 10. For the neutralizing antibody assay, Vero cells are plated in 96-well plates at a density of 12,500 cells per well in 100 Ml of FBS supplemented with DMEM and incubated overnight at 37°C in a 5% CO2 humidiﬁed incubator. 11. The following day, each serum sample is diluted 1/50 in serum-free medium (see Note 22). 12. 100 Ml of the diluted serum preparation is plated into an empty 96-well microtiter plate. 13. In each subsequent row, 50 Ml of serum-free DMEM is plated. 14. 50 Ml from the 1/50 dilution is transferred and serially diluted (1:2) across the plate. 50 Ml is discarded from the last row. 15. Subsequently, 2 × 105 PFU VSV per 50 Ml serum-free DMEM is added to each well (see Note 23). A negative control, where virusfree DMEM is used, and a positive control, where no serum is put in the initial 1/50 dilution, should also be included. A schematic representation of a typical dilution setup is shown in Fig. 4a. 16. The mixtures of serum dilutions and virus are incubated for 1 h at 37°C. 17. Media is removed from 96-well plates containing Vero cells, and the contents of the 96-well plate containing virus:serum mixtures are immediately transferred. 18. Cells and virus:serum mixtures are incubated at 37°C in a 5% CO2 humidiﬁed incubator for 1 h. 19. 100 Ml of DMEM containing FBS is then added to all of the wells, and the plates are further incubated for 48 h. 20. After this time, CPEs can be visualized or quantiﬁed by a variety of methods, including visual inspection (cell rounding), Coomassie blue stain, or using metabolic dyes, such as MTT or Alamar Blue®. 21. Antibody titers in a given serum sample can be determined by assessing the dilution at which 50% CPE is observed. Figure 4b shows the progression of neutralizing antibody titers collected form Balb/C mice injected intravenously with 1 × 107 PFU of VSV$51. In this case, the same virus was used for the neutralization antibody assay.

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Fig. 4. VSV-neutralizing antibody titers following intravenous administration of VSV$51 in mice. (a) Typical experimental setup for determining neutralizing antibody titers. (b) Balb/C mice were injected with 1 × 107 PFU of VSV$51 (in 100 Ml PBS). Serum was collected over time by saphenous vein bleed, and VSV-neutralizing antibody titers were determined using the method described in Subheading 3.5. In this case, Alamar Blue® metabolic dye was used to assess Vero cell viability. Five mice were included in each group and error bars represent standard error. Note that in this experiment, neutralizing antibody titers remained at their high plateau for at least 56 days post infection.

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4. Notes 1. In general, high-titer seed stocks tend to yield more viral progeny. 2. Typically, we use 10 × 150-mm petris which are sufﬁcient for routine use, although more stock may be necessary for in vivo experiments. 3. Brief vortexing of stocks prior to dilution is always recommended as concentrated viruses may deposit or clump. Vortexing after each dilution is also recommended. 4. Low-volume infection is necessary in order to optimize virus adsorption. During this step, it is important that cells do not dry and so the best results are obtained when plates are shaken lightly every 5 min. 5. Longer incubation times give higher yields, but also increase the number of defective particles and potential cellular contaminants. 6. The ﬁltrate contains VSV crude that can be titered (see Subheading 3.3) and used in in vitro experiments if so desired. It is nonetheless recommended to further purify and concentrate the virus by ultracentrifugation and better yet to purify the virus on gradient. 7. At this stage, if so desired, the virus pellet can be resuspended in 1 ml PBS solution and frozen at −80°C for use at a later time. This puriﬁed crude can be titered and used directly in vitro or in vivo. Because some contaminants can remain in puriﬁed crude, best results are obtained following puriﬁcation on either sucrose or optiprep gradient. 8. Because sucrose can be toxic when injected in mice, we use OptiPrep® (Iodixinol) in order to be able to use the same virus stock for in vitro and in vivo experiments. Puriﬁcation on optiprep gradient yields the highest titers when initiated from fresh puriﬁed crude (as opposed to frozen) since each freeze– thaw cycle of virus stocks leads to approximately two- to ﬁvefold decreases in titers. 9. Obtaining a homogenous virus preparation without clumps is critical at this stage since clumps may lead to multiple bands to extract in the gradient. To this end, it is recommended to allow the resuspended pellet to disperse overnight at 4°C. Also, rigorous aspiration and vortexing may help. Ultimately, virus and protein aggregates may not be dispersed completely, and thus it is more important to collect the homogenous band from the gradient.

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10. Alternately, 12-well plates can be used in which case ~2.5 × 105 Vero cells in 1 ml can be used. To keep consistent titers between formats, infection volumes should be adjusted accordingly and considered in the calculation of titers. 11. Usually, one in ten dilutions are used, although this can be further reﬁned if more precision is required. Importantly, best results are obtained when tips are changed after each dilution step as the virus can stick to the tips and carry over to more diluted samples, leading to overestimations in titers. 12. This must be done fairly quickly since the agarose/DMEM mixture congeals rapidly as it cools. If several six-well plates need to be done simultaneously, it is best to do several small batches and make small quantities of agarose/DMEM mixtures each time. 13. After 24 h, viral plaques should be clearly visible by eye and can be counted directly. To facilitate the counting of plaques, we recommend ﬁxing and staining Vero monolayers. 14. There exist several variants of the CT26 cell line. The CT26 cell line that can be obtained directly from ATCC is very resistant to VSV$51 infection. Some CT26 strains that have been subcloned, for example the CT26 clone 25 and CT26-lacZ strains, can be more sensitive to VSV$51. 15. Doses above 1 × 109 can lead to hind limb paralysis if the mice have not been previously immunized with the virus. 16. Typically, VSV replication can be observed at early time points (as early as 8–10 h) in the liver and spleen of mice, but this signal is transient. In contrast, replication in subcutaneous tumors increases over 24–72 h. 17. Do not drench tissues directly in ethanol as this may inactivate virus present at the surface. 18. For viral plaque assays starting from tissue homogenates, 2× DMEM media should be supplemented with penicillin– streptomycin in order to minimize the potential for contamination. 19. Up to 150 Ml of sample can be collected at once using this method; however, serum can be easily obtained from as little as 20 Ml. 20. Care should be taken not to collect any blood cells at this step. 21. Serum can be kept at 4°C for extended periods of time (easily up to a year). 22. It is important that the serum does not contain red blood cells as this leads to abnormally high viability values if a metabolic assay is used (e.g., spin down tubes ﬁrst and take serum from the top).

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23. The quantity of VSV used at this step depends on the expected range of antibody titer and the desired precision. For example, if low antibody titers are expected, small quantities of VSV should be used to increase precision. If a wide range of titers between samples is expected, more VSV is better as long as the quantity does not exceed the maximum antibody titer.

Acknowledgments This work was supported in part by the Canadian Institutes of Health Research, the Ontario Institute for Cancer research, and the Fonds de Recherche en Santé du Québec. References 1. Rose, J., Whitt, M. (2001) Rhabdoviridae: The viruses and Their Replication, 4th ed., Lippincott Williams and Wilkins, New York. 2. Lichty, B. D., Power, A. T., Stojdl, D. F., and Bell, J. C. (2004) Vesicular stomatitis virus: re-inventing the bullet. Trends Mol Med 10, 210–6. 3. Stojdl, D. F., Lichty, B., Knowles, S., Marius, R., Atkins, H., Sonenberg, N., and Bell, J. C. (2000) Exploiting tumor-speciﬁc defects in the interferon pathway with a previously unknown oncolytic virus. Nat Med 6, 821–5. 4. Stojdl, D. F., Lichty, B. D., tenOever, B. R., Paterson, J. M., Power, A. T., Knowles, S., Marius, R., Reynard, J., Poliquin, L., Atkins, H., Brown, E. G., Durbin, R. K., Durbin, J. E., Hiscott, J., and Bell, J. C. (2003) VSV strains with defects in their ability to shutdown innate immunity are potent systemic anti-cancer agents. Cancer Cell 4, 263–75. 5. Obuchi, M., Fernandez, M., and Barber, G. N. (2003) Development of recombinant vesicular stomatitis viruses that exploit defects in host defense to augment speciﬁc oncolytic activity. J Virol 77, 8843–56. 6. Segura, M. M., Garnier, A., and Kamen, A. (2006) Puriﬁcation and characterization of retrovirus vector particles by rate zonal ultracentrifugation. J Virol Methods 133, 82–91. 7. Cantin, R., Diou, J., Belanger, D., Tremblay, A. M., and Gilbert, C. (2008) Discrimination between exosomes and HIV-1: puriﬁcation of

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both vesicles from cell-free supernatants. J Immunol Methods 338, 21–30. Le Boeuf, F., Diallo, J. S., McCart, J. A., Thorne, S., Falls, T., Stanford, M., Kanji, F., Auer, R., Brown, C. W., Lichty, B. D., Parato, K., Atkins, H., Kirn, D., and Bell, J. C. (2010) Synergistic Interaction Between Oncolytic Viruses Augments Tumor Killing. Mol Ther 18 (5), 888–95. Nguyen, T. L., Abdelbary, H., Arguello, M., Breitbach, C., Leveille, S., Diallo, J. S., Yasmeen, A., Bismar, T. A., Kirn, D., Falls, T., Snoulten, V. E., Vanderhyden, B. C., Werier, J., Atkins, H., Vaha-Koskela, M. J., Stojdl, D. F., Bell, J. C., and Hiscott, J. (2008) Chemical targeting of the innate antiviral response by histone deacetylase inhibitors renders refractory cancers sensitive to viral oncolysis. Proc Natl Acad Sci USA 105, 14981–6. Parato, K. A., Senger, D., Forsyth, P. A., and Bell, J. C. (2005) Recent progress in the battle between oncolytic viruses and tumours. Nat Rev Cancer 5, 965–76. Stanford, M. M., Breitbach, C. J., Bell, J. C., and McFadden, G. (2008) Innate immunity, tumor microenvironment and oncolytic virus therapy: friends or foes? Curr Opin Mol Ther 10, 32–7. Ikeda, K., Ichikawa, T., Wakimoto, H., Silver, J. S., Deisboeck, T. S., Finkelstein, D., Harsh, G. R., Louis, D. N., Bartus, R. T., Hochberg, F. H., and Chiocca, E. A. (1999) Oncolytic virus therapy of multiple tumors in the brain requires suppression of innate and elicited antiviral responses. Nat Med 5, 881–7.

Chapter 11 Oncolytic Measles Virus Retargeting by Ligand Display Pavlos Msaouel, Ianko D. Iankov, Cory Allen, Stephen J. Russell, and Evanthia Galanis Abstract Despite signiﬁcant advances in recent years, treatment of metastatic malignancies remains a signiﬁcant challenge. There is an urgent need for development of novel therapeutic approaches. Virotherapy approaches have considerable potential, and among them measles virus (MV) vaccine strains have emerged as a promising oncolytic platform. Retargeted MV strains deriving from the Edmonston vaccine lineage (MV-Edm) have shown comparable antitumor efﬁcacy to unmodiﬁed strains against receptor expressing tumor cells with improved therapeutic index. Here, we describe the construction, rescue, ampliﬁcation, and titration of fully retargeted MV-Edm derivatives displaying tumor speciﬁc receptor binding ligands on the viral surface in combination with H protein CD46 and SLAM entry ablating mutations. Key words: Oncolytic measles virus, Measles retargeting, Virotherapy, Measles engineering

1. Introduction Oncolytic measles virus has emerged as a very promising oncolytic platform. In contrast to the wild-type measles virus, which can cause potentially serious disease, the Edmonston measles virus (MV-Edm) is an attenuated vaccine strain that has an excellent safety proﬁle. Our group and others have demonstrated the therapeutic potential of MV-Edm derivatives against a variety of solid tumors and hematologic malignancies including ovarian cancer (1, 2), glioblastoma multiforme (3), multiple myeloma (4, 5), lymphoma (6), breast cancer (7, 8), prostate cancer (9, 10), and hepatocellular carcinoma (11). MV-Edm strains exhibit considerable local bystander oncolytic effect achieved via the massive cell–cell fusion of infected cells with neighboring uninfected cells. Of note, attachment of measles virus

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to the target cells and fusion are mediated by two distinct envelope glycoproteins: the hemagglutinin (H) protein and the fusion (F) protein, respectively. Measles virus enters the cells via interaction of the H glycoprotein with two known receptors: CD46 (12, 13) a regulator of complement activation that is found on all human nucleated cells but overexpressed in tumors and SLAM (14–16), the signaling lymphocyte activating molecule, which is expressed on activated T and B cells and macrophages. Receptor recognition by the H protein induces conformational changes of the F protein that result in fusion and viral entry (17). Thus, viral binding speciﬁcity is determined by the measles H protein (18). This allows retargeting strategies to focus on H protein modiﬁcation without affecting the potent fusogenic capacity of the virus. Indeed, a number of studies from our laboratory and others have conﬁrmed that retargeted MV-Edm derivatives retain the potent oncolytic activity of the parent strain against cells or xenografts expressing the target receptors (19–25). A variety of ligands have been successfully displayed on H, including single-chain antibodies against EGFRvIII (22), EGFR (20, 21), CD38 (20), folate receptor alpha (FRalpha) (23), HER-2/neu (26), CD20 (27), echistatin (an RGD-containing peptide binding to integrins AvB3 and A5B1) (28), and cytokines such as IL-13 (19). An alternative MV-Edm targeting approach has also been developed utilizing protease-cleavable linkers that are inserted into the F protein to restrict proteolytic maturation of F at sites where the protease is abundantly secreted (29). It is, thus, now possible to reprogram MV-Edm speciﬁcity at different levels. Although ongoing clinical trials using engineered oncolytic MV-Edm strains have not demonstrated dose limiting toxicity to date (30), viral retargeting may address any future toxicity concerns associated with higher viral doses and use of potent therapeutic transgenes (where even tighter speciﬁcity might be desired), overcome variability in viral receptor expression by tumor cells, and increase the efﬁcacy of targeted delivery following systemic administration. In the following sections, we describe the methods employed in our laboratory for rescue and propagation of MV-Edm derivatives reprogrammed to speciﬁcally target cancer cells by displaying alternate receptor binding ligands on the viral surface. The ﬁrst step to MV-Edm retargeting is the introduction of H protein mutations that ablate viral recognition and entry via the natural measles receptors CD46 and SLAM. Several H mutations that ablate entry through CD46 or SLAM have been identiﬁed (31, 32). We routinely use the combination of a single CD46 ablating mutation at position 481 (Y481A) and a single SLAM ablating mutation at position 533 (R533A) (20, 22, 31, 32) shown by Nakamura et al. (32) to effectively ablate entry via both CD46 and SLAM with comparable fusogenicity to unablated strains.

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Fig. 1. Schematic representation of the retargeted His-tagged MV-GFP-HAA strain. The Y481A and R533A mutations in the hemagglutinin (H) gene ablate CD46 and SLAM interaction, respectively (HAA). The target receptor-binding ligand is displayed on the C-terminus of the ablated HAA protein. The ligand is ﬂanked by SﬁI (encoding AAQPA amino-acid sequence) and NotI (encoding AAA amino-acid sequence) restriction sites. A 6-histidine peptide (H6) is attached to the C-terminus displayed ligand via a short linker sequence (AAARGS). H6 facilitates the “pseudoreceptor” interaction, which allows propagation of the retargeted strain using Vero cells (Vero-aHis) generated to stably express a membrane-anchored single-chain antibody that binds to the H6 tag allowing virus entry and propagation. The FLAG-tag inserted after the start codon at the N-terminus of HAA is used to facilitate detection by Western immunoblotting. Fxa is a factor Xa protease cleavage sequence (IEGR) that links the HAA glycoprotein with the target receptor ligand peptide and can be used to determine the recombinant nature of the chimeric HAA (37). Thus, factor Xa protease cleavage allows removal of the peptide ligand, resulting in an HAA protein with similar molecular weight as that of unmodiﬁed virus. PacI/SpeI restriction enzymes are used to subclone the chimeric HAA into the full-length infectious clone of the MV-NSe strain, deriving from MV-Edm. The virus also contains the gene encoding green ﬂuorescent protein (eGFP) in position 1 (N nucleoprotein gene, P phosphoprotein gene, M matrix protein gene, F fusion protein gene, L large protein gene, * stop codons; standard one-letter abbreviations are used to denote amino-acid residues) (Modiﬁed from Nakamura T et al., Nature Biotechnology 23(2):209–14, 2005).

To facilitate viral rescue and allow in vitro and in vivo visualization of infection, the virus may also contain the enhanced green ﬂuorescent protein (eGFP) gene at position 1 (Fig. 1). All retargeted viruses are derived from the Edmonston-Nse strain and are rescued using the pseudoreceptor STAR (Six-His Tagging And Retargeting) system (20). This system uses the 293-3-46 helper cells generated by Radecke et al. (33) to stably express the measles nucleocapsid (N) and phosphoprotein (P) proteins and T7 RNA polymerase for viral rescue (Fig. 2), and Vero-aHis cells for viral propagation. The protocol we describe here includes construction of the full length retargeted measles construct by cloning of the target receptor peptide ligand at the C terminus of H, rescue and propagation of the virus as well as preparation, puriﬁcation, and titration of viral stocks. The targeting ligands introduced at the C-terminus of the H protein are coupled with a 6-histidine tag (Fig. 1), which allows propagation of retargeted MV-Edm derivatives using Vero-aHis cells. This cell line was generated by Nakamura et al. (20) to express a membraneanchored single-chain antibody that binds to the 6-histidine tag allowing virus entry and propagation of the retargeted strain.

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Fig. 2. Rescue procedure for retargeted measles strains. The helper 293-3-46 cells are transfected with the full-length vector expressing the retargeted ablated infectious clone as well as the pEMC-La and pSC6-N plasmids expressing measles L and N proteins, respectively. The cells are incubated at 37°C CO2 for 16–20 h and then heat-shocked at 44°C for 2 h. The plate is returned to the 37°C CO2 incubator for approximately 3 days and then overlain on Vero-aHis cells. Binding of the single chain antibody expressed on the cell surface of Vero-aHis (pseudoreceptor) with the 6-histidine tag on the C-terminus of the recombinant H protein allows for entry of the retargeted virus and propagation in Vero-aHis cells. The plates are examined for the characteristic cytopathic effect (syncytia formation) and the infectious virus clone is subsequently harvested (modiﬁed from Nakamura T et al., Nature Biotechnology 23(2):209–14, 2005).

2. Materials 2.1. Construction of the Retargeted Measles Construct

1. Oligonucleotide primers for ampliﬁcation of the targeting ligands. The primers should be designed to ﬂank the ligand with SﬁI/NotI digestion sites.

2.1.1. Targeting Ligand Ampliﬁcation

2. Standard polymerase chain reaction (PCR) reagents. We routinely use the Taq polymerase for PCR ampliﬁcation, which allows the use of the TA cloning system. 3. Standard reagents and apparatus for agarose gel electrophoresis.

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2.1.2. Gel Fragment Puriﬁcation

1. QIAEX Gel Extraction Kit (Qiagen, Almada, CA).

2.1.3. Cloning of the Peptide Ligand

1. Invitrogen TA Cloning Kit (Invitrogen, Carlsbad, CA).

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2. 10 mM Tris–HCl, pH 8.5 or H2O with pH 7.0–8.5.

2. Lysogeny broth (LB) plates containing 100 Mg/ml ampicillin (store at 4°C). 3. LB medium containing 100 Mg/ml of ampicillin (store at 4°C).

2.1.4. Plasmid Isolation and Analyzation

1. QIAprep Miniprep kit (Qiagen, Almada, CA). 2. 10 mM Tris–HCl, pH 8.5 or H2O (pH 7.0–8.5). 3. Digestion enzymes and reagents used to verify DNA sequence were bought from New England Biolabs (Ipswich, MA). Store at −20°C. 4. DNA sequencer (Perkin-Elmer, Foster City, CA).

2.1.5. Subcloning of the Peptide Ligand into the pCG-HAA-H6 Vector

1. pCG-HAA-H6 shuttle vector developed by Dr. T. Nakamura (20). Store at −20°C. 2. NEBuffer 2 10× (New England Biolabs, Ipswich, MA) (10× concentrated stock of 50 mM NaCl, 10 mM Tris–HCl, 10 mM MgCl2, 1 mM dithiothreitol (DTT), pH 7.9). Store at −20°C. 3. BSA 100× (New England Biolabs) (100× concentrated stock of 20 mM KPO4 (pH 7.0), 50 mM NaCl, 0.1 mM EDTA and 5% (w/v) glycerol). Store at −20°C. 4. NotI and SﬁI digestion enzymes (New England Biolabs). Store at −20°C. 5. 10× ligation buffer (Invitrogen) (a 10× concentrated stock of 60 mM Tris–HCl, pH 7.5, 60 mM MgCl2, 50 mM NaCl, 1 mg/ml bovine serum albumin, 70 mM B-mercaptoethanol, 1 mM ATP, 20 mM dithiothreitol (DTT), 10 mM spermidine). Store at −20°C. 6. T4 DNA Ligase (4.0 Weiss units/Ml) (Invitrogen). Store at −20°C.

2.1.6. Propagation and Puriﬁcation of Transfection-Grade Plasmid DNA

1. QIAprep Midiprep kit (Qiagen). 2. LB medium containing 100 Mg/ml of ampicillin. Store at 4°C. 3. 99.7% (v/v) pure isopropanol, 70% (v/v) ethanol solution. 4. H2O (pH 7.0–8.5) or 10 mM Tris–HCl, pH 8.5.

2.1.7. Ligation into Full-Length Plasmid

1. The p(+) MV-eGFP(−Spe) plasmid encodes the full-length infectious clone of the MV-NSe strain, deriving from MV-Edm. The green ﬂuorescent protein gene (eGFP) has been inserted as an additional transcription unit upstream of the measles N

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gene (Fig. 1). The vector has been modiﬁed (34) to contain unique PacI and SpeI digestion sites that facilitate cloning of the modiﬁed H cDNAs. Store at −20°C. 2. NEBuffer 4 10× (New England Biolabs) (10× concentrated stock of 50 mM potassium acetate, 20 mM Tris–acetate, 10 mM magnesium acetate, 1 mM DTT, pH 7.9). Store at −20°C. 3. BSA 100× (New England Biolabs) (100× concentrated stock of 20 mM KPO4 (pH 7.0), 50 mM NaCl, 0.1 mM EDTA and 5% glycerol). Store at −20°C. 4. PacI and SpeI digestion enzymes (New England Biolabs). Store at −20°C. 5. 10× ligation buffer (Invitrogen) (10× concentrated stock of 60 mM Tris–HCl, pH 7.5, 60 mM MgCl2, 50 mM NaCl, 1 mg/ml bovine serum albumin, 70 mM B-mercaptoethanol, 1 mM ATP, 20 mM dithiothreitol (DTT), 10 mM spermidine) Store at −20°C. 6. T4 DNA Ligase (4.0 Weiss units/Ml) (Invitrogen). Store at −20°C. 2.2. Rescue, Propagation, and Titration of Retargeted MV Strains

1. 293-3-46 helper cell line generated by Radecke et al. (33) to stably express the T7 RNA polymerase as well as measles N and P proteins. 2. Vero-aHis cell line developed by Nakamura et al. (20) to stably express a membrane-anchored single-chain antibody recognizing the 6-histidine peptide (20, 35). Vero-aHis are derived from the Vero (African green monkey kidney) cell line and are grown in DMEM supplemented with 10% (v/v) FBS and 100 U/ml penicillin–streptomycin (see Note 1). 3. Dulbecco’s Modiﬁed Eagle’s Medium (DMEM) supplemented with 10% (v/v) fetal bovine serum. The DMEM may also contain 100 U/ml penicillin–streptomycin and 1.2 mg/ml G418 when indicated. Store at 4°C. 4. Opti-MEM I Reduced Serum Medium (Invitrogen) supplemented with 100 U/ml penicillin–streptomycin as indicated. The Opti-MEM used in all protocols described herein does not contain fetal bovine serum (FBS). Store at 4°C. 5. Standard cell culture equipment and materials.

2.2.1. Transfection of 293-3-46 Helper Cells

1. ProFection Mammalian Transfection System–Calcium Phosphate kit (Promega). 2. pEMC-La plasmid expressing the measles large (L) protein developed by Radecke et al. (33). Store at −20°C. 3. pSC6-N plasmid expressing the measles N protein developed by Radecke et al. (33). Store at −20°C.

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4. p(+) MV-eGFP(−Spe) plasmid engineered to contain the chimeric fully retargeted measles H as described in Subheading 3.1.7.

3. Methods 3.1. Construction of the Retargeted Measles Construct

The targeting ligand is subcloned into the C terminus of the measles H gene. To facilitate cloning, the pCG-HAA-H6 plasmid contains SﬁI/NotI restriction sites (Fig. 1) that are used to insert the ligand cDNA (see Note 2).

3.1.1. Peptide Ligand Ampliﬁcation

1. Begin by PCR amplifying the target receptor ligand. Any vector, tissue, cell line, etc. that expresses the ligand of interest can be used for PCR ampliﬁcation. Primers are designed to ﬂank the ligand with SﬁI/NotI digestion sites (see Note 2). 2. The PCR product is ran on a 1% agarose gel and visualized by ethidium bromide staining. The fragment is then excised and extracted from the agarose gel with a sterile scalpel and placed into a clean 1.5-ml microcentrifuge tube.

3.1.2. Gel Fragment Puriﬁcation

We routinely use the QIAEX II Gel Extraction Kit for puriﬁcation of ligands 500–3,000 bp in size. 1. Prepare a water bath at 50°C. 2. In a clean 1.5-ml microcentrifuge tube mix 10 Ml of the DNA adsorption resin (QIAEX II) with 650 Ml of QX1 buffer. Then, add the mix into the tube containing the gel fragment. 3. Incubate the tube at 50°C for 10–30 min. This solubilizes the agarose and adsorb the DNA to the resin, which appears as a pellet at the bottom of the tube. 4. Remove the supernatant (being careful not to disturb the pellet) with a pipette. Then, wash the pellet with 750 Ml of Buffer QX1, resuspend the pellet by vortexing, and do a quick spindown for 30 s. Decant the supernatant using a pipette without disturbing the pellet. All supernatant should be removed to clear any residual agarose contaminants. 5. Wash the pellet with 500 Ml of wash buffer (buffer PE) and do a quick spin-down for 30 s. Again remove the supernatant using a pipette without disturbing the pellet. All supernatant should be removed to clear any residual salt contaminants. 6. Air-dry the pellet until it becomes white. 7. For DNA elution, add 20 Ml of 10 mM Tris–HCl, pH 8.5 or H2O with pH 7.0–8.5 and vortex until all of the pellet is resuspended. 8. Incubate at room temperature for 10–15 min (up to 30 min).

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9. Centrifuge for 1 min. The puriﬁed DNA now is contained in the supernatant. Carefully place the supernatant into a sterile tube. It can now be used for cloning into the TA cloning vector or stored at −20°C until use. 3.1.3. Cloning of the Ligand

These instructions assume the use of the Invitrogen TA cloning kit containing the pCR2.1 cloning vector but are easily adaptable to other cloning systems. The TA cloning kit (Invitrogen) allows subcloning into the pCR2.1 vector without enzymatic digestion of the PCR product or vector. 1. Ligation of the puriﬁed DNA into the pCR2.1 cloning vector: Prepare the 10 Ml ligation reaction by mixing 1 Ml of the puriﬁed DNA, with 1 Ml of the pCR2.1 vector (25 ng/Ml), 1 Ml of the 10× Ligation Buffer, 1 Ml of the T4 DNA ligase, and 6 Ml of sterile water. This step should be executed on ice; the 10× ligation buffer is especially sensitive to temperature variations. Always do a quick spin of the ligation buffer before use. 2. The ligation reaction (ligated pCR2.1 vector) is then incubated at 14°C for at least 1 h but preferably overnight; store at −20°C until ready for transformation. 3. Transformation of the ligand vector into competent Escherichia coli: Our laboratory routinely uses the One Shot® TOP10 E. coli cells included into the TA cloning kit, but other competent strains may be used. 4. Thaw one vial (50 M) of frozen One Shot® Competent E. coli. Thawing should always be performed on ice. 5. Immediately after thawing, add 2.5 Ml of the ligated vector into the E. coli vial. Preferably do not pipette the mix because this may harm the bacteria. 6. Incubate the vial on ice for 30 min. The remaining ligated vector can be stored at −20°C. 7. The vial should then be placed at 42–50°C for 30 s to heatshock the bacteria. 8. Immediately place the vial on ice for 2 min. 9. Add 250 Ml of SOC medium (which has been thawed at room temperature) to the vial. 10. The vial should then be placed in a shaking incubator and shaken horizontally (~10 × g) at 37°C for 30–60 min. 11. Plate 50 Ml of the vial on an LB agar plate containing 100 Mg/ml ampicillin. Alternatively, one may spread 10 Ml of the vial into an LB agar plate, 50 Ml into a second plate, and the rest of the transformation vial on a third plate. This ensures that at least one plate has well-spaced colonies (see Note 3).

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12. The LB plate/plates should then be incubated at 37°C overnight. The next day, bacterial colonies should be visible on at least one plate. 13. The transformed colonies may then be analyzed or the plates can be stored at 4°C for up to 2 weeks. 14. To analyze the presence of the insert, six colonies may be picked for plasmid isolation. 15. Each colony should be placed in a tube with 1.5 ml LB medium containing 100 Mg/ml of ampicillin and then shaken (~10 × g) at 37°C for 4–5 h. 16. Centrifuge at ~10,000 × g in a conventional, tabletop microcentrifuge. The bacterial cells appear as a pellet at the bottom of the tube. Remove the supernatant by inverting the tube. The bacteria can now be used for plasmid isolation or stored at −20°C. 3.1.4. Plasmid Isolation and Analysis

1. Our laboratory routinely employs the QIAprep Miniprep kit for plasmid DNA puriﬁcation, but this can be exchanged for any other appropriate plasmid puriﬁcation method. Before starting, Buffers P2 and N3 should be checked for salt precipitation. Any such salts should be redissolved by warming to 37°C. The P2 buffer should not be vigorously shaken. Also ensure that RNAse A and Lyseblue have been added to the P1 buffer. The pelleted bacteria in each of the six tubes should ﬁrst be resuspended in 250 Ml of resuspension buffer (buffer P1) and vortexed until no pellet or bacterial clumps are visible. 2. Add to the tubes 250 Ml of lysis buffer (buffer P2). The bacterial suspension should then turn blue due to the dissolving effect of buffer P2 on the Lyseblue reagent contained in the P1 buffer. Gently shake the tube until the solution is homogeneously colored blue and let it rest at room temperature for no more than 5 min. 3. Add to the tubes 350 Ml of buffer N3 or 4.2 M Gu–HCl, 0.9 M potassium acetate, pH 4.8 and mix by inverting the tube until the suspension becomes completely colorless. Let it rest at room temperature for 5–10 min. 4. Transfer the tubes to a microcentrifuge and centrifuge at ~10,000 × g for 1.5 min. The supernatant contains the plasmid. 5. Apply the supernatant of each tube to a spin column and wait for ~10 min. 6. Centrifuge the columns at ~500 × g for 1.5 min in a microcentrifuge and just before the end increase to 10,000 × g. The ﬂow-through should then be discarded.

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7. Add 500 Ml of buffer PB to each spin column and centrifuge at ~10,000 × g for 1 min. Again the ﬂow-through should be discarded. 8. Add 500 Ml wash buffer (buffer PE) to each spin column and centrifuge at 10,000 × g in a microcentrifuge for 1 min. The ﬂow-through should always be discarded before proceeding to the next step. 9. Centrifuge again at 10,000 × g for 2 min to clear any residual wash buffer. 10. Each spin column should be placed in a sterile 1.5-ml microcentrifuge tube. The plasmid DNA can be eluted by applying 50 Ml of 10 mM Tris–HCl, pH 8.5 or H2O with pH 7.0–8.5 to the center of the column. Let it rest for 5 min at room temperature and then centrifuge at ~2,000 × g in a microcentrifuge for 2 min. 11. Quick spin at 10,000 × g in a microcentrifuge. The tube now contains the eluted plasmid DNA, which may be stored at −20°C. 12. The sequence of the ligated plasmid should now be veriﬁed. We usually perform analytical digestions of all six isolated plasmids and then perform full sequencing of at least one of the plasmids that appear to have the correct sequence according to the analytical digestion. After conﬁrming the correct sequence of the ligated plasmid, we proceed with subcloning into the pCG-HAA-H6 vector. 3.1.5. Subcloning of the Targeting Ligand into the pCG-HAA-H6 Vector

The pCG-HAA-H6 shuttle vector (provided by Dr. T. Nakamura) (20) encodes a modiﬁed measles H sequence that incorporates both a CD46 (Y481A) and SLAM (R533A) ablating mutation. As previously noted, it also includes unique SﬁI and NotI restriction sites (Fig. 1) that can be used to insert the ligand cDNA. 1. Prepare a 20 Ml digestion reaction for each of the two plasmids (pCR2.1 containing the peptide ligand cDNA and pCG-HAAH6) by mixing 0.5–1.0 Mg of the corresponding plasmid DNA with 2 Ml NEBuffer 2 10×, 0.5 Ml of BSA 100×, 0.5 Ml of NotI, and sterile water as needed to reach a volume of 20 Ml. Each reaction should be incubated at 37°C for ~2 h and then 0.5 Ml SﬁI should be added and the temperature raised to 50°C for 40 min. 2. The digestion product from the pCR2.1-ligand reaction can be ran on a 1% agarose gel and visualized by ethidium bromide staining. After conﬁrming the correct size of the fragment, it can be excised and gel-puriﬁed as described in Subheading 3.1.2. The digested pCG-HAA-H6 vector is similarly visualized and puriﬁed.

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3. The gel-puriﬁed fragment is then ligated to the SﬁI/NotIdigested pCG-HAA-H6 vector. For this purpose, we typically use the T4 DNA Ligase (Invitrogen) in our laboratory, but this may be substituted for any other effective ligation procedure. A 20 Ml ligation reaction is prepared by mixing 2 Ml of the T4 reaction buffer 10×, 2 Ml of the T4 ligase, 5 Ml of the gel-puriﬁed open pCG-HAA-H6 vector, 5 Ml of the gel-puriﬁed SﬁI/NotI-digested fragment, and 6 Ml of sterile water. Incubate overnight at 14°C. 4. The ligated pCG-HAA-H6 plasmid containing the targeting ligand can now be transformed into competent E. coli and propagated as described in Subheading 3.1.3. Plasmid isolation and analysis is performed as described in Subheading 3.1.4. 3.1.6. Propagation and Puriﬁcation of Transfection-Grade pCG-HAA-H6 Plasmid DNA

Before proceeding to constructing the full-length plasmid, it is important that transfection-grade preparations of the ligated pCGHAA-H6 plasmid are produced. Occasionally, ligation of the reengineered H cDNA into the full-length plasmid may fail and we have observed that plasmid quality can be a critical factor in resolving such issues. After conﬁrming the correct sequence of the ligated pCG-HAA-H6 by analytical digestions and DNA sequencing, the plasmid should be transformed into competent E. coli and plated as described in Subheading 3.1.3. For plasmid DNA puriﬁcation we have found the QIAprep Midiprep kit to yield excellent highquality transfection-grade DNA. Numerous competitive systems are available from other commercial sources. 1. Before starting, the P2 Buffer should be checked for SDS precipitation and the P3 buffer should be prechilled at 4°C. Also ensure that RNAse A and Lyseblue have been added to the P1 buffer. A single colony from a fresh plate should be inoculated into 3 ml of LB medium containing 100 Mg/ml ampicillin, to prepare a starter culture. 2. Place the tube in a shaking incubator (~10 × g) at 37°C for approximately 9 h. If the LB medium is still clear after 9 h (due to slow bacterial growth) then lower the temperature to 30°C and leave the tube overnight in the shaking incubator. 3. Inoculate 25–50 Ml of the starter culture into 50 ml of LB medium (100 Mg/ml ampicillin), shake at 37°C for approximately 5 h and then change the temperature to 30°C for approximately 5 h. 4. Add ampicillin to a total concentration of 200 Mg/ml and continue incubation at 30°C overnight. 5. Centrifuge at 6,000 × g for 15 min at 4°C. 6. The bacterial cells get pelleted at the bottom of the tube. Discard the supernatant by tube inversion. The bacteria may then be used for plasmid isolation or stored at −20°C.

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7. Resuspend the bacteria in 4 ml of resuspension buffer (buffer P1) and vortex until no pellet or bacterial clumps are visible. 8. Add 4 ml of lysis buffer (buffer P2). The bacterial suspension should then turn blue due to the dissolving effect of buffer P2 on the Lyseblue reagent contained in the P1 buffer. Gently shake the tube until the solution is homogeneously colored blue and let it stay at room temperature for no more than 5 min. 9. Add 4 ml of prechilled buffer P3 and mix by inverting the tube until the suspension becomes completely colorless. Let it stay at room temperature for 10 min. 10. During this time, equilibrate a QIAGEN-tip 100 column by applying 4 ml of equilibration buffer (Buffer QBT). The column should be allowed to empty by gravity ﬂow and the ﬂowthrough should be discarded. 11. Pour the bacterial suspension into the barrel of the QIAﬁlter syringe (QIAﬁlter Midi Cartridge) and let it stay at room temperature for another 10 min. The outer nozzle of the syringe should be capped. 12. Remove the cap from the syringe; insert the syringe plunger (provided in the kit) and press it to ﬁlter the bacterial lysate into the equilibrated QIAGEN-tip. 13. Allow the lysate to pass through the column (may take a few minutes) and discard the ﬂow-through (approximately 10 ml volume). 14. Wash the column twice with 10 ml of wash buffer (Buffer QC). Discard the ﬂow-through each time. 15. Place a clear 50-ml tube below the column and elute the plasmid DNA by applying 5 ml of elution buffer (buffer QF). 16. Precipitate the eluted DNA by adding 3.5–4 ml of isopropanol mix gently (do not vortex) and immediately centrifuge at t15,000 × g for 30 min at 4°C. Alternatively, DNA precipitation can be accomplished by centrifuging at 10,000–11,000 × g for 1 h at 4°C. 17. Carefully remove the supernatant and wash with 25 ml of 70% ethanol. 18. Centrifuge at 8,000 × g for 5 min at 4°C. 19. Carefully remove all of the supernatant without disturbing the DNA pellet. 20. Allow the pellet to air-dry for 5–10 min and redissolve the DNA with 100 Ml of 10 mM Tris–HCl, pH 8.5 or H2O with pH 7.0–8.5. To recover all DNA, vortexing may be performed to resuspend the DNA from the tube walls. 21. Determine DNA yield by UV spectrophotometry at 260 nm (see Note 4).

11 3.1.7. Subcloning of the Retargeted H into of the Full-Length MV-Edm Infectious Clone

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The ablated H protein displaying the targeting ligand is subsequently subcloned into the plasmid p(+) MV-eGFP (−Spe) that encodes the full-length infectious clone of the MV-Edm NSe strain and also contains the eGFP gene at position 1. This vector has been modiﬁed to contain a unique SpeI restriction site (34). Since both the pCG-HAA-H6 and the full-length p(+) MV-eGFP (−Spe) plasmids contain unique PacI and SpeI digestion sites, these can be used to subclone the ablated chimeric H cDNA into the full-length plasmid. 1. Prepare a 20 Ml digestion reaction for each of the two plasmids (pCG-HAA-H6 containing the targeting ligand cDNA and p(+) MV-eGFP (−Spe)) by mixing 0.5–1.0 Mg of each plasmid DNA with 2 Ml NEBuffer 4 10×, 0.5 Ml of BSA 100×, 0.5–1 Ml of PacI, 0.5–1 Ml of SpeI, and sterile water as needed to reach a volume of 20 Ml. Each reaction should be incubated at 37°C for ~3–4 h. 2. The product from the chimeric pCG-HAA-H6 digestion can be ran on a 1% agarose gel and visualized by ethidium bromide staining. After conﬁrming the correct size of the fragment it can be excised and gel-puriﬁed as described in Sub-heading 3.1.2. 3. The gel-puriﬁed fragment is subsequently ligated to the PacI/ SpeI-digested p(+) MV-eGFP (−Spe) vector, as described in Subheading 3.1.5. A 20 Ml ligation reaction is prepared by mixing 2 Ml of the T4 reaction buffer 10×, 2 Ml of the T4 ligase, 2 Ml of the gel-puriﬁed Pacl/Spel digested p(+) MV-eGFP (−Spe) vector, 8 Ml of the gel-puriﬁed PacI/SpeI-digested fragment, and 6 Ml of sterile water. Incubate overnight at 14°C. 4. The ligated full-length p(+) MV-eGFP (−Spe) plasmid containing the chimeric ablated H can now be transformed into competent E. coli and propagated as described in Subheading 3.1.3. Due to the vector size this transformation reaction may be less efﬁcient but at least a few colonies should be visible on the agar plate. Plasmid isolation and analysis is then performed as described in Subheading 3.1.4. After conﬁrming the correct sequence of the full-length infectious clone, the plasmid is propagated and puriﬁed in transfection-grade quality as described in Subheading 3.1.6.

3.2. Rescue, Propagation, and Titration of the Retargeted Virus

To rescue the retargeted MV-Edm derivative, a modiﬁed version of the rescue system developed by Radecke et al. (33) is utilized. The helper cells 293-3-46 are derived from the human embryonic kidney 293 cell line and engineered to stably express the measles N and P proteins as well as the T7 RNA polymerase (33). To rescue the virus (Fig. 2), the 293-3-46 cells are cotransfected with a plasmid expressing the measles large (L) protein (pEMC-La), a plasmid expressing the measles N protein (pSC6-N) and the full-length

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p(+) MV-eGFP (−Spe) plasmid containing the chimeric ablated H(see Note 5). Rescue of MV-Edm strains capable of interacting with the natural measles CD46 receptor on the cell surface of 2933-46 cells can easily be visualized because of the formation of giant mononuclear cell aggregates (syncytia) due to cell to cell fusion. On the other hand, fully retargeted strains are incapable of fusing the 293-3-46 cells, since these cells in most cases are not expressing the receptor recognized by the engineered H. Thus, the expression of the eGFP gene at position 1 of the retargeted strains is an effective way to visualize initial virus propagation in the helper cells. As the next step, overlaying the infected 293-3-46 cells onto Vero-aHis cells (Fig. 2) results in syncytia formation via interaction of the anti-His single chain antibody, displayed on the surface of the Vero-aHis cells, with the 6-histidine tag at the C-terminus of the retargeted H. Unless otherwise speciﬁed, all following steps should be performed in a tissue culture hood. 3.2.1. 293-3-46 Cell Culture and Plating (see Note 6)

1. The 293-3-46 helper cells are grown as monolayers in Dulbecco’s Modiﬁed Eagle’s Medium (DMEM) supplemented with 10% fetal bovine serum along with 100 U/ml penicillin– streptomycin and 1.2 mg/ml G418; cells are grown on T75 ﬂasks and passaged when 70–80% conﬂuent, using an 1:4–1:8 split rate. 2. The helper cells should be plated in a six-well plate 24 h before transfection. After splitting a T75 ﬂask, prepare 1:20, 1:40, and 1:80 cell dilutions of the aspirated cells and plate each dilution in two wells of a six-well plate (use a total of 2 ml per well). Incubate the plate at a 37°C CO2 incubator to allow the cells to attach and grow. It is important that on the day of transfection the monolayer consists of isolated cells (there should be as little cell clumping as possible) evenly spread throughout the well at 50–60% conﬂuency. Plating three different dilutions in two wells each ensures optimal cell density at the time of transfection.

3.2.2. Transfection of 293-3-46 Helper Cells (see Note 7)

1. At least 4 h prior to transfection the growth medium is removed and replaced with DMEM, with 10% FBS. The medium should be prewarmed in a water bath at 37°C prior to being added to the cells. 2. In our experience, calcium phosphate-mediated transfection has shown excellent results for viral rescue. Our laboratory typically uses the ProFection Mammalian Transfection System– Calcium Phosphate kit (Promega, Madison, WI). For a onewell transfection, 5 Mg of the full-length p(+) MV-eGFP (−Spe), 50 Mg of the pEMC-La plasmid, and 250 Mg of the pSC6-N provide optimum efﬁciency. The plasmids are solubilized in a sterile 15-ml tube containing 175 Ml of nuclease-free water.

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3. Add 25 Ml of 2 M CaCl2 to the tube and mix well. Let it rest at room temperature for 2–5 min. 4. In a separate sterile 15-ml tube add 200 Ml of 2× HBS. 5. Gently vortex the second tube containing the 2× HBS. While continuing vortexing slowly add the content of the ﬁrst tube dropwise to the second tube. Incubate the tube at room temperature for 30 min. 6. Add the transfection solution to the well containing the helper cells dropwise and at various locations around the well. Gently swirl the six-well plate to evenly distribute the transfection precipitate over the helper cells. 7. Incubate the plate in a 37°C CO2 incubator for 16–20 h. 8. Replace growth medium with DMEM 10% FBS containing penicillin–streptomycin (G418 is no longer required) to avoid any unintentional contamination during the next steps. 9. Heat shock the plate in a water bath at 44°C for 2 h. 10. Return the plate to a 37°C CO2 incubator. 3.2.3. Vero-aHis Cell Overlay

Approximately 3 days after the heat-shock, if the rescue is successful, isolated green 293-3-46 cells can be visualized under the ﬂuorescent microscope due to eGFP expression (see Note 8). Even if no green ﬂuorescent cells are visible, the helper cells should be overlaid onto Vero-aHis cells at this time. 1. Plate one or two 10 cm tissue culture plates with 50–60% conﬂuent Vero-aHis cells (in 5–10 ml of DMEM, 10% FBS, 100 U/ml penicillin–streptomycin). 2. Allow >3 h for the Vero-aHis to attach to the plate. 3. Scrape the 293-3-46 helper cells from the surface of the transfected well into the overlying medium using a sterile tissue culture cell scraper. 4. Slowly add the helper cell suspension dropwise and at various locations to the Vero-aHis plate. 5. Return the plates to a 37°C CO2 incubator. 6. Examine the plates daily under the microscope to identify the formation of syncytia which indicate the rescue of infectious viral particles (see Note 9).

3.2.4. Viral Clone Harvesting

1. As soon as one or more syncytia are formed, harvesting of each syncytium (corresponding to a different viral clone) should commence. Plate a six-well plate with 50–60% conﬂuent VeroaHis cells in 1 ml Opti-MEM containing 100 U/ml penicillin– streptomycin. Prepare one well for each visible syncytium. 2. Allow >3 h for the Vero-aHis to attach to the well surface. 3. Use a marker pen to circle the location of each syncytium on the undersurface of the 10 cm tissue culture plate.

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4. Remove the growth medium from the tissue culture plate. 5. Use a standard micropipette with a 10 Ml tip to take 5 Ml of Opti-MEM and repeatedly pipette the Opti-MEM to the location of the syncytium. The goal is to detach the syncytium and resuspend it into the Opti-MEM. Collect the Opti-MEM and add it to a Vero-aHis well on the six-well plate and place on a 37°C CO2 incubator for 2–3 h. Repeat this step for every visible syncytium which should be harvested and placed into a separate well corresponding to a distinct viral clone. 6. 2–3 h later, add 1 ml of DMEM 10%FBS, 100 U/ml penicillin–streptomycin to each well and return the plate to a 37°C CO2 incubator. 7. Two to three days later, wells are covered with syncytia. 8. Plate two 10 cm tissue culture plate with 50–60% conﬂuent Vero-aHis cells for each well. 9. Allow >3 h for the Vero-aHis to attach to the plate surface. 10. Remove the growth medium from each 10 cm plate and add 5 ml of Opti-MEM, 100 U/ml penicillin–streptomycin. 11. Remove the growth medium from each well and add 1 ml of Opti-MEM. 12. Scrape the Vero-aHis from each well into the overlying OptiMEM using a sterile tissue culture cell scraper. 13. Place the suspension of scraped cells/syncytia into a cryovial and vortex for 1 s. 14. Snap-freeze the cryovial by submersion in liquid N2. 15. Once completely frozen, thaw the cryovial using a water bath set to 37°C. When the suspension is half-thawed, vortex for 1 s and then return the cryovial to the water bath. 16. Following complete thawing, vortex for 1 s and then repeat the cycle of snap-freezing/thawing (steps 14 and 15). This releases the infectious viral particles into the supernatant. 17. Centrifuge at ~3,250 × g, for 5 min at 4°C to pellet the cell debris. Collect the cell lysate (supernatant) containing the virus. It may now be stored at a −80°C or used to proceed immediately to the additional puriﬁcation steps. 18. Slowly add the cell lysate of each well drop wise and at various locations around two 10 cm Vero-aHis dishes. 19. Incubate the dishes to a 37°C CO2 incubator for 2–3 h. 20. Add 5 ml of DMEM 10% FBS, 100 U/ml penicillin– streptomycin to each plate. 21. Return the plates to a 37°C CO2 incubator. 22. Approximately 2–3 days later each plate should be covered with syncytia.

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23. Plate two to three 15 cm tissue culture plates with 50–60% conﬂuent Vero-aHis cells corresponding to each infected 10 cm tissue culture plate. 24. Allow >3 h for the Vero-aHis to attach to the plate surface. 25. Remove the growth medium from each 15 cm plate and add 10 ml of Opti-MEM. 26. Remove the growth medium from each 10 cm plate and add 5–6 ml of Opti-MEM. 27. Scrape the Vero-aHis from each 10 cm plate into the overlying medium using a sterile tissue culture cell scraper. 28. Place the suspension of scraped cells/syncytia into a 50-ml tube and vortex for 1 s. Perform the two cycles of freeze-thawing as described in steps 14–17 to collect a cleared cell lysate containing the virus. 29. Slowly add the cell lysate of each 10 cm plate dropwise and at various locations around two to three 15 cm Vero-aHis plates. 30. Incubate the 15 cm plates to a 37°C CO2 incubator for 2–3 h. 31. Add 10 ml of DMEM 10% FBS, 100 U/ml penicillin– streptomycin to each plate. 32. Return the 15 cm plates to a 37°C CO2 incubator. 33. Two to three days later, wells are covered with syncytia. 34. Remove the growth medium from each 15 cm plate and add 8–10 ml of Opti-MEM. 35. Scrape the Vero-aHis from each 15 cm plate into the overlying medium using a sterile tissue culture cell scraper. 36. Place the suspension of scraped cells/syncytia into a 50-ml tube and vortex for 1 s. Perform the two cycles of freezing– thawing as described in steps 14–17 to collect a cleared cell lysate containing the virus. 37. The virus preparation may now be aliquotted into cryovials, stored at −80°C and titered. One of the aliquots can be marked to be used later for titration (see Note 10). 3.2.5. Measles Virus Titration

1. The following titration method is used to calculate the 50% tissue culture infective dose (TCID50) of a viral aliquot. First, prepare a 96-well plate by plating each well of the ﬁrst 8 columns with 100 Ml of 104 Vero-aHis (DMEM 10% FBS, 100 U/ ml penicillin–streptomycin). One or more additional columns of the 96-well plate may also be plated and used as noninoculated control wells (see Note 11). 2. Incubate the plate in a 37°C CO2 incubator.

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3. The next day, add 900 Ml of Opti-MEM (100 U/ml penicillin– streptomycin) in each of eight 1.5 ml microcentrifuge tubes. These tubes are used to prepare serial dilutions (10−1 to 10−8) of the viral aliquot. Place the tubes in a rack immersed in ice and label them −1 to −8 from left to right. 4. Thaw the cryovial marked for virus titration and add 100 Ml of the virus preparation into the ﬁrst 1.5-ml tube (labeled −1). Mix well by pipetting and inverting the tube. This tube contains the ﬁrst (1:10) dilution of the viral preparation. 5. Replace the pipette or tip and add 100 Ml of the 1:10 (10−1) dilution to the second tube (labeled −2) and mix well by pipetting and inversion. This second tube corresponds to the second (1:100) dilution of the viral preparation. Repeat this process to prepare all dilutions. Remember to always use a fresh pipette or tip before transferring 100 Ml of previous virus dilution to next dilution. 6. The serial dilutions should now be inoculated (50 Ml/well) to the Vero-aHis in the 96-well plate. Each dilution should be inoculated to all wells of one corresponding column (total of eight columns) starting from the highest (10−8) to the lowest (10−1) dilution. 7. Place the 96-well plate to a 37°C CO2 incubator. 8. Three to four days later when a fully conﬂuent monolayer of Vero-aHis cells is seen in control wells, all wells (inoculated with different virus dilutions) are examined under microscope for the presence of syncytia. An inoculated well is considered positive if the presence of even a single syncytium is detected (see Note 12). 9. Mark the syncytia-positive inoculated wells on the plate. The TCID50 is equal to 10X where X = absolute value of the exponent corresponding to the highest dilution where all wells in the column are positive + 1.8 + the fraction of positive wells past this dilution (see Note 13).

4. Notes 1. Vero-aHis cells should typically be passaged on a subcultivation ratio of 1:8–1:10 every 3–4 days. 2. The following considerations apply in choosing the strategy for targeting ligand cloning: (a) the primers used for ampliﬁcation of the ligand should also include SﬁI/NotI digestion sites necessary for subcloning the ampliﬁed fragment into the pCG-HAAH6 plasmid (Fig. 1); the presence of Sﬁ, NotI, PacI, or SpeI restriction sites inside the ligand coding sequence itself may

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complicate cloning. (b) For efﬁcient viral transcription to occur, it is critical that the genome size of the engineered retargeted virus is a multitude of six nucleotides (rule of six) (33, 36). 3. The pCR2.1 vector also encodes a lacZA fragment that allows for blue-white screening of bacterial colonies by supplementation of X-Gal into the LB agar plate. Due to the high efﬁciency of the cloning system in our experience, we do not typically use blue-white screening when selecting bacterial colonies. One may nevertheless use this feature when needed. 4. Before proceeding to the more complicated and time-consuming process of rescuing the reengineered virus, the chimeric pCG-HAA-H6 containing the targeting ligand may now be used for cotransfection along with plasmid expressing the MV-Edm F protein to conﬁrm that the chimeric ablated H is expressed onto the cell surface at a proper conformation that allows fusion of cells expressing the target receptor as well as of Vero-aHis cells. 5. The original rescue system by Radecke et al. (33) required the transfection of only two plasmids (pEMC-La and the fulllength vector) into the 293-3-46 cells. The additional cotransfection of the N expressing plasmid during rescue is a modiﬁcation used at the Galanis laboratory, which in our experience has signiﬁcantly increased the rescue efﬁcacy for both retargeted and nonretargeted cloned MV-Edm strains (19). 6. To increase the likelihood of a successful rescue, it is important that a healthy culture of 293-3-46 helper cells is maintained. Furthermore, plating before transfection must yield wellspaced, well-attached helper cells to optimize plasmid uptake. 7. We recommend that at least one additional well is used to simultaneously rescue a nonablated unmodiﬁed MV-Edm strain expressing eGFP (MV-eGFP) as a positive control conﬁrming that the helper cells are in good condition. 8. While this is a good indicator that viral rescue is progressing smoothly, not all of these green cells are able to assemble infectious virus particles. 9. Syncytia may appear at different time points up to 10 days after transfection of the 293-3-46 helper cells. Syncytia formation beyond 10 days signiﬁcantly increases the likelihood that the rescued virus is a mutated strain. Sequencing and other characterization techniques, including Western immunoblotting for detection of the correct size viral H protein (19), should be performed to verify that one or more of the rescued viral clones represents the correct retargeted strain. 10. Each freeze–thaw cycle likely decreases the viral titer. To determine the correct titer of the stored viral preparation, this should be aliquotted before transferring to −80°C. One aliquot can be

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thawed and titrated at a later time point, as described in Subheading 3.2.5. The calculated titer corresponds to that of each of the remaining aliquots when thawed for the ﬁrst time. 11. When preparing viral stocks, we typically prefer to titrate twice (in two 96-well plates inoculated simultaneously) and use the mean TCID50 calculated by these two titrations. 12. The cytopathic effect (syncytia formation) of MV-Edm derivatives is relatively easy to recognize under light microscopy even in cases when only a single syncytium is present. The expression of eGFP by infected cells further facilitates detection. By deﬁnition, no syncytia should be detectable on the noninoculated control wells. The presence of cytopathic effect in these wells may indicate contamination during titration or, worse, contamination of the prime Vero-aHis culture. 13. For example, if all wells up to and including the 10−5 dilution are syncytia-positive, three wells in the 10−6 are positive, one well in the 10−7 dilution is positive, and all wells in the 10−8 dilution are negative, then the TCID50 is equal to 105+1.8+3/8+1/8+0/8 = 107.3, which is approximately equal to 2 × 107 TCID50. References 1. Peng, K. W., TenEyck, C. J., Galanis, E., Kalli, K. R., Hartmann, L. C., and Russell, S. J. (2002) Intraperitoneal therapy of ovarian cancer using an engineered measles virus, Cancer research 62, 4656–4662. 2. Hasegawa, K., Pham, L., O’Connor, M. K., Federspiel, M. J., Russell, S. J., and Peng, K. W. (2006) Dual therapy of ovarian cancer using measles viruses expressing carcinoembryonic antigen and sodium iodide symporter, Clin Cancer Res 12, 1868–1875. 3. Phuong, L. K., Allen, C., Peng, K. W., Giannini, C., Greiner, S., TenEyck, C. J., Mishra, P. K., Macura, S. I., Russell, S. J., and Galanis, E. C. (2003) Use of a vaccine strain of measles virus genetically engineered to produce carcinoembryonic antigen as a novel therapeutic agent against glioblastoma multiforme, Cancer research 63, 2462–2469. 4. Peng, K. W., Ahmann, G. J., Pham, L., Greipp, P. R., Cattaneo, R., and Russell, S. J. (2001) Systemic therapy of myeloma xenografts by an attenuated measles virus, Blood 98, 2002–2007. 5. Dingli, D., Peng, K. W., Harvey, M. E., Greipp, P. R., O’Connor, M. K., Cattaneo, R., Morris, J. C., and Russell, S. J. (2004) Imageguided radiovirotherapy for multiple myeloma using a recombinant measles virus expressing

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the thyroidal sodium iodide symporter, Blood 103, 1641–1646. Grote, D., Russell, S. J., Cornu, T. I., Cattaneo, R., Vile, R., Poland, G. A., and Fielding, A. K. (2001) Live attenuated measles virus induces regression of human lymphoma xenografts in immunodeﬁcient mice, Blood 97, 3746–3754. McDonald, C. J., Erlichman, C., Ingle, J. N., Rosales, G. A., Allen, C., Greiner, S. M., Harvey, M. E., Zollman, P. J., Russell, S. J., and Galanis, E. (2006) A measles virus vaccine strain derivative as a novel oncolytic agent against breast cancer, Breast cancer research and treatment 99, 177–184. Iankov, I. D., Msaouel, P., Allen, C., Federspiel, M. J., Bulur, P. A., Dietz, A. B., Gastineau, D., Ikeda, Y., Ingle, J. N., Russell, S. J., and Galanis, E. (2009) Demonstration of antitumor activity of oncolytic measles virus strains in a malignant pleural effusion breast cancer model, Breast cancer research and treatment 2010 Aug;122(3), 745–54. Msaouel, P., Iankov, I. D., Allen, C., Morris, J. C., von Messling, V., Cattaneo, R., Koutsilieris, M., Russell, S. J., and Galanis, E. (2009) Engineered measles virus as a novel oncolytic therapy against prostate cancer, The Prostate 69, 82–91. Msaouel, P., Iankov, I. D., Allen, C., Aderca, I., Federspiel, M. J., Tindall, D. J., Morris, J. C.,

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Koutsilieris, M., Russell, S. J., and Galanis, E. (2009) Noninvasive imaging and radiovirotherapy of prostate cancer using an oncolytic measles virus expressing the sodium iodide symporter, Mol Ther 17, 2041–2048. Blechacz, B., Splinter, P. L., Greiner, S., Myers, R., Peng, K. W., Federspiel, M. J., Russell, S. J., and LaRusso, N. F. (2006) Engineered measles virus as a novel oncolytic viral therapy system for hepatocellular carcinoma, Hepatology (Baltimore, Md) 44, 1465–1477. Dorig, R. E., Marcil, A., Chopra, A., and Richardson, C. D. (1993) The human CD46 molecule is a receptor for measles virus (Edmonston strain), Cell 75, 295–305. Naniche, D., Varior-Krishnan, G., Cervoni, F., Wild, T. F., Rossi, B., Rabourdin-Combe, C., and Gerlier, D. (1993) Human membrane cofactor protein (CD46) acts as a cellular receptor for measles virus, Journal of virology 67, 6025–6032. Tatsuo, H., Ono, N., Tanaka, K., and Yanagi, Y. (2000) SLAM (CDw150) is a cellular receptor for measles virus, Nature 406, 893–897. Hahm, B., Arbour, N., Naniche, D., Homann, D., Manchester, M., and Oldstone, M. B. (2003) Measles virus infects and suppresses proliferation of T lymphocytes from transgenic mice bearing human signaling lymphocytic activation molecule, Journal of virology 77, 3505–3515. Schneider-Schaulies, S., Bieback, K., Avota, E., Klagge, I., and ter Meulen, V. (2002) Regulation of gene expression in lymphocytes and antigenpresenting cells by measles virus: consequences for immunomodulation, Journal of molecular medicine (Berlin, Germany) 80, 73–85. Yanagi, Y., Takeda, M., and Ohno, S. (2006) Measles virus: cellular receptors, tropism and pathogenesis, The Journal of general virology 87, 2767–2779. Nussbaum, O., Broder, C. C., Moss, B., Stern, L. B., Rozenblatt, S., and Berger, E. A. (1995) Functional and structural interactions between measles virus hemagglutinin and CD46, Journal of virology 69, 3341–3349. Allen, C., Paraskevakou, G., Iankov, I., Giannini, C., Schroeder, M., Sarkaria, J., Schroeder, M., Puri, R. K., Russell, S. J., and Galanis, E. (2008) Interleukin-13 displaying retargeted oncolytic measles virus strains have signiﬁcant activity against gliomas with improved speciﬁcity, Mol Ther 16, 1556–1564. Nakamura, T., Peng, K. W., Harvey, M., Greiner, S., Lorimer, I. A., James, C. D., and Russell, S. J. (2005) Rescue and propagation of fully retargeted oncolytic measles viruses, Nature biotechnology 23, 209–214.

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measles virus strains in the treatment of cancer: an overview, Current opinion in molecular therapeutics 11, 43–53. 31. Vongpunsawad, S., Oezgun, N., Braun, W., and Cattaneo, R. (2004) Selectively receptor-blind measles viruses: Identiﬁcation of residues necessary for SLAM- or CD46-induced fusion and their localization on a new hemagglutinin structural model, Journal of virology 78, 302–313. 32. Nakamura, T., Peng, K. W., Vongpunsawad, S., Harvey, M., Mizuguchi, H., Hayakawa, T., Cattaneo, R., and Russell, S. J. (2004) Antibody-targeted cell fusion, Nature biotechnology 22, 331–336. 33. Radecke, F., Spielhofer, P., Schneider, H., Kaelin, K., Huber, M., Dotsch, C., Christiansen, G., and Billeter, M. A. (1995) Rescue of measles viruses from cloned DNA, The EMBO journal 14, 5773–5784.

34. Hadac, E. M., Peng, K. W., Nakamura, T., and Russell, S. J. (2004) Reengineering paramyxovirus tropism, Virology 329, 217–225. 35. Douglas, J. T., Miller, C. R., Kim, M., Dmitriev, I., Mikheeva, G., Krasnykh, V., and Curiel, D. T. (1999) A system for the propagation of adenoviral vectors with genetically modiﬁed receptor speciﬁcities, Nature biotechnology 17, 470–475. 36. Kolakofsky, D., Pelet, T., Garcin, D., Hausmann, S., Curran, J., and Roux, L. (1998) Paramyxovirus RNA synthesis and the requirement for hexamer genome length: the rule of six revisited, Journal of virology 72, 891–899. 37. Peng, K. W., Donovan, K. A., Schneider, U., Cattaneo, R., Lust, J., and Russell, S. J. (2003) Oncolytic measles viruses displaying a singlechain antibody against CD38, a myeloma cell marker, Blood 101, 2557–2562.

Chapter 12 Exploring Host Factors that Impact Reovirus Replication, Dissemination, and Reovirus-Induced Cell Death in Cancer Versus Normal Cells in Culture Maya Shmulevitz and Patrick W.K. Lee Abstract Oncolytic viruses, such as reovirus, offer a promising approach to cancer treatment. Concurrently, oncolytic viruses provide a valuable tool for deciphering unique attributes of cancer cells that support superior virus replication, cell death, or virus dissemination. Through our studies on various cancer cell lines, as well as isogenic cells with and without transformation by oncogenic Ras, we have identiﬁed at least four steps of virus replication that can be augmented in transformed cells. Ras transformation can support efﬁcient reovirus uncoating during entry, production of progeny with high infectious capacity, and reovirus-induced apoptosis. Furthermore, Ras transformation also precludes interferon production following reovirus infection, permitting enhanced cell-to-cell virus spread. Methods that measure the efﬁciency of reovirus replication and dissemination described in this chapter can be used in combination with assorted cell culture systems to better understand the host factors that regulate reovirus oncolysis. Key words: Oncolytic virus, Reovirus, Plaque titration, Immunohistochemistry, Apoptosis, Cell-to-cell spread, MAPK/ERK, Ras

1. Introduction Reovirus is a naturally occurring human virus found to preferentially replicate in cancer cells and possess oncolytic activity both in vitro and in vivo (1–4). While in vivo studies using various murine cancer models and human clinical trials are ongoing to show the efﬁcacy of reovirus as a cancer therapy, we continue to explore the molecular basis of reovirus oncolysis in cell culture systems. Using isogenic cell lines (with and without mutated forms of oncogenes, such as Ras), we found that several steps of reovirus replication

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and spread are enhanced in Ras-transformed cells (5), including (1) reovirus disassembly necessary for entry and onset of infection, (2) production of progeny virions with high infectious capacity, and (3) induction of apoptosis and release of reovirus. Our recent studies show that cell-to-cell spread of reovirus is also greatly enhanced in transformed cells, owing to the impaired ability of transformed cells to express and respond to interferon beta (IFN-E) and establish an antiviral state (6, 7). Understanding the host and viral factors involved in reovirus oncolysis assists in developing combination therapies, allows for best application of reovirus as a therapeutic, and sheds light on other viral- or drug-based cancer therapies. This chapter describes the various methods used to discern the effects of host factors on the efﬁciency of reovirus infection in cell culture systems. Reovirus replication, apoptosis-mediated release, and cell-to-cell spread can be easily monitored with the methods described in this chapter, including immunohistochemistry (IHC) of reovirus-infected cells (Subheading 3.2), plaque titration of reovirus (Subheading 3.3), and ﬂow cytometric analysis of reovirusinduced cell death (Subheading 3.4). Using these straightforward methods, reovirus replication and spread can be compared between various cell lines and under various tissue culture conditions, thereby providing important information on cellular and viral factors that promote preferential virus replication in transformed or cancer cells. For example, efﬁciency of reovirus replication and spread can be monitored in cells from various cancer origins, with or without oncogene activation, inhibitors of cell signalling, host gene complementation, siRNA-mediated knock-down, or following infection with various reovirus mutants.

2. Materials 2.1. Preparation of Reovirus-Infected Samples

1. Cell culture medium according to ATCC. 2. Dulbeco’s modiﬁed Eagle’s medium (DMEM). 3. Phosphate-buffered saline (PBS): 137 mM NaCl, 2.7 mM KCl, 4.3 mM Na2HPO4, 1.47 mM KH2PO4. 4. U0126 (MEK1/2 inhibitor, Calbiochem): Make stock solution of 10 mg/mL (24.8 mM) by dissolving 1 mg in 100 PL of dimethyl sulfoxide (DMSO). Aliquot and store at −20°C for 1 month. Dilute to 10 PM (ﬁnal concentration) in media immediately before use. 5. Z-VAD(OMe)-FMK (Calbiochem): Make stock solution of 20 mM (1 mg/107PL) in DMSO and store at −20°C for up to 6 months. Dilute to 20 PM (ﬁnal concentration) in media immediately before use.

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6. Reovirus serotype T3 Dearing (ATCC): Alternatively, reovirus stocks can be produced in a level 2 biosafety facility (8). 2.2. Immunohistochemistry of Reovirus-Infected Cells

1. Methanol, pre-cooled at −20°C. 2. Antibody block solution: PBS with 0.1% triton X-100 and 3% (w/v) of fraction V bovine serum albumin (BSA). Store at 4°C. 3. Monoclonal antibody supernatants against reovirus serotype T3 V3 protein (9) (4F2, Developmental Studies Hybridoma Bank), stored at 4°C for 1–2 months or diluted to 50% glycerol for long-term storage at −20°C. 4. Secondary antibody: Horseradish peroxidise (HRP)-conjugated goat anti-mouse antibody (Jackson ImmunoResearch) reconstituted with 50% glycerol according to datasheet. 5. Sodium azide. 6. 100 mM Tris–HCl, pH 9.5. 7. Wash solution: PBS with 0.1% triton X-100, store at 4°C. 8. BCIP/NBT substrate working solution: BCIP/NBT Alkaline Phosphatase Substrate Kit IV (Vector Laboratories). Immediately before use, add two drops each of reagents 1, 2 and 3–5 mL of 100 mM Tris–HCl, pH 9.5 buffer. Mix between additions of each reagent. 9. Stop solution: PBS containing 5 mM EDTA. 10. 4,6-diamidino-2-phenylindole (DAPI, Molecular Probes): Stock solutions of 5 mg/mL (14.3 mM) DAPI in deionized water can be stored at −20°C. Dilute to the working concentration (0.5 Pg/mL) in PBS immediately before use.

2.3. Plaque Titration of Reovirus

1. 10× RIPA: 50 mM Tris, 150 mM NaCl, 5% (w/v) sodium deoxycholate, 10% NP40. Allow to mix with stir bar 6 h overnight until detergents are dissolved. 2. 1× RIPA: 50 mM Tris, 150 mM NaCl, 0.5% (w/v) sodium deoxycholate, 1% NP40. 3. Protease inhibitor cocktail (Sigma). 4. L929 cells (ATCC). 5. L929 growth medium: Minimal essential media (MEM), 10% horse serum (HS), 1× antibiotic–antimycotic. 6. L929 seeding medium: MEM with 10% fetal bovine serum (FBS) and 1× antibiotic–antimycotic. 7. Methyl cellulose (MC) overlay: Add 5 g MC (Sigma) to 215 mL of water (it does not dissolve) and autoclave for 20 min with a stir bar (it creates a gelatinous white gloop). Let stir overnight at 4°C to dissolve. Add 250 mL 2× MEM (GIBCO), 25 mL FBS, 5 mL 100× antibiotic–antimycotic, and 5 mL

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100× sodium pyruvate. Store at 4°C; pre-warm to 37°C prior to titration. 8. Crystal violet (Sigma): 1% (w/v) in 50% ethanol/50% water. 9. 96-well storage plates (Fisher) for routine large-scale plaque titrations. 2.4. Reovirus-Induced Apoptosis

1. PBS with 5 mM EDTA. 2. Annexin V PE (BD Pharmingen). 3. 7-Amino-acitomycin D (7AAD, BD Pharmingen). 4. 10× binding buffer: 0.1 M Hepes (pH 7.4), 1.4 M NaCl, 25 mM CaCl2. Store at 4°C. Dilute to 1× solution with deionized water immediately before use.

3. Methods Reovirus replication and spread can be evaluated in different cancer cell lines under various conditions to reveal the basis of reovirus oncolysis. When exploring reovirus oncolysis in a new cell line in culture, IHC can be used initially to determine the dose of reovirus necessary to infect the desired number of cells for subsequent experiments (follow steps 3a and 2a in Subheadings 3.1 and 3.2, respectively) (Fig. 1a). The amount of virus needed to infect various cell types varies depending on factors such as availability of receptors and uptake of virus (Fig. 1a; compare H1299, A549, and U2OS cells). Once the optimal reovirus dose for the desired level of infection is determined, IHC can be applied to monitor the extent of viral cell-to-cell spread over several rounds of infection (follow steps 3b and 2b in Subheadings 3.1 and 3.2, respectively). For example, cell-to-cell spread of reovirus (monitored by IHC) in U20S cells (a human osteosarcoma cell line), U2OS cells transduced with K-RasV12, and A549 cells (a Human lung adenocarcinoma epithelial cell line) is reduced by the MEK/ERK inhibitor U0126 (10), supporting a role for activated MEK/ERK signalling in promoting reovirus dissemination in these cells (Fig. 1b) (7). Note that although A549 cells were more conducive to virus uptake (Fig. 1a), they did not support efﬁcient virus dissemination when compared to U2OS (Fig. 1b), suggesting that cellular factors affecting post-entry steps of virus infection help dictate susceptibility to reovirus. An alternative means of assessing the extent of reovirus replication and spread is by measuring the titres of reovirus produced by cells (follow Subheadings 3.1 with step 3c and 3.3). Total titres (cell associated and released) reﬂect the ability of reovirus to complete the various steps of virus replication, from cell binding to assembly

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Fig. 1. MEK/ERK signalling supports reovirus dissemination in A549 and U20S cells. (a) A549, H1299, and U2OS cells were exposed to serial dilutions of reovirus at indicated multiplicity of infection (MOI, based on titres obtained using L929 cells). Bright-ﬁeld microscopy shows reovirus-infected cells (black) versus uninfected cells (grey) following IHC staining with reovirus V3-speciﬁc anti-reovirus monoclonal antibody supernatants. MOIs corresponding to approximately 50% infected cells are indicated (white numbers). (b) A549, U2OS, and K-RasG12V expressing U20S cells (1.25 × 105 cells/cm2) were infected with reovirus (MOI of 0.5, 10, and 10, respectively) and incubated for 18, 48, or 72 h in the presence of no inhibitor (untreated), MEK1/2 inhibitor (U0126, 10 PM), or polyclonal rabbit anti-reovirus antiserum (2.5 Pl/mL). IHC shows the extent of reovirus cell-to-cell spread over time. A549 cells have a K-RasG12S mutation, but presumably lack downstream host factors that promote robust reovirus replication and cell-to-cell spread. (c) DAPI staining showed similar cell numbers between conditions within a single time point. Immunoﬂuorescence comparison of DAPI-stained A549 cells with and without treatment with U0126 or reovirus antibodies at 72 hpi is provided as an example.

of progeny virions. Figure 2 shows that total reovirus titres are increased by Ras transformation even in the ﬁrst round (24 h) of virus replication. If titres are increased by a speciﬁc oncogene or cell culture condition, then the precise step of virus replication that is enhanced can be explored with speciﬁc assays for every step of the reovirus life cycle (as was done previously with H-Ras- and non-transformed NIH3T3 cells (5)). Released titres reﬂect the

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Fig. 2. Reovirus titres are enhanced within the ﬁrst round of replication in HRasV12-transformed relative to non-transformed NIH3T3 cells. In lieu of the heterogeneity of cancer cell lines, we commonly use isogenic cell lines with and without oncogenic transformation to study host factors that promote reovirus replication. Cells are transduced with retrovirus made from pBabe-puro vector (referred to as “non-transfomed cells”) or the same vector expressing a constitutively active H-Ras-V12 oncogene (referred to as “Ras-transformed” cells). The photograph shows crystal violet-stained plaques formed on L929 cells infected with total lysates collected at 16 and 24 h post infection from H-RasV12-transformed and nontransformed NIH3T3 cells exposed to the same reovirus dose. Tenfold serial dilutions used are indicated. Number of plaques is indicated in brackets for corresponding dilutions. Final titres are 7.8 × 106, 9.4 × 107, 1.9 × 107, and 2.7 × 108 (left-to-right columns). A complete 48-h time course was previously published (5).

extent of cell death and release of progeny virions, and are presented as a fraction of total titres (i.e. two samples with similar total titres, but different released titres are likely to have different extents of cell death). Cell death of cancer cells is the ultimate desirable consequence of infection by an oncolytic virus. The extent of reovirus-induced cell death and apoptosis can be assessed directly using ﬂow cytometry analysis of cells following staining with Annexin V and 7AAD (follow Subheadings 3.1 with step 3d and 3.4). Cells staining positive for both Annexin V and 7AAD are considered dead. For example, the extent of reovirus-induced apoptosis was measured in H-Ras- versus non-transformed cells and found to be enhanced by activated Ras signalling (Fig. 3) (5). To look exclusively at cell death caused by apoptosis, a time course can be used to evaluate the numbers of cells that stain positive for Annexin V but negative for 7AAD (apoptotic but not dead). Alternatively, ZVAD can be included as a control to indicate the extent of death attributed directly to caspasemediated apoptosis (Fig. 3).

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Fig. 3. Ras transformation of NIH3T3 mouse ﬁbroblasts enhances reovirus-induced apoptosis. H-RasV12- or non-transformed NIH3T3 cells (1 × 105 cells/cm2) were exposed to media alone or reovirus at a dose sufﬁcient to infect 60–70% of cells (according to IHC and ﬂow cytometric quantiﬁcation) in the presence or absence of 20 PM (ﬁnal ) ZVAD-fmk. Extent of cell death (upper right quadrant) and early apoptosis (lower right quadrant) was measured at 24 h post infection with Annexin-PE and 7AAD. Numbers indicate the percent of cells (out of 10,000 total) within that quadrant. A more comprehensive analysis of reovirus-induced cell death was previously published (5).

3.1. Preparation of Reovirus-Infected Samples

1. Seed cells to be 70–90% conﬂuent the following day in 12-well plates in 1 mL (see Notes 1 and 2) using tissue culture conditions suggested by ATCC. Cell numbers for seeding must be determined empirically, but can be estimated based on surface area and doubling time. 2. Infect cells on the following day. Aspirate tissue culture media and wash cells once with PBS (see Note 3). Discard PBS and add 100 PL of diluted reovirus per well. Be sure to include mock-infected controls (DMEM only or UV-inactivated reovirus, see Note 4). (a) For initial determination of how much reovirus to use for a given cell line: Determine the titre of the reovirus stock on L929 cells using plaque titration method (Subheading 3.3). Dilute reovirus stock in DMEM (no serum) at a starting multiplicity of infection (MOI) of 100 (i.e. 100 plaque forming units (pfus) per cell in 100 PL DMEM). Make 11 serial one-half dilutions in DMEM (no serum) and apply 100 PL of each dilution to a separate well of cells. Depending on results, procedure may require repetition with higher or lower dilutions of virus to obtain ideal infection conditions. (b) For monitoring reovirus cell-to-cell spread by IHC: Use reovirus dose predetermined to infect approximately 5–10%

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of cells. Low infection ensures that spread to neighbouring cells shows observable differences over several rounds of replication. (c) For monitoring reovirus titres: Use low reovirus dose (5–10% infected cells) if titres will be monitored over several rounds of replication to assess dissemination, but high reovirus dose (50–80% infected cells) if titres will be monitored to assess the efﬁciency of a single round of virus replication. (d) For monitoring reovirus-induced apoptosis: Use reovirus dose predetermined to infect approximately 50–80% of cells. This ensures sufﬁcient infected cells to produce notable extent of apoptosis. 3. Swish the plate several times to coat cells with a wave of media. Place plates in a 37°C incubator (5% CO2) for 1 h, swishing every 10 min. 4. Aspirate the virus, starting at most dilute samples, and replace with complete media speciﬁc for that cell type (according to ATCC). This allows synchronous infection. 5. Cells can be treated with various agonists or inhibitors immediately following infection to assess the effects of these compounds on virus replication and subsequent spread. To determine the role of the MEK/ERK pathway on reovirus replication, cells were treated with 10 PM (ﬁnal) of U0126 immediately following infection for the duration of the experiment (Fig. 1b). Similarly, cells were treated with 20 PM (ﬁnal) of ZVAD-fmk (inhibitor of caspase-mediated apoptosis) as a control to limit the extent of virus-induced apoptosis (Fig. 3) (see Note 5). 6. Incubate cells in the 37°C incubator (5% CO2) until you are ready to collect samples for titres, IHC, or apoptosis assays. 3.2. Immunohistochemistry of Reovirus-Infected Cells

1. Fix cells at desired time points post infection. Aspirate the media, wash one time with PBS, and incubate cells with 1 mL/ well of ice-cold methanol for 5 min. (a) For initial determination of how much reovirus to use for a given cell line: Fix cells at 18 hpi to monitor the number of cells infected in the ﬁrst round of reovirus replication. (b) For monitoring reovirus cell-to-cell spread: Stop cells at 18 hpi (ﬁrst round of replication), but also at 48, 72, and 96 hpi to monitor subsequent rounds of reovirus release and reinfection. 2. Aspirate the methanol, wash once with PBS, and incubate with 1 mL block solution for at least 30 min. Cells can be stored at 4°C for 1–4 weeks or used immediately for subsequent IHC. For long-term storage, add 0.05% sodium azide (preservative).

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3. Aspirate blocking solution and add 400 PL per well of monoclonal antibody supernatants against reovirus V3 diluted 1:100 in block solution (see Note 6). Incubate for 1 h at room temperature. 4. Aspirate primary antibodies and wash cells three times with wash solution for 15 min per wash. 5. Incubate cells for 1 h with 400 PL of secondary antibody diluted in block solution (1:1,000). 6. Discard secondary antibody and wash cells three times with wash solution for 15 min per wash. 7. Discard wash solution and add 400 PL of fresh BCIP/NBT substrate working solution per well. 8. Incubate the cells at room temperature until desired staining intensity develops (10–30 min). 9. Stop the reaction with 1 mL stop solution; incubate for 5 min. 10. (Optional) For ﬂuorescent detection of nuclei to show all (infected and uninfected) cells, add 1 ml of DAPI diluted to 0.5 Pg/ml in PBS, incubate for 1 min at room temperature, and rinse two times with PBS. 11. Phase-contrast microscopy can be used to visualize reovirus-infected (dark stained) and uninfected (unstained) cells (Fig. 1a, b). DAPI-stained nuclei can be visualized by immunoﬂuorescence microscopy (Fig. 1c, absorption and emission maximum at 358 and 461 nm, respectively). 12. Cells can be stored in PBS, water, or dried for long-term storage. 3.3. Plaque Titration of Reovirus

1. Prepare samples for plaque titration: (a) Puriﬁed reovirus or (b) Total titres (see Note 7): To generate lysates for total (cell associated and released) titres, remove 1/10th volume of the media from reovirus-infected cells and replace with 1/10th ﬁnal volume of 10× RIPA (with protease inhibitor cocktail). Using a pipet (p1000) tip, scrape cells and collect in a microcentrifuge tube. Flash freeze lysates in liquid nitrogen and store at −80°C. (c) Released titres: The amount of reovirus released from cells can be measured directly from the media of reovirusinfected cells. Remove media and centrifuge at 1,000 × g for 5 min to remove dead or ﬂoating cells. Transfer and ﬂash freeze media in liquid nitrogen and store at −80°C. 2. Seed L929 cells in 24-well plates to be 85–100% conﬂuent on the day of titration. We routinely use one conﬂuent (100%) 150-cm2 dish to seed six to eight plates, with 500 PL of cells

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per well, 24 h prior to titration. Substitute media with 10% FBS rather than 10% HS (see Note 8). 3. On the day of titration, dilute puriﬁed reovirus or lysates in PBS. Until the titres can be accurately estimated, make eight serial 1:10 dilutions and use dilutions from 10−3 to 10−8. For large-scale experiments, 8–12 samples can be conveniently diluted (50 PL sample in 450 PL PBS) in 96-well storage plates (1–2 mL/well capacity) using a multipipette. 4. Aspirate the media from one plate of L929 cells, starting with the wells that receive the lowest to highest amount of reovirus (see Note 9). Using the same pipette tip for a given sample, add 50 PL of the diluted virus to the corresponding well, starting at the lowest to highest amount of virus. 5. After one plate is complete, swish the plate to ensure that cells are coated with a “wave” of liquid. Put the plate aside until all titrations are complete, but do not allow it to stand more than 10 min without a swish. 6. When all virus dilutions have been added, place 24-well plates in the 37 C incubator (5% CO2) for 1 h, swishing every 10 min. 7. Without removing the virus (see Note 10), add 1 mL of 1% methyl cellulose overlay per well. 8. Incubate plates for 7 days at 37°C (5% CO2). Importantly, plates cannot be disturbed during incubation, as the assay relies on the inability of virus to diffuse large distances. 9. To stain plaques, aspirate the methyl cellulose. Add approximately 500 PL of 100% methanol to cells and let sit for 5–20 min. Remove methanol and replace with enough 1% crystal violet solution to cover the bottom of each well. Let sit for 5 min. Pour the concentrated crystal violet solution back into the container for reuse (see Note 11), and then rinse the plate with slow-running tap water several times until all residual crystal violet is removed. Plates can be stored indeﬁnitely when dry. 10. Count the plaques in the wells, where the most number of plaques can be accurately distinguished. Where possible, count wells containing 40–200 plaques. Calculate the titre using the formula: titre (pfu/mL) = number of plaques × virus dilution factor × 20 (to account for the addition of 50 PL and recalculate to 1 mL). An example is provided in Fig. 2. 3.4. Reovirus-Induced Apoptosis

1. When measuring reovirus-induced apoptosis, be sure to include controls to assess background apoptosis of uninfected cells and background apoptosis of cells treated with the various agonists or inhibitors. Also, include a positive control of cells infected with

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very high doses of reovirus to set ﬂow cytometry parameters. Two to three wells of a 12-well plate (with the same condition) can be pooled for apoptosis analysis if cell numbers are low. 2. At 24 h post infection (see Note 12), remove and keep media (containing dead or detached cells). Store media and subsequent collections in a 15-mL plastic centrifuge tube on ice. 3. Wash cells with 1 ml PBS containing 5mM EDTA. Pool the wash with media collected in step 2. 4. Add 1 mL PBS with 5 mM EDTA and incubate at room temperature for 10 min (see Note 13). 5. Using a pipet-aid and a 10-mL pipet loaded with 2 mL of PBS, spray cells to dissociate from plate. Repeat up to ﬁve times until all cells are dissociated. Apply the same technique for all wells (even if cells dissociate faster) to ensure consistency. 6. Add cells from step 5 to the 15-mL tubes from steps 2 and 3, and pellet cells by centrifugation at 500 × g for 7 min at 4°C. 7. Gently wash cells once with 10 mL of cold PBS, and then pellet cells by centrifugation at 500 × g for 7 min at 4°C. 8. Re-suspend cells gently in 1× Annexin binding buffer at 1 × 106 cells/mL. 9. Transfer 100 PL of the solution (1 × 105 cells) to an FACS analysis tube. For positive control samples (i.e. highly infected and apoptotic), make two tubes with 100 PL each, labelled “Annexin-only” or “7AAD-only”. 10. Add 5 PL of Annexin V-FITC or Annexin V-PE (see Note 14) directly to the cell suspension in every tube, except the 7AADonly tube. Flick tubes gently to mix. 11. Add 10 PL of 7AAD directly into the cell suspension in every tube, except the Annexin-only tube, followed by gently mixing. 12. Allow cells to incubate for 15 min in the dark at room temperature. 13. Add 400 PL of binding buffer to each tube, and analyze by ﬂow cytometry within 1 h. 14. These instructions assume the use of the FACScan ﬂow cytometer (Becton Dickinson), but can be adapted to other ﬂow cytometry formats. Using the positive-control Annexin-only and 7AAD-only samples, parameters are set to produce two distinguishable cell populations in FL2 (for Annexin-PE) and FL3 (for 7AAD) channels. The negative control (uninfected, untreated, healthy cells) is used to conﬁrm that parameters produce only a single population with low ﬂuorescence for both channels. Once the parameters are set, samples are processed to count 10,000 cells per condition.

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15. Results are analyzed using one of several FACS analysis software (examples: FCS express 3 from De Novo Softwares, WinDMI Version 2.8 from the Scripps Research Institute, and BD CellQuest from BD Biosciences). An example is provided in Fig. 3.

4. Notes 1. Methods can be adapted to 24-well plates by using 1/2 volumes at every step. 2. For cells that are highly transformed and do not grow in monolayers, pre-coat the tissue culture plates with gelatin to encourage cell-surface adhesion. Solubilize Gelatin (Sigma) at 37°C, add 1 mL per well, and then collect for reuse. Aspirate residual gelatine and allow plates to dry for over 2 h, uncovered, in a tissue culture hood prior to cell seeding. 3. Reovirus is inactivated two- to tenfold by 10% horse serum and newborn calf serum. If using these serums, it is necessary to wash cells with PBS prior to virus infection. When using cells grown in FBS, the PBS wash is optional. 4. Mock-infected samples can either contain no virus or UV-inactivated reovirus. For the latter, diluted reovirus is treated with 6 × 9,999 PJ of UV light using the UV Stratalinker 1800 (Stratagene). Complete inactivation can be conﬁrmed by plaque titration (Subheading 3.3). 5. When using IHC or titres to follow reovirus cell-to-cell spread, neutralizing monoclonal antibody supernatants against reovirus V1 protein (G5, Developmental Studies Hybridoma Band) (11) can be added post infection to limit reovirus cellto-cell spread in negative-control samples. The amount of antibody necessary to neutralize reovirus and prevent cell-to-cell spread must be determined empirically. Premix 1 × 109 pfu of reovirus diluted in 50 PL (ﬁnal) DMEM (no serum) with increasing volumes of antibody in 50 PL (ﬁnal) of DMEM. Incubate at 37°C for 60 min and check reovirus titre according to Subheading 3.3. The amount of antibody sufﬁcient to cause t90% reduction in reovirus titre should be added to the postinfection media for neutralization in experiments. 6. Alternatively, IHC can be performed using polyclonal antibodies raised in rabbits against reovirus (diluted 1:10,000 in step 5 of Subheading 3.2). These antibodies are not commercially available, but can be generated for routine use (12). When using rabbit serum, use HRP-conjugated goat anti-rabbit secondary antibody (Jackson ImmunoResearch).

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7. The presence of detergent kills L929 cells at dilutions less than 10−2. Therefore, the detergent lysis method for collecting total lysates can only be used for titres expected to be higher than 10−3 pfu/mL. When titres are expected to be lower, we recommend using the freeze–thaw method. Scrape cells into the media (for total) or PBS (for cell associated) with a pipette tip and subject to three cycles of freeze (−80°C) and thaw (37°C) before titration. 8. Reovirus titres are reduced two- to tenfold by 10% horse serum and normal calf serum. Thus, compare titres from total lysates only when media constituents are identical. For L929 cells during titration, it is easiest to maintain cells in HS as suggested by ATCC but to seed them in FBS (limited inhibitory effect on reovirus titres) for plaque assays. This precludes having to wash cells with PBS prior to titration. 9. Cell drying is a common problem during reovirus infection, and especially during plaque assays. To avoid this, turn off the fan in the tissue culture hood, empty media from wells in the same order that virus is added, and work swiftly. When media is insufﬁcient to cover the cells, plates should be “swished” every 5–10 min. 10. When infecting L929 cells for plaque titration assays, the virus does not need to be removed from cells, since the overlay prevents further infection. When infecting cells for experiments (without overlays), however, it is important to remove the reovirus, thus ensuring synchronous infection. 11. Crystal violet is toxic to aquatic organisms. We, therefore, collect and continually reuse the concentrated solution without any visible loss in plaque clarity. 12. A single round of reovirus replication from entry to virus release is approximately 24 h in several cell lines (e.g. L929, NIH3T3, HCT116) under moderate infection conditions (50–80% reovirus-infected cells). For a new cell line, the length of a single round of replication must ﬁrst be determined empirically by IHC (to visualize cell-to-cell spread) or titres of released virus. 13. Using PBS with 5 mM EDTA to dissociate cells works well for some cells (e.g. NIH3T3). If cells do not dissociate easily with this method, use 100 PL of 0.05% trypsin–EDTA or trypsin– EDTA diluted 50:50 with PBS/5 mM EDTA. Collect cells immediately following signs of dissociation using culture media (with 10% serum) instead of PBS in step 5. Trypsin removes cells more readily, but can provide inaccurate results with some cell types. 14. Annexin V FITC (BD Pharmingen) can be used as an alternative to Annexin V-PE. Using FITC eliminates the need for compensation during ﬂow cytometry capture or analysis, as there is minimal overlap in ﬂuorescence with 7AAD.

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through negative regulation of RIG-I signaling. Cancer Res 70: 4912–21. Mendez I.I., Hermann L.L., Hazelton P.R., Coombs K.M. (2000) A comparative analysis of freon substitutes in the puriﬁcation of reovirus and calicivirus. J Virol Methods 90: 59–67. Virgin H.W., Mann M.A., Fields B.N., Tyler K.L. (1991) Monoclonal antibodies to reovirus reveal structure/function relationships between capsid proteins and genetics of susceptibility to antibody action. J Virol 65: 6772–81. Duncia J.V., Santella J.B., III, Higley C.A. et al. (1998) MEK inhibitors: the chemistry and biological activity of U0126, its analogs, and cyclization products. Bioorg Med Chem Lett 8: 2839–44. Tyler K.L., Mann M.A., Fields B.N., Virgin H.W. (1993) Protective anti-reovirus monoclonal antibodies and their effects on viral pathogenesis. J Virol 67: 3446–53. Virgin H.W., Bassel-Duby R., Fields B.N., Tyler K.L. (1988) Antibody protects against lethal infection with the neurally spreading reovirus type 3 (Dearing). J Virol 62: 4594–604.

Chapter 13 Analysis of Three Properties of Newcastle Disease Virus for Fighting Cancer: Tumor-Selective Replication, Antitumor Cytotoxicity, and Immunostimulation Philippe Fournier, Huijie Bian, József Szeberényi, and Volker Schirrmacher Abstract Newcastle disease virus (NDV), a bird paramyxovirus, is an antitumor agent which has shown beneﬁts to cancer patients. Its antineoplastic efﬁcacy appears to be associated with three properties of the virus: 1. Selective replication in tumor cells. This feature can be studied at the RNA level, for example by RT-PCR, and at the protein level by immunochemistry. 2. Oncolytic properties (of some strains). The use of cultures of tumor cell lines represents a selective model to study direct viral oncolysis at the cellular level. The capacity of NDV to lyse tumor cells can be analyzed in vitro using cytotoxic assays based on the WST1 chemical reagent. The endoplasmic reticulum stress, which is induced by infection with the oncolytic NDV strain MTH-68/H and which plays an important role in the viral oncolytic effects, can be analyzed by Western blotting using speciﬁc monoclonal antibodies. Such stress appears as a key component of NDV cytotoxicity. 3. Immunostimulatory capacity. We describe an in vitro test called “Tumor Neutralisation Assay” which allows the analysis of bystander antitumor immune effects induced in human peripheral blood mononuclear cells by NDV. There are two variants, one for oncolytic NDV strains and the other one for nonlytic NDV strains. NDV may use several mechanisms to exert its tumor-killing action: direct cytotoxicity against cancer cells but also nonspeciﬁc as well as active-speciﬁc antitumor immune responses from the host organism. All the methods described here allow to evaluate the different oncolytic and immunostimulatory capacities of various strains of NDV. They are crucial to harness optimal antitumor activity by appropriate combinations of virus strains and application regimens. Key words: Newcastle disease virus, Tumor/cancer, transformed cells, hepatic stellate cells, Selective replication, oncolytic, Viral oncolysis, Apoptosis, Endoplasmic reticulum stress, Cytotoxicity, Immunostimulatory action or immunostimulation, Type I interferon, Tumor vaccination, Immunotherapy

David H. Kirn et al. (eds.), Oncolytic Viruses: Methods and Protocols, Methods in Molecular Biology, vol. 797, DOI 10.1007/978-1-61779-340-0_13, © Springer Science+Business Media, LLC 2012

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List of Abbreviations D-SMA A490 ATV-NDV BSA cDNA CT ELISA ELISPOT ER HN HSC HU IFN IRF-3 IRF-7 MOI NDV pAb PBMC PBS PCR PHA RIG-I rpm RPNI RT-PCR SDS STAT Tet TGI TNA UV

D-Smooth muscle actin Absorbance at 490 nm Autologous tumor vaccine with NDV Bovin serum albumin Complementary DNA Threshold cycle Enzyme-linked immunosorbent assay Enzyme-linked immunosorbent spot Endoplasmic reticulum Hemagglutinin protein of NDV Hepatic stellate cell Hemagglutination unit Interferon Interferon regulatory factor 3 Interferon regulatory factor 7 Multiplicity of infection Newcastle disease virus Polyclonal antibodies Peripheral blood mononuclear cell Phosphate-buffered saline Polymerase chain reaction Phytohemagglutinin Retinoic acid-inducible gene I Rotation per minute Roswell park memorial institute medium Real-time PCR Sodium dodecyl sulfate Signal transducers and activation of transcription Tetracycline Tumor growth inhibition Tumor-neutralization assay Ultraviolet

1. Introduction Although Newcastle disease virus (NDV) can induce fatal respiratory diseases in birds (e.g., chicken pest), this avian paramyxovirus is not a human pathogen (1, 2). NDV is an enveloped virus of 100–300-nm diameter with a negative-sense single-stranded RNA genome of roughly 16,000 nucleotides. This viral RNA contains six genes encoding six major polypeptides, among them two surface proteins: the hemagglutinin– neuraminidase (HN) protein (74 kDa) and the fusion (F) protein (67 kDa). These two proteins are involved during NDV infection,

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respectively, in the binding of the virus to a host cell’s surface via ubiquitously expressed sialic acid containing receptors and in the fusion of the viral and the host cell membrane. This membrane fusion event allows the viral genome to enter the cytoplasm of the host cell. It is important to mention that this viral genome replicates only in the cytoplasm and, not having a DNA intermediate, virus-speciﬁc sequences do not integrate into the genome of the infected cell. At the beginning of viral infection, the negativestranded RNA genome is transcribed within the cytoplasm into messenger RNAs (positive) and translated to viral proteins (ﬁrst step of viral infection). The nucleocapsid as “antigenome” is then used as a template for viral replication (second step of viral infection). This virus can then produce progeny particles that can eventually infect other cells. One main property which is important for classifying NDV viral strains is their virulence. They are either lytic and show, then, a capacity of multicylic replication in tumor cells. Or they are nonlytic and have a monocyclic replication pattern in tumor cells (3). There is a long history of the use of both types of NDV strains for treating cancer in man. Therefore, there exists now an extensive safety database for NDV (4). It shows a high tolerability for humans. When exposed to oncolytic forms, the maximal tolerated dose is around at least 3.3 × 109 infectious particles when the virus is applied by the intravenous root and 4.3 × 1012 infectious particles by the intratumoral root (5). Since Cassel et al. made their pioneering studies in the 1960s (6), both types of NDV strains have been used for treating different types of cancer, such as melanoma or glioblastoma. NDV treatment following various settings showed beneﬁts to cancer patients (for more details, see review (7)). Different concepts based on the use of NDV have been developed. All of them are based on the three following features presented by NDV. 1. Selectivity of replication in tumor cells. A very interesting feature of NDV as antitumor agent is its tumor selectivity of replication as shown for mouse and human cells (8, 9). We present a protocol allowing the quantiﬁcation of both strands of viral RNA products within normal cells (for example, peripheral blood mononuclear cells (PBMCs)) and tumor cells (for example, established tumor cell lines). NDV has been also shown recently to be able to replicate efﬁciently in activated hepatic stellate cells (HSCs) when compared to nonactivated HSCs (10). This results in cell death of the activated HSCs. This observation is important since activated HSCs are responsible for liver ﬁbrosis and are also involved in the development of hepatocarcinoma. The application of NDV via the tail vein in a hepatic ﬁbrosis mouse model has been shown to lead to suppression of (1) collagen deposition and (2) A-smooth

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muscle actin (A-SMA) expression (10). A protocol of in vivo colocalization of NDV and A-SMA is provided. 2. Oncolytic activity. Oncolytic strains of NDV can be very efﬁcient in inducing tumor cell lysis (11) in contrast to nonlytic NDV strains (12). We present a protocol based on the use of the WST1 chemical reagent to reveal the cell viability allowing an easy determination of the oncolytic potential of the virus. MTH-68/H replicates in a multicyclic fashion in certain tumor cell lines (as HeLa cells) and may, thus, be considered velogenic. However, since it has been attenuated, some think it is mesogenic. Viral replication of this strain within tumor cells has been reported recently to induce a p53-independent, endoplasmic reticulum (ER) stress-mediated cytotoxicity in tumor cell lines (13). We, here, highlight a protocol of Western blotting experiments allowing the analysis of the induction of such cellular stress at the protein level. An increased phosphorylation of the endoplasmic reticulum (ER)-resident alpha subunit of the eukaryotic initiation factor 2 of translation (eIF2A) – which is a soluble cytosolic initiation factor – preceded by the activation of the protein kinase R (PKR)-like endoplasmic reticulum kinase (PERK) is characteristic for stress induced by viral replication. 3. Immunostimulatory activities. NDV has also very interesting immune-stimulating properties (14). We have reported recently that the antitumor effects of oncolytic NDV strains are not entirely due to direct antitumor effects of the virus and involve also host-mediated antitumor mechanisms (12). To analyze these bystander effects, we incubated PBMC with UV-inactivated MTH-68/H for several days and added these precultured PBMCs on the top of a monolayer of tumor cells. By this way, we could demonstrate and quantify host immune cell-mediated antitumor activity. A detailed setting of such in vitro experiments is given. We present also a variant protocol of these in vitro assays which we called “Tumor Neutralization Assays” (TNAs) (15). It can be performed only with lentogenic (nonlytic) NDV strains. This assay is based on the inhibition of the growth of a monolayer of tumor cells by human PBMC to which tumor cells – which were modiﬁed by inactivation through G-irradiation and infection by the nonlytic strain NDV Ulster – were added. This system allowed the quantiﬁcation of the bystander effects induced in vitro in PBMC by the virus-modiﬁed tumor cells. The three main features we describe here for NDV make this virus a particularly suitable vector for cancer viroimmunotherapy, although oncolytic and immunostimulatory properties appear to vary according to the NDV strain used.

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2. Materials 2.1. Cell Culture and NDV Viruses

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Medium Cell culture media (RPMI 1640 and Dulbecco’s modiﬁed Eagle’s medium (DMEM)) were supplemented with FCS, 2 mM L-glutamine, 10 mM HEPES, 100 U/ml penicillin, and 100 Mg/ml streptomycin (all purchased from Gibco Invitrogen, Karlsruhe, Germany). All the reagents for the cell culture were purchased from Gibco Life Technologies (Karlsruhe, Germany), unless otherwise mentioned.

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Cells All cell lines were obtained from the tumor cell bank of the German Cancer Research Center (DKFZ, Heidelberg, Germany). BT-20 (human breast cancer cell line); MCF-7 (human breast cancer cell line); Hela (human cervical carcinoma); NCH82 and NCH89 (human glioblastoma); NB Kelley, SK, Shep, WT (all as neuroblastoma and Wilm’s tumor cell lines); and Stomach Ca (stomach carcinoma cell line) were grown in DMEM + 10% FCS with supplements. The Jurkat cell line sorted for CD28 expression (JCD28) and the U937 promyelomonocytic cell line were cultured in RPMI-1640 medium with 10% FCS. The peripheral blood mononuclear cells (PBMC) (see Note 1) are cultured in RPMI1640 medium containing 2% FCS, 100 Mg/ml streptomycin, 100 U/ml penicillin, 2.5 Mg/ml fungizone, 2 mM L-glutamine, and 10 mM HEPES.

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Viruses (see Notes 2 and 3) MTH-68/H is a nonpathogenic, attenuated virus isolated from the Hertfordshire strain of NDV (16). It was kindly provided by Dr G. Noss (Malsch, Germany). For its puriﬁcation, a sucrose concentration of 50% (w/w) was used (12). The oncolytic NDV-Italien was obtained in 1986 from Dr H.D. Klenk (University Giessen, Germany). The nonlytic, avirulent Ulster NDV strain was provided originally by Dr P. Russell (London, England) and was used for the generation of the tumor vaccine ATV-NDV (see Note 4). NDFL-enhanced green ﬂuorescence protein (EGFP) (see Note 5) is derived from the nonlytic strain La Sota, in which the gene sequence for the EGFP has been inserted (17). This virus was kindly provided by Dr B. Peeters (Animal Sciences Group, Wageningen University and Research Centre, Division of Infectious Diseases, Lelystad, The Netherlands). HUJ, a lentogenic NDV strain, was kindly given from Theravir (Jerusalem, Israel) as lyophilized.

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2.2. Quantiﬁcation of Positive- and Negative-Strand Viral RNA by RT-PCR

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RNA-extraction kit (Qiagen, Hilden, Germany).

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SuperScript II RNase H Reverse Transcriptase Kit (Invitrogen, Karlsruhe, Germany).

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RNAse H (Invitrogen, Karlsruhe, Germany).

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Taq DNA-polymerase, PCR-buffer, a deoxynucleotide mix (all provided by Sigma, Deisenhofen, Germany).

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Primers –

For HN of NDV: Sense primer HN1 5c-GAGGCTAGCACACCTAGCGACCTT Antisense primer HN2 5c-CTGGCAGCGTAAGACTCG

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For glyceraldehyde 3-phosphate dehydrogenase (GAPDH): Sense primer G1 5c-AGTCAGCCGCATCTTCTTTTGC Antisense primer G2 5c-CTCCTGGAAGATGGTGGGA

2.3. Colocalization Assay of NDV and a-SMA

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PCR-thermocycler (MJ Research Inc., Watertown, USA).

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SPF Kunning mice.

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Carbon tetrachloride (CCl4)/peanut oil solution (20% v/v).

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BSA.

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Tween 20.

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Triton X-100.

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Fluorescent microscope.

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Sirius red (Maixin Bio, Fuzhou, China).

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2.4. WST1 Assay

Two primary antibodies: Mouse anti-A-SMA (Abcam, Cambridge, UK) and chicken anti-NDV serum (US Biological, Swampscott, MA, USA).

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Secondary antibodies: Alexa Flour 594 goat anti-mouse IgG (Molecular Probes, Eugene, OR, USA) and rabbit anti-chicken IgY-FITC (US Biological, Swampscott, MA, USA).

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DAPI (Molecular Probes, Eugene, OR, USA).

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Anti-actin antibody (Oncogene, Merck, Budapest, Hungary).

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Tetracycline (Tet) (Sigma–Aldrich, Hungary, Budapest, Hungary).

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Anisomycin (Sigma–Aldrich, Hungary, Budapest, Hungary).

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WST-1 reagent (Roche, Hungary, Budapest, Hungary).

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2.5. Analysis by Western Blot of the Endoplasmic Reticulum Stress Induced by NDV Infection

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Human glioblastoma cell line: LNZTA3WT4 cells expressing the wild-type p53 protein from a Tet-repressible promoter: in the presence (Tet+) or absence (Tet−) of Tet, they behave as p53− and p53+ cells, respectively.

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ELISA reader (PerkinElmer Wallac, Freiburg, Germany).

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Protein-assay-solution kit (Bio-Rad, Munich, Germany).

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Nonidet P40 lysis-buffer (Boehringer, Mannheim, Germany).

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Transfer device (Bio-Rad, Munich, Germany).

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Polyvinylidene diﬂuoride membranes (Amersham Pharmacia Biotech, Uppsala, Sweden).

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Nonfat-milk powder (Merck, Darmstadt, Germany).

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Enhanced chemiluminescence detection kit (Amersham Pharmacia Biotech, Uppsala, Sweden).

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Polyclonal antibodies (pAbs) speciﬁc for caspase-8 (Cell Signaling, Beverly, MA, USA).

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PAb speciﬁc for cleaved caspase-3 (Cell Signaling, Beverly, MA, USA).

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PAb speciﬁc for cleaved caspase-9 (Cell Signaling, Beverly, MA, USA).

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2.6. Tumor Growth Inhibition or TumorNeutralization Assay for Lytic and Nonlytic NDV Strains

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PAb speciﬁc for eIF2A (Cell Signaling, Beverly, MA, USA).

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PAb speciﬁc for phospho-eIF2 (Ser51) (Cell Signaling, Beverly, MA, USA).

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PERK- and phospho-PERK-speciﬁc (Thr981) antibodies from Santa Cruz Biotechnology (Santa Cruz, CA, USA).

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Caspase-12-speciﬁc pAb (MBL Laboratories, Nagoya, Japan).

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pAb speciﬁc for phospho-MDM2 (Cell Signaling, Beverly, MA, USA).

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pAb speciﬁc for p53 (Cell Signaling, Beverly, MA, USA).

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pAb speciﬁc for phospho-PTEN (Cell Signaling, Beverly, MA, USA).

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pAb speciﬁc for PTEN (Cell Signaling, Beverly, MA, USA).

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48-well plate for cell culture.

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96-well plate (round bottom) for cell culture.

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MTS (MTS CellTiter 96® AQueous, Promega, Mannheim, Germany).

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PHA (Sigma–Aldrich, Schnelldorf, Germany).

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Sodium dodecyl sulfate (SDS), 10% (Merck, Darmstadt, Germany).

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ELISA reader (PerkinElmer Wallac, Freiburg, Germany).

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Phyto-hemagglutinin (PHA).

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3. Methods 3.1. Analysis of the Selective Replication of NDV in Tumor and Transformed Cells 3.1.1. Quantiﬁcation of Positive- and Negative-Strand Viral RNA by RT-PCR Principle

This assay allows to compare the quantity of positive- and negativestrand viral RNA in tumorigenic and nontumorigenic cells after infection by NDV.

Protocol

1. Cells (of different origin) are washed twice with serum-free medium. Then, 1 × 106 cells are coincubated with 10 hemagglutination unit (HU) (see Note 2) of NDV in a volume of 100 Ml in a CO2 incubator at 37°C for 1 h. The cell–virus suspensions are shaken every 15 min. 2. Finally, the cells are washed twice with PBS (see Note 6). 3. To study virus replication within the cytoplasm, cells are then further incubated in suitable growth media for 1, 12, 24, and 48 h. 4. At each time point, samples of the virus-carrying cells are removed and processed for further analysis. 5. RNA is extracted from aliquots (1 × 107 cells/aliquot) using an RNA-extraction kit (see Note 7). 6. RNA concentration is measured at 260 nm. The ratio A260/ A280 is used to determine the purity of the RNA sample. 7. For the synthesis of cDNA from 1–5 Mg RNA, we used the SuperScript II Rnase H Reverse Transcriptase Kit according to the manufacturer’s instructions. 8. The obtained cDNA is puriﬁed by digesting the remaining RNA with RNAse H. 9. For ampliﬁcation of the speciﬁc cDNA segments of interest, 50-Ml aliquots are produced by mixing Taq DNA-polymerase, PCR-buffer, a deoxynucleotide mix, 3.5 Ml sense primer and antisense primer, and 10 ng of template cDNA. 10. To ensure that exclusively the negative strand or positive strand of NDV(HN)-RNA is ampliﬁed, use the sense primer 5c-GAGGCTAGCACACCTAGCGACCTT for ampliﬁcation of the negative-strand NDV(HN)-RNA and the antisense primer 5c-CTGGCAGCGTAAGACTCG for the positivestrand NDV(HN)-RNA.

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11. GAPDH is used as control with the following primers: sense primer 5c-AGTCAGCCGCATCTTCTTTTGC; antisense primer 5c-CTCCTGGAAGATGGTGGGA. 12. The samples are ampliﬁed in the PCR-thermocycler using the following parameters: 33 cycles; denature at 94°C for 55 s; anneal primers at 55°C for 60 s; extend at 72°C for 90 s. 13. The ampliﬁed DNA fragments are being evaluated by agarose gel electrophoresis. Observations and Conclusions

Here, we compared the production of HN-speciﬁc positive and negative NDV-RNA in PBMC from healthy donors and in two breast cancer cell lines (BT-20 and MCF-7) after coincubation with live and UV-inactivated NDV. The coincubation of the different cell types with UV-inactivated NDV led to no measurable production of NDVRNA. Upon coincubation with live NDV, in all tested cells, the amount of positive-strand RNA increased within 1 h. With regard to negative-strand RNA, we observed an increase in the two breast cancer cell lines BT-20 and MCF-7, 12 h after coincubation with live NDV, while in PBMC negative-strand RNA production remained weak and transitory (see Fig. 1). The PBMCs appear to have established an antiviral state, enabling them to block the replication cycle of NDV downstream of the positive-strand NDV-RNA production.

Fig. 1. Difference in positive- and negative-strand NDV(HN)-RNA production. Panel (a) shows kinetics of positive- and negative-strand HN-RNA in nontumorigenic cells (PBMC) and tumor cells (BT-20, MCF-7). RT-PCR of HN-speciﬁc RNA was performed after coincubation of the cells with live NDV Ulster for 1, 12, 24, and 48 h (0 h: uninfected control). RNA was extracted and cDNA of the NDV-HN-RNA sequences produced as described in the main text. The product of the RT-PCR was fractionated by agarose gel electrophoresis and analyzed under UV light. Panel (b) shows quantitative changes in the PCR band luminosity during 48 h of NDV infection of different cell types (Reproduced from ref. 9).

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3.1.2. Analysis of Replication of NDV in Hepatic Stellate Cells by Colocalization Studies of NDV and A-SMA Principle

Activated HSCs (ﬁbroblasts) are the crucial factor responsible for liver ﬁbrosis and involved in the development of hepatocellular carcinoma by interaction with tumor cells. This assay allows to visualize the selective replication of NDV in activated HSCs compared with quiescent HSCs.

Protocol

1. For induction of liver ﬁbrosis, mice are injected i.p. with 100 Ml carbon tetrachloride (CCl4)/peanut oil solution (20% v/v) twice a week for up to 8 weeks (see Note 8). 2. Each mouse is treated with 1,000 HU (see Note 2) of NDV Italien in a 200-Ml volume via the tail vein for one time or three times with a 24-h interval. 3. The mouse is sacriﬁced 24 h after the last injection of NDV. 4. Cryostat sections of liver tissues are ﬁxed with cold acetone for 20 min at −20°C. 5. After washing with PBS, the sections are permeabilized with PBS containing 0.1% Triton X-100 for 5 min. 6. Then, the sections are blocked with 5% BSA for 20 min at room temperature and two primary antibodies, mouse anti-ASMA (diluted 1:50) and chicken anti-NDV serum (diluted 1:200), are added to the slides and incubated for 1 h at 37°C. 7. Following washing with PBST (0.2% Tween-20 in PBS) three times, the sections are incubated with secondary antibodies, Alexa Flour 594 goat anti-mouse IgG (diluted 1:800), and rabbit anti-chicken IgY-FITC (diluted 1:200) for 30 min at 37°C in the dark (see Note 9). 8. After washing three times with PBST, the sections are stained with DAPI (1:5,000 dilution) for 2 min. 9. NDV virions and A-SMA expression can be visualized under a ﬂuorescence microscope (Fig. 2).

Observations and Conclusions

NDV was injected to normal and hepatic ﬁbrosis mice in parallel to investigate whether the regression of liver ﬁbrosis resulted from the NDV replication in activated HSCs. By double immunoﬂuorescence, NDV was detected in perisinusoidal spaces and colocated with A-SMApositive cells in liver tissues from CCl4-induced mice, which indicated the selective replication in activated HSCs. No speciﬁc deposition of virions was observed in normal mice. Importantly, parenchymal liver cells were found to be spared from virus infection both in normal and CCl4-induced mice after NDV injection (see Fig. 2A). The deposition of collagen ﬁbers in hepatic tissue allows to evaluate the progress of liver ﬁbrosis. We detected that excessive collagen ﬁbers were

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Fig. 2. Colocalization of A-smooth muscle actin (A-SMA) and NDV (A) and repression of hepatic ﬁbrosis (B) after NDV application. (A) Colocalization of A-SMA and NDV by immunohistochemistry. Hepatic ﬁbrosis in mice was induced by carbon tetrachloride (CCl4) for 8 weeks and then mice were injected with NDV Italien via the tail vein. Colocalization of A-SMA and NDV in (a) ﬁbrotic liver and (b) normal liver tissues by immunoﬂuorescence (×1,000). (B) Repression of hepatic ﬁbrosis by NDV in mice induced with carbon tetrachloride. Hepatic ﬁbrosis in mice was induced by CCl4 for 8 weeks and then mice were injected with NDV Italien (1,000 HU) via the tail vein once (marked 1+) or three times (marked 3+). Normal mice that were inoculated with physiological saline were subjected to the same injections as parallel controls. All the mice were killed at 24 h post injection. (a) The whole livers were excised for gross morphology. (b) Collagen ﬁbers were stained pink by Sirius red staining (×200). (c) The expression of A-SMA was detected by Western blotting. Both ﬁgures reproduced from ref. 10.

accumulated in the CCl4-induced mouse liver compared with the PBS group in which collagen ﬁbers deposited only around the blood vessel. These elevated collagen ﬁbers were suppressed by NDV injection (three times; see Fig. 2B(b)). Western blot analysis (Fig. 2B(c)) showed that A-SMA expression in the induced ﬁbrotic liver tissue was dramatically enhanced (lane 1). After treatment with NDV one (lane 2)

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or three times (lane 3), the expression of A-SMA in the ﬁbrotic liver decreased considerably (see Fig. 2B(c)). 3.2. Analysis of the Oncolytic Properties of NDV 3.2.1. WST1 Cell Viability Test Principle

This assay uses the WST-1 reagent (see Note 10) to detect cytotoxic effects on cells upon their infection with NDV. It is based on the reduction of WST-1 by viable cells. This reaction produces a soluble formazan salt. This dye can be quantiﬁed in a microplate using an ELISA plate reader. The absorbance directly correlates with the cell number.

Protocol

1. Cells are grown in tissue culture grade, 24-well plates, with 1 ml culture medium in each well. 2. They are then infected with MTH-68/H for 72 h. For positive apoptosis control, cells are treated for 24 h with 1 Mg/ml anisomycin and for negative control, they are treated with vehicle. 3. WST-1 is added directly to the cells (1/10th of the culture volume) (see Note 11). 4. Absorbance at 450 nm can be measured using an ELISA plate reader following a short incubation at 37°C (see Note 12).

Observations and Conclusions

The human glioblastoma cell line LNZTA3WT4 (18, 19) has two useful features important for studies with MTH-68/H. First, due to genomic rearrangements, it does not have endogenous wild-type p53 expression but produces p53 protein encoded by a transfected cDNA under the control of a Tet-repressible promoter: p53 is expressed in the absence of Tet in the medium but not in its presence. The data presented in Fig. 3 clearly show that MTH-68/H-induced tumor cell death is independent of the presence of functional p53.

3.2.2. Analysis by Western Blotting of the Endoplasmic Reticulum Stress Induced by NDV Infection Principle

This speciﬁc Western blotting allows the detection of virus-induced stress at the level of the endoplasmic reticulum.

Protocol

1. 5 × 106 cells were centrifuged at 537 × g for 2 min, resuspended in Nonidet P40 lysis-buffer, and put on ice for 15 min. 2. The cells lysates were cleared of insoluble material by centrifugation at 22,673 × g for 15 min at 4°C. 3. The protein concentration was measured using a protein-assaysolution kit.

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Fig. 3. p53-independent cytotoxicity of oncolytic NDV (MTH-68/H) in a human glioblastoma cell line. (a) WST-1 analysis of cell viability in the presence (ﬁlled squares) or absence (open squares) of 1 Mg/ml tetracycline (Tet) at various MOIs. Untreated and anisomycin-treated cultures were used as negative and positive controls, respectively. (b) Western blot analysis of proteins of the p53 pathway in LNZTA3WT4 cells in the presence (Tet+) or absence (Tet−) of 1 Mg/ml Tet. Tet strongly reduced the expression of the p53 protein as well as the expression and phosphorylation of p53-regulated MDM-2 and PTEN proteins. An antiactin antibody was used as loading control. UV irradiation was used as a positive control to stimulate the p53 pathway. (Reproduced from ref. 13 with permission from the American Society for Microbiology.) http:// jvi.asm.org/content/vol81/issue6/images/large/zjv006078879002a.jpeg.

4. Protein (100 Mg) for each cell extract is resolved by SDSpolyacrylamide gel electrophoresis in 10 or 18% gels. 5. The electroblotting was carried out in transfer buffer with a device for Western blot using a constant current of 30 mA per gel for 90 min (see Note 13). 6. The membrane was blocked by saturation with PBS containing 5% nonfat-milk powder and 0.5% Tween20 for 30 min at room temperature on a shaker.

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7. The primary antibody was diluted in milk–TTBS buffer (5% nonfat milk, 10 mM Tris buffer, 150 mM NaCl, 0.5% Tween20) and the blots were incubated in this solution at 4°C overnight. 8. After washing the blots three times with TTBS, they were incubated for 1 h in milk–TTBS buffer containing the secondary horseradish peroxidase (HRP)-conjugated antibody. 9. Blots were washed again three times and a Western blotting luminol reagent (enhanced chemiluminescence (ECL) reagent detection system) was added. The speciﬁc protein bands were then visualized in a darkroom by exposing light-sensitive photopaper to the luminous blots. Observations and Conclusion

MTH-68/H infection induces apoptotic cell death in several tumor cell lines (13). Interestingly, caspase-8, mediator of the extrinsic pathway of apoptosis, is not activated by MTH-68/H in PC12 cells (13), and the activation of caspase-9, a key element of the mitochondrial pathway of apoptosis, is a relatively late event during virus infection (Fig. 4). In contrast, stimulation of two endoplasmic

Fig. 4. Cellular stress induced upon NDV infection studied by Western blot analysis of apoptosis pathways in MTH-68/H-infected PC12 cells. (a) MTH-68/H-induced activation of caspase-3 and -12 precedes activation of caspase-9. (b) Early activation of the endoplasmic reticulum resident PERK enzyme precedes (c) increased phosphorylation of the translation initiation factor elF2A that leads to virus-induced shutdown of protein synthesis in the infected cells. (Reproduced from ref. 13 with permission from the American Society for Microbiology).

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reticulum-resident enzymes, phosphorylation of PERK, and proteolytic activation of caspase 12 take place early after MTH-68/H infection and precede the phosphorylation of the translation initiation factor eIF2A (Fig. 4). Caspase-12 and PERK are important proteins of the unfolded protein response of virus-infected cells that lead to endoplasmic reticulum stress, shutdown of protein synthesis by the inactivation of eIF2A, and ultimately to cell death (for a review, see ref. 20). 3.3. Determination of the Immunostimulatory Properties of NDV 3.3.1. TNA with Oncolytic NDV Strains Principle

This assay allows the evaluation of immune modulatory properties of NDV. For this purpose, NDV-infected irradiated MCF7 human mammary carcinoma-derived vaccine cells are coincubated with PBMC on an MCF-7 monolayer. After 6 days, viable cell staining with MTS (3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)-2H-tetrazolium) is per-formed and tumor growth inhibition (TGI) can be calculated.

Protocol

1. PBMCs are infected with different doses of UV-inactivated NDV strains (see Notes 3 and 14). 2. PBMCs are incubated in six-well plates (5 × 106 cells per well) for 3 days containing DMEM supplemented with 5% FCS. 3. Afterward, the PBMCs are transferred as triplicates to 48-well plates on an MCF-7 monolayer of 2.8 × 104 per well at 3.8 × 105 PBMCs per well. 4. After incubation at 37°C for 4 days, the plates are developed with MTS (see Note 15). 5. Measurements of the optical density at 490 using an ELISA reader. 6. Percentage of TGI is determined using the following calculations. TGI = 1 − ((A490 experimental well − A490 background)/ (A490 positive control − A490 background)) × 100

Observations and Conclusion

MTH-68/H (UV)-treated PBMCs (1,000 and 100 HU per 10(7) cells) showed a signiﬁcant augmentation of the %TGI compared to PBMCs without virus. Italien (UV) and Ulster (UV) strains of NDV did not induce a signiﬁcant TGI effect in vitro (see Fig. 5).

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MTH-68/H (UV) 10 00

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Fig. 5. Immune modulatory properties of different NDV strains. Human PBMCs were treated with NDV (UV) at different doses and placed in six-well plates at 5 × 106 cells per well. After 3 days, the PBMCs were transferred in triplicates onto MCF-7 monolayers at a concentration of 3.8 × 105 cells per well. Tumor growth inhibition was determined after four additional days by development with MTS. *p < 0.05. (Reproduced from ref. 12).

3.3.2. TNA (Bystander Assay) with Nonlytic NDV Strains Principle

We established a TGI assay in which NDV-preactivated effector cells are coincubated with a monolayer of live MCF-7 human breast carcinoma cells for 3 days. After this period, the effector cells are washed off. The remaining live tumor target cells are then quantiﬁed by staining them with MTS and reading light absorption at 490 nm using an ELISA reader.

Protocol

1. Seed live, adherent, growing tumor cells into 96-well roundbottom plates (U-bottom); for MCF7, take 3,000 cells/well in culture medium (see Note 16). 2. Incubate the plate for 4 h (or overnight) to allow cell adhesion to the bottom of the well. 3. Prepare the tumor cell vaccine by irradiation and NDV modification of the tumor cells (see Note 4). 4. Prepare PBMCs or other cells as for example T cells (see Note 1). 5. Adjust all cells needed for the assay to their respective cell numbers. 6. Give cells (vaccine cells (750 cells per well) with the indicated amounts of bsAbs (see Note 17) and PBMCs (effector cells)) into the microtiter plate to the adherent tumor cells so that the ﬁnal volume per well is 200 Ml (see Note 18).

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7. Take six wells for growth control (the wells contain only live, adherent, tumor cells). Experimental wells are performed in triplicate. 8. For positive control, take PHA, and for all other groups pipet triplicates. 9. Incubate plate at 37°C for 5–7 days (see Note 19). The incubation time depends on the growth of the tumor cells as observed in the control wells. 10. When control wells are conﬂuent, remove supernatants from the different wells (see Note 20). 11. Shake out plate carefully. 12. Wash the plate 2–3× with PBS 200 Ml/well under microscopic control to make sure that all effector cells are washed away and only adherent cells are left in the wells of the plate (see Note 21). 13. Then, add 100 Ml RPMI 1 5% FCS and 20 Ml MTS solution per well; as background control, give medium and MTS in 3–6 empty wells and incubate the plate for 60–120 min at 37°C in the dark (until the color changes from yellow to brown but not dark brown). 14. When the color switches from yellow to brown, the reaction in which the MTS tetrazol is converted to a formazan product is stopped by the addition of 20 Ml 10% SDS per well. 15. Measure the optical density at 490 nm of each well using an ELISA reader (see Note 22). 16. The background is taken from six control wells containing only medium and MTS solution. TGI is calculated according to the following formula: TGI = 1 − ((A490 experimental well − A490 background)/ (A490 positive control − A490 background)) × 100. Observations and Conclusions

Figure 6 shows the percentage of TGI of different tumor cell lines obtained with tumor vaccine generated from various tumor cell lines. We can observe a strong stimulation of the PBMC toward the different tumor cells upon coaddition of bsHN-CD3 and bsHN-CD28 to the tumor vaccine ATV-NDV.

4. Notes 1. Human PBMCs and lymphocytes: Heparinized samples from the peripheral blood of healthy donors are subjected to Ficoll density gradient centrifugation (Pharmacia, Uppsala, Sweden). The peripheral blood mononuclear cells (PBMC) are then harvested from the interphase. To obtain the lymphocyte fraction of PBMC, monocytes were ﬁrst depleted by an adhesion step.

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Fig. 6. Vaccine cell dependency of the TNA effect. The tumor growth inhibition (TGI) shown corresponds to 5-day TNA with PBMCs as effector cells. For modiﬁcation of vaccine cells, we used per 107 cells 1.0 Mg bsHN-CD3 and 7.3 Mg bsHN-CD28 (see Note 23). After 5 days of incubation, the nonadherent cells were removed and the adherent cells stained by addition of MTS solution. Triplicates were evaluated in an ELISA reader. Shown are means and standard deviations of three independent experiments. Shown are the means of four experiments with four different donors of PBMC (#1 to # 4) using MCF-7 cells (a), stomach carcinoma cells (b), or NHC82/NHC89 (c) as targets. Signiﬁcant (p < 0.05) vaccine cell typedependent differences were only seen with U937 cells compared to the three carcinoma lines, which behaved similarly. The columns that contain only two colors indicate that, in these groups, the results obtained with vaccine-NDV-bsHN-CD3 were identical to those obtained with vaccine-NDV. Shown are the means and standard deviations of three independent experiments. (Adapted partially from ref. 16).

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2. All viruses were propagated in embryonated chicken eggs, harvested from the allantoic ﬂuid, puriﬁed by ultracentrifugation as described previously (14) – unless speciﬁed – and cryopreserved in aliquots at −70°C. Quantiﬁcation was performed by hemagglutination (HA) assay. The smallest virus concentration leading to visible agglutination of sheep erythrocytes is deﬁned as 1 hemagglutination unit. 3. For the production of UV-inactivated formulations, NDVUlster and NDFL-EGFP were exposed to UV-light (254 nm, 2 mW/cm2, 7 cm distance) for 5 min. 4. To prepare the ATV-NDV vaccine, the tumor cells are ﬁrst inactivated by G-irradiation. This led to the loss of the ability of the cells to divide, where the cells remained vital and could still be cultured for at least another 3 days without dying. The required amount of cells was irradiated in suspension (in 15- or 50-ml centrifuge tube) with a 137 cesium source (Gammacell 1000; Atomic Energy of Canada, Chalk River, Canada). For mouse and human cells, a dose of respectively 100 and 200 Gy is used. In vitro infection of tumor cells with NDV strain Ulster. Cell suspensions were washed twice with FCSfree RPMI-1640 medium, and 1 × 106 cells were incubated with 10 HU of NDV in a ﬁnal volume of 100 Ml for 1 h at 37°C in a CO2 incubator. During the incubation, cells were shaken every 15 min. The cells were then washed twice with medium (250 × g, 5 min) before their use in assays. 5. The reverse genetics system allows the modiﬁcation of the RNA genome of NDV. 6. This step is to remove nonabsorbed virus particles. The celladsorbed virus can be visualized by FACS analysis using NDVspeciﬁc antibodies as shown in ref. 14. 7. During the extraction procedure for RNA, the operator must always wear disposable gloves, masks, and caps to prevent RNase contamination and Trizol reagent contact with the skin. Use sterile disposable plasticware and automatic pipettes reserved for RNA work to prevent cross-contamination with RNases from shared equipment. Disposable plasticware can be treated by deionized water containing diethyl pyrocarbonate (DEPC). 8. This assay can also be performed in vitro. 9. From incubating ﬂuorescein-labeled IgG to all later steps, the cells should be kept dark to avoid ﬂuorescence quenching. 10. WST-1 is a water-soluble tetrazolium salt that can be used for cell-proliferation or cell-viability assays. The rate of WST-1 cleavage by mitochondrial dehydrogenases correlates with the number of viable cells in the culture. The simple protocol is ideal for screening large numbers of samples in 96-well plates.

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WST-1 is a safe alternative to tritiated thymidine uptake and is a signiﬁcant improvement over XTT or MTS reagents. The advantages of WST-1 include safety, water solubility of the end product, stability of the reagent, scalability, and sensitivity. 11. When using WST-1, care should be taken to check for formation of precipitates in the reagent stock. Incubation at 37°C for 10 min followed by gentle agitation is always sufﬁcient to resolve the problem. It is important to have at least one well with medium alone (without cells) for blank control. 12. WST-1 added to the well should give background less than 0.2 absorbance units. The background value should be subtracted from the experimental wells for optimal results. Although WST-1 has a relatively wide linear range, the signal could become saturated after prolonged incubation. The problem could be avoided by measuring absorbance at several time points for the initial experiments. We found that optimization of the incubation time is necessary for every new application or cell type. In addition, it is important to remove any bubbles from the culture wells. A short exposure to warm air (from a hair dryer) is effective for that purpose. 13. Avoid air bubbles during the protein transfer to the membrane. Use a glass rod to extrude air bubbles. 14. When using live virus of strains Italien and MTH-68/H for vaccine production, a high antitumor effect with TGI up to 90% is observed irrespective of the presence of PBMC (data not shown). To exclude direct oncolytic effects of the virus in the TNA, we performed further experiments with UV-inactivated virus. 15. The reagents comprise the tetrazolium compound (3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4sulfophenyl)-2H-tetrazolium), inner salt; MTS and an electron-coupling reagent (phenazine methosulfate (PMS)). MTS is bioreduced by cells into a formazan product that is soluble in tissue culture medium. The absorbance of the formazan at 490 nm can be measured directly from 96-well assay plates without additional processing. The conversion of MTS into aqueous, soluble formazan is accomplished by dehydrogenase enzymes found in metabolically active cells. The quantity of formazan product as measured by the amount of 490-nm absorbance is directly proportional to the number of living cells in culture. 16. Use the medium in which the cells are used to grow; for each tumor cell line to be tested, one has to ﬁrst determine the amount of cells to be seeded per well of the 96-well plate. This depends on the size of the tumor cells and the rate of their division.

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17. We designed, produced, and tested bispeciﬁc molecules able to bind via one arm to NDV (of the vaccine cells) and via the other to the CD3− or CD28− molecules of human T cells for their activation. Such a vaccine was observed to induce strong antitumor activity in human lymphocytes in such in vitro TNA assays since activated cells were able to destroy very efﬁciently monolayers of tumor cells in these in vitro assays. 18. The ratio effector to target cell has to be deﬁned for each cell line to prevent or minimize allogeneic effects. 19. The microtiter wells are then wrapped in cling ﬁlm to prevent evaporation of medium and incubated in a CO2 incubator at 37°C. The time of incubation depends on the time of conﬂuence of the tumor cell monolayer in control wells without effector cells. 20. These supernatants (about 100 Ml/well) can be kept as froze for further eventual investigations (as for example cytokine determination by ELISA). 21. The plates have to be carefully washed (twice in PBS). 22. It is also possible to stop the reaction with 20 Ml 10% SDS to store the plate and to measure at a later time point. 23. The proteins bsHN-CD3 and bsHN-CD28 correspond to bispeciﬁc antibodies binding to the HN protein of NDV via one arm and to CD3 or CD28, respectively, via the other arm. The production, puriﬁcation and biochemical characterization of these proteins have been described in (15). Two permanent producing CHO cell lines (CHO-242 [bsHN-CD3] and CHO-290 [bsHN-CD28]) were established in dihydrofolate reductase (dhfr)-deﬁcient CHO cells and cultured in alphaMEM medium containing 5% dialysed fetal calf serum (dFCS), 2 mM L-glutamine, 100 U/ml penicillin, 100 Mg/ml streptomycin, and a concentration of methothrexate of 10 mM. BsHN-CD3 was puriﬁed from the production medium by immobilized metal afﬁnity chromatography (IMAC) and bsHN-CD28 by an anti-E Tag column.

5. Conclusions The characterization of the properties of several strains of NDV: MTH-68, HUJ, and Ulster as shown in Table 1 have been performed using various methods (some of them being described in this manuscript). Although the NV strains show diverse oncolytic and immunostimulatory properties, NDV has interesting antineoplastic features based on the three following properties.

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Table 1 Comparison of the oncolytic (A) and immunostimulatory (B) properties of the three NDV strains: MTH-68/H, HUJ, and Ulster NDV strains

(A) Oncolytic activity (in HU/well) MTS assayb Cervical Ca (Hela) Glioblastoma (NCH89) Crystal violet assayc Cervical Ca (Hela) Glioblastoma (NCH89) (B) Immune stimulatory properties (in pg/HU) Interferon-A induction assayd

HUJ 86500 HU/mla

Ulster

MTH68/H

80–100 100–200

10 2–3.5

1 × 10−4 – 1.5 × 10−3 1 × 10−4 – 1 × 10−3

537–2162 259.5

10–25 n.t.

2.5 × 10−6 12.8 × 10−6

0,18

0,18

200

n.t. not tested a The amount of NDV particles has been quantiﬁed by real-time reverse transcriptase PCR (RRT-PCR) based on the viral M gene. RNA was extracted from virus suspension via the QIAamp® Viral RNA kit (Qiagen GmbH, Hilden, Germany). cDNA was then synthesized using SuperScript II RNase H- reverse transcriptase and random hexamers (both from Invitrogen, Karlsruhe, Germany). RRT-PCR reactions were performed on a GeneAmp® 5700 sequence detection system (Applied Biosystems GmbH, Weiterstadt, Germany), all primers and probes were synthesized by MWG-Biotech AG (Munich, Germany). For absolute quantiﬁcation of the M gene of NDV, RRT-PCR was performed in a 25 Ml reaction mixture containing 18.5 Ml master mix (qPCR Core Kit, Eurogentec, Liege, Belgium), 300 nM of each primer for the M gene (sense 5c-AGT GAT GTG CTC GGA CCT TC-3c, antisense 5c-CCT GAG GAG AGG CAT TTG CTA-3c) and 100 nM probe for the M gene (5c-[FAM] TTC TCT AGC AGT GGG ACA GCC TGC [TAMRA]-3c). The samples were dilutions of viral cDNA or of a plasmid containing the M gene, named PCI-Neo-M (kindly provided by Dr B. Peeters, ID Lelystadt, The Netherlands). A common threshold was set during the exponential ampliﬁcation period to determine a speciﬁc threshold cycle (CT) for each sample. By correlating the CT values with the HU titer initially used for RNA extraction on the one hand and with the copy number of the M-plasmid on the other hand, M gene copy number per HU could be determined b On a 96-well plate, NDV strains were serially diluted and the Hela cell line was added at a concentration of 50,000 cells per well at a total volume of 100 Ml in their respective medium. After 48 h of incubation at 37°C, the cells were washed. And then, 200 Ml RPMI was added with 20 Ml MTS (MTS CellTiter 96® AQueous, Promega, Mannheim, Germany) to the wells and the plate was incubated for 15–60 min at 37°C in the dark. When the color switched from yellow to brown, reaction was stopped by addition of 20 Ml 10% SDS and the plates were analyzed in an ELISA at 490 nm. Percent of tumor growth inhibition (TGI) for each sample was calculated from the absorbance values at 490 nm (A490) according to [1 − (A490sample − A490background)/(A490negative control − A490background)] × 100. The number indicated in the table corresponds to the amount of virus that shows 50% of TGI. (1) Cervical Ca (Hela cell line); (2) Glioblastoma (NCH89) c Virus dilution endpoint assay with crystal violet staining. Several dilutions of NDV strains were added onto a monolayer of cells [here, (1) Cervical Ca (Hela) and (2) Glioblastoma (NCH89)] starting with a dose of 50 HU per well. After 6 days incubation in 5% CO2 atmosphere at 37°C, the medium was removed and the wells were stained with crystal violet. After 30 min at 37°C, the crystal violet was removed by aspiration. The number indicated in the table corresponds to the lowest amount of virus showing cell lysis after crystal violet staining d In this assay, PBMCs were put into 96-well plates at a concentration of 1 × 106 cells per well, and NDV (HUJ and Ulster strains) was added at different concentrations. After a 24-h incubation at 37°C, the supernatants were removed and freed from cells by centrifugation. ELISA for IFN-A was performed with an interferon (IFN)-A module set (Bender MedSystems, Vienna, Austria) as recommended by the manufacturer. The number indicated in the table corresponds to the amount of IFN produced in ng per HU of virus

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1. Selective replication in tumor and transformed cells: Through all the studies performed, NDV showed selective replication in human and mouse tumor cells and transformed cells. NDV, like other viruses, has developed immune escape mechanisms in its natural host, the birds. The capability to evade the host’s immune system is exerted via the V protein, which interferes with STAT-mediated interferon signals (21). This immune evasion mechanism of NDV, however, is species restricted and does not apply to mammalian cells. The feature of selective replication in tumor cells is directly related to the capacity of the cells which are in contact with NDV to mount an interferon (IFN) response. Nontumorigenic cells exert a capacity to thwart NDV infection by establishing an early antiviral state via a strong type I IFN response and a functional IFN-signaling cascade that prevent viral genome ampliﬁcation. In contrast, tumor cells have a weaker type I IFN response. The impairment of crucial antiviral pathways occurring after tumorigenesis appears to be rather common. That makes tumor cells highly susceptible to NDV infection and to oncolytic effects. We reported recently an inverse correlation between the expression of four antiviral genes (RIG-I, IRF-3, IRF-7, and IFN-B) and the susceptibility of cells (normal cells and tumor cells) to infection with NDV (22). We observed that the step leading to the generation of a full-length viral antigenome which is the template for production of progeny genomes is prevented in nontumorigenic cells (9). 2. Oncolytic activities: Some NDV strains (as MTH-68/H or Italien) have strong oncolytic capacity. They show oncolytic activities in vitro and in immunoincompetent animals. The attenuated, puriﬁed MTH68/H strain was reported to have beneﬁcial effects in patients with advanced cancer (23, 24). Since this strain exerts direct cytotoxicity in vitro against several tumor cell lines, it was concluded that direct cytotoxicity and oncolysis are key factors of antitumor activity of this virus strain (13). Oncolytic NDV strains are already being applied to cancer patients. Phase I/II studies have shown a good tolerance of cancer patients to systemic application even of oncolytic NDV, such as PV701 (5). There is an assumption that the oncolytic activities of NDV account for its antineoplastic activity. However, this is not corroborated by our data from in vivo tumor models (12). We observed that a virus-resistant tumor cell line could be induced to tumor regression by application of the oncolytic NDV strain MTH-68/H. This argues against oncolysis as the main mechanism of in vivo activity of this virus. Moreover, we have shown that lysates of NDV-Ulster-infected cells are processed by dendritic cells which then can potently stimulate autologous T cells from cancer patients,

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much better than lysate from noninfected cells (25). In contrast to these host-mediated immune mechanisms, direct oncolytic effects after systemic virus application would require successful targeting of the virus to the tumor. Our previous studies in animals, however, revealed that only a small fraction of the injected NDV is found at the tumor site after systemic application (26). In addition, we observed stronger antitumor effects when the virus was applied locally than when given systemically (27, 28). These observations suggest that oncolytic events may be important but, without support from the host’s immune system, these effects may not translate into improvement of overall survival. 3. Immunostimulatory activities: NDV has strong immune-activating effects. Before IFN-A was cloned and available in recombinant form, natural IFN-A was obtained from NDV stimulated by NDV. Lytic strains of NDV have been used for obtaining viral oncolysates upon tumor cell infection. Such products have been used as adjunctive immunotherapeutic agents in the postsurgical management of high-risk melanoma patients. Clinical studies performed in prospective settings suggest that this treatment is associated with prolonged survival (29). Nonlytic NDV strains also have immunostimulatory properties. By infecting patient-derived tumor cells with the nonlytic Ulster strain of NDV, we developed the autologous tumor vaccine modiﬁed by NDV infection (called ATV-NDV). Over the last 10 years, our group at the German Cancer Research Center (DKFZ, Heidelberg, Germany) was involved in studying the interactions of NDV with tumor cells and with cells of the immune system and led to the design and clinical evaluation of the ATV-NDV vaccine. It was found to be efﬁcient and a safe cancer vaccine with pleiotropic immune stimulatory properties (14, 30). We analyzed extensively the augmented immunostimulatory properties of tumor cells upon their infection with NDV (7, 14). This modiﬁcation of tumor cells with NDV was shown to introduce into the tumor cells’ danger signals (31) by way of doublestranded RNA (32, 33) and T-cell costimulatory activity (34) via the viral cell surface-expressed HN protein (35). NDV-modiﬁed tumor cells can upregulate MHC and cell adhesion molecules important for cognitive interactions with tumor antigen-speciﬁc T cells. Type I IFNs and chemokines (36), such as RANTES and IFN-G-inducible protein 10 (IP-10), are produced by NDV-infected tumor cells and activate the host’s immune system (30, 33, 37). Antitumor immune bystander effects have been analyzed extensively by us in many TNA experiments (15, 38). The ATV-NDV vaccine showed in cancer patients to be capable to activate preexisting antitumor memory T cells as observed by delayed-type hypersensitivity (DTH) responses (25). Signiﬁcant improvement of patient survival was reported in clinical trials of antitumor vaccination

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with ATV-NDV vaccine for patients with breast carcinoma (39). This was true also even for very aggressive tumors refractory to standard treatments, such as glioblastoma and head and neck squamous cell carcinoma (40, 41). Recently, results of a randomized clinical study in colorectal cancer patients have shown the effectiveness of the ATV-NDV vaccine in terms of long-term (10-year) survival (42). This result conﬁrms previous clinical data in a dozen of clinical studies suggesting that postoperative vaccination of cancer patients with this vaccine had beneﬁcial effects (43). The ATV-NDV vaccine acts at the interface between innate and adaptive immunity. When combined with bispeciﬁc antibodies introducing CD3 and CD28 ligands at the surface of the tumor vaccine, such vaccine showed further augmented antitumor activity in the TNA (15). It has also been shown capable to reactivate efﬁciently patient-derived memory T cells in an autologous setting as observed by IFN-G ELISPOT responses (44). In conclusion, NDV appears as a promising antitumoral therapeutic agent. In vitro and in vivo data obtained so far (some following the technical protocols presented in this chapter) encourage the use of NDV for cancer treatment. However, much more research has to be performed in order to decipher more precisely at the cellular and molecular levels the complex antitumor mechanisms induced by NDV.

Acknowledgments The authors are grateful to many colleagues at the German Cancer Research Center (DKFZ, Heidelberg, Germany), at the Department of Medical Biology, Medical School, University of Pécs, Pécs, Hungary, at the United Cancer Research Institute, Alexandria, Virginia, and at Cell Engineering Research Centre and Department of Cell Biology, Fourth Military Medical University, Xi’an, China, but also all the clinicians worldwide who have been implicated in clinical trials testing NDV as therapeutic agent. The authors would like to thank Drs. L.K. Csatary, Zs. Fábián, and M. Pap for their contributions in various phases of the research work on oncolytic MTH-68/H supported by the United Cancer Research Institute, Alexandria, Virginia, USA. The aim of all these studies at the DKFZ was to acquire the basic scientiﬁc knowledge necessary for the development of cancer immunovirotherapies using NDV. The authors would like to thank previous colleagues and especially Claudia Haas and Leonidas Apostolidis for their contribution, respectively, in the development of the in vitro tumor-neutralization assays based on nonlytic and lytic NDV strains. We would also like to thank Annette Arnold for her excellent technical help throughout the project.

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This work was supported by many grants over the last 10 years. We thank the IOZK Köln (www.iozk.de) for support and for clinical application of NOV to cancer patients. At the time when this manuscript is written, P.F. is supported by the Christian BerndtStiftung, Rostock, Germany. References 1. Cassel WA, Garrett RE. Newcastle disease virus as an antineoplastic agent. Cancer 1965; 18:863–868. 2. Csatary LK. Viruses in the treatment of cancer. Lancet 1971; ii:825. 3. Alexander DJ, Allan WH. Newcastle disease virus pathotypes. Avian Path. 1974; 3(4): 269–278. 4. Lorence RM, Roberts MS, Groene WS, Rabin H. Replication-competent, oncolytic Newcastle disease virus for cancer therapy. In: Hern aiz Driever P, Rabkin SD, eds. Replicationcompetent viruses for cancer therapy. Monographs in virology, Vol. 22. Basel, Switzerland: Karger, 2001:160–82. 5. Pecora AL, Rizvi N, Cohen GI, Meropol NJ, Sterman D, Marshall JL, Goldberg S, Gross P, O’Neil JD, Groene WS, Roberts MS, Rabin H, Bamat MK, Lorence RM. Phase I trial of intravenous administration of PV701, an oncolytic virus, in patients with advanced solid cancers. J. Clin. Oncol. 2002; 20:2251–66. 6. Cassel WA, Murray DR, Olkowski ZL. Newcastle disease virus oncolysate in the management of stage III malignant melanoma. In: Sinkovics JG, Horvath JC, eds. Viral therapy of human cancer. New York: Marcel Dekker, 2005; 577–689. 7. Schirrmacher V, Fournier P. Newcastle Disease Virus : a promising vector for viral therapy of cancer. In: Harrington KJ, Pandha HS and Vile RG, eds. Viral therapy of cancer, vol 542. Wiley Publishers, 2008:171–186. 8. Reichard KW, Lorence RM, Cascino CJ, Peeples ME, Walter RJ, Fernando MB, Reyes HM, Greager JA. Newcastle disease virus selectively kills human tumor cells. J. Surg. Res. 1992; 52(5): 448–453. 9. Fiola C, Peeters B, Fournier P, Arnold A, Bucur M, Schirrmacher V. Tumor-selective replication of Newcastle Disease Virus: association with defects of tumor cells defence. Int. J. Cancer 2006; 119(2): 328–38. 10. Li YL, Wu J, Wei D, Zhang DW, Feng H, Chen ZN, Bian H. Newcastle disease virus represses the activation of human hepatic stellate cells and reverses the development of hepatic ﬁbrosis in mice. Liver Int. 2009; 29(4):593–602.

11. Sinkovics JG, Horvath JC. Newcastle disease virus (NDV): brief history of its oncolytic strains. J. Clin. Virol. 2000; 16:1–15. 12. Apostolidis L, Schirrmacher V, Fournier P. Host mediated anti-tumor effect of oncolytic Newcastle disease virus after locoregional application. Int. J. Oncol. 2007; 31(5):1009–19. 13. Fábián Z, Csatary CM, Szeberényi J, Csatary LK. p53-independent endoplasmic reticulum stress-mediated cytotoxicity of a Newcastle disease virus strain in tumor cell lines. J. Virol. 2007; 81(6):2817–30. 14. Schirrmacher V, Haas C, Bonifer R, Ahlert T, Gerhards R, Ertel C. Human tumor cell modiﬁcation by virus infection: an efﬁcient and safe way to produce cancer vaccine with pleiotropic immune stimulatory properties when using Newcastle Disease Virus. Gene Ther. 1999; 6: 63–73. 15. Czeglédi A, Wehmann E, Lomniczi B. On the origins and relationships of Newcastle disease virus vaccine strains Hertfordshire and Mukteswar, and virulent strain Herts’33. Avian Pathol. 2003; 32, 271–276. 16. Haas C, Lulei M, Fournier P, Arnold A, Schirrmacher V. T-cell triggering by CD3- and CD28-binding molecules linked to a human virus-modiﬁed tumor cell vaccine. Vaccine. 2005; 23(19):2439–53. 17. Bian H, Fournier P, Moormann R, Peeters B, Schirrmacher V. Selective gene transfer in vitro to tumor cells via recombinant Newcastle disease virus. Cancer Gene Ther. 2005; 12:295–303. 18. Van Meir EG, Polverini PJ, Chazin VR, Su Huang HJ, de Tribolet N, Cavenee WK. Release of an inhibitor of angiogenesis upon induction of wild type p53 expression in glioblastoma cells. Nat. Genet. 1994; 8(2):171–6. 19. Van Meir EG, Kikuchi T, Tada M, Li H, Diserens AC, Wojcik BE, Huang HJ, Friedmann T, de Tribolet N, Cavenee WK. Analysis of the p53 gene and its expression in human glioblastoma cells. Cancer Res. 1994; 54(3):649–52. 20. Xu C, Bailly-Maitre B, Reed JC. Endoplasmic reticulum stress: cell life and death decisions. J. Clin. Invest. 2005; 115: 2656–2664.

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21. Horvath CM. Weapons of STAT destruction. Interferon evasion by paramyxovirus V protein. Eur. J. Biochem. 2004; 271:4621–8. 22. Wilden H, Fournier P, Zawatzky R, Schirrmacher V. Expression of RIG-I, IRF3, IFN-beta and IRF7 determines resistance or susceptibility of cells to infection by Newcastle Disease Virus. Int. J. Oncol. 2009; 34(4): 971–82. 23. Csatary LK, Gosztonyi G, Szeberenyi J, Fabian Z, Liszka V, Bodey B, Csatary CM. MTH-68/H oncolytic viral treatment in human high-grade gliomas. J. Neurooncol. 2004; 67: 83–93. 24. Csatary LK, Moss RW, Beuth J, Torocsik B, Szeberenyi J, Bakacs T. Beneﬁcial treatment of patients with advanced cancer using a Newcastle disease virus vaccine (MTH-68/H). Anticancer Res. 1999; 19:635–638. 25. Bai L, Koopmann J, Fiola C, Fournier P and Schirrmacher V. Dendritic cells pulsed with viral oncolysates potently stimulate autologous T cells from cancer patients. Int. J. Oncol. 2002; 21:685–694. 26. Bian H, Wilden H, Fournier P, Peeters B and Schirrmacher V. In vivo efﬁcacy of systemic tumor targeting of a viral RNA vector with oncolytic properties using a bispeciﬁc adapter protein. Int. J. Oncol. 29: 1359–1369, 2006. 27. Phuangsab A, Lorence RM, Reichard KW, Peeples ME, Walter RJ. Newcastle disease virus therapy of human tumor xenografts: antitumor effects of local or systemic administration. Cancer Lett. 2001; 172:27–36. 28. Schirrmacher V, Griesbach A and Ahlert T. Antitumor effects of Newcastle disease virus in vivo: Local versus systemic effects. Int. J. Oncol. 2001; 18:945–952. 29. Batliwalla FM, Bateman BA, Serrano D, Murray D, Macphail S, Maino VC, Ansel JC, Gregersen PK, Armstrong CA. A 15-year follow-up of AJCC stage III malignant melanoma patients treated postsurgically with Newcastle disease virus (NDV) oncolysate and determination of alterations in the CD8 T cell repertoire. Mol. Med. 1998; 4(12):783–94. 30. Washburn B, Schirrmacher V. Human tumor cell infection by Newcastle disease virus leads to upregulation of HLA and cell adhesion molecules and to induction of interferons, chemokines and ﬁnally apoptosis. Int. J. Oncol. 2002; 21:85–93. 31. Matzinger P. The danger model: a renewed sense of self. Science 2002; 296:301–5. 32. Alexopoulou L, Holt AC, Medzhitov R, Flavell RA. Recognition of double-stranded RNA and activation of NK-k B by Toll-like receptor 3. Nature 2001; 413:732–8. 33. Fournier P, Zeng J, Schirrmacher V. Two ways to induce innate immune responses in human

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Chapter 14 Next-Generation Oncolytic Vaccinia Vectors Steve H. Thorne Abstract Oncolytic vaccinia viruses have made some impressive advances over the last 5 years, with a range of different backbones displaying signiﬁcant antitumor responses in preclinical models, and some exciting clinical results being reported against liver cancers. Because the virus is capable of rapid spread within the tumor, has evolved to spread relatively undetected within the blood stream, does not integrate into the host cell chromosome, and can infect almost any cell type, it is well-suited to the requirements for a successful oncolytic. In addition, the extensive clinical use of this virus means that contraindications to its use are known, and approved and experimental antivirals are available. Furthermore, because the virus has a large array of virulence genes whose deletion may target different properties of the cancer cell, and a large cloning capacity allowing for insertion of multiple transgenes, the possibilities for further development of novel and next-generation oncolytic vectors are multitude. Key words: Vaccinia, Oncolytic virus, Virotherapy, Tumor targeting, Replication selective

1. Introduction Many of the adaptations that a virus induces within a cell subsequent to infection are similar to the hallmarks of a malignant cell, including loss of cell cycle control, evasion of apoptotic cell death, and a breakdown in ability to produce or respond to immunomodulatory factors. It is, therefore, not surprising that the root causes of some cancers are known to be chronic viral infections. However, this also means that many viral virulence genes or functional domains are redundant within cancer cells that are already adapted for viral replication and so their deletion results in viral strains that are attenuated in normal tissues, but not tumors. It is this strategy that is most effective for creating tumor-targeting strains of vaccinia, as the virus does not interact extensively with the host cell translational machinery and does not have deﬁned cellular uptake

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receptors, so the use of tumor- or tissue-speciﬁc promoters driving an essential viral gene or alterations of viral tropism are unlikely to be effective targeting strategies. However, vaccinia is a large virus with an extensive array of genes involved in many aspects of viral virulence (1, 2), including limiting the immune response targeting the virus, preventing apoptosis of infected cells, or activating cell-signaling pathways. Because many of these viral genes are likely to be redundant in certain cancer cells, their deletion can produce a tumor-speciﬁc or tumor-selective vaccinia strain. As such, deletions in several of these genes have been shown to display tumor-targeting potential (these include viral thymidine kinase (TK), vaccinia growth factor (VGF), vaccinia type I interferonbinding protein (B18R), dsRNA-binding protein (E3L), and viral serpins (B13R, B22R)) (3, 4). However, the tumor-targeting potential of deletions in many more remain unexplored, and so 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. Further reﬁnements in the tumor speciﬁcity and increases in tumor cell destruction can then be achieved through expression of combinations of transgenes from the viruses. Here, we focus on the steps involved in design and testing of a novel oncolytic vector based on vaccinia virus and cover areas of preclinical development.

2. Materials 2.1. Construction of Oncolytic Vaccinia Strains

1. Cloning plasmids, such as pSC-65 (5). 2. Cell lines CV-1, 143B TK-, and HeLa (ATCC). 3. 10-cm tissue culture plate. 4. Lipid vesicle transfection reagent (such as lipofectamine (Invitrogen), with specialized media if necessary (e.g., OptiMEM, Gibco)). 5. Cell scrapers. 6. Dry ice: Methanol bath (add dry ice to an insulated bucket, and then carefully add methanol until a slurry is formed). 7. Water bath 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.

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1. Six-well plates. 2. Overlay medium for plaque assay (per 100 ml): 50 ml 2× DMEM, 2 ml FBS, 48 ml 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. Ninety-six-well plates. 2. MTS assay reagent (Promega). 3. ELISA plate reader.

2.4. Bioluminescence Imaging

1. D-Luciferin, 30 mg/ml in PBS, ﬁlter sterilized (Xenogen, part of Caliper LIfe Sciences). 2. Mice, typically obtained from commercial vendors (e.g., Jackson laboratories or Charles River). 3. IVIS imaging system or equivalent whole animal imaging system (Caliper Life Sciences) with heated (37°C) stage and appropriate image analysis software (e.g., Living Image, Xenogen, part of Caliper Life Sciences.). 4. 70% ethanol. 5. Anesthetic (ideally inhaled, such as isoﬂurane, but may be injected, such as avertin). 6. Black-walled, clear-bottom, 96-well plates. 7. Luciferase assay system (Promega).

3. Methods The initial design of an oncolytic vaccinia construct depends on the cancer phenotype that is to be targeted. Consideration should be given to both the parental vaccinia (or related poxvirus) strain (see Note 1) as well as the choice of gene deletions when designing a backbone vector. The location and promoter of any foreign transgene is also signiﬁcant, as this affects transgene expression levels (6). 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 gene. A variety of methods are possible to disrupt or delete nonessential viral genes, and vaccinia strains containing deletions in several of these virulence genes have been previously constructed by a variety of investigators. However, as methods, especially for recombinant selection, are nonstandard, they are not extensively covered here.

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Instead, we describe a protocol for the insertion of a foreign transgene into the viral TK gene by homologous recombination as an example (see Note 2). 3.1. Construction of Oncolytic Vaccinia Virus

1. Standard cloning strategies can be used to insert the transgene of choice into the multiple cloning site of a cloning plasmid (see Note 3). 2. Grow a CV-1 cell monolayer until 80% conﬂuent in a 10-cm tissue culture dish (achieved by splitting a 90% conﬂuent plate 1:2 overnight). 3. Infect with target virus (e.g., vaccinia virus containing all required deletions, except for the thymidine kinase deletion) at a multiplicity of infection (MOI) of 0.05 plaque forming units (PFUs)/cell. Assume the number of cells at time of plating to be present in the dish. 4. After a 2-h incubation at 37°C, transfect the cell layer with the cloning plasmid using commercial transfection reagent (see Note 4). 5. Leave CV-1 cells at 37°C, 5% CO2 for 48 h, and then scrape cells into media. 6. Freeze/thaw cells and media three times (by transferring between a dry ice/methanol bath and a 37°C water bath). Spin (300 × g, 5 min) to remove large cellular debris. The crude viral lysate can then be stored at −80°C until needed. 7. 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 volume of fresh 5 mg/ml BrdU (see Note 5). 8. After a 48-h incubation, again collect cells and media and freeze/thaw three times. 9. Add 10 Ml/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 Ml media/well overnight). Dilute virus 1:4 every row down the plate (i.e., take 50 Ml from each well in top row and add to corresponding well in row directly below; repeat down plate. Keep bottom row uninfected). Add 1:100 volume of 5 mg/ml BrdU to all wells. 10. Incubate 96-well plates for 48 h (37°C; 5% CO2), and then examine wells individually under inverted microscope. Mark wells with a single visible plaque (if no wells contain only one plaque, select wells with smallest numbers of plaques). Ideally, when a secondary marker, such as a reporter gene, is available for selection of successful recombination (see Note 5), this can then be further applied to the wells containing a single plaque to conﬁrm recombination (to differentiate from spontaneous

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TK mutation). Alternatively, the secondary selection marker may be used directly instead of BrDU; this avoids possible complications due to the mutagenic nature of this chemical. Collect media and cells from up to 10 of the single plaque wells, and repeat steps 11 and 12 for two (or more) further rounds of plaque puriﬁcation. 11. It is necessary to conﬁrm correct insertion of transgene by PCR and sequencing of DNA collected from infected cells. Primers should be designed to ﬂank transgene insertion site. 12. Expand ﬁnal puriﬁed plaque by addition of crude viral lysate to HeLa cells. Observe cells under inverted microscope every 24 h for signs of cytopathic effect (i.e., cell rounding) and when this is evident, aspirate media, add 10 mM Tris–HCl, pH 9.0, and scrape cells into this buffer. Freeze/thaw three times as before; spin crude viral lysate (300 × g, 5 min) to pellet cellular debris and retain supernatant. Expansion can be performed in steps, with virus from a single plaque expanded in a well of a six-well plate, then a T-25 ﬂask, a T-150 ﬂask, and then multiple T-150 ﬂasks (or roller bottle or cell factory apparatus) in order to produce enough virus for preclinical testing. Virus can be further puriﬁed if desired (especially if it is to be used in vivo) and should be titered by plaque assay (see below). 3.2. Screening Selectivity of Oncolytic Virus In Vitro

3.2.1. One-Step Viral Growth Curve Assayed by Plaque Assay

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. Nontumor-bearing animals may be used to ascertain safety of the constructs. 1. A panel of tumor and “normal” (primary or immortalized, nontransformed) cell lines should be grown to 80% conﬂuence in six-well plates (see Note 6). Preliminary experiments need to be performed to ascertain the correct split ratios to obtain cells at the correct conﬂuence. Cells should be chosen to represent the tumor type and phenotypic properties that are to be targeted. 2. Aspirate media and add 1 ml of appropriate media/well with 2% FBS (heat inactivated). Then, add virus at an MOI of 1.0 PFU/cell. 3. Wait for 2 h, then aspirate media again, and add 3 ml/well of fresh media (with 2% FBS). 4. At 12-h intervals (up to 96 h), scrape cells from one well of each six-well plate into media and collect cells and media. Freeze/thaw three times as before, and 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).

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5. Plate BS-C-1 cells in six-well plates (2 × 106 cells/well) overnight. Aspirate media and add 900 Ml of media with 2% FBS per well. Sonicate dilutions of crude viral lysate for 30 s at full power. Add 100 Ml of dilutions 2–6 of the crude viral lysate to ﬁve wells of the six-well plate (leave last well uninfected as 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 overlay and adding 1 ml/well of crystal violet solution. 8. Leave at room temperature for 2 h before aspirating crystal violet and inverting plates to dry. 9. Count plaques in all wells that display between 10 and 150 plaques, and based on the dilution used calculate initial concentration of virus (PFU/ml). Ratio of normal to tumor viral replication gives a value for the selectivity of the virus (Fig. 1a). The oncolytic construct should be compared to wild-type virus. 3.2.2. Determination of Viral-Mediated Cytopathic Effect

1. Seed 96-well plates with 200 Ml of media with tumor or “normal” cell lines (see Note 7) and incubate overnight to allow cells to attach (preliminary experiments are needed to determine correct seeding density to give cells 80–90% conﬂuence).

Fig. 1. (a) Comparison of replication of a candidate oncolytic viral strain in a tumor and normal cell line. Viral titers determined at times up to 72 h after infection. (b) Relative cytopathic effect of a candidate oncolytic viral stain in a tumor and normal cell line (read by viability assay 72 h post infection). EC50 values (the viral titer (PFU/cell) required to reduce cell viability by 50%) are compared between cell lines. *Therapeutic index of the candidate strain.

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Do not seed the outer wells on the plate; instead, ﬁll these with 200 Ml 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 using 1:4 dilutions (i.e., remove 50 Ml/well and add to corresponding well in next column). Leave ﬁnal column of cells uninfected. Keep outer wells with PBS only. Perform each dilution series in triplicate. 3. Leave plates at 37°C for 48, 72, or 96 h, then aspirate media, add fresh media with no serum to each well, and add MTS (Promega) according to manufacturer’s guidelines. 4. Read the MTS color change after 2 h using an ELISA plate reader. 5. Plot cell survival for the dilutions of virus used by deﬁning the uninfected cells as 100% viable and PBS control wells as 0% viable. A value for percentage viability can, therefore, be determined for all treatment wells. The PFU/cell needed to reduce viability by 50% can, therefore, be determined from the plot produced. Ratios of these EC50 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). 3.3. In Vivo Screening Using Bioluminescence Imaging

Noninvasive whole animal bioluminescence imaging provides several advantages over traditional animal models. The ability to repeatedly image the same animals means that they become their own internal controls, so increasing sensitivity of the assay and reducing the number of animals required to run experiments. In addition, data produced during whole animal imaging can be used to deﬁne time points and tissues selected for ex vivo analysis (such as immunohistochemistry, qPCR of viral genomes, etc.), so more accurately deﬁning the biodistribution of infection, timings of therapeutic responses, and any unexpected infection of nontumor tissues. 1. Bioluminescence imaging requires viral strains constructed to express luciferase as a reporter gene (see Note 7). These should be ﬁrst checked in vitro to ascertain the levels of bioluminescence produced. Alternatively, experiments may be run with luciferase-labeled tumor cell lines (and unlabeled virus) in order to follow the therapeutic effects of the virus, especially if orthotopic or metastatic tumor models are to be used. 2. Consideration should also be given to the animal model to be used. Because of the depth limitations of optical imaging, the ideal species to use is the mouse. Choice of mouse strain may be limited by the availability of cell lines, but in general white mice are preferable (as the pigmentation in the skin of black or

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brown mice further adsorbs emitted light). Some normally dark mice (e.g., C57B/6) are commercially available as albinos. Animal fur also diffracts light, and so shaving or use of depilatory cream may enhance weak signal. Alternatively, athymic nu-/nu- or similar hairless mice may be used. Pilot experiments should always be run in vivo to ﬁrst characterize the model (see Note 8). 3. Deliver luciferin substrate to mice. Imaging of insect luciferase typically requires injection of 200 Ml of 30 mg/ml luciferin intraperitoneally 5–20 min prior to imaging; coelentrazine for Renilla luciferase is typically injected intravenously (tail vein or retino-orbital injection) with 100 Ml of 5 mg/ml injected no longer than 2 min prior to imaging. 4. Anesthetize mice. The exact order of delivery of substrate and anesthesia depends on the system being used. The choice of anesthetic may also be important. As mice take longer to recover from injectable anesthetics, fewer imaging time points can be used. Inhaled anesthetics (such as isoﬂurane, 2%) are, therefore, typically preferable; however, there is some evidence that these may produce a low level of background luminescence and so may not be suitable for detection of very weak signal, especially from the lungs. 5. Arrange 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 for imaging spine, left side for imaging spleen). Imaging stage should be wiped with 70% ethanol prior to use, and a piece of black paper placed on the stage to protect it. Dividers may be used between the animals (if multiple animals are to be imaged at once) to prevent strong signal from one animal reﬂecting off its neighbor. All nonessential materials should be removed from the imaging chamber. The stage should be heated to 37°C, and animals should be checked for level of anesthesia for 30 s prior to closing the chamber (see Note 9). 6. Take image. Several variables may be adjusted to improve the quality of the image, including height of stage; length of exposure; and binning and aperture setting (see Note 10). In order 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 3D reconstructions, whereby animals are imaged consecutively with ﬁlter sets to only allow light from deﬁned wavelengths to pass, and the relative levels of blue and red light emitted can be used to determine the depth of a signal with known spectra (as blue light is absorbed more extensively than red light).

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7. Recover mice. Once imaging is complete, mice should be recovered on a heated stage, until capable of self-righting. 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 to 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. Bioluminescence signal from ﬁreﬂy luciferase remains strong for up to 45 min postmortem, and so individual organs can be reimaged ex vivo to verify the tissue of origin of any signal (although the signal no longer remains quantiﬁable). In addition, it should be remembered that bioluminescent signal represents 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 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, including strains used during the smallpox eradication program (e.g., Wyeth/New York City Board of Health, Lister, Copenhagen, Tian Tan, USSR) that have the advantage of extensive previous use in human subjects (by intradermal delivery); other strains have been developed as laboratory strains (e.g., Western Reserve, IHD-J), and the majority of our knowledge on the effects of gene deletions has been obtained from these. Many virulence genes are only expressed or functional in a subset of the strains available, and so 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 most common method for inserting a foreign transgene into vaccinia involves homologous recombination into the viral thymidine kinase gene. 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, so making this gene deletion tumor targeting.

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3. A number of different cloning plasmids are available 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 that need to be incorporated. However, many vaccinia promoters have been well-described, and 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. There is also a wide range of promoter strengths available. The pSC65 plasmid, for example, contains both the natural viral p7.5 promoter and the synthetic pSE/L promoter. 4. A variety of transfection reagents and protocols are available. CaCl2 transfection works 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 transfection according to manufacturer’s guidelines. 5. 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, GFP, lacZ) allows selection of viral plaques that express these genes (but again, this is not 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). 6. Tumor cells used in these assays should be selected based on the desired phenotypic or tissue target for the virus; “normal” control 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 to hepatocellular cancers). Cells may be grown under specialized conditions to further examine methods of selectivity (e.g., pretreated with IFN; serum starved; etc.). 7. The choice of luciferase enzyme incorporated for viral labeling is important. In order 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, as light below this wavelength is adsorbed. The most

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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 (7). Other insect luciferases, such as those of the click beetle or railroad worm, produce a variety of spectra. Several luciferases have also been cloned from marine organisms, such as Renilla and Aequorea, but these produce blue–green light (peak of 475 nm). However, the advantage of Renilla luciferase is that it utilizes a different substrate (coelentrazine) to the insect luciferases (luciferin) and so dual bioluminescence imaging is possible. However, the coelentrazine substrate is unstable meaning that a much higher background signal is produced. Both substrates are nontoxic, are rapidly distributed throughout the body, and can cross most membranes, including cellular membranes and the blood–brain barrier. 8. Protocols for all planned experiments should ﬁrst be approved by institutional review boards. 9. Many plastics and other materials (including animal bedding, chow, and dander) produce phosphorescence, and so careful selection of any item used in the imaging chamber is important. In particular, anesthesia nose cones, paper used to protect the stage, plastic dividers, and animal shields should always be carefully tested, and all bedding, dander, and chow should be cleaned from the stage before imaging. Use of a specialized alfalfa-free imaging diet reduces background from the GI tract. 10. 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 so shorter exposure times should be tried and smaller binning used. If the original image produces weak or no signal, then increased exposure times (up to 5 min) and increased binning may be tried. References 1. Smith GL, Symons JA, Khanna A, Vanderplasschen A, Alcami A. 1997 Vaccinia virus immune evasion. Immunol Rev;159:137–54. 2. Buller RM, Palumbo GJ. 1991 Poxvirus pathogenesis. Microbiol Rev;55:80–122. 3. Thorne SH, Hwang TH, Kirn DH. 2005 Vaccinia virus and oncolytic virotherapy of cancer. Curr Opin Mol Ther;7:359–65. 4. Thorne SH, Kirn DH. 2004 Future directions for the ﬁeld of oncolytic virotherapy: a perspective on the use of vaccinia virus. Expert Opin Biol Ther;4:1307–21.

5. Chakrabarti S, Sisler JR, Moss B. 1997 Compact, synthetic, vaccinia virus early/late promoter for protein expression. Biotechniques;23:1094–7. 6. Coupar BE, Oke PG, Andrew ME. 2000 Insertion sites for recombinant vaccinia virus construction: effects on expression of a foreign protein. J Gen Virol;81:431–9. 7. Zhao H, Doyle TC, Coquoz O, Kalish F, Rice BW, Contag CH. 2005 Emission spectra of bioluminescent reporters and interaction with mammalian tissue determine the sensitivity of detection in vivo. J Biomed Opt;10:41210.

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Chapter 15 Evaluation of Innate Immune Signaling Pathways in Transformed Cells Joshua F. Heiber and Glen N. Barber Abstract Oncolytic viruses, the use of viruses to treat cancer, is emerging as a new option for cancer therapy. Oncolytic viruses, of both DNA and RNA origin, exhibit the ability to preferentially replicate in and kill cancer cells plausibly due to defects in innate immune signaling or translation regulation that are acquired during cellular transformation. Here, we review concepts and assays that describe how to analyze signaling pathways that govern the regulation of Type I IFN production as well as the induction of interferonstimulated antiviral genes, events that are critical for mounting an effective antiviral response. The following procedures can be used to assess whether innate immune pathways that control antiviral host defense are defective in tumor cells – mechanisms that may help to explain viral oncolysis. Key words: Innate immunity, Interferon, Oncolytic, Host defense, STING

1. Introduction That natural virus infections occasionally coincided with the regression of certain types of tumors in patients suffering from such maladies is relatively well documented and has led to a spate of studies designed to utilize viruses as oncolytic agents for the treatment of malignant disease (1). However, the mechanisms that favorably enable the replication of viruses in tumor cells compared to normal cells still remain to be clariﬁed. Such reasons could involve the accumulation of genetic mutations in innate signaling pathways important for mediating antiviral activity. In cancer cells, these defects can lead to a reduction or inactivation of the type 1 interferon (IFN-D/E) response, critical for upregulating numerous antiviral genes, making such cells highly susceptible to viral replication and

David H. Kirn et al. (eds.), Oncolytic Viruses: Methods and Protocols, Methods in Molecular Biology, vol. 797, DOI 10.1007/978-1-61779-340-0_15, © Springer Science+Business Media, LLC 2012

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subsequent cellular lysis (2). In order to fully explore the mechanisms of oncolytic virus action, it is necessary to understand the architecture of innate immune signaling cascades in normal cells and then evaluate their plausible deregulation in cancer cells. Only in the last few years has it become clear that the host has evolved a number of pathways to detect and respond to foreign pathogens such as viruses and bacteria (3–5). However, it is likely that important components of innate signaling still remain to be identiﬁed, which will make the reasons underlining oncolysis potentially elusive for some time. Nevertheless, progress has been made toward understanding these processes, and now these signaling cascades including their components can start to be analyzed in cancer cells. The reasons underlining viral oncolysis are particularly intriguing since numerous virus types replicate in a wide variety of cancer cells, making the responsible defective pathway potentially a major reason behind tumorigenesis (6). These recently unraveled pathways may include the Toll-Like Receptor (TLR) pathway, the RIG-I Like Helicase (RLH) pathway, and recently the STING regulated pathway (SRP), a brief description of which follows. 1.1. Toll-Like Receptor Signaling

The triggering of innate immune responses follows the detection of pathogen speciﬁc molecules (pathogen-associated molecular patterns- PAMPS) by cellular pattern-recognition receptors (PRRs). For example, bacteria have cell wall components comprising lipopolysaccharides that can trigger innate immune responses, while viral nucleic acids are also known to be potent PAMPs (5, 7). A key innate immune pathway that is highly conserved and able to recognize a wide array of pathogens is the Toll pathway, ﬁrst identiﬁed in Drosophila melanogaster. Activation of D. melanogaster Toll leads to the fat body cells triggering, via dMyD88, tube, and pelle, the production of antimicrobial peptides (AMPs) and the subsequent secretion of the peptides into the hemolymph. Another pathway, referred to as IMD (for immunodeﬁciency) utilizes Drosophila RIP1 and dFADD to trigger the production of a different set of antimicrobial peptides (8–10). Both pathways culminate in the activation of NF-NB related transcription factors, Relish and Dorsalrelated Immune Factor (DIF), respectively, and the transcription of hundreds of genes including those encoding the effector AMPs directed against the intruding microorganisms. However, these pathways are geared toward the recognition of bacteria, and almost nothing is known of how ﬂies respond to virus infection (11). Nevertheless, these pathways have been preserved in mammalian cells and there are 13 TLRs currently known in human (12). Signiﬁcantly, four TLRs (TLR 3, 7/8, and 9) are considered associated with the detection of viral infection (5). For example, TLR-3 is found in macrophages, myeloid dendritic cells, ﬁbroblasts, and epithelial cells, is located in the endoplasmic reticulum, and senses extrinsic dsRNA species (13, 14). TLR-7/8 is similarly associated

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with the endoplasmic reticulum, though predominantly in plasmacytoid dendritic cells (pDCs) and is responsible for detecting single-stranded RNA species (15, 16). By contrast, TLR 9 recognizes unmethylated DNA species (CpG DNA) and is similarly found in pDCs (15, 17). These TLRs reside in endosomes and can trigger the production of IFN-D/E in either a MyD88 dependent (TLR7/9) or independent (TLR-3) manner (18). Upon recognition of extrinsic dsRNA captured in endosomes, TLR-3 activates the signal adaptor TIR domain-containing adapter inducing interferon-E (TRIF), which mediates association with TNF receptor-associated factor 6 (TRAF6) and 3 (19, 20). TRIF complexes then activate tank binding kinase 1 (TBK-1) leading to phosphorylation of interferon regulatory factor 3 (IRF-3) and IRF-7. IRF3 or 7 dimerize, facilitating translocation to the nucleus and activation of IFN-E transcription (19). TRIF also interacts with RIP1, which in association with TRAF6 activates the IKK complex, which signals through TGF-E activated kinase 1 (TAK-1, leading to dissociation of the IKK complex from NF-NB (21). IRF3/7 and NF-NB are both required for the robust transcriptional activation of type I IFN. In the case of TLR7 and 9, recognition of the appropriate PAMP leads to the recruitment of MyD88, initiating the phosphorylation of interleukin-1 receptor-associated kinase 4 (IRAK4) (22, 23). IRAK4 then activates both IRAK1 and TRAF-6 and 3, which triggers IRF7 leading to production of type I IFN (24). Experiments revealed using IRF-7−/− mice have emphasized that IRF-7 is a master regulator of type I IFN production. For example, ﬁbroblasts and DC’s from IRF-7−/− knockout mice have shown that such cells do not produce IFN-D/E in response to stimulation with poly(I:C) or virus infection underscoring the necessity of TBK-1 phosphorylation of IRF-7 in type I IFN signaling (25). Of course, since TLR7/8 and 9 are predominantly associated with the production of type I in certain types of dendritic cells, it is unlikely that these PRRs are deregulated in cancers not originating from these types of cells. However, the TLR3 pathway is utilized in a wider range of cell types and may indeed be suppressed in various tumor types. 1.2. The RLH Pathway

A second key pathway that is responsible for producing type I IFN in response to RNA virus infection, in all cells but pDCs (which are TLR7 dependent), is referred to as the RLH (RIG-like helicase) pathway (26). Activation of this cascade relies on two DExD/H helicases referred to as retinoic acid inducible gene-I (RIG-I) and melanoma differentiation antigen 5 (MDA-5). RIG-I recognizes 5c-triphosphates on viral RNAs, such as those encoded by negativestranded viruses such as vesicular stomatitis virus (VSV) and Newcastle Disease Virus (NDV). By contrast, MDA-5 recognizes longer RNAs (>1 kb) such as those encoded by positive-stranded viruses such as the picornavirus encephalomyocarditis virus

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(EMCV) (27, 28). Binding of viral RNA leads to a conformational change in the helicases which exposes N-terminal Caspase Activation and Recruitment Domains (CARD), which facilitate a CARDCARD interaction with a second CARD carrying mitochondrially associated protein referred to as Interferon-E Promoter Stimulator 1 (IPS-1/VISA/MAVS/CARDIF) (29, 30). IPS-1 is then able to mediate the phosphorylation of TRAF 3 and trigger phosphorylation of TBK1 and IKKi, leading to the phosphorylation of IRF-3 and 7 (20, 31). Concomitantly, IPS-1 also activates the NF-NB pathway through Fas Associated Death Domain (FADD) and Receptor Interacting Protein 1 (RIP-1) (32, 33). This is important, since as mentioned earlier, IRF-3, NF-NB, and AP-1 are required for maximal production of IFN-E. Even though the RLH and TLR pathways are activated by unique PAMPs, it is important to remember that they both signal through TBK-1 and IRF-3/7, as well as the NF-NB pathway. Thus, the RLH pathway is largely responsible for recognizing RNA viruses and triggering the production of type I IFN. Despite this progress, little was known relating to the recognition of DNA pathogens, such as parasites and DNA viruses, by the host innate immune system. A number of DNA viruses are known to remain latent in the cell for many years and viruses such as human papilloma virus (HPV) and Epstein-Barr Virus (EBV) are known to contribute toward tumorigenesis (34). However, a molecule referred to as STING (Stimulator of interferon genes) has recently been shown to be responsible for regulating the production of type I IFN by intracellular DNA, as described next. 1.3. The STING Regulated Pathway

The Interferon stimulatory DNA (ISD) pathway is the least well characterized of the signaling pathways that lead to the production of type I IFN. ISD signaling plays a critical role in responding to DNA viral infections and may be defective in many types of cancer (35). Current data have indicated the existence of cytosolic DNA sensors such as a molecule referred to as DAI, which may indeed play a role in the production of type I IFN (36). However, a key intracellular DNA sensor(s) responsible for triggering the production of type I IFN has not yet been discovered. Nevertheless, a recently identiﬁed essential component for DNA-mediated IFN production has been found and is referred to as STING (Stimulator of Interferon Genes). STING is an endoplasmic reticulumassociated protein that is indispensible for activation of IFN-E in response to a variety of DNA pathogens such as herpes simplex virus (HSV) and some RNA viruses such as VSV (implicating a role for STING in regulating RIG-I activity). Upon activation STING associates with TBK-1, which can activate IRF-3, IRF-7, and NF-NB responsible for driving expression of Type I IFN (37, 38). Given the importance of STING in governing this pathway, it is possible that defects in the STING regulated pathway (SRP) could

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explain the ability of DNA viruses such as vaccinia virus, HSV, and adenovirus to mediate viral oncolysis. Further research will undoubtedly reveal other components of this pathway that could be deregulated in a wide variety of tumors. Needless to say, defects in these pathways may lead to the suppression of type I IFN production and other key antiviral genes activated through this primary innate immune signaling cascade. This may enable viruses to replicate in an uncontrolled way leading to cellular lysis. Presumably, viruses that infect normal cells trigger these potent host defense mechanisms and suffer deleterious consequences to their replication. However, it is plausible that the primary innate immune pathway remains intact in many cancer types and it may be that the JAK/STAT pathway responsible for triggering the production of antiviral genes may be defective (39). In addition, key IFN-inducible genes, such as PKR, themselves may be defective. In this light, many of the key components of the RLH and TLR as well as STING pathways, such as TLR3, RIG-I, MDA5, and IRF7, are IFN-inducible themselves (40). Thus, a defect in JAK/STAT signaling would prevent the production of such molecules that may enable the efﬁcient production of type I IFN in the infected cell. Given this, we also include a section of the analysis of the JAK/STAT pathway. 1.4. JAK/STAT Signaling

IFN-D/E can be secreted from most cell types, and TLR7dependent pDCs are able to produce type I IFN in very high quantities. Type I IFN can act in an autocrine and paracrine manner and bind to and activate the IFN-D/E receptor (IFNAR), which is composed of two subunits IFNAR1 and IFNAR2 (41, 42). Activation of the IFNAR results in a triggering of the JAK-STAT signaling cascade. After engagement with type I IFN, JAK1 associates with the intracellular domain of IFNAR, as well as TYK2 (43). Additionally, JAK1 has tyrosine kinase activity, which allows it to phosphorylate the phosphotyrosine-binding domain (SH2) on SH2 containing STATs. Phosphorylated STATs then homo- or heterodimerize (42). Although STAT-1 and STAT-2 are commonly activated by type I IFNs, additional STAT molecules including STAT-3, -4, -5, and -6 can also be activated by this pathway (44). Predominantly, hetero- or homodimerized STAT-1/2 associate with an additional molecule, IRF9, prior to the complex (IFNstimulated gene factor 3 – ISGF3) translocating into the nucleus. After translocation into the nucleus ISGF3 binds to IFN-stimulated response elements (ISRE) or IFN-activated sites (GAS) in the promoters of more than 100 IFN stimulated genes (ISGs) such as PKR, Mx, OAS, IFI6, and ISG15 which results in their transcriptional activation (45). Furthermore, IFN activated JAK-STAT signaling can activate additional pathways including the p38 mitogen activated protein kinase (MAP kinase) and the phosphatidylinositol 3 kinase (PI3K) pathways (45, 46). Thus, a defect in

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this pathway would suppress the induction of type I IFN itself, since TLR3, RIG-I/MDA5, and IRF7 as well as IRF9 are IFNinducible, as well as a plethora of important IFN-inducible antiviral genes such as PKR (40). While it is highly plausible that key IFNregulated antiviral gene products could be mutated in numerous cancers, this prospect has been difﬁcult to assess due to the large number of genes induced by IFN. As mentioned, key IFN-inducible genes that could be damaged in cancer include the following: PKR, 2,5 OAS, MX1/2, RNase L, ISG15, IFI6, CASP3, STAT-1, CFB, IRF7, TAP2, IL6, RANTES, and IL15 (40). It is also noteworthy that other elements of the innate immune response that does not directly involve IFN could also be damaged in cancer. For example, all viruses require translation, and conceivably ribosomal checkpoints normally responsible for preventing the unscheduled translation of viral mRNAs could also be commonly defective in cancer cells. For example, Ras-transformed MEFs or telomerase-immortalized human ﬁbroblasts are highly susceptible to VSV infection due to aberrant eIF2B-guanine nucleotide exchange rendering these cells insensitive to the phosphorylation of eIF2B by PKR in response to viral infection (47). Another defect in cancer cells that could facilitate increased viral replication may occur in RNA trafﬁcking as shown by knockdown of NFAR, a molecule that is important for regulating RNA export from the nucleus (48). Additionally, cancer cells could also subvert the IFN response through translational repression of genes that lead to the expression of IFN such as IRF7, which can be repressed at the RNA level by 4E-BPs, a negative regulator of IFN signaling, leading to increased viral replication (49). Furthermore, IFN can activate transcription of p53 through an ISRE in its promoter and ﬁrst intron. p53 is a putative regulator of cell cycle, apoptosis, and senescence and is commonly mutated in cancer, which can inhibit its function and lead to deregulated replication (50). p53 can also positively feedback and lead to an increase in IFN production by upregulating TLR3 (14). IFN can also regulate micro RNAs (miRNA) such as mir-122, which plays a role in the establishment of an antiviral state, or mir-21, which is downregulated by STAT3, and contributes to IFNs antitumor effects observed in glioma (51, 52). Moreover, cancer cells can inhibit the transcription of tumor suppressors and other genes by altering the methylation status of the promoters or genomic sequences of these genes, thereby preventing their production (53). 1.5. Innate Immune Defects in Cancer

A variety of evidence now indicates that innate immune pathways may be suppressed in cancer. For example, documented defects include but are not limited to alternative spliced isoforms of IRF-3 in hepatocellular carcinoma (54), CpG methylation of IRF-7 in gastric and lung cancers (55), mutations of CYLD in skin appendage tumors (56), reduced ability to phosphorylate STAT-1 and

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activate transcription of IFN stimulated genes (ISGs) in melanoma (57, 58), mutations in JAKs leading to constitutive activation in childhood ALL (59), and TRAF-3 mutations in multiple myeloma and some B-cell lymphomas (60, 61). As a result, many cancer cells are unable to mount an effective immune response against infection while surrounding normal tissues are well protected by an intact innate immune system. However, although TBK-1 is a critical regulator of type I IFN signaling this kinase also appears to have an unexpected role in the oncogenesis. Recently, it has been illustrated using siRNA knockdown that TBK-1 is required for KRAS-driven tumorigenesis in lung cancer. In this KRAS model, TBK-1 speciﬁcally activates NF-NB signaling leading to production of antiapoptotic proteins such as BCL-XL, which facilitates tumor formation (62). Thus, it appears that important components of the type I IFN pathway such as TBK-1 have multiple functions that can include the production of IFN or the targeting of other transcription factors that activate a different set of genes such as those involves with cell death. In any event, these ﬁndings have lead to the concept that TBK-1 targeting for suppression by drugs will prove to be an effective tumor therapy. While such data indicates that TBK-1 knockdown does inhibit KRAS-dependent tumor growth it may also suppress the general antiviral response in healthy cells. This would make normal cells highly susceptible to viral infection. The assays described in this chapter can be used to determine where defects in type I IFN signaling occur though the pathways emphasized above. These assays include IFN-D/E luciferase assays used to assess activation of the IFN-D/E promoter, western blotting for native and active phosphorylated forms of signaling proteins, ELISA to determine if IFN-D/E proteins are being produced and secreted, qRT-PCR to evaluate whether transcription of downstream targets regulated by type I IFN is occurring, and immunostaining for IRF-3 or 7 to determine if these transcription factors are translocating to the nucleus. Furthermore, it is possible establish if defects occur up or downstream of TBK-1 or IFN-D/E production by activating each pathway speciﬁcally. This can be done by transfecting cDNA plasmids; TRIF (TLR), IPS-1 (RLH), STING (ISD), TBK-1(IRFs, JAK-STAT), nucleotide analogs; Poly (I:C) (TLR), Poly (dA:dT) (ISD), pretreating cells with recombinant IFN-D/E protein (JAK-STAT), or infecting cells with DNA or RNA viruses.

2. Materials 2.1. Activation of Innate Signaling Pathways

1. 293-T cells (or other cancer cell line). 2. Opti-MEM reduced serum media (GIBCO). 3. Lipofectamine 2000 (INVITROGEN).

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4. Plasmids: pCDNA-3.1 (vector control), pCDNA-3.1-IPS-1, pCDNA-3.1-TRIF, pCDNA-3.1-RIG-I, pCDNA-3.1-STING, pCDNA-3.1-TBK-1, poly (I:C), poly (dA:dT). 5. Recombinant Vesicular Stomatitis Virus expressing Green Fluorescent Protein (VSV-GFP). 2.2. Cell Culture

1. 293-T cells: Dulbecco’s Modiﬁed Eagle Medium (DMEM) (Gibco) supplemented with 10% fetal bovine serum (FBS) (Gibco) and 1% antibiotic/antimycotic solution (Gibco). 2. 0.05% trypsin-EDTA (Gibco). 3. Sterile phosphate buffer saline (PBS) (Gibco).

2.3. SDSPolyacrylamide Gel Electrophoresis

1. RIPA Buffer: 50 mM Tris–HCl pH 7.8, 150 mM NaCl, 5 mM EDTA, 1% NP-40 (IGEPAL), 0.1% SDS, 0.75% sodium deoxycholate, 1 mM DTT, 10 mM NaF, deionized water, and protease inhibitor tablet (complete) 1 tablet per 50 mL (Roche), .1 mM PMSF (add PMSF just before use) store aliquots at −20°C. 2. Separating buffer 1.5 M Tris–HCl pH 8.8. 3. Stacking buffer 0.5 M Tris–HCl pH 6.8. 4. 40% Stock (19:1 Ratio) acrylamide/bis-acrylamide (Sigma). 5. 10% SDS solution (Gibco). 6. N,N,N,N c-Tetramethyl-ethylenediamine Science).

(TEMED)

(EM

7. 10% ammonia persulfate (APS) in water (Store at −20°C). 8. dH2O. 9. 100% isopropanol. 10. 6× SDS sample loading buffer: 1.21 g SDS, 6 mg bromophenol blue, 4.7 mL glycerol, 1.5 mL 0.5 M Tris–HCl pH 6.8, 2.1 mL dH2O; shake and warm until dissolved, then add 0.93 g DTT, aliquot and store at −20°C. 11. Running buffer (10×): Tris–HCl 121 g, HEPES 238 g, SDS 10 g, dH2O to 1 L. 12. Precision plus protein standard kaleidoscope (Bio-Rad). 13. Coomassie plus protein assay reagent (Thermo Scientiﬁc). 14. Albumin standard (Thermo Scientiﬁc). 2.4. Western Blotting for Innate Signaling Proteins

1. Transfer buffer (1×): Tris Base 3 g, glycine 14.4 g, 200 mL methanol, dH2O to 1 L. 2. BioTrace polyvinylidene ﬂuoride transfer membrane (PVDF) 0.45um (Pall Corp.). 3. 100% methanol.

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4. PBS-T: PBS (Gibco) with 0.1% TWEEN-20. 5. Blocking buffer: 5% (w/v) nonfat dry milk in PBS-T. 6. Primary antibody dilution buffer: PBS-T with 1%(w/v) bovine serum albumin (BSA) (Sigma). 7. Primary antibodies: see Table 1 below. 8. Secondary antibody: goat anti-mouse IgG or goat anti-rabbit IgG (per the primary antibody) conjugated to horseradish peroxidase (Santa Cruz). 9. Detection reagents: SuperSignal West Pico stable peroxide solution and luminol enhancer solution (Thermo Scientiﬁc). 10. X-ray ﬁlm (Kodak). 11. X-ray ﬁlm cassette (BIO-RAD). 2.5. IFN-a /b Luciferase Reporter Assay

1. Opti-MEM reduced serum media (GIBCO). 2. Lipofectamine 2000 (INVITROGEN). 3. Fireﬂy luciferase reporter plasmid (pIFN-D/E-luc). 4. Renilla luciferase transfection control plasmid (pRLTK). 5. pCDNA-3.1, pCDNA-3.1-IPS-1, pCDNA-3.1-TRIF, pCDNA-3.1-RIG-I, poly (I:C), poly (dA:dT). 6. Recombinant vesicular stomatitis virus expressing green ﬂuorescent protein (VSV-GFP). 7. Luciferase assay system (PROMEGA). 8. Luminometer (Turner Designs).

2.6. ELISA for hIFN-a /b

1. Verikine human IFN-D/E ELISA kit (PBL-Interferon Source). 2. Supernatants from treated cells. 3. Microplate reader.

2.7. qRT-PCR for mRNA Levels of Type I IFN-Induced Proteins

1. RNeasy kit (Qiagen). 2. QIAshredder (Qiagen). 3. Thin-walled PCR tubes. 4. AMV reverse transcriptase (Promega). 5. 10× reverse transcriptase buffer (Roche). 6. dNTP (Promega). 7. 25 mM MgCl2. 8. Random hexamer (or oligo d(T)) (Promega). 9. RNase inhibitor (Promega). 10. PCR-grade H2O. 11. Taqman Probe (ABI).

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Table 1 Primary antibodies for Western blotting Antibody

TLR pathway

TLR 3

Ú

ebiosciences

TLR 7

Ú

Alexis

TLR 8

Ú

Abcam

TLR 9

Ú

Abcam

MyD88

Ú

Stressgen

IRAK 1

Ú

Santa Cruz

IRAK 4

Ú

Upstate

TRAF 6

Ú

Santa Cruz

RIP-1

Ú

p38 MAPK

Ú

IKKi

Ú

Ú

Ú

Calbiochem

IkBa

Ú

Ú

Ú

Santa Cruz

pIkBa

Ú

Ú

Ú

Cell Signaling

IkBb

Ú

Ú

Ú

Santa Cruz

NF-kB

Ú

Ú

Ú

Cell Signaling

TBK-1

Ú

Ú

Ú

Abcam

IRF 3

Ú

Ú

Ú

Zymed

pIRF 3

Ú

Ú

Ú

Abcam

IRF 7

Ú

Ú

Ú

Ú

Abcam

RIG-I

Ú

Ú

Ú

Cell Signaling

MDA-5

Ú

Abcam

IPS-1

Ú

Alexis

TRAF 3

Ú

Abcam

TANK

Ú

Santa Cruz

FADD

Ú

Upstate

pFADD

Ú

Cell Signaling

Fas

Ú

Upstate

TRIF STING JAK 1

RLH pathway

SRP pathway

Jak/Stat

ISGs

Ú

Company

Abcam Ú

Ú

Santa Cruz

Alexis Ú

IMGENEX Ú

Biosource

(continued)

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Table 1 (continued) Antibody

TLR pathway

RLH pathway

SRP pathway

Jak/Stat

ISGs

Company

STAT 1

Ú

Ú

Upstate

pSTAT-1

Ú

Ú

Upstate

STAT 2

Ú

Upstate

pSTAT-2

Ú

Upstate

STAT 3

Ú

Millipore

STAT 5

Ú

Millipore

IRF 9

Ú

Abcam

Tyk2

Ú

Sigma

Gab

Ú

Santa Cruz

PI3K

Ú

Santa Cruz

CBP/p300

Ú

Upstate

IFN-D

Ú

Ú

Ú

Ú

Abcam

IFN-E

Ú

Ú

Ú

Ú

Abcam

p53

Ú

Santa Cruz

PKR

Ú

Santa Cruz

MX1

Ú

Santa Cruz

RNase L

Ú

Millipore

ISG15

Ú

Cell Signaling

IFIT3

Ú

Abcam

RANTES

Ú

Abcam

12. LightCycler FastStart DNA master hybridization probes (Roche). 13. 18S RNA probe. 14. DNA-free (Ambion). 15. Capillary cooling block. 16. 20 Pl light cycle capillaries (Roche). 17. PCR machine (Eppendorf). 18. qRT machine (Light Cycler) (Roche). 19. Light cycler carousel centrifuge (Roche).

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2.8. Immunostaining for IRF-3, -7 Nuclear Translocation

1. Sterile PBS. 2. Bovine serum albumin (BSA) (Sigma). 3. Blocking buffer 1% BSA in PBS. 4. Triton X-100 0.2% in PBS. 5. Formaldehyde 4% in DMEM. 6. Primary antibody: IRF-3, -7 (Santa Cruz). 7. Fluorophore-conjugated secondary antibody (Sigma). 8. 4c,6-diamidino-2-phenylindole (DAPI). 9. Microscope slides (VWR). 10. Mounting media: ProLong gold antifade reagent (Invitrogen).

3. Methods 3.1. Activation of Innate Signaling Pathways

It is possible to activate these pathways either by transfection (step 2), pretreatment with recombinant IFN-D/E (step 3), or VSV infection (step 4). 1. In a 12-well cell-culture plate seed 2 × 105 HEK-293 T cells (or other cancer cell line) and allow to adhere overnight (O/N). 2. Prepare the transfection in 1.5-mL eppendorf tubes. Each transfection will require two tubes. In the ﬁrst tube add 250 PL serum free Optimem and the needed amount of Lipofectamine 2000 (~1 Pl/Pg DNA). In the second tube, add 250 PL serum free Optimem and the plasmid cDNA or the DNA or RNA analog to be transfected (pCDNA3.1-IPS-1, TRIF, STING, RIG-I, TBK, poly (I:C), poly (dA:dT)). Allow the tubes to sit separated at room temperature (RT) for 10 min. Then, combine and mix the tubes by inverting and incubate at room temperature for 20 min. After the incubation, remove the media from the cells and wash one time with PBS. Then, plate the transfection mix and incubate for 3–4 h at 37°C. Following this incubation, remove the transfection from the cells and add complete media. 3. To activate cells by pretreating with IFN-D/E prepare a stock of complete media with IFN-D/E add such that the concentration is 100–500 U/mL. Remove media form the plate and add 1 mL of IFN-D/E complete media. 4. In order to prepare the virus for infection ﬁrst count one well of cells. This will allow calculation of the multiplicity of infection (MOI), which generally ranges from 0.01 to 10 for VSV. After counting the cells remove a stock of puriﬁed, titered virus from −80°C and calculate the amount of viral stock needed (see Note 1). Add the amount of virus stock to 300 Pl serum

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free media. Remove the complete media from the cells, wash one time with PBS, and plate the infections. Incubate for 1 h at 37°C. Then, remove the virus infection wash two times with PBS and add complete media. 5. Incubate for desired time 4–48 h at 37°C and use for the following assays. 3.2. Preparation of Samples for Western Blotting

1. Seed 293 T cells in 12-well plates at a density of 2 × 105 cells/ well and allow to adhere O/N. 2. Activate the cells as previously described in Subheading 3.1. Transfect with cDNA, nucleotide analogs, or infect with virus or treat with 100–500 U/mL of recombinant human IFN-D/E. 3. After desired incubation period generally 4–24 h prepare cells for lysis by collecting the supernatant in a 1.5-mL tube and freezing at −80°C washing one time with cold PBS (see Note 2). 4. Lyse the cells by adding 50–100 Pl (depending on protein concentration desired and number of gels to run) of ice-cold RIPA buffer. Rock plates for 5 min at 4°C and transfer lysate to 1.5-mL eppendorf tube and centrifuge for 2 min at 13,000 × g. Keep samples on ice. 5. Quantify the amount of protein in each sample using Coomassie Plus Protein Assay reagent and a spectrophotometer reading at 595 nm absorbance. First prepare a standard curve using albumin standard. Make six serial dilutions from a beginning stock of 2 Pg/mL. Add 2 Pl of each dilution to 1 mL Coomassie and read at 595 nm absorbance (see Note 3). After generating the standard curve, read each sample by adding 2 Pl of sample to 1 mL of Coomassie reagent, vortex brieﬂy, add to cuvette, and read at 595 nm absorbance. 6. Prepare the samples by adding 20 Pg of total protein to 5 Pl of loading buffer. Boil the sample at 100°C for 5 min. Then, centrifuge the sample from 2 min at 13,000 RPM. The samples are now ready for SDS-polyacrylamide gel electrophoresis (SDS-PAGE).

3.3. SDS-PAGE

1. This protocol is designed for use with the BIORAD MINIProtean gel system. It is, however, easily adaptable to other formats including precast and larger gels. It is essential that the gel casting plates are cleaned prior to use with dH2O and 70% ethanol. Dry the plates with a Kimwipe. 2. Prepare a 1.0-mm thick 10% gel by mixing 4.8 mL dH2O with 2.5 mL 40% acrylamide/bis-acrylamide solution, 2.5 mL 1.5 M Tris–HCl pH 8.8, 100 Pl 10% SDS, 100 Pl 10% APS, and 10 Pl TEMED for a total of 10 mL, enough for two gels. Pour the gel leaving space for the stacking gel and overlay with 100% isopropanol. The gel should polymerize in 15–20 min.

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3. Pour off the isopropanol, and wash the top of the gel with water. Remove the remainder of the water using a kimwipe or blotting paper. 4. Prepare a 5% stacking gel by mixing 3 mL dH2O with 625 Pl 40% acrylamide/bis-acrylamide solution, 1.25 mL 0.5 M Tris– HCL pH 6.8, 50 Pl 10% SDS, and 50 Pl 10% APS, 8 Pl TEMED for a total of 10 mL, enough for two gels. Pour the stacking gel to the top of the glass plate and insert the comb. The stacking gel should polymerize in 15–20 min. 5. Prepare the running buffer by diluting 100 mL of the 10× running buffer with 900 mL dH2O. 6. Once the stacking gel has set, carefully remove the comb and use a squirt bottle to wash the wells and remove bubbles with dH2O. 7. Assemble the gel unit. Be sure to check that the casting plates form a good seal with the rubber insert. 8. Add the running buffer to the inner and outer chambers of the gel unit and load the sample to each well. Include one well for prestained molecular weight marker. 9. Complete the assembly of the gel unit and connect to a power supply. Set the power supply to constant volts and run the gel at 80–100 V. The dye front (blue) can be run off the gel if desired. 3.4. Western Blotting for Innate Immune Proteins

1. After the samples have been separated by SDS-page the protein needs to be transferred to a PVDF membrane electrophoretically. Cut pieces of PVDF membrane just larger than the gel 8 mm × 6 mm and soak in 100% methanol for at least 30 min (see Note 4). 2. These directions assume the use of a BIO-RAD transfer system. Set up a tray of transfer buffer that is large enough to accommodate the transfer cassette with the foam pad and two pieces of blotting paper submerged on the anode (clear plastic) side. Place the PVDF membrane on top of the submerged blotting paper. 3. Disconnect and disassemble the gel unit from the power supply and remove and discard the stacking gel. Rinse the separating gel with 1× transfer buffer (in the tray) and place on top of the PVDF membrane. Check to make sure there are no bubbles between the gel and the membrane. If bubbles are present they can be rolled out using a sterile serological pipette or by gently guiding them using a ﬁnger. 4. Carefully place two more pieces of blotting paper that has been prewet in 1× transfer buffer on top of the gel, taking care to not introduce bubbles into the resulting sandwich. Place the second foam pad on top of the blotting paper and close the transfer cassette.

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5. Place the cassette into the transfer tank such that the PVDF membrane is between the gel and the anode. This orientation is critical for the capture of proteins, if placed in backward proteins will be run into the buffer and be lost. 6. Place an ice block into the transfer box and ﬁll with 1× transfer buffer (see Note 5). 7. Put the lid on the tank and set the power supply for constant amperes. Run the transfer at 240 mA for 90–110 min depending on the protein target size. 8. Once the transfer is complete and the cassette has been removed and disassembled remove the PVDF from the gel. The molecular weight ladder should be clearly visible on the membrane. 9. Place the membrane in 10 mL of blocking solution and shake for 1 h at RT on a rocking or shaking platform. 10. Discard the blocking solution and wash the membrane 3× for 5 min with PBS-T. 11. Add the primary antibody in PBS-T/1% BSA. Incubate overnight at 4°C on a rocking platform. Dilutions generally range from 1:250 to 1:1,000 depending on which primary antibody is being used. 12. Remove the primary antibody and wash the membrane 3× for 5 min with 10 mL PBS-T. 13. Prepare the secondary antibody in blocking buffer at a 1:5,000 dilution. After the ﬁnal wash add 10 mL to the membrane and incubate for 1 h at RT on a rocking or shaking platform. 14. Discard the secondary antibody and wash the membrane 3× for 10 min each with PBS-T. 15. After the ﬁnal wash prepare the ECL reagents by mixing equal volumes of substrate and luminol enhancer. Remove excess PBS-T by removing the membrane and blotting one corner with a paper towel. Add the mixed ECL reagents to the membrane and shake by hand for 1 min to ensure complete coverage. 16. Remove the blot from the ECL reagents and blot onto a paper towel to remove excess reagent. Place the membrane into the leaves of a plastic sheet protector and place into a ﬁlm cassette. 17. Take the cassette and X-ray ﬁlm to a dark room for developing. Place a piece of X-ray ﬁlm into the cassette for a suitable exposure time (see Note 6). After the exposure develop the ﬁlm. 3.5. IFN-a /b Luciferase Assay

1. In a 12-well cell-culture plate, seed 2 × 105 HEK-293-T cells allow to adhere overnight (O/N). 2. Treat cells as previously described in Subheading 3.1.

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3. After the desired incubation time prepare a 1× solution of Cell Culture Lysis Reagent (CCLR) (PROMEGA) from the 5× stock solution with PBS. Remove the media from the cells and wash one time with PBS (see Note 7). Add 100 Pl 1× CCLR to each well and rock the plate for 5 min at 4°C. Then, transfer the lysate to 1.5-mL eppendorf tubes and centrifuge for 2 min at 13,000 RPM. 4. After preparing the sample turn on the luminometer (Turner Designs TD 20/20) to run the luciferase assay. First read the ﬁreﬂy luciferase by adding 10 Pl of the cell lysate with 10 Pl ﬁreﬂy luciferase substrate in a 0.5-mL tube. Mix by pipetting up and down three to four times and then placing the tube in the luminometer. Set the luminometer to wait for 2 s and then integrate (read) the signal for 10 s. The value it returns represents the amount the IFN-E promoter was induced. Next, read the renilla luciferase by adding 10 Pl of lysate to 10 Pl renilla luciferase substrate and mix by pipetting three to four times. Then, read the as described above. This value represents the transfection control (see Note 8). 3.6. ELISA for IFN-a /b

1. Treat the cells as previously described in Subheading 3.1. After the desired incubation time remove and collect the supernatant into 1.5-mL eppendorf tubes. Centrifuge the supernatants to remove any cells. These can then be stored at −80°C if the ELISA is not being run immediately after collection. 2. Prepare 1× wash buffer from the 10× wash concentrate and prepare a 1:10 working stock of the mouse IFN-E standard by pipetting 10 Pl mouse IFN-E standard (Vial B) in 90 Pl of sample diluent (Bottle C). From this working stock (50,000 pg/ mL) prepare the standard curve, with the ﬁrst dilution being 1,000 pg/mL and serial diluting 1:1 in assay diluent to 15.6 pg/mL and one blank. Prepare fresh dilutions of the standard curve for each assay run. 3. Determine the number of precoated wells needed to run the assay. Add precisely 100 Pl of interferon samples and standard curve prepared in the previous step in individual wells of the microplate in duplicate (see Note 9). 4. Cover the plate with a plastic plate sealer (included in the kit) and incubate for 60 min at RT. After the 60 min incubation was the plate 3× with at least 250 Pl wash solution blotting well after the ﬁnal wash. 5. During the incubation, prepare the antibody solution. Dilute the Antibody Concentrate with Concentrate Diluent, refer to the Certiﬁcate of Analysis (COA) to determine the correct amounts of each reagent to use. Store unused undiluted antibody concentrate at 2–8°C for future use.

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6. Add 100 Pl of the antibody solution prepared using the COA to each well. Cover the plate with a plastic plate sealer (included in the kit) and incubate for 60 min at RT. After the 60 min incubation was the plate 3× with at least 250 Pl wash solution blotting well after the ﬁnal wash. 7. During the incubation prepare the HRP solution. Dilute the HRP Concentrate with HRP conjugate Diluent, refer to the Certiﬁcate of Analysis (COA) to determine the correct amounts of each reagent to use. Store unused undiluted antibody concentrate at 2–8°C for future use. 8. Add 100 Pl of the HRP solution prepared using the COA to each well. Cover the plate with a plastic plate sealer (included in the kit) and incubate for 60 min at RT. After the 60 min incubation was the plate 3× with at least 250 Pl wash solution blotting well after the ﬁnal wash. 9. During the HRP incubation period warm the TMB substrate solution to RT. Add 100 Pl of the TMB substrate solution to each well. Incubate for 15 min at RT in the dark. Do not seal the plate during this step. 10. Add 100 Pl of Stop solution to each well. Mix gently using caution to avoid bubbles as this can affect results. Using a microplate reader, determine the absorbance at 450 nm within 5 min of adding the stop solution. Unknown sample concentrations can be determined by extrapolation off the standard curve. 3.7. qRT-PCR for mRNA Levels of Type I IFN-Induced Proteins

1. Treat the cells as previously described in Subheading 3.1. 2. Harvest the mRNA using the mRNeasy kit form Qiagen. Determine the yield using a spectrophotometer (see Note 10). 3. First cDNA must be synthesized from the RNA template. Label the required number of thin walled PCR tubes on a cooling block or ice. Add up to 1 Pg of total RNA in a total volume of 5 Pl. Include one tube with only water as a negative control (see Note 11). Denature the RNA for by placing the tubes in a thermocycler at 65°C for 10 min. 4. During the RNA denaturation prepare the cDNA master mix. For 20 Pl of cDNA (enough for 10 qRT reactions) add 3.2 Pl PCR-grade water, 2 Pl 10× RT buffer, 0.8 Pl AMV RT, 2 Pl dNTP, 4 Pl 25 mM MgCl2, 2 Pl Random hexamer (or oligo d(T)), and 1 Pl RNase inhibitor (see Note 12). 5. Spin PCR tubes quickly in a bench top centrifuge and add 15 Pl of the master mix to each tube. Mix by pipetting up and down a few times, do not vortex. Perform the cDNA synthesis in a thermocycler with the following program. 10 min 25°C, 60 min 42°C, 5 min 99°C (denature), Hold at 4°C. Use 2 Pl of the reaction product as template for qRT-PCR in the light cycler.

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6. If using Taqman probes, use the LightCycler FastStart DNA Master Hybridization Probes from Roche to set up the PCR mix. Each kit contains three tubes labeled 1b and one tube labeled 1a. To make the PCR mix add 10 Pl of enzyme (tube 1a) to a tube of buffer mix (tube 1b). Make the PCR master by mixing the following components: PCR mix 4 Pl, taqman probe 1 Pl, 18S rRNA probe 1 Pl, and H2O 12 Pl (see Note 13). 7. Take the capillary cooling block out of the refrigerator and place the required number of 20 Pl light cycler capillaries in to the slots. Add 2 Pl cDNA sample to each capillary tube, taking care not to jam the tip in too far and break the capillary. Apply 18 Pl of the PCR master mix to each capillary and mix by pipetting. Place the capillaries onto the carousel provided for the light cycler and spin down the capillaries to get the samples to the bottom using the light cycler the LightCycler carousel centrifuge. 8. Load the carousel in the LightCycler and execute the program. The run takes approximately 1.5 h. 3.8. Immunostaining for IRF-3, -7 Nuclear Translocation

1. On the day before treatment seed cells in slide chamber plates or on 15 mm cover slips in a 10-cm cell-culture plate (Confocal microscope). Alternatively, this can be done in 12-well plates (Fluorescent microscope). 2. Treat the cells as previously described in Subheading 3.1. After the desired incubation time (1–4 h), wash the cells 2× for 5 min with PBS. 3. Fix the cells with 4% formaldehyde for 15 min at 37°C. Wash 3× for 5 min with PBS. 4. Permeabilize the cells with 0.2% Triton X100 for 5 min at 4°C. Wash the cells 3× with PBS. 5. Add blocking buffer and incubate overnight at 4°C. Remove the blocking buffer and add primary antibody (mouse antiIRF-3) in PBS with 1% BSA and incubate for 1 h at RT. During this incubation remove the mounting media from the freezer. 6. After the primary antibody incubation wash 3× for 5 min with PBS. Next add a secondary antibody that is conjugated to a ﬂuorophore (mouse anti-FITC green or CY3 red) Incubate at RT for 30 min. Wash 3× for 5 min with PBS. 7. Add DAPI 1 Pg/mL to stain the nucleus. Incubate for 10 min at RT. Wash 3× for 5 min with PBS. 8. Mount slides and allow to dry for ~12 h (see Note 14). 9. Store in the dark (or wrap in tin foil) to prevent photobleaching and analyze by confocal microscopy within 24 h of staining.

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In conclusion, it is highly plausible that defects in innate immune signaling pathways involving the production of type I IFN, or JAK/STAT pathway, or in key IFN-inducible genes themselves, may enable the replication of viruses to proceed. Research into these processes will undoubtedly shed light into mechanisms of oncolysis, which may be frequently defective in cancer, and so bring to light important causes of tumorigenesis. Understanding these processes will also enable the design of new oncolytic agents that could exhibit more speciﬁc and enhanced antitumor activity (6). Further unraveling of these important signaling cascades, thus, affords a multitude of opportunities to combat malignant disease.

4. Notes 1. The equation used to determine the amount of virus needed for infection of cells (# of cells × Multiplicity of Infection (MOI))/Virus Titer = mL viral stock needed. 2. If cells were infected with virus many cells may be ﬂoating in supernatant at later time points. These supernatants should be spun down to pellet and remove the cells prior to storage. 3. It is best to add the 2 Pl of sample to the Coomassie just prior to mixing and reading. As samples sit for a longer period of time in the Coomassie reagent, the color change becomes more pronounced and higher OD readings are returned. 4. This can be done while the SDS-page gel is running. 5. Alternatively, place the transfer box in a bucket and surround it with ice or run the transfer in a 4°C cold room. 6. Exposure times can range from 5 s to 10 min depending on the strength of signal Additionally, if the has a strong signal creating a “dirty” blot insert two pieces of ﬁlm (one on top of the other) and expose. This can reduce the background observed on the ﬁlm. Furthermore, to maintain the orientation of the ﬁlm make a crease in the top left hand corner of the ﬁlm. 7. If cells were infected with virus many cells may be ﬂoating in the supernatant. These cells should also be pelleted and lysed. 8. To normalize the data, divide the ﬁreﬂy value by the renilla value. You can then determine the fold induction by dividing the normalized values by either the mock infected or vector transfected control. 9. Samples may need to be diluted 1:5 or 1:10 with assay diluent so that upon completion of the assay the reading will fall within the detectable range of the standard curve. Values can then be multiplied by the dilution factor to determine the concentration.

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10. If the RNA will be used to assess IFN-E mRNA levels the samples need to be treated with DNase I (DNA-free kit Ambion) to remove the genomic DNA. This will greatly reduce the background during assessment of IFN-E mRNA. 11. As low as 0.1 Pg of RNA is acceptable to use for the cDNA synthesis reaction and in some cases can yield better results. 12. This mix is for 1 cDNA reaction and can be scaled up for as many reactions as needed. If preparing master mix for multiple reactions make 5–10% additional mix to prevent running out when aliquotting it to tubes. 13. This master mix is for one reaction but can be scaled up for as many reactions as needed with a particular probe. 14. More than one primary antibody can be used, useful for studying colocalization, as long as the primary antibodies come from different species and each secondary has a different ﬂuorophore. References 1. Barber GN. (2004) Vesicular stomatitis virus as an oncolytic vector Viral Immunol 17, 516–527. 2. Barber GN. (2005) VSV-tumor selective replication and protein translation Oncogene 24, 7710–7719. 3. Choi MK, Wang Z, Ban T, Yanai H, Lu Y, et al. (2009) A selective contribution of the RIG-Ilike receptor pathway to type I interferon responses activated by cytosolic DNA Proc Natl Acad Sci USA 106, 17870–17875. 4. Yoneyama M, Fujita T. (2009) RNA recognition and signal transduction by RIG-I-like receptors Immunol Rev 227, 54–65. 5. Blasius AL, Beutler B. (2010) Intracellular tolllike receptors Immunity 32, 305–315. 6. Obuchi M, Fernandez M, Barber GN. (2003) Development of recombinant vesicular stomatitis viruses that exploit defects in host defense to augment speciﬁc oncolytic activity Journal of Virology 77, 8843–8856. 7. Akira S, Uematsu S, Takeuchi O. (2006) Pathogen recognition and innate immunity Cell 124, 783–801. 8. Wang L, Ligoxygakis P. (2006) Pathogen recognition and signalling in the Drosophila innate immune response Immunobiology 211, 251–261. 9. Silverman N, Maniatis T. (2001) NF-kappaB signaling pathways in mammalian and insect innate immunity Genes & Development 15, 2321–2342. 10. Leclerc V, Reichhart J-M. (2004) The immune response of Drosophila melanogaster Immunol Rev 198, 59–71.

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INDEX A Adenovirus.............................. 20–33, 35–51, 53, 56–57, 68, 69, 75, 80, 91, 94, 97–108, 111–125, 221 Apoptosis................................................164, 165, 168–170, 172–174, 188, 190, 206, 222 Autophagy...................................... .112, 115–116, 120–123

B BAC. See Bacterial artificial chromosome (BAC) Bacterial artificial chromosome (BAC)...............3, 5, 10–12, 14–16, 18 Banding.................................. 24, 30–31, 40, 43, 44, 50, 101 B-cell................................................................130, 142, 223 Biodistribution ........................21, 23, 31, 32, 37, 47, 49, 53, 54, 211, 213 Brain cancer .........................................18, 97–108, 111–125 Breast cancer....................................................141, 181, 185

C Capsid................................................... 2, 21–33, 35, 37, 41, 43, 47, 50, 127 CAR. See Coxsackie-adenovirus receptor (CAR) CD46........................................... 37, 43, 142, 143, 150, 154 CD133.............................................................105, 112, 119 Cell carrier .................................................99–101, 106–108 Colorectal................................................................ . 76, 201 Coxsackie-adenovirus receptor (CAR) .................22, 28, 36, 37, 43, 66, 69, 73, 77, 114, 120 Cytotoxicity ....................... 66, 112, 115, 120, 177–202, 207

D Detargeting ......................................... 21, 22, 26, 32, 43–44 Dopamine D2................................................................... 90

E E1................................................................... .30, 41–46, 74 E3........................................................................ . 36, 38–46 Edmonston ............................................................. 141, 143 EGFR. See Epidermal growth factor receptor (EGFR) Endoplasmic reticulum (ER) ..................................180, 183, 188–191, 218–220

Epidermal growth factor receptor (EGFR) .................... 142 Explant..................................................................... .. 65–77

F Fusion (F) protein......................................22, 142, 143, 178

G GBM. See Glioblastoma multiforme (GBM) GFP. See Green fluorescent protein (GFP) Gene therapy ............................................. 35, 36, 89, 91, 98 Glioblastoma multiforme (GBM) ....................98, 111–113, 116–117 Green fluorescent protein (GFP)............................5, 10, 14, 15, 74, 80, 130, 143, 145, 214, 224, 225

H Hemagglutinin (H) protein ............................................ 142 Hepatic stellate cells ........................................179, 186–187 Hepatocellular ................................... 37, 141, 186, 214, 222 Herpes simplex virus (HSV) ............................1–18, 66, 75, 90, 220, 221 Hexon........ .................22, 26–28, 44, 48, 68, 73, 75–77, 104 HSV. See Herpes simplex virus (HSV)

I IFN. See Interferon (IFN) Imaging bioluminescence..................... 42, 49, 207, 211–213, 215 luminescence............................................................. 212 optical.. ............................................................... 80, 211 PET.......................................................... .80, 89, 90, 93 SPECT................................................................. 80, 90 Immunity, innate ............................... 47, 135, 201, 217–235 Immunization ....................................................... 56–57, 98 Immunohistochemistry (IHC).. .....................69, 71–73, 75, 76, 164–167, 169–171, 174, 175, 187, 211 Immunosuppression ................................................... 54, 56 Integrin................... 22, 27, 28, 66, 69, 73, 77, 114, 120, 142 Interferon (IFN) ......................... 39, 40, 128, 164, 198–201, 206, 214, 217, 219–223, 225, 227–229, 231–233, 235, 236 Intraperitoneal ............................. 56, 57, 60, 61, 92, 93, 134

David H. Kirn et al. (eds.), Oncolytic Viruses: Methods and Protocols, Methods in Molecular Biology, vol. 797, DOI 10.1007/978-1-61779-340-0, © Springer Science+Business Media, LLC 2012

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ONCOLYTIC VIRUSES 240 Index Intratumoral ..................................................36, 54, 92, 179 Intravenous ...................................... 49, 54, 92, 93, 137, 179 Iodixanol..................................................................... 21–33

L Luciferase.. ..................................... 32, 49, 79–86, 130, 207, 211–215, 223, 225, 231–232 Luciferin.... ..........................38, 49, 51, 80–82, 86, 129, 130, 134, 207, 212, 215 Lymphoma ............................................................. 141, 223

M Macrophage .......................................................48, 142, 218 MAPK............................................................................ 226 Measles...... ..................................................90, 91, 141–160 Myeloma................................................................. 141, 223

N Neural stem cells (NSCs) ..........................97–108, 111, 112 Neurosphere ............................ 112, 113, 116–117, 121, 124 Neutralizing antibody (NAb)........................54, 57–58, 129, 130, 135–137 Newcastle disease virus (NDV) ............................. 178–195, 197–201, 219 Norepinephrine ................................................................ 90

O ONYX–015 ................................................................ 36, 65 Oral gavage ........................................................... 54, 60–61 Ovarian cancer.......................................................... 65, 141

P P53................ .......................36, 180, 183, 188, 189, 222, 227 Peripheral blood mononuclear cells (PBMC)... .............................179–181, 185, 191–194, 196, 198 Plaque assay................ ........................ 47, 50, 75, 130, 132–134, 139, 175, 207, 209–210, 213 purification ........................................15–16, 45–47, 209 Polyethylene glycol (PEG) ............ 23, 29, 37, 38, 47–48, 50 pRb................. .............................................................. 38, 39 Prostate cancer.......................................................... 75, 141

R Ras.................. ................................. 163, 164, 167–169, 222 Real-time PCR (RT-PCR) .......................... 32, 38, 49, 124, 182, 184, 185, 198, 223, 225–227, 233–234 Recombination homologous .............................. 2, 23–28, 33, 39, 40, 42, 44, 45, 208, 213 site specific.................................................................. 10 Reovirus.................................................................. 163–175

Retargeting ............................................................. 141–160 Retinoic acid-inducible gene I (RIG-I) ......................... 199, 218–222, 224–226, 228 RGD................................................................112, 122, 142 RIG-I. See Retinoic acid-inducible gene I (RIG-I) RT-PCR. See Real-time PCR (RT-PCR)

S Selectivity........................................ 36, 39, 40, 43, 179–180, 209–211, 214 Signaling lymphocyte activating molecule (SLAM) .........................................142, 143, 150 Signal transducers and activation of transcription (STAT) .......................... 199, 221–223, 227, 235 SLAM. See Signaling lymphocyte activating molecule (SLAM) A-Smooth muscle actin (A-SMA)..........................180, 182, 186–188 Sodium iodide symporter (NIS) ........................... 80, 89–95 Somatostatin type 2 .......................................................... 90 STAT. See Signal transducers and activation of transcription (STAT) Stem cells...................................................97–108, 111–125 Stimulator of interferon genes (STING)....................... 218, 220–221, 223, 224, 226, 228 STING. See Stimulator of interferon genes (STING) Syrian hamster .................................................48, 53–62, 66

T Targeted delivery ...................................................... 99, 142 T-cell................. ....................... 80, 192, 197, 199–201, 223, 228, 229, 231 TCID50...... .............................. 69, 73–75, 77, 157, 158, 160 Thymidine kinase (TK) ......................................2, 3, 80, 90, 206–209, 213, 214 TNA. See Tumor neutralisation assay (TNA) Toll like receptor (TLR) ..........................218–223, 226, 227 Toxicity................... .. 21, 49, 51, 53, 54, 66, 74, 98, 134, 142 Transgene... ...............................5, 10–12, 14, 15, 18, 36, 38, 39, 41–43, 49, 142, 206–209, 213, 214 Tumor neutralisation assay (TNA).. ...............180, 191–192, 194, 196, 197, 200, 201

V Vaccinia.......................................................... .205–215, 221 Vaccinia growth factor (VGF) ........................................ 206 Vesicular stomatitis virus (VSV).................90, 91, 127–140, 219, 220, 222, 224, 225, 228 Virus amplification .........................1, 4, 6, 14, 25, 70, 144, 199 cytotoxicity .................112, 115, 120, 180, 189, 199, 207 infection.............................66, 69, 72, 75, 76, 80, 98, 99, 112, 143, 164, 169, 174, 179, 186, 205, 217–220, 222, 223, 229

ONCOLYTIC VIRUSES 241 Index propogation ................................ 45, 101, 127–140, 143, 144, 153–158 purification ........................................8–10, 30, 127–140 replication .....................38, 134, 164, 166, 167, 170, 184 titering. ............................................................... 15, 129 titration ......................................................6–8, 157–158

W WST1...............................180, 182–183, 188, 189, 195, 196

Y Yeast.......................................................................... . 21–33