Human Cytomegaloviruses: Methods and Protocols (Methods in Molecular Biology, 2244) 1071611100, 9781071611104

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Methods in Molecular Biology 2244

Andrew D. Yurochko Editor

Human Cytomegaloviruses Methods and Protocols Second Edition

METHODS

IN

MOLECULAR BIOLOGY

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

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

For over 35 years, biological scientists have come to rely on the research protocols and methodologies in the critically acclaimed Methods in Molecular Biology series. The series was the first to introduce the step-by-step protocols approach that has become the standard in all biomedical protocol publishing. Each protocol is provided in readily-reproducible step-bystep fashion, opening with an introductory overview, a list of the materials and reagents needed to complete the experiment, and followed by a detailed procedure that is supported with a helpful notes section offering tips and tricks of the trade as well as troubleshooting advice. These hallmark features were introduced by series editor Dr. John Walker and constitute the key ingredient in each and every volume of the Methods in Molecular Biology series. Tested and trusted, comprehensive and reliable, all protocols from the series are indexed in PubMed.

Human Cytomegaloviruses Methods and Protocols Second Edition

Edited by

Andrew D. Yurochko Department of Microbiology and Immunology, Louisiana State University Health Sciences Center—Shreveport, Shreveport, LA, USA

Editor Andrew D. Yurochko Department of Microbiology and Immunology Louisiana State University Health Sciences Center—Shreveport Shreveport, LA, USA

ISSN 1064-3745 ISSN 1940-6029 (electronic) Methods in Molecular Biology ISBN 978-1-0716-1110-4 ISBN 978-1-0716-1111-1 (eBook) https://doi.org/10.1007/978-1-0716-1111-1 © Springer Science+Business Media, LLC, part of Springer Nature 2021 This work is subject to copyright. All rights are reserved by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Humana imprint is published by the registered company Springer Science+Business Media, LLC, part of Springer Nature. The registered company address is: 1 New York Plaza, New York, NY 10004, U.S.A.

Preface Human cytomegalovirus infection is of great concern to the medical community because of the key role the virus plays in significant acute and chronic human disease in all stages of life, from newborns to seniors. This new edition on human cytomegaloviruses in the Methods of Molecular Biology series documents the biology and pathogenesis of the virus along with the approaches currently being utilized to investigate the molecular aspects of viral infection and how these new research studies are leading to new approaches to mitigate disease. Thus, the goal for the book is to be a scientific reference for basic and clinical scientists, medical personnel, and the lay public on the modern approaches, techniques, and models to study human cytomegalovirus infection and disease. Shreveport, LA, USA

Andrew D. Yurochko

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

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1 Overview of Human Cytomegalovirus Pathogenesis. . . . . . . . . . . . . . . . . . . . . . . . . Heather L. Fulkerson, Maciej T. Nogalski, Donna Collins-McMillen, and Andrew D. Yurochko 2 Distinct Properties of Human Cytomegalovirus Strains and the Appropriate Choice of Strains for Particular Studies . . . . . . . . . . . . . . . . . . . . . . . . . Giada Frascaroli and Christian Sinzger 3 Using Diploid Human Fibroblasts as a Model System to Culture, Grow, and Study Human Cytomegalovirus Infection . . . . . . . . . . . . . Elizabeth A. Fortunato 4 Using Primary Human Cells to Analyze Human Cytomegalovirus Biology. . . . . Emma Poole, Ian Groves, Sarah Jackson, Mark Wills, and John Sinclair 5 Human Hematopoietic Long-Term Culture (hLTC) for Human Cytomegalovirus Latency and Reactivation . . . . . . . . . . . . . . . . . . . . . . Megan Peppenelli, Jason Buehler, and Felicia Goodrum 6 Collection and Isolation of CD14+ Primary Human Monocytes Via Dual Density Gradient Centrifugation as a Model System to Study Human Cytomegalovirus Infection and Pathogenesis . . . . . . . . . . . . . . . Bailey S. Mosher, Heather L. Fulkerson, and Andrew D. Yurochko 7 Stable and Inducible Gene Knockdown in Primary Human Fibroblasts: A Versatile Tool to Study the Role of Human Cytomegalovirus Host Cell Factors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Anne-Charlotte Stilp, Patrick Ko¨nig, Myriam Scherer, and Thomas Stamminger 8 Construction of Human Cytomegalovirus Mutants with Markerless BAC Mutagenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ˇ icˇin-Sˇain M. Zeeshan Chaudhry, Martin Messerle, and Luka C 9 Methods for Studying the Function of Cytomegalovirus GPCRs. . . . . . . . . . . . . . Christine M. O’Connor and William E. Miller 10 Analysis of Cytomegalovirus Glycoprotein and Cellular Receptor Interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jamil Mahmud and Gary C. Chan 11 Antibody-Independent Quantification of Cytomegalovirus Virion Protein Incorporation Using HiBiT . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Iris K. A. Jones and Daniel N. Streblow 12 Using a Phosphoproteomic Screen to Profile Early Changes During HCMV Infection of Human Monocytes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Liudmila S. Chesnokova and Andrew D. Yurochko

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A Generally Applicable CRISPR/Cas9 Screening Technique to Identify Host Genes Required for Virus Infection as Applied to Human Cytomegalovirus (HCMV) Infection of Epithelial Cells . . . . . . . . . . . . . . . . . . . . . Xiaofei E and Timothy F. Kowalik Quantitative Electron Microscopy to Study HCMV Morphogenesis . . . . . . . . . . Clarissa Read, Paul Walther, and Jens von Einem Detection of Cytomegalovirus Interleukin 10 (cmvIL-10) by Enzyme-Linked Immunosorbent Assay (ELISA) . . . . . . . . . . . . . . . . . . . . . . . . . Vivian P. Young, Margarette C. Mariano, Lionel Faure, and Juliet V. Spencer Techniques for Characterizing Cytomegalovirus-Encoded miRNAs . . . . . . . . . . . Nicole L. Diggins, Lindsey B. Crawford, Hillary M. Struthers, Lauren M. Hook, Igor Landais, Rebecca L. Skalsky, and Meaghan H. Hancock Development of a huBLT Mouse Model to Study HCMV Latency, Reactivation, and Immune Response . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Lindsey B. Crawford and Patrizia Caposio Rodent Models of Congenital Cytomegalovirus Infection . . . . . . . . . . . . . . . . . . . Berislav Lisnic´, Jelena Tomac, Djurdjica Cekinovic´, Stipan Jonjic´, and Vanda Juranic´ Lisnic´

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Recent Approaches and Strategies in the Generation of Anti-human Cytomegalovirus Vaccines. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 403 Suresh B. Boppana and William J. Britt

Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Contributors SURESH B. BOPPANA • Department of Pediatrics, The University of Alabama at Birmingham, Birmingham, AL, USA; Department of Microbiology, The University of Alabama at Birmingham, Birmingham, AL, USA WILLIAM J. BRITT • Department of Pediatrics, The University of Alabama at Birmingham, Birmingham, AL, USA; Department of Microbiology, The University of Alabama at Birmingham, Birmingham, AL, USA; Department of Neurobiology, The University of Alabama at Birmingham, Birmingham, AL, USA JASON BUEHLER • Department of Immunobiology, BIO5 Institute, The University of Arizona, Tucson, AZ, USA PATRIZIA CAPOSIO • Vaccine and Gene Therapy Institute, Oregon Health and Science University, Portland, OR, USA DJURDJICA CEKINOVIC´ • Department for Histology and Embryology and Center for Proteomics, Faculty of Medicine, University of Rijeka, Rijeka, Croatia; Department of Infectious Diseases, Faculty of Medicine, University of Rijeka, Rijeka, Croatia GARY C. CHAN • Department of Microbiology & Immunology, SUNY Upstate Medical University, Syracuse, NY, USA M. ZEESHAN CHAUDHRY • Department of Vaccinology, Helmholtz Centre for Infection Research, Braunschweig, Germany LIUDMILA S. CHESNOKOVA • Department of Microbiology & Immunology, Center for Molecular and Tumor Virology, Feist-Weiller Cancer Center, Louisiana State University Health Sciences Center—Shreveport, Shreveport, LA, USA ˇ ICˇIN-SˇAIN • Department of Vaccinology, Helmholtz Centre for Infection Research, LUKA C Braunschweig, Germany; Center for Individualized Infection Medicine (CIIM), A Joint Venture of HZI and MHH, Braunschweig, Germany DONNA COLLINS-MCMILLEN • Department of Immunobiology, BIO5 Institute, University of Arizona, Tucson, AZ, USA LINDSEY B. CRAWFORD • Vaccine and Gene Therapy Institute, Oregon Health and Sciences University, Beaverton, OR, USA NICOLE L. DIGGINS • Vaccine and Gene Therapy Institute, Oregon Health and Sciences University, Beaverton, OR, USA XIAOFEI E • Department of Microbiology and Physiological Systems, University of Massachusetts Medical School, Worcester, MA, USA LIONEL FAURE • Department of Biology, Texas Woman’s University, Denton, TX, USA ELIZABETH A. FORTUNATO • Department of Biological Sciences, University of Idaho, Moscow, ID, USA GIADA FRASCAROLI • Heinrich Pette Institute, Leibniz Institute for Experimental Virology, Hamburg, Germany HEATHER L. FULKERSON • Department of Microbiology & Immunology, Center for Molecular and Tumor Virology, Feist-Weiller Cancer Center, Louisiana State University Health Sciences Center—Shreveport, Shreveport, LA, USA; Center for Cardiovascular Diseases and Sciences, Louisiana State University Health Sciences Center—Shreveport, Shreveport, LA, USA

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FELICIA GOODRUM • BIO5 Institute, The University of Arizona, Tucson, AZ, USA; Department of Immunobiology, The University of Arizona, Tucson, AZ, USA IAN GROVES • Department of Medicine, Addenbrookes Hospital, University of Cambridge, Cambridge, UK MEAGHAN H. HANCOCK • Vaccine and Gene Therapy Institute, Oregon Health and Sciences University, Beaverton, OR, USA LAUREN M. HOOK • The Vaccine and Gene Therapy Institute, Oregon Health and Science University, Beaverton, OR, USA SARAH JACKSON • Department of Medicine, Addenbrookes Hospital, University of Cambridge, Cambridge, UK IRIS K. A. JONES • The Vaccine and Gene Therapy Institute, Oregon Health and Science University, Beaverton, OR, USA STIPAN JONJIC´ • Department for Histology and Embryology and Center for Proteomics, Faculty of Medicine, University of Rijeka, Rijeka, Croatia PATRICK KO¨NIG • Institute of Virology, Ulm University Medical Center, Ulm, Germany TIMOTHY F. KOWALIK • Department of Microbiology and Physiological Systems, University of Massachusetts Medical School, Worcester, MA, USA IGOR LANDAIS • Vaccine and Gene Therapy Institute, Oregon Health and Sciences University, Beaverton, OR, USA BERISLAV LISNIC´ • Department for Histology and Embryology and Center for Proteomics, Faculty of Medicine, University of Rijeka, Rijeka, Croatia VANDA JURANIC´ LISNIC´ • Department for Histology and Embryology and Center for Proteomics, Faculty of Medicine, University of Rijeka, Rijeka, Croatia JAMIL MAHMUD • Department of Microbiology & Immunology, SUNY Upstate Medical University, Syracuse, NY, USA MARGARETTE C. MARIANO • Department of Biology, Texas Woman’s University, Denton, TX, USA MARTIN MESSERLE • Institute of Virology, Hannover Medical School, Hannover, Germany WILLIAM E. MILLER • Department of Molecular Genetics, Biochemistry, and Microbiology, University of Cincinnati College of Medicine, Cincinnati, OH, USA BAILEY S. MOSHER • Department of Microbiology & Immunology, Center for Molecular and Tumor Virology, Feist-Weiller Cancer Center, Louisiana State University Health Sciences Center—Shreveport, Shreveport, LA, USA MACIEJ T. NOGALSKI • Department of Molecular Biology, Princeton University, Princeton, NJ, USA CHRISTINE M. O’CONNOR • Genomic Medicine, Lerner Research Institute, Cleveland Clinic, Cleveland, OH, USA MEGAN PEPPENELLI • BIO5 Institute, The University of Arizona, Tucson, AZ, USA EMMA POOLE • Department of Medicine, Addenbrookes Hospital, University of Cambridge, Cambridge, UK CLARISSA READ • Institute of Virology, Ulm University Medical Center, Ulm, Germany; Central Facility for Electron Microscopy, Ulm University, Ulm, Germany MYRIAM SCHERER • Institute of Virology, Ulm University Medical Center, Ulm, Germany JOHN SINCLAIR • Department of Medicine, Addenbrookes Hospital, University of Cambridge, Cambridge, UK CHRISTIAN SINZGER • Institute of Virology, Ulm University Medical Center, Ulm, Germany REBECCA L. SKALSKY • Vaccine and Gene Therapy Institute, Oregon Health and Sciences University, Beaverton, OR, USA

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JULIET V. SPENCER • Department of Biology, Texas Woman’s University, Denton, TX, USA THOMAS STAMMINGER • Institute of Virology, Ulm University Medical Center, Ulm, Germany ANNE-CHARLOTTE STILP • Institute of Virology, Ulm University Medical Center, Ulm, Germany DANIEL N. STREBLOW • The Vaccine and Gene Therapy Institute, Oregon Health and Science University, Beaverton, OR, USA HILLARY M. STRUTHERS • Vaccine and Gene Therapy Institute, Oregon Health and Sciences University, Beaverton, OR, USA JELENA TOMAC • Department for Histology and Embryology and Center for Proteomics, Faculty of Medicine, University of Rijeka, Rijeka, Croatia JENS VON EINEM • Institute of Virology, Ulm University Medical Center, Ulm, Germany PAUL WALTHER • Central Facility for Electron Microscopy, Ulm University, Ulm, Germany MARK WILLS • Department of Medicine, Addenbrookes Hospital, University of Cambridge, Cambridge, UK VIVIAN P. YOUNG • Department of Biology, Texas Woman’s University, Denton, TX, USA ANDREW D. YUROCHKO • Department of Microbiology and Immunology, Louisiana State University Health Sciences Center—Shreveport, Shreveport, LA, USA

Chapter 1 Overview of Human Cytomegalovirus Pathogenesis Heather L. Fulkerson, Maciej T. Nogalski, Donna Collins-McMillen, and Andrew D. Yurochko Abstract Human cytomegalovirus (HCMV) is a betaherpesvirus with a global seroprevalence of 60–90%. HCMV is the leading cause of congenital infections and poses a great health risk to immunocompromised individuals. Although HCMV infection is typically asymptomatic in the immunocompetent population, infection can result in mononucleosis and has also been associated with the development of certain cancers, as well as chronic inflammatory diseases such as various cardiovascular diseases. In immunocompromised patients, including AIDS patients, transplant recipients, and developing fetuses, HCMV infection is associated with increased rates of morbidity and mortality. Currently there is no vaccine for HCMV and there is a need for new pharmacological treatments. Ongoing research seeks to further define the complex aspects of HCMV pathogenesis, which could potentially lead to the generation of new therapeutics to mitigate the disease states associated with HCMV infection. The following chapter reviews the advancements in our understanding of HCMV pathogenesis in the immunocompetent and immunocompromised hosts. Key words Human Cytomegalovirus, Viral Pathogenesis, Immunocompetent, Immunocompromised, Vascular Disease, Oncogenesis, Congenital Infection, Transplant Recipients, AIDS Patients

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Introduction Human cytomegalovirus (HCMV) is a prevalent infectious agent that globally affects the health of the human population. In a simple sense, HCMV pathogenesis can be broken down to that observed in immunocompetent hosts and to that observed in immunocompromised hosts [1, 2]. HCMV pathogenesis in immunologically normal/healthy individuals is usually considered less severe when compared to the morbidity and mortality seen in immunocompromised individuals. Severe complications such as pneumonia, retinitis, hepatitis, encephalitis, and disseminated HCMV disease with multiorgan involvement are extremely rare in immunologically healthy people [1–4]. The majority of HCMV infections in the immunocompetent are asymptomatic [1]; however, primary infection can result in a mononucleosis-like syndrome [3]. In addition,

Andrew D. Yurochko (ed.), Human Cytomegaloviruses: Methods and Protocols, Methods in Molecular Biology, vol. 2244, https://doi.org/10.1007/978-1-0716-1111-1_1, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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data from both clinical and experimental studies now define a potentially strong role for HCMV infection in the development and/or severity of inflammatory cardiovascular diseases and to the development of certain types of cancers [2, 5–12]. In immunocompromised individuals, HCMV infection can result in severe disease [1, 2]. For example, in patients undergoing immunosuppressive therapies, such as solid organ and stem cell transplant recipients and cancer patients undergoing chemotherapy, and in patients with acquired immunodeficiency syndrome (AIDS), HCMV infection is of significant clinical concern. In addition, HCMV is one of the leading infectious agents causing congenital infection [13–15]. Thus, individuals with suppressed or underdeveloped immune systems are prone to severe disease following primary HCMV infection or reactivation of latent virus. This review chapter focuses briefly on some of the consequences of HCMV infection in the immunocompetent and the immunocompromised.

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Pathogenesis in Immunocompetent Hosts

2.1 Infectious Mononucleosis

The most common clinical manifestation of HCMV infection in the immunocompetent host is a self-limiting febrile illness that resembles the infectious mononucleosis resulting from infection with Epstein–Barr Virus (EBV) [3, 16]. The clinical picture of HCMV mononucleosis is typically indistinguishable from EBV mononucleosis, with the exception that pharyngitis, adenopathy, and splenomegaly occur less commonly with HCMV infection [1, 3, 16, 17]. In addition, HCMV mononucleosis is a heterophile-negative mononucleosis accounting for approximately 10% of mononucleosis diagnoses [1, 3, 16]. Fever, malaise, myalgia, headache, and fatigue are the most commonly experienced signs and symptoms of the disease [1, 3, 16–18]. A smaller number of patients can present with splenomegaly, hepatomegaly, adenopathy, and a rash [1, 3, 16, 17]. Laboratory tests commonly reveal lymphocytosis, activated or atypical lymphocytes, and abnormal liver function [1, 3, 19].

2.2 Viral Role in Vascular Disease

Mounting evidence suggests that HCMV infection is an etiological/coetiologic agent in the development and/or severity of inflammatory cardiovascular diseases [7, 8, 10, 11]. Beginning in the 1980s, several studies established a potential link between HCMV infection and the development of atherosclerosis [20– 23]. A prospective study conducted from 1987 to 1992 examined medical records, including anti-HCMV antibody titers taken more than a decade earlier from patients with carotid intimal–medial thickening (IMT) and their age- and gender-matched controls [23]. Investigators observed higher anti-HCMV antibody titers in

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those patients with IMT than in the control group, consistent with HCMV infection being a possible factor/cofactor in the development of disease [23]. HCMV nucleic acid and antigen have also been detected in tissue sections taken from atherosclerotic lesions [10, 22, 24, 25]. A link has also been established between HCMV infection and coronary restenosis [26, 27]. For example, HCMV seropositivity and higher HCMV IgG titers were shown to correlate with the incidence of restenosis following surgical intervention to treat atherosclerosis [26]. Studies have also linked HCMV infection to transplant vascular sclerosis [8, 28, 29]; and it has been reported that patients with coronary artery disease have higher blood serum levels of C-reactive protein, a marker of the inflammatory response, that correlates with HCMV seropositivity, suggesting that inflammation resulting from HCMV infection may serve as a risk factor for vascular disease [30]. Given that HCMV infection has been linked to the development of cardiovascular disease, several studies have now demonstrated a correlation between HCMV seropositivity and increased risk of cardiovascular mortality among the aging population [31–33]. HCMV infects endothelial cells, smooth muscle cells, and monocytes in vivo and in vitro, all of which, when aberrantly activated, can directly contribute to the development of cardiovascular disease [34–40]. Additionally, studies have shown that HCMV alters host cell metabolism, specifically the lipidome, which has previously been linked to atherosclerotic progression [41–43]. HCMV infection can also lead to an overall increase in cholesterol and triglycerides within the host and to an accumulation of these lipids in arterial walls [10]. In fibroblasts, US28 has been shown to disrupt host lipid rafts, ultimately resulting in increased cholesterol efflux [43]. HCMV infection of vascular smooth muscle cells results in an increase of cellular cholesterol through increased expression of HMG-CoA synthase and reductase [41]. It has also been demonstrated that monocytes can migrate into arterial tissue, differentiate into macrophages, and engulf oxidized low-density lipoproteins, becoming the foamy macrophages that accumulate in arteries and atherosclerotic lesions [35, 44– 46]. Because HCMV infection of monocytes has been shown to alter many of these processes [34, 47, 48], it is plausible to hypothesize that viral infection of monocytes and the resulting biological changes in these cells contributes to atherosclerotic disease. The same idea holds true for HCMV infection of vascular endothelial cells and smooth muscle cells [8, 36, 49–51]. Animal studies in various rodent models have demonstrated (essentially confirming Koch’s postulates) that CMV infection is linked to endothelial damage, monocyte infiltration, foam cell accumulation, and vascular disease [40, 52–56]. Thus, both clinical and experimental studies have provided strong evidence that CMV infection promotes vascular disease at almost every stage of the disease process

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(enhancement of the proinflammatory response, vascular injury, increased migration and proliferation of smooth muscle cells, migration of monocytes into lesions, formation of foamy macrophages, plaque development and other biological changes consistent with a role in vascular disease) [10, 11]. 2.3 Possible Viral Role in Oncogenesis

The potential relationship between HCMV infection and cancer has been debated for decades, and during that time, HCMV has been argued to be associated with a variety of malignancies [6, 9– 12]. Seroepidemiological studies, as well as detection of viral nucleic acids and/or antigens in malignant tissues, are suggestive of an etiological role for HCMV in the development of various types of cancers [6, 57–61]. However, it remains unclear whether HCMV itself is an oncogenic virus in the classical sense [6]. HCMV does possess many of the molecular hallmarks of the small DNA tumor viruses (altering p53 and Rb function, inducing cellular proliferation, enhancing cellular survival, etc.), suggesting at least from a molecular standpoint that it could be an etiologic agent in tumor development and/or progression [62–71]. Studies have also shown that, like classical oncogenic viruses, HCMV has the ability to transform cells [72–74]. For example, it was demonstrated that HCMV US28 can transform NIH-3T3 cells and promote tumor growth in nude mice [72], and that the HCMV strain DB can transform human mammary epithelial cells in vitro and the injection of these cells into NSG mice promotes tumor growth [73]. It has also been suggested that HCMV infection might produce an environment of a “smoldering inflammation,” which in turn could mechanistically promote oncogenesis in a similar manner to that defined for the hepatitis viruses [75]. In keeping with this idea, it has been proposed that “oncomodulation” could describe HCMV’s effect on tumor cells; in that HCMV could infect tumor cells and enhance their malignancy, thereby promoting tumor progression without being an oncogenic virus per se [72, 76–81]. The body of evidence supporting a possible role for the virus in tumor growth continues to expand, with investigators focusing on both clinical and experimental aspects of HCMV cancer research. HCMV genome and antigen indicative of a persistent low level of replication have been detected in tumor cells, but not in the surrounding tissue, of a variety of malignancies including colorectal cancers [58], malignant gliomas [57, 61, 82, 83], prostate cancers [59], and breast cancers [60]. HCMV has also been proposed to be a coetiologic agent in the development of certain types of cancers [84]. A recent study aimed at establishing the clinical relevance of HCMV infection in malignant glioblastomas, grouped patients based on the level of HCMV-infected tumor cells and uncovered a relationship between the level of infection and the patient’s life expectancy: patients with low level HCMV infection outlived those with higher levels of HCMV infection [5, 85]. There are multiple

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molecular mechanisms, which are proposed to contribute to HCMV-induced oncomodulation. A recent review provides a more comprehensive discussion of data in support of and against the oncogenic potential of HCMV and the mechanisms by which viral proteins could alter the molecular and functional properties of HCMV-infected tumor cells, as well as the tumor microenvironment [86]. Experimental evidence suggests that HCMV alters the cell cycle and inhibits apoptosis in infected cancer cells, thereby promoting proliferation and survival of the cells [6]. HCMV infection also appears to influence invasion, migration, and endothelial adhesion of malignant cells, potentially contributing to metastatic complications in HCMV-infected patients [6, 79]. In addition, HCMV infection has been shown to promote angiogenesis, a process that is central to the initiation and progression of malignancies [6, 36, 80, 87]. Infection with HCMV also diminishes cancer cell immunogenicity and causes chromosomal abnormalities in infected cells [6, 88]. Nevertheless, the relationship between HCMV infection and cancer is unresolved and remains an important question in the area of HCMV pathogenesis. Additional research is needed to define if and how HCMV infection modulates and/or is an etiologic agent in tumor initiation and/or progression.

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Pathogenesis in Immunocompromised Hosts

3.1 Congenital Infection

HCMV is the leading cause of congenital viral infections in the USA [14, 15]. Worldwide, the rate of congenital HCMV infection averages at about 5–7 per 1000 live births, In some areas such as Latin America, Africa, and many Asian countries, the rate of congenital HCMV infection is higher at about 10–30 out of 1000 live births [89]. Congenital HCMV infection occurs when the virus crosses the placenta, thus allowing transmission of the virus from mother to baby. This process can occur following primary infection, in which a seronegative mother is infected by HCMV for the first time, or following a nonprimary infection—mothers seropositive prior to conception transmit virus to the fetus [1, 13]. Nonprimary infection is a term that includes seropositive mothers that are infected by new, serologically distinct strains of HCMV as well as seropositive mothers in which latent virus reactivates [89, 90]. It is estimated that about 13% of seronegative mothers develop a primary infection during pregnancy, and the transmission rate of virus to the fetus is approximately 30% [13, 91–95]. Previously, it had been argued that the transmission rate of virus to fetus in mothers with nonprimary infections is about 1%. However, newer observations suggest that this argument is not sufficiently supported by the available data [90]. These studies seeking to quantify the viral transmission rate to the fetus during nonprimary infections may have suffered from limitations such as potential patient enrollment bias as well as methodological inadequacies in accurately

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identifying and classifying mothers as having nonprimary infections [90]. Thus, the rate of viral transmission to the fetus during nonprimary infection of the mother remains unclear and requires further rigorous investigation. Congenitally infected neonates are categorized based on symptomology at birth: symptomatic (~10%) and asymptomatic (~90%). Of the symptomatic infants, 30–40% will suffer severe symptoms associated with congenital HCMV infection [15, 89, 90]. Congenitally infected babies can have a multisymptomatic disease affecting many organ systems and ranging from pneumonia, through gastrointestinal and retinal diseases, to central nervous system (CNS) diseases [15, 96–99]. Congenital HCMV may also manifest as a hematologic disease with for example, thrombocytopenia and hemolytic anemia being commonly observed abnormalities [1]. Additionally, congenital HCMV infection may result in jaundice, hepatitis, hepatosplenomegaly, petechiae, and thrombocytopenia in the infected neonate [15, 99]. Although symptoms generally subside a few weeks after birth, the disease can be severe for some newborns, even leading to neonatal death in a small percentage of cases [13, 93, 96, 100]. Greater than 50% of cases of symptomatic congenital HCMV infection present with abnormalities in the CNS [14, 15, 93, 101, 102]. These abnormalities often cause a range of neurological symptoms, such as mental retardation, diminished motor skills, sensorineural hearing loss, and vision loss [1, 13, 15, 100, 102, 103]. CNS sequelae are also present in congenitally infected newborns that were asymptomatic at birth [13, 15, 101, 104, 105]. Newer reports suggest that about 10% of asymptomatic infants are at risk of developing sensorineural hearing loss [15, 89, 106, 107]. Overall, 8–10% of all congenitally infected infants suffer from sensorineural hearing loss [90]. Based on the prevalence and severity of disease, congenital HCMV infection is considered to be a leading cause of CNS damage in children [1, 13, 101]. Longstanding dogma for HCMV congenital infection has been that symptomatic congenital HCMV infection presents with greater disease severity and occurs more often if the mother was exposed to a primary infection during pregnancy [101, 108]. Many have long argued that preconceptual maternal immunity to HCMV decreases viral transmission to the fetus and decreases the severity of disease outcome in the fetus. However, more recent findings demonstrate that nonprimary maternal infections can lead to symptomatic congenital HCMV infection of the fetus [15, 90, 109] and that similar rates of neurodevelopmental sequelae exists between babies infected via primary or nonprimary maternal infections [15, 90, 110]. How maternal immunity affects viral transmission and severity of disease in congenitally infected neonates remains unclear. Investigators are currently working toward further understanding this important facet of congenital HCMV infection with the hope of developing an effective vaccine or other preventative measures.

HCMV Pathogenesis

3.2 Infection of Infants

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While an infant’s immune system is developing, it is programmed to be suppressive to cytotoxicity. The immune system does not finish maturing completely until teen age [111]. Thus, infants have a lower ability to mount effective immune responses to pathogens. Nevertheless, maternal antibodies transferred through the placenta and antibodies transferred through breast milk generally serve to protect newborns from infections during their early life. In regards to HCMV infection, newborns can be infected in utero (as discussed above), via intrapartum transmission, or via postnatal infection [102]. It is estimated that HCMV is shed in around 10–15% of women from the vagina or cervix [1, 112, 113] and that the rate of intrapartum transmission from virus shedding mothers is reported to be around 50% [114]. Human breast milk is also considered to be a common means of postnatal mother-toinfant transmission of viruses [115]. The rate of newborn HCMV infection via this route strongly correlates with the length of time that the baby is nursed. It has been thought that approximately 40–60% of breast-fed infants will be infected by HCMV at the end of their first year of life if fed by seropositive mothers [102, 116]. This high rate of transmission is consistent with studies that have documented that up to ~95% of tested milk samples are positive for HCMV DNA [102, 115, 117, 118]. Although full-term newborns usually do not present with significant disease if infected early in life, there have been reported cases of hepatomegaly, elevated hepatic enzymes and inflammation of lung tissue [119–121]. The risk of complications, however, rises in preterm infants. For example, there is an association between HCMV disease manifestation and premature infants of seronegative mothers that received blood from seropositive donors. In these reported cases, symptoms suggested multiorgan dysfunction [122, 123]. In addition, it has been found that preterm neonates are infected at a higher rate than full-term infants if nursed by seropositive mothers [116, 118]. In preterm infants, the risks of complications associated with infection include thrombocytopenia, hepatosplenomegaly, distending bowels neutropenia, apnea and bradycardia, liver dysfunction, sepsis syndrome, and a mononucleosis-like disorder [115, 118, 124]. Some studies have linked early postnatal HCMV infection of preterm infants with negative long-term neurological/cognitive developmental outcomes, while others have found no such association [115]. Limiting preterm newborns to blood products from seronegative donors or by eliminating leukocytes from the transfused blood reduces the transfusion-acquired HCMV disease in these infants [123, 125, 126]. Although the health related issues resulting from HCMV infection via intrapartum or postnatal transmission are not usually associated with adverse outcomes in full-term babies, the role of infected infants in the epidemiology of HCMV spread is significant, as they can shed the virus for years, thereby enhancing viral

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transmission [1, 13, 94, 113]. A newer meta-analysis has shown that 32% of children attending day care facilities are HCMVinfected and there is a significant association between day care attendance and HCMV infection [127]. This point may be especially relevant to pregnant women who have older children in day care, as it increases their risk of exposure to the virus. 3.3 Infection of Immunocompromised Hosts

HCMV is considered to be one of the most common opportunistic pathogens seen in immunocompromised patients. These patients are at risk for viral-mediated disease as a result of a primary infection, reinfection (of an already seropositive host) and reactivation of latent virus. It has been documented that the stronger the suppression of the immune system, the greater the risk for HCMVmediated disease [1]. Allogeneic stem cell transplant patients and AIDS patients are characterized as having the most severe disease manifestations. HCMV infection and disease is also seen in solid organ transplant and cancer patients undergoing immunosuppressive therapy [1]. Clinical manifestations in these patients can range from a short febrile illness to multiple organ system involvement. Although the investigation of the impact HCMV infection has on immunocompromised patients is complex, studies performed on those patients have allowed a better understanding of HCMV infection, immune control of the virus and viral-mediated diseases. For example, these studies have provided evidence about the infection (or multiple infection) of patients with new strains of virus [128–132], the importance of humoral and cell mediated immunity in limiting HCMV infection [13, 133–136] and evidence that because of the seemingly unique properties of HCMV infection the development of an efficacious vaccine remains a complex and difficult task [13, 137–139]. Additionally, the investigation and generation of new antiviral drugs has been influenced by the need for better management of HCMV infection in immunocompromised patients [140, 141].

3.3.1 Infection of Transplant Recipients

Complications resulting from HCMV infection of transplant recipients significantly increase the overall cost and length of hospitalization for the recipient [142–145]. CMV disease in transplant patients is a term defined by the detection of virus within the patient alongside clinical signs and symptoms. Reactivation of latent virus within the allograft is the leading cause of CMV disease in solid organ transplant recipients, while reactivation of virus in the seropositive recipient is the leading cause of CMV disease in stem cell transplant patients; although, the recipients could potentially develop a primary infection through contact with a person who is actively shedding virus [146, 147]. Typically, the virus is detected in the blood or excretions at around 4–8 weeks posttransplantation and CMV disease, consequently, manifests within the first 3 months posttransplantation (early onset CMV disease). A lower percentage

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of patients develop CMV disease after the first 3 months posttransplantation (late onset CMV disease) [1, 146–148]. Infection of transplant recipients with HCMV results in both direct and indirect clinical manifestations. Direct clinical outcomes of HCMV infection arise from viral replication, dissemination, and invasion of peripheral organs [147, 149]. These direct clinical manifestations can be sorted into two groups: (1) CMV syndrome in solid organ transplant recipients and (2) tissue-invasive CMV disease. CMV syndrome includes general flu-like symptoms, malaise, fever, thrombocytopenia, and leucopenia. Tissue-invasive CMV disease can involve multiple organs, especially the allograft, leading to hepatitis, pneumonitis, colitis, nephritis, and retinitis. The indirect effects of HCMV infection of transplant patients are thought to arise from the immunomodulatory capability of the virus and include the following: increased rate of bacterial and fungal infections, increased rate of cardiovascular disease, increased rate of allograft rejection in solid organ transplant patients, increased rate of graft-versus-host disease in stem cell transplant patients, as well as the overall increase in mortality in transplant patients [146, 147, 149]. Over the past 2–3 decades, researchers have developed a better understanding of CMV disease in transplant recipients and have thus sought to find ways to prevent or mitigate the direct and indirect manifestations of HCMV infection in these vulnerable patients. To attempt to reduce the risk of CMV disease in transplant patients, hospitals now screen both the donor and recipient for HCMV. Donor/recipient serostatus has been linked to recipient risk of contracting HCMV infection post-transplantation. In solid organ transplant patients, the highest risk group consists of donor positive–recipient negative HCMV serostatus. In stem cell transplants, the highest risk group is that of recipients seropositive for HCMV [146, 147, 149]. There are currently two main preventative measures to combat CMV disease: (1) antiviral prophylaxis and (2) preemptive therapy (PET). Antiviral prophylaxis involves the administration of HCMV antivirals to the patient during times of high risk without testing to determine whether HCMV is present in the bloodstream. Standard PET protocol monitors the patient weekly for the presence of HCMV in the blood with the intended purpose of administering antivirals before the onset of CMV disease. Doctors choose between the preventative measures on a patient-to-patient basis after evaluating the overall risk of the patient to develop CMV disease posttransplantation [146, 147, 149–154]. By having these protocols in place, the rates of HCMV disease and infection-related deaths have been significantly reduced [155]. In fact, in allogenic stem cell transplant recipients the early onset of CMV disease has been reduced from 60% to about 5%. However, late onset CMV disease still remains a significant problem for these patients [146].

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The severity of complications caused by HCMV infection in transplant recipients has set the stage for the improved management of HCMV disease. In general, pretransplant donor and recipient screening, as well as post-transplant screening for the presence of HCMV are performed, along with the preemptive and prophylactic administration of antivirals. However, as mentioned above, late-onset HCMV disease remains a significant problem and is responsible for the high mortality seen in these patients [146, 153, 154, 156–159]. 3.3.2 Infection in AIDS Patients

4

HCMV infection is considered a leading opportunistic pathogen in AIDS patients and has been associated with progression of HIV infection [1, 13, 160–163]. The serological prevalence of HCMV is evident in nearly all adults and about half of the children seropositive for HIV infection [1]. It was estimated that approximately 40% of adults and about 10% of children with AIDS showed manifestations of HCMV disease before the introduction of highly active antiretroviral therapy (HAART) [1, 164, 165]. Common manifestations of HCMV disease in AIDS patients are retinitis, esophagitis and colitis; case reports have also documented encephalitis, neuropathy, polyradiculoneuritis, pneumonitis, gastritis, and liver dysfunction [165]. HIV has also been demonstrated to be a significant risk factor for the transmission of HCMV from mother to fetus in utero (congenital infection) [166]. Because of the use of HAART, incidence of each of these pathologies has significantly decreased in treated patients [13, 167–169]. Nevertheless, there is evidence that HCMV infection remains an independent predictor of morbidity and mortality in AIDS patients [162, 170–176]. HCMV infection has long been attributed to the progression of HIV infection and morbidity in these patients, although mechanistically it remains unclear how HCMV may affect the outcome of HIV-infected patients (outside of HCMV’s role as an opportunistic pathogen). Some potential examples for how HCMV may alter the course of infection include transactivation of the HIV promoter [160], changes in Fc receptor expression [161], altered immune activation [163], and increased T cell senescence [175–178]. Regardless of the mechanism for how HCMV may affect the outcome of HIV-infected patients, it seems that these two pathogens possess a partnership. Therefore, even in the day of HAART, HCMV remains an important pathogen in AIDS patients.

Conclusions As we expand our knowledge about HCMV infection and the associated pathologies, we gain a greater appreciation for the unique aspects of the virus, as well as the long-term human consequences associated with that infection. HCMV remains an

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important human pathogen due to (1) its high seroprevalence and widespread geographic dissemination in the human population and (2) because of the multitude of disease pathologies caused by or associated with infection. HCMV-mediated disease can loosely be grouped into the diseases observed in immunocompromised individuals and the diseases observed in immunocompetent individuals. In immunocompromised people, HCMV infection can cause severe disease and significantly affect a variety of organ systems. In healthy people, HCMV infection has generally been thought of as benign/ asymptomatic; however, with the association of viral infection with cardiovascular diseases and now potentially with several cancers, perhaps this idea needs to be revisited. Overall, HCMV is a complex pathogen with an interesting and diverse pathobiology. Additional studies into the basic aspects of HCMV biology, as well as the specific mechanisms that directly cause human disease, are needed. In addition, new pharmacological treatment options and the generation of an effective vaccine are still needed to counter the morbidity and mortality associated with viral infection.

Acknowledgments This work was supported by grants from the National Institutes of Health NIAID P01 AI127335, AI056077, P30GM110703, P20GM121288, P20GM121307, and a Malcolm Feist Predoctoral Fellowship. References 1. Mocarsk ES Jr et al (2013) Cytomegaloviruses. In: Knipe DM, Howley PM (eds) Fields Virology. Lippincott Williams & Wilkins, Philadelphia, pp 1960–2014 2. Britt W (2008) Manifestations of human cytomegalovirus infection: Proposed mechanisms of acute and chronic disease. In: Stinksi MF, Shenk T (eds) Human Cytomegaloviruses. Springer-Verlag, Berlin, pp 417–470 3. Bravender T (2010) Epstein-Barr virus, cytomegalovirus, and infectious mononucleosis. Adolesc Med State Art Rev 21(2):251–264 4. Eddleston M et al (1997) Severe cytomegalovirus infection in immunocompetent patients. Clin Infect Dis 24(1):52–56 5. So¨derberg-Naucle´r C (2008) HCMV microinfections in inflammatory diseases and cancer. J Clin Virol 41(3):218–223 6. Michaelis M, Doerr HW, Cinatl J (2009) The story of human cytomegalovirus and cancer: increasing evidence and open questions. Neoplasia 11(1):1–9

7. Caposio P, Orloff SL, Streblow DN (2011) The role of cytomegalovirus in angiogenesis. Virus Res 157:204–211 8. Streblow DN et al (2008) Mechanisms of cytomegalovirus-accelerated vascular disease: induction of paracrine factors that promote angiogenesis and wound healing. Curr Top Microbiol Immunol 325:397–415 9. Bai B et al (2016) Human cytomegalovirus infection and colorectal cancer risk: a metaanalysis. Oncotarget 7(47):76735–76742 10. Du Y, Zhang G, Liu Z (2018) Human cytomegalovirus infection and coronary heart disease: a systematic review. Virol J 15(1):31 11. Lebedeva AM et al (2018) Cytomegalovirus infection in cardiovascular diseases. Biochemistry (Mosc) 83(12):1437–1447 12. McFaline-Figueroa JR, Wen PY (2017) The viral connection to glioblastoma. Curr Infect Dis Rep 19(5):5

12

Heather L. Fulkerson et al.

13. Manicklal S et al (2013) The "silent" global burden of congenital cytomegalovirus. Clin Microbiol Rev 26:86–102 14. Ross SA, Boppana SB (2005) Congenital cytomegalovirus infection: outcome and diagnosis. Semin Pediatr Infect Dis 16:44–49 15. Boppana SB, Ross SA, Fowler KB (2013) Congenital cytomegalovirus infection: clinical outcome. Clin Infect Dis 57(Suppl 4): S178–S181 16. Klemola E et al (1970) Infectiousmononucleosis-like disease with negative heterophil agglutination test. Clinical features in relation to Epstein-Barr virus and cytomegalovirus antibodies. J Infect Dis 121 (6):608–614 17. Jordan MC et al (1973) Spontaneous cytomegalovirus mononucleosis. Clinical and laboratory observations in nine cases. Ann Intern Med 79(2):153–160 18. Nolan N et al (2017) Primary cytomegalovirus infection in immunocompetent adults in the United States—A case series. IDCases 10:123–126 19. Wreghitt TG et al (2003) Cytomegalovirus infection in immunocompetent patients. Clin Infect Dis 37(12):1603–1606 20. Adam E et al (1987) High levels of cytomegalovirus antibody in patients requiring vascular surgery for atherosclerosis. Lancet 2 (8554):291–293 21. Hendrix MG et al (1990) High prevalence of latently present cytomegalovirus in arterial walls of patients suffering from grade III atherosclerosis. Am J Pathol 136(1):23–28 22. Melnick JL et al (1983) Cytomegalovirus antigen within human arterial smooth muscle cells. Lancet 2:644–647 23. Nieto FJ et al (1996) Cohort study of cytomegalovirus infection as a risk factor for carotid intimal-medial thickening, a measure of subclinical atherosclerosis. Circulation 94 (5):922–927 24. Gyorkey F et al (1984) Herpesviridae in the endothelial and smooth muscle cells of the proximal aorta in atherosclerotic patients. Exp Mol Pathol 40:328–339 25. Beyaz MO et al (2019) Evaluation of the relationship between plaque formation leading to symptomatic carotid artery stenosis and cytomegalovirus by investigating the virus DNA. Arch Med Sci Atheroscler Dis 4: e19–e24 26. Zhou YF et al (1996) Association between prior cytomegalovirus infection and the risk of restenosis after coronary atherectomy. N Engl J Med 335:624–630

27. Speir E et al (1994) Potential role of human cytomegalovirus and p53 interaction in coronary restenosis. Science 265(5170):391–394 28. Hussain T et al (2007) Positive pretransplantation cytomegalovirus serology is a risk factor for cardiac allograft vasculopathy in children. Circulation 115:1798–1805 29. So¨derberg-Naucle´r C, Emery VC (2001) Viral infections and their impact on chronic renal allograft dysfunction. Transpantation 71:SS24–SS30 30. Zhu J et al (1999) Cytomegalovirus in the pathogenesis of atherosclerosis: the role of inflammation as reflected by elevated C-reactive protein levels. J Am Coll Cardiol 34(6):1738–1743 31. Savva GM et al (2013) Cytomegalovirus infection is associated with increased mortality in the older population. Aging Cell 12 (3):381–387 32. Spyridopoulos I et al (2016) CMV seropositivity and T-cell senescence predict increased cardiovascular mortality in octogenarians: results from the Newcastle 85+ study. Aging Cell 15(2):389–392 33. Simanek AM et al (2011) Seropositivity to cytomegalovirus, inflammation, all-cause and cardiovascular disease-related mortality in the United States. PLoS One 6(2):e16103 34. Chan G et al (2012) Human cytomegalovirus induction of a unique signalsome during viral entry into monocytes mediates distinct functional changes: a strategy for viral dissemination. J Leukoc Biol 92:743–752 35. Hansson GK, Libby P (2006) The immune response in atherosclerosis: a double-edged sword. Nat Rev Immunol 6(7):508–519 36. Bentz GL, Yurochko AD (2008) HCMV infection of endothelial cells induces an angiogenic response through viral-binding to the epidermal growth factor receptor and the β1 and β3 integrins. Proc Natl Acad Sci U S A 105:5531–5536 37. Falk E (2006) Pathogenesis of atherosclerosis. J Am Coll Cardiol 47(8 Suppl):C7–C12 38. Lusis AJ (2000) Atherosclerosis. Nature 407 (6801):233–241 39. Streblow DN et al (1999) The human cytomegalovirus chemokine receptor US28 mediates vascular smooth muscle cell migration. Cell 99:511–520 40. Streblow DN, Orloff SL, Nelson JA (2001) Do pathogens accelerate atherosclerosis? J Nutr 131:2798–2804 41. Li L et al (2016) Lipid metabolism in vascular smooth muscle cells Infuenced by HCMV

HCMV Pathogenesis infection. Cell Physiol Biochem 39 (5):1804–1812 42. Kohno S et al (2018) Lipidomic insight into cardiovascular diseases. Biochem Biophys Res Commun 504(3):590–595 43. Low H et al (2016) Cytomegalovirus restructures lipid rafts via a US28/CDC42-mediated pathway, enhancing cholesterol efflux from host cells. Cell Rep 16(1):186–200 44. Jaipersad AS et al (2014) The role of monocytes in angiogenesis and atherosclerosis. J Am Coll Cardiol 63(1):1–11 45. Moroni F et al (2019) The role of monocytes and macrophages in human atherosclerosis, plaque Neoangiogenesis, and Atherothrombosis. Mediat Inflamm 2019:7434376 46. Hilgendorf I, Swirski FK, Robbins CS (2015) Monocyte fate in atherosclerosis. Arterioscler Thromb Vasc Biol 35(2):272–279 47. Chan G, Nogalski MT, Yurochko AD (2012) Human cytomegalovirus stimulates monocyte-to-macrophage differentiation via the temporal regulation of caspace 3. J Virol 92:10714–10723 48. Collins-McMillen D et al (2017) Human cytomegalovirus utilizes a nontraditional signal transducer and activator of transcription 1 activation cascade via signaling through epidermal growth factor receptor and integrins to efficiently promote the motility, differentiation, and polarization of infected monocytes. J Virol 91(24):e00622–e00617 49. Botto S et al (2011) IL-6 in human cytomegalovirus secretome promotes angiogenesis and survival of endothelial cells through the stimulation of survivin. Blood 117 (1):352–361 50. Zhou YF et al (1999) The immediate early gene products of human cytomegalovirus increase vascular smooth muscle cell migration, proliferation, and expression of PDGF beta-receptor. Biochem Biophys Res Commun 256(3):608–613 51. Khoretonenko MV et al (2010) Cytomegalovirus infection leads to microvascular dysfunction and exacerbates hypercholesterolemiainduced responses. Am J Pathol 177 (4):2134–2144 52. Streblow DN et al (2005) Rat cytomegalovirus-accelerated transplant vascular sclerosis is reduced with mutation of the chemokine-receptor R33. Am J Transplant 5:436–442 53. Orloff SL et al (2011) Cytomegalovirus latency promotes cardiac lymphoid neogenesis and accelerated allograft rejection in

13

CMV naı¨ve recipients. Am J Transplant 11:45–55 54. Streblow DN et al (2008) The role of angiogenic and wound repair factors during CMV-accelerated transplant vascular sclerosis in rat cardiac transplants. Am J Transplant 8:277–287 55. Gombos RB et al (2013) Vascular dysfunction in young, mid-aged and aged mice with latent cytomegalovirus infections. Am J Physiol Heart Circ Physiol 304:H183–H194 56. Tang-Feldman YJ et al (2013) Murine cytomegalovirus (MCMV) infection upregulates P38 MAP kinase in aortas of Apo E KO mice: a molecular mechanism for MCMVinduced acceleration of atherosclerosis. J Cardiovasc Transl Res 6:54–64 57. Bhattacharjee B, Renzette N, Kowalik TF (2012) Genetic analysis of cytomegalovirus in malignant gliomas. J Virol 86:6815–6824 58. Harkins L et al (2002) Specific localisation of human cytomegalovirus nucleic acids and proteins in human colorectal cancer. Lancet 360(9345):1557–1563 59. Samanta M et al (2003) High prevalence of human cytomegalovirus in prostatic intraepithelial neoplasia and prostatic carcinoma. J Urol 170(3):998–1002 60. Harkins LE et al (2010) Detection of human cytomegalovirus in normal and neoplastic breast epithelium. Herpesviridae 1(1):8 61. Cobbs CS et al (2002) Human cytomegalovirus infection and expression in human malignant glioma. Cancer Res 62:3347–3350 62. Hwang ES et al (2009) Human cytomegalovirus IE1-72 protein interacts with p53 and inhibits p53-dependent transactivation by a mechanism different from that of IE2-86 protein. J Virol 83:12388–12398 63. Shen Y, Zhu H, Shenk T (1997) Human cytomegalovirus IE1 and IE2 proteins are mutagenic and mediate "hit-and-run" oncogenic transformation in cooperation with the adenovirus E1A proteins. Proc Natl Acad Sci U S A 94:3341–3345 64. Xiaofei E et al (2011) An E2F1-mediated DNA damage response contributes to the replication of human cytomegalovirus. PLoS Pathog 7:e1001342 65. Castillo JP et al (2005) Human cytomegalovirus IE1-72 activates ataxia telangiectasia mutated kinase and a p53/p21-mediated growth arrest response. J Virol 79:11467–11475

14

Heather L. Fulkerson et al.

66. Hume AJ, Kalejta RF (2009) Regulation of the retinoblastoma proteins by the human herpesviruses. Cell Div 4:1 67. Iwahori S et al (2015) Molecular determinants for the inactivation of the retinoblastoma tumor suppressor by the viral cyclindependent kinase UL97. J Biol Chem 290 (32):19666–19680 68. Iwahori S, Kalejta RF (2017) Phosphorylation of transcriptional regulators in the retinoblastoma protein pathway by UL97, the viral cyclin-dependent kinase encoded by human cytomegalovirus. Virology 512:95–103 69. Iwahori S et al (2017) Human cytomegalovirus-encoded viral cyclindependent kinase (v-CDK) UL97 phosphorylates and inactivates the retinoblastoma protein-related p107 and p130 proteins. J Biol Chem 292(16):6583–6599 70. Spector DH (2015) Human cytomegalovirus riding the cell cycle. Med Microbiol Immunol 204(3):409–419 71. Steingruber M et al (2019) Cyclins B1, T1, and H differ in their molecular mode of interaction with cytomegalovirus protein kinase pUL97. J Biol Chem 294(15):6188–6203 72. Maussang D et al (2006) Human cytomegalovirus-encoded chemokine receptor US28 promotes tumorigenesis. Proc Natl Acad Sci U S A 103(35):13068–13073 73. Kumar A et al (2018) The human cytomegalovirus strain DB activates oncogenic pathways in mammary epithelial cells. EBioMedicine 30:167–183 74. Moussawi FA et al (2018) The transcriptome of human mammary epithelial cells infected with the HCMV-DB strain displays oncogenic traits. Sci Rep 8(1):12574 75. Soroceanu L, Cobbs CS (2011) Is HCMV a tumor promoter? Virus Res 157(2):193–203 76. Cinatl J Jr et al (1996) Modulatory effects of human cytomegalovirus infection on malignant properties of cancer cells. Intervirology 39(4):259–269 77. Barami K (2010) Oncomodulatory mechanisms of human cytomegalovirus in gliomas. J Clin Neurosci 17:819–823 78. So¨derberg-Naucle´r C, Johnsen JI (2012) Cytomegalovirus infection in brain tumors: a potential new target for therapy? Onco Targets Ther 1:739–740 79. Yang Z et al (2019) Latent cytomegalovirus infection in female mice increases breast cancer metastasis. Cancers (Basel) 11(4):447

80. Krenzlin H et al (2019) Cytomegalovirus promotes murine glioblastoma growth via pericyte recruitment and angiogenesis. J Clin Invest 130:1671–1683 81. Heukers R et al (2018) The constitutive activity of the virally encoded chemokine receptor US28 accelerates glioblastoma growth. Oncogene 37(30):4110–4121 82. Mitchell DA et al (2008) Sensitive detection of human cytomegalovirus in tumors and peripheral blood of patients diagnosed with glioblastoma. Neuro-Oncology 10(1):10–18 83. Scheurer ME et al (2008) Detection of human cytomegalovirus in different histological types of gliomas. Acta Neuropathol 116 (1):79–86 84. Shen CY et al (1993) High rate of concurrent genital infections with human cytomegalovirus and human papillomaviruses in cervical cancer patients. J Infect Dis 168(2):449–452 85. Rahbar A et al (2012) Low levels of human cytomegalovirus infection in glioblastoma multiforme associates with patient survival; -a case-control study. Herpesviridae 3:3 86. Herbein G (2018) The human cytomegalovirus, from oncomodulation to oncogenesis. Viruses 10(8):408 87. Dumortier J et al (2008) Human cytomegalovirus secretome contains factors that induce angiogenesis and wound healing. J Virol 82:6524–6535 88. Fortunato EA, Spector DH (2003) Viral induction of site-specific chromosome damage. Rev Med Virol 13:21–37 89. Fowler KB, Boppana SB (2018) Congenital cytomegalovirus infection. Semin Perinatol 42(3):149–154 90. Britt WJ (2018) Maternal immunity and the natural history of congenital human cytomegalovirus infection. Viruses 10(8):405 91. Adler SP (2005) Congenital cytomegalovirus screening. Pediatr Infect Dis J 24 (12):1105–1106 92. Adler SP (2011) Screening for cytomegalovirus during pregnancy. Infect Dis Obstet Gynecol 2011:1–9 93. Nigro G, Adler SP (2011) Cytomegalovirus infections during pregnancy. Curr Opin Obstet Gynecol 23:123–128 94. Marshall BC, Adler SP (2009) The frequency of pregnancy and exposure to cytomegalovirus infections among women with a young child in day care. Am J Obstet Gynecol 200:163.e1–163.e5

HCMV Pathogenesis 95. Mussi-Pinhata MM et al (2018) Seroconversion for cytomegalovirus infection during pregnancy and fetal infection in a highly seropositive population: "the BraCHS study". J Infect Dis 218(8):1200–1204 96. Vancikova Z, Dvorak P (2001) Cytomegalovirus infection in immunocompetent and immunocompromised individuals—a review. Curr Drug Targets Immune Endocr Metabol Disord 1(2):179–187 97. Mussi-Pinhata MM et al (2009) Birth prevalence and natural history of congenital cytomegalovirus infection in a highly seroimmune population. Clin Infect Dis 49:522–528 98. Naing ZW et al (2016) Congenital cytomegalovirus infection in pregnancy: a review of prevalence, clinical features, diagnosis and prevention. Aust N Z J Obstet Gynaecol 56 (1):9–18 99. Dreher AM et al (2014) Spectrum of disease and outcome in children with symptomatic congenital cytomegalovirus infection. J Pediatr 164(4):855–859 100. Boppana SB et al (1992) Symptomatic congenital cytomegalovirus infection: neonatal morbidity and mortality. Pediatr Infect Dis J 11(2):93–99 101. Fowler KB et al (1992) The outcome of congenital cytomegalovirus infection in relation to maternal antibody status. N Engl J Med 326(10):663–667 102. Nassetta L, Kimberlin D, Whitley R (2009) Treatment of congenital cytomegalovirus infection: implications for future therapeutic strategies. J Antimicrob Chemother 63:862–867 103. Bale JF Jr, Blackman JA, Sato Y (1990) Outcome in children with symptomatic congenital cytomegalovirus infection. J Child Neurol 5(2):131–136 104. Williamson WD et al (1990) Asymptomatic congenital cytomegalovirus infection. Audiologic, neuroradiologic, and neurodevelopmental abnormalities during the first year. Am J Dis Child 144(12):1365–1368 105. Saigal S et al (1982) The outcome in children with congenital cytomegalovirus infection. A longitudinal follow-up study. Am J Dis Child 136(10):896–901 106. Yamamoto AY et al (2019) Contribution of congenital cytomegalovirus (cCMV) to permanent hearing loss in a highly seropositive population: "the BraCHS study". Clin Infect Dis 70(7):1379–1384 107. Boppana SB, Fowler KB (2017) Insight into long-term neurodevelopmental outcomes in

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asymptomatic congenital cmv infection. Pediatrics 140(5):e20172526 108. Stagno S et al (1986) Primary cytomegalovirus infection in pregnancy. Incidence, transmission to fetus, and clinical outcome. JAMA 256(14):1904–1908 109. Britt W (2015) Controversies in the natural history of congenital human cytomegalovirus infection: the paradox of infection and disease in offspring of women with immunity prior to pregnancy. Med Microbiol Immunol 204 (3):263–271 110. Townsend CL et al (2013) Long-term outcomes of congenital cytomegalovirus infection in Sweden and the United Kingdom. Clin Infect Dis 56(9):1232–1239 111. Ygberg S, Nilsson A (2012) The developing immune system—from foetus to toddler. Acta Paediatr 101(2):120–127 112. Stagno S et al (1982) Maternal cytomegalovirus infection and perinatal transmission. Clin Obstet Gynecol 25(3):563–576 113. Cannon MJ, Hyde TB, Schmid DS (2011) Review of cytomegalovirus shedding in bodily fluids and relevance to congenital cytomegalovirus infection. Rev Med Virol 21:240–255 114. Reynolds DW et al (1973) Maternal cytomegalovirus excretion and perinatal infection. N Engl J Med 289(1):1–5 115. Hamprecht K, Goelz R (2017) Postnatal cytomegalovirus infection through human Milk in preterm infants: transmission, clinical presentation, and prevention. Clin Perinatol 44(1):121–130 116. Dworsky M et al (1983) Cytomegalovirus infection of breast milk and transmission in infancy. Pediatrics 72(3):295–299 117. Jim WT et al (2004) Transmission of cytomegalovirus from mothers to preterm infants by breast milk. Pediatr Infect Dis J 23 (9):848–851 118. Vochem M et al (1998) Transmission of cytomegalovirus to preterm infants through breast milk. Pediatr Infect Dis J 17(1):53–58 119. Chiba S et al (1975) Primary cytomegalovirus infection and liver involvement in early infancy. Tohoku J Exp Med 117(2):143–151 120. Stagno S et al (1981) Infant pneumonitis associated with cytomegalovirus, chlamydia, pneumocystis, and Ureaplasma: a prospective study. Pediatrics 68(3):322–329 121. Kumar ML et al (1984) Postnatally acquired cytomegalovirus infections in infants of CMV-excreting mothers. J Pediatr 104 (5):669–673

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Heather L. Fulkerson et al.

122. Ballard RA et al (1979) Acquired cytomegalovirus infection in preterm infants. Am J Dis Child 133(5):482–485 123. Yeager AS et al (1981) Prevention of transfusion-acquired cytomegalovirus infections in newborn infants. J Pediatr 98 (2):281–287 124. Hamprecht K et al (2001) Epidemiology of transmission of cytomegalovirus from mother to preterm infant by breastfeeding. Lancet 357(9255):513–518 125. Eisenfeld L et al (1992) Prevention of transfusion-associated cytomegalovirus infection in neonatal patients by the removal of white cells from blood. Transfusion 32 (3):205–209 126. Gilbert GL et al (1989) Prevention of transfusion-acquired cytomegalovirus infection in infants by blood filtration to remove leucocytes. Neonatal Cytomegalovirus Infection Study Group. Lancet 1 (8649):1228–1231 127. Zheng QY et al (2019) Cytomegalovirus infection in day care centres: a systematic review and meta-analysis of prevalence of infection in children. Rev Med Virol 29(1): e2011 128. Chou SW (1986) Acquisition of donor strains of cytomegalovirus by renal-transplant recipients. N Engl J Med 314(22):1418–1423 129. Drew WL et al (1984) Multiple infections by cytomegalovirus in patients with acquired immunodeficiency syndrome: documentation by southern blot hybridization. J Infect Dis 150(6):952–953 130. Grundy JE et al (1988) Symptomatic cytomegalovirus infection in seropositive kidney recipients: reinfection with donor virus rather than reactivation of recipient virus. Lancet 2 (8603):132–135 131. Vinuesa V et al (2017) The impact of virus population diversity on the dynamics of cytomegalovirus DNAemia in allogeneic stem cell transplant recipients. J Gen Virol 98 (10):2530–2542 132. Cudini J et al (2019) Human cytomegalovirus haplotype reconstruction reveals high diversity due to superinfection and evidence of within-host recombination. Proc Natl Acad Sci U S A 116(12):5693–5698 133. Snydman DR et al (1987) Use of cytomegalovirus immune globulin to prevent cytomegalovirus disease in renal-transplant recipients. N Engl J Med 317(17):1049–1054 134. Walter EA et al (1995) Reconstitution of cellular immunity against cytomegalovirus in

recipients of allogeneic bone marrow by transfer of T-cell clones from the donor. N Engl J Med 333(16):1038–1044 135. Sissons JG, Wills MR (2015) How understanding immunology contributes to managing CMV disease in immunosuppressed patients: now and in future. Med Microbiol Immunol 204(3):307–316 136. Klenerman P, Oxenius A (2016) T cell responses to cytomegalovirus. Nat Rev Immunol 16(6):367–377 137. Plotkin SA et al (1984) Towne-vaccineinduced prevention of cytomegalovirus disease after renal transplants. Lancet 1 (8376):528–530 138. Plotkin SA, Boppana SB (2018) Vaccination against the human cytomegalovirus. Vaccine 37(50):7437–7442 139. McCormick AL, Mocarski ES (2015) The immunological underpinnings of vaccinations to prevent cytomegalovirus disease. Cell Mol Immunol 12(2):170–179 140. Erice A et al (1997) Antiviral susceptibilities and analysis of UL97 and DNA polymerase sequences of clinical cytomegalovirus isolates from immunocompromised. patients. J Infect Dis 175(5):1087–1092 141. Griffiths P (2019) New vaccines and antiviral drugs for cytomegalovirus. J Clin Virol 116:58–61 142. Legendre CM et al (2000) Valaciclovir prophylaxis of cytomegalovirus infection and disease in renal transplantation: an economic evaluation. Transplantation 70 (10):1463–1468 143. Mauskopf JA et al (2000) Cost-effectiveness model of cytomegalovirus management strategies in renal transplantation. Comparing valaciclovir prophylaxis with current practice. PharmacoEconomics 18(3):239–251 144. Webb BJ et al (2018) The clinical and economic impact of cytomegalovirus infection in recipients of hematopoietic stem cell transplantation. Transpl Infect Dis 20(5):e12961 145. Restelli U et al (2019) Cost-effectiveness analysis of the use of letermovir for the prophylaxis of cytomegalovirus in adult cytomegalovirus seropositive recipients undergoing allogenic hematopoietic stem cell transplantation in Italy. Infect Drug Resist 12:1127–1138 146. Pande A, Dubberke ER (2019) Cytomegalovirus infections of the stem cell transplant recipient and hematologic malignancy patient. Infect Dis Clin N Am 33(2):485–500

HCMV Pathogenesis 147. Ramanan P, Razonable RR (2013) Cytomegalovirus infections in solid organ transplantation: a review. Infect Chemother 45 (3):260–271 148. Eid AJ, Razonable RR (2010) New developments in the management of cytomegalovirus infection after solid organ transplantation. Drugs 70:965–981 149. Hodowanec AC et al (2019) Treatment and prevention of CMV disease in transplant recipients: current knowledge and future perspectives. J Clin Pharmacol 59(6):784–798 150. Hebart H et al (1998) Management of cytomegalovirus infection after solid-organ or stem-cell transplantation. Current guidelines and future prospects. Drugs 55(1):59–72 151. Prentice HG, Kho P (1997) Clinical strategies for the management of cytomegalovirus infection and disease in allogeneic bone marrow transplant. Bone Marrow Transplant 19 (2):135–142 152. Boeckh M, Geballe AP (2011) Cytomegalovirus: pathogen, paradigm, and puzzle. J Clin Invest 121:1673–1680 153. Ljungman P, Hakki M, Boeckh M (2011) Cytomegalovirus in hematopoietic stem cell transplant recipients. Hematol Oncol Clin North Am 25:151–169 154. Kowalsky S, Arnon R, Posada R (2013) Prevention of cytomegalovirus following solid organ transplantation: a literature review. Pediatr Transplant 17:499–509 155. Ljungman P (1996) Cytomegalovirus infections in transplant patients. Scand J Infect Dis Suppl 100:59–63 156. Akalin E et al (2003) Cytomegalovirus disease in high-risk transplant recipients despite ganciclovir or valganciclovir prophylaxis. Am J Transplant 3(6):731–735 157. Limaye AP et al (2004) Late-onset cytomegalovirus disease in liver transplant recipients despite antiviral prophylaxis. Transplantation 78(9):1390–1396 158. Razonable RR et al (2001) Allograft rejection predicts the occurrence of late-onset cytomegalovirus (CMV) disease among CMV-mismatched solid organ transplant patients receiving prophylaxis with oral ganciclovir. J Infect Dis 184(11):1461–1464 159. Boeckh M et al (2003) Late cytomegalovirus disease and mortality in recipients of allogeneic hematopoietic stem cell transplants: importance of viral load and T-cell immunity. Blood 101(2):407–414 160. Barry PA et al (1990) Cytomegalovirus activates transcription directed by the long

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terminal repeat of human immunodeficiency virus type 1. J Virol 64(6):2932–2940 161. McKeating JA, Griffiths PD, Weiss RA (1990) HIV susceptibility conferred to human fibroblasts by cytomegalovirus-induced Fc receptor. Nature 343(6259):659–661 162. Griffiths PD (2006) CMV as a cofactor enhancing progression of AIDS. J Clin Virol 35:489–492 163. Ostrowski MA et al (1998) Effect of immune activation on the dynamics of human immunodeficiency virus replication and on the distribution of viral quasispecies. J Virol 72 (10):7772–7784 164. Gallant JE et al (1992) Incidence and natural history of cytomegalovirus disease in patients with advanced human immunodeficiency virus disease treated with zidovudine. The Zidovudine Epidemiology Study Group. J Infect Dis 166(6):1223–1227 165. Cheung TW, Teich SA (1999) Cytomegalovirus infection in patients with HIV infection. Mt Sinai J Med 66(2):113–124 166. Adachi K et al (2018) Congenital cytomegalovirus and HIV perinatal transmission. Pediatr Infect Dis J 37(10):1016–1021 167. Deayton J et al (1999) Loss of cytomegalovirus (CMV) viraemia following highly active antiretroviral therapy in the absence of specific anti-CMV therapy. AIDS 13(10):1203–1206 168. O’Sullivan CE et al (1999) Decrease of cytomegalovirus replication in human immunodeficiency virus infected-patients after treatment with highly active antiretroviral therapy. J Infect Dis 180(3):847–849 169. Jacobson MA et al (2001) Cytomegalovirus (CMV)-specific CD4+ T lymphocyte immune function in long-term survivors of AIDSrelated CMV end-organ disease who are receiving potent antiretroviral therapy. J Infect Dis 183:1399–1404 170. Spector SA et al (1999) Cytomegalovirus (CMV) DNA load is an independent predictor of CMV disease and survival in advanced AIDS. J Virol 73:7027–7030 171. Deayton JR et al (2004) Importance of cytomegalovirus viraemia in risk of disease progression and death in HIV-infected patients receiving highly active antiretroviral therapy. Lancet 363:2116–2121 172. Hodowanec AC et al (2019) Increased CMV IgG antibody titer is associated with non-AIDS events among virologically suppressed HIV-positive persons. Pathog Immun 4(1):66–78

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173. Zicari S et al (2019) Immune Activation, Inflammation, and non-AIDS Co-morbidities in HIV-infected patients under long-term ART. Viruses 11(3):200 174. Maidji E et al (2017) Replication of CMV in the gut of HIV-infected individuals and epithelial barrier dysfunction. PLoS Pathog 13 (2):e1006202 175. Christensen-Quick A et al (2017) Cytomegalovirus and HIV persistence: pouring gas on the fire. AIDS Res Hum Retrovir 33(S1): S23–S30

176. Gianella S, Letendre S (2016) Cytomegalovirus and HIV: a dangerous pas de Deux. J Infect Dis 214(Suppl 2):S67–S74 177. Dock JN, Effros RB (2011) Role of CD8 T cell replicative senescence in human aging and in HIV-mediated immunosenescence. Aging Dis 2:382–397 178. Effros RB (2016) The silent war of CMV in aging and HIV infection. Mech Ageing Dev 158:46–52

Chapter 2 Distinct Properties of Human Cytomegalovirus Strains and the Appropriate Choice of Strains for Particular Studies Giada Frascaroli and Christian Sinzger Abstract Human cytomegalovirus is routinely isolated by inoculating fibroblast cultures with clinical specimens suspected of harboring HCMV and then monitoring the cultures for cytopathic effects characteristic of this virus. Initially, such clinical isolates are usually strictly cell-associated, but continued propagation in cell culture increases the capacity of an HCMV isolate to release cell-free infectious progeny. Once cell-free infection is possible, genetically homogenous virus strains can be purified by limiting dilution infections. HCMV strains can differ greatly with regard to the titers that can be achieved, the tropism for certain cell types, and the degree to which nonessential genes have been lost during propagation. As there is no ideal HCMV strain for all purposes, the choice of the most appropriate strain depends on the requirements of the particular experiment or project. In this chapter, we provide information that can serve as a basis for deciding which strain may be the most appropriate for a given experiment. Key words Cytomegalovirus, Isolate, Strain, Tropism, Titer

1

Introduction In the 50 years since the discovery of the original HCMV strain (AD169) [1], numerous different strains have been propagated and used for HCMV research in various cell culture systems. With increasing knowledge about phenotypic and genotypic differences between these strains, it has become clear that the choice of the virus strains used for experiments will greatly influence the results of a certain project. It may be particularly relevant to decide whether recent clinical HCMV isolates or established laboratory strains should be used, which is the most suitable strain for a given cell type, or whether a BAC-cloned HCMV strain that can be easily modified by genetic manipulation will be advantageous. This chapter is designed to facilitate this decision-making process by describing the distinct features of the various HCMV strains that are available and discussing their advantages and disadvantages for certain experimental approaches.

Andrew D. Yurochko (ed.), Human Cytomegaloviruses: Methods and Protocols, Methods in Molecular Biology, vol. 2244, https://doi.org/10.1007/978-1-0716-1111-1_2, © Springer Science+Business Media, LLC, part of Springer Nature 2021

19

20

2 2.1

Giada Frascaroli and Christian Sinzger

Materials Plasticware

1. Sterile 1.0 ml cryovials. 2. Sterile 1.5 ml microtubes. 3. Sterile 15 ml conical tubes. 4. 96-well flat bottom cell culture plates. 5. 6-well cell culture plates. 6. 25 cm2 tissue culture flasks. 7. Sterile syringe filters (pore size 0.45 μm).

2.2 Solutions and Culture Media

1. Cell culture grade H2O. 2. Dulbecco’s Modified Eagle Medium with 5% fetal bovine serum, 2.4 mmol/l glutamine and 50 mg/l gentamicin (DMEM5). 3. Minimum Essential Medium with 5% fetal bovine serum, 2.4 mmol/l glutamine, and 50 mg/l gentamicin (MEM5). 4. 2
MEM with 10% fetal bovine serum, 4.8 mmol/l glutamine, and 100 mg/l gentamicin. 5. RPMI 1640 medium containing 10% human serum (CMV seronegative), 50 mg/l gentamicin, 5 IU/ml heparin, and 50 μg/ml endothelial cell growth supplement (RPMI10). 6. Trypsin–EDTA (with 0.05% trypsin).

2.3

Other Chemicals

1. Cell culture grade agarose. 2. Mitomycin C.

2.4

Cells

1. Human foreskin fibroblasts (HFF). 2. Human umbilical vein endothelial cells (HUVEC).

2.5

Equipment

1. Cell culture incubator with CO2 and humidification. 2. Centrifuge with a swingout rotor and racks suitable for centrifugation of 15 ml conical tubes at 3000
g and microtiter plates at 300
g. 3. Water bath. 4. Microwave oven. 5. Inverted light microscope, ideally equipped for phase contrast.

Distinct Properties of Human Cytomegalovirus Strains

3 3.1

21

Methods HCMV Strains

3.1.1 Clinical Isolates

HCMV can be isolated from various specimens of patients with acute infection, including urine, throat washings, bronchoalveolar lavage, leukocytes, and biopsies from affected organs. This is a routine procedure in most virological laboratories, and human fibroblasts are the cell of choice for these isolations. Virus recovered from a human specimen and passaged in culture is regarded as a clinical isolate until phenotypic alteration occurs (e.g., release of cell-free progeny is an indicator of adaptation to cell culture; see Subheading 3.1.2). Isolation of HCMV from Clinical Specimens (e.g., urine). 1. Dilute 2 ml of the urine sample with 2 ml of DMEM5 in a 15 ml conical tube (see Note 1). 2. Centrifuge for 10 min at 3000
g. 3. Filter supernatant through a 0.45 μm filter to remove bacteria and fungi. 4. Inoculate multiple subconfluent HFF cultures (e.g., duplicates or triplicates) with the filtered supernatant (e.g., 100 μl per 15,000 cells in a 96-well microculture plate). 5. Centrifuge plates at 300
g for 30 min. 6. Incubate for additional 30 min at 37  C with 5% CO2 in an incubator. 7. Remove inoculum and wash cells with 200 μl of DMEM5. 8. Add 200 μl of DMEM5 and incubate cells at 37  C with 5% CO2 in an incubator until a focal cytopathic effect becomes detectable under the inverted light microscope, typically within 1 or 2 weeks after inoculation. 9. Detach cells of infected cultures with Trypsin/EDTA and coculture with uninfected fibroblasts for passaging of the isolate. Advantages of clinical isolates are that (1) they are phenotypically and genotypically closely related to the viruses replicating within patients including maintenance of wild-type cell tropism, (2) they are well suitable for correlative studies between viral genetic polymorphisms and clinical symptoms, and (3) they may serve as sources for the identification of new viral genes or gene variants lost in laboratory strains due to the selective pressure in cell culture. (4) Additionally, clinical isolates are an ideal material for analyses of natural genetic variability present in HCMV. On the other hand, it may be disadvantageous that isolates are not plaque purified, hence are not genetically pure [2] and do not usually yield high-titer cell-free viral stocks. Low passage clinical isolates usually grow

22

Giada Frascaroli and Christian Sinzger

strictly cell-associated with almost no infectivity released into the supernatant [3, 4]. This means that the virus has to be propagated by coculture of infected and uninfected cells, and synchronous infection of cultured cells with cell-free virus preparations is not possible. Thus, these low passage clinical isolates may not be useful for downstream techniques that require synchronous infection such as analysis of gene expression kinetics in cell lysates by immunoblotting. To obtain cell-free infectivity from recent clinical isolates, release of “intracellular virus” from infected cells by ultrasonication, freeze–thaw lysis, or mechanical disruption has been suggested. However, reasonable virus titers can only be achieved by these methods when the isolate is capable of releasing at least small amounts of infectious virus progeny into the supernatant (unpublished observation). A further drawback is that clinical isolates are genetically unstable due to the selective pressure for replicative fitness under certain cell culture conditions. Viral genes necessary for interaction with the host immune system or for replication in specialized cell types and tissues may be lost if they are not beneficial in the cell type used during isolation. For example, clinical isolates rapidly lose the replication inhibitor RL13 during propagation in various cell types. Isolates that have lost RL13 acquire the increased capacity to release infectious progeny into the supernatant (see Note 2) [5]. 3.1.2 Established Laboratory Strains with Restricted Tropism

It is commonly assumed that once an HCMV isolate is fully adapted for growth in fibroblast cell culture and there is no further selective pressure against any of the viral genes, the genome will remain relatively stable during continued propagation. This is supposedly the case when—after an initial increment of progeny virus titers during cell-free passages in fibroblasts—no further increase occurs. At this point, a genetically homogenous HCMV strain may be generated by plaque purification, which relies on the assumption that a single viral genome can cause the formation of a focus of infected cells or “plaque” in an otherwise uninfected monolayer. To increase the chance of picking a single genome and hence the chance for selection of a homogenous strain, the procedure is repeated three times, before the resulting strain is considered “purified” (see Note 3). A number of highly productive HCMV strains now widely used as standard strains by many researchers were established in this way. The most frequently used of these “laboratory strains” are AD169 and Towne [1, 6]. Other laboratory strains like Toledo or TR [7–9] have also been proven to be very useful for HCMV research. Plaque Purification. 1. Seed HFFs in a 6-well plate at a cell density of 150,000 cells per well in MEM5.

Distinct Properties of Human Cytomegalovirus Strains

23

2. The next day, add serial tenfold dilutions of virus to the cell cultures (e.g., 103 to 108 fold dilutions of virus in medium) and incubate for 1 h. 3. Prepare 0.6% agarose solution by adding 0.6 g agarose to 100 ml H2O. Boil the solution in a microwave oven and be sure to account for evaporative loss of fluid. Let cool to 60  C and add 100 ml of twofold concentrated cell culture medium (e.g., 2
MEM with 10% fetal bovine serum, 4.8 mmol/l glutamine, and 100 mg/l gentamicin). Let the agarose– medium solution cool to 37  C and maintain at this temperature in a water bath. 4. When the incubation with virus from step 1 is finished, remove supernatants and replace with 2 ml of the agarose–medium solution. Incubate for 5–12 days or until plaques are easily visible. The amount of time required for this incubation is variable depending on the growth properties of the virus being used. 5. The day before harvesting plaques, seed HFFs in 25 cm2 tissue culture flasks at a density of 300,000 cells per flask. 6. Under a phase contrast microscope, identify plaques of infected cells, scratch them with a pipette tip, and aspirate 20–30 μl. Transfer aspirate immediately to 1 ml of MEM5 and add to HFFs in 25 cm2 flasks. 7. Incubate for 7–14 days until plaques have formed in the inoculated culture and passage infected cultures by detaching cells with trypsin–EDTA solution and reseeding them in the appropriate medium. 8. When >90% of HFFs show a late stage cytopathic effect (CPE) (see Note 4), wait two additional days, harvest plaque-purified virus by clarifying the supernatants from cell debris. 9. Store aliquots of virus at 80  C until further use. 10. Repeat steps 1–8 twice to increase the chance of yielding a genetically pure “clonal” virus. Besides highly efficient growth yielding high titers of viral progeny, another important advantage of these strains is that they have been widely distributed allowing one to compare experimental results with published data obtained under similar conditions. For example, in the case of the AD169 strain, many viral genes have been cloned from this strain and subsequently analyzed in isolation, yielding an enormous amount of information useful for various aspects of molecular HCMV research. It is also advantageous that the genomic sequences are available for many of these established strains allowing for genotype–phenotype comparison, cloning of viral genes, and so on. One caveat to consider is that derivatives of established

24

Giada Frascaroli and Christian Sinzger

strains may exist, such as the genotypically and phenotypically distinct variants varATCC, varUK, and varUC of strain AD169 and varS and varL of strain Towne [10]. An obvious disadvantage of these well-established strains is the loss of genomic material due to extensive adaptation for growth in fibroblasts, which almost inevitably results in the potential loss of important functions not required for optimal growth in fibroblasts. HCMV strains such as AD169 and Towne that have been extensively propagated in fibroblasts have a restricted host cell range, that is, besides fibroblasts these strains can infect smooth muscle cells, hepatocytes, and trophoblast cells but they can only inefficiently infect endothelial cells, epithelial cells, macrophages, and dendritic cells. This phenotypic change occurs after about 20 passages and is due to functional disruption of the UL128 gene locus, that is, at least one of the three proteins pUL128, pUL130, and pUL131A is deleted or expressed in a nonfunctional form [3, 5, 11–14]. As a consequence, strains AD169 and Towne are inadequate for infection of endothelial cells, epithelial cells, and cells derived from peripheral blood monocytes. 3.1.3 Established Laboratory Strains with Extended Tropism

The host cell restriction of HCMV strains observed during extensive passaging in fibroblasts can be prevented by propagation in endothelial cell cultures, which have the capability to preserve the natural broad cell tropism of HCMV [14]. Although there is considerable interstrain variation, almost all clinical HCMV isolates have an extended cell tropism represented by their ability to form foci both in fibroblast and in endothelial cell cultures [3]. During further propagation in fibroblasts, the strict cell association is usually lost around passage number 10, presumably due to disruption of the replication inhibitor RL13. Disruption of RL13 allows the resulting virus strains to infect a broad range of cell types in a cellfree mode for a limited number of passages until the abovementioned disruption of the UL128 locus occurs, restricting the host cell range [3, 5]. A convenient way to preserve the extended cell tropism of newly isolated virus strains is to “transfer” the virus to endothelial cell cultures shortly after isolation. Transfer of cell-associated HCMV isolates from fibroblasts to endothelial cells. 1. Propagate the virus isolate in HFFs until 10% of the cells exhibit a CPE (see Note 4). 2. To induce terminal differentiation and abolish mitotic activity, treat fibroblast cultures with 2
106 mol/l mitomycin C for 48 h. Wash cultures and incubate in medium without mitomycin C for 48 h. Repeat treatment with mitomycin C (see Note 5).

Distinct Properties of Human Cytomegalovirus Strains

25

Fig. 1 HCMV strains differ with regard to their productivity. Fibroblast cultures were infected with the respective strain and washed extensively on day 3 after infection to remove residual input virus. Progeny virus was collected on day 7 after infection and stored at 80  C until determination of virus titers. Titers of infectivity were determined by infection of fibroblast cultures with dilution series of the various virus preparations and detection of viral IE antigens by indirect immunofluorescence 24 h after infection. The result of a representative experiment is shown

3. Coculture aliquots of the terminally differentiated isolateinfected HFF cultures with HUVECs at an HFF–HUVEC ratio of 1:10 in RPMI10. Propagate cultures until the desired CPE is reached. For an alternative approach, in case the isolate releases cellfree infectivity, see Note 6. Typical strains that have been adapted to endothelial cell cultures shortly after primary isolation and thus have preserved the extended tropism are VHL/E [14] and TB40/E [3]. Both have the additional advantage that they produce high titers (see Note 7), in contrast to other endotheliotropic HCMV strains like VR1814 (Figs. 1 and 2). The extended tropism also includes high infectivity for epithelial cells and monocytederived macrophages and dendritic cells [15]. 3.1.4 BAC-Cloned Strains

To facilitate mutation of viral genes in the context of replicating virus, HCMV genomes of strains AD169, Towne, VR1814, TR, PH, Toledo, VHL/E, and TB40/E have been cloned into F1 plasmid vectors that allow for amplification, mutation, and selection of the viral DNA as bacterial artificial chromosomes (BACs) in

26

Giada Frascaroli and Christian Sinzger

Fig. 2 HCMV strains differ with regard to their endothelial cell (EC) tropism. Fibroblast cultures were infected with the respective strain and washed extensively on day 3 after infection to remove residual input virus. Progeny virus was collected on day 7 after infection and stored at 80  C until determination of virus titers. The relative EC tropism of the various virus preparations was determined by simultaneous infection of human umbilical vein endothelial cell (HUVEC) cultures and fibroblast (HFF) cultures with dilution series of the various virus preparations and detection of viral IE antigens by indirect immunofluorescence 24 h after infection. The relative EC tropism was calculated as the infection efficiency in HUVECs/HFFs. The result of a representative experiment is shown

bacteria [7, 16–21]. Briefly, in a first step, HCMV genomes replicating in infected fibroblasts are recombined with a cassette containing the bacterial origin of replication, a resistance gene for selection in fibroblasts and a resistance gene for selection in bacteria. In a second step, the selected recombined viral genomes are purified and transferred into bacteria, where they can then be amplified and mutated. In a third step, the mutated replicationcompetent viral genomes are transfected into fibroblasts, and successfully transfected fibroblasts will then reconstitute the BAC-cloned virus. More detailed protocols regarding isolation and genetic manipulation of viral BACs can be found in other chapters in this volume. As the size of the HCMV genome is already at the packaging limit, insertion of additional DNA, such as the BAC cassette, will reduce the replicative fitness of the virus [22] unless viral genes are deleted. In BAC clones generated according to the method developed by Borst et al., the viral open reading frames US2–6, known to be nonessential for replication in cell

FJ616285

–

–

KX544841

GU179289

AY446894 NC_006273

Towne

TR

TB40/E

VHL/E

VR1814

Merlin

Based on pAL1111(RL13, UL128 repaired)

BAC: IRS1, US1-6

UL135-147 BAC: US2-6

RL6, RL13, (UL141) BAC: IRS1, US1-6

UL97nt1771-1782 BAC: US2-5

pAL1128

GU179001

High

High

Low

Low

Low

High

High

Medium

Based on TowneBAC(UL130 repaired) High

UL130; UL133-146 BAC: IRS1; US1-12

RL13, UL128

AC146907

MK425187

EF999921

AC146906

AY315197

Based on BADwt(UL131 repaired)

References

Extended [23]

Restricted [5, 23]

Extended [7, 19]

Extended [14, 21]

Extended [3, 20]

Extended [7, 9]

Extended [32]

Restricted [16, 44]

Extended [31]

Restricted [10, 18]

High

KU317610

RL5A, RL13, (UL42-43) UL131A;UL133-150 pHB5: US2-6

Productivity Tropism

GenBank accession no. Genes known to be altered/deleted

pAL1111

FIX-BAC

VHL/EBAC19

TB40-BAC4

TR-BAC

TowneUL130rep

TowneBAC

BADrUL131

BADwt pHB15 pHB5

GenBank accession no BAC-clone

AD169 AC146999 (varATCC)

Strain

Table 1 Properties of selected HCMV strains and their BAC clones

Distinct Properties of Human Cytomegalovirus Strains 27

28

Giada Frascaroli and Christian Sinzger

culture, are replaced with the BAC cassette [18]. The resulting genomes are still about 4.8 kb larger than the wild-type genomes, and this may be one reason for the slightly decreased titers obtained by BAC-derived virus when compared to wild-type virus (compare AD169 and pHB5 in Fig. 1). In BAC clones of strain Towne, generated with the method developed by Marchini et al., IRS1 and US12 are truncated and US1 to US11 are deleted [16]. Attempts have been made to generate HCMV-BAC clones without deletion of viral sequences. In this approach, a BAC cloning cassette flanked by LoxP sites was introduced between the US28 and US29 open reading frames of HCMV by homologous recombination in fibroblasts, and the successfully recombined genomes were then transferred into Escherichia coli where mutagenesis techniques can be applied [22, 23]. If desired, the BAC cassette can be excised during reconstitution of virus in fibroblasts by cotransfection of Cre recombinase expression plasmid, resulting in a mostly intact HCMV carrying only the desired mutations. An alternative approach places the BAC cassette into an essential virus gene, relying on removal of the cassette as the essential gene is repaired during reconstitution after transfection into fibroblasts [24]. However, self-excising BAC cassettes have the disadvantage that the genomes of the reconstituted viruses cannot be retransferred into bacteria as they no longer contain the F1-plasmid sequences. A further caveat comes from the possibility that introduction of additional sequences during the initial homologous recombination step in fibroblasts may lead to undesired deletions as a consequence of the increased genome size. When working with HCMV strains reconstituted from BACs, it is essential to consider which genes are present or missing due to the respective BAC cloning procedure or due to strain-specific polymorphisms in the genetic backbone (see Table 1). 3.1.5 Genetically Repaired BAC-Derived Strains

While many cell culture-adapted HCMV strains are available as BAC clones, BAC-cloned clinical isolates (see Note 2) have not been reported. Obviously, the first step of the BAC cloning procedure, that is, recombination of viral genomes with the BAC cassette in infected fibroblasts, will put a strong selective pressure against cell-associated genomes with intact RL13, thus promoting selection of BAC clones with mutations in RL13, which can spread more efficiently due to the capacity to release higher titers of infectious virus. To overcome this limitation, RL13 has been repaired in an existing BAC clone of the HCMV strain Merlin originally derived from the urine of a congenitally infected infant [25]. The rapid appearance of disabling RL13 mutations already during the reconstitution of repaired genomes in fibroblasts emphasized how strong is the abovementioned selective pressure against RL13 [23]. Only when expression of the intact RL13 gene was conditionally

Distinct Properties of Human Cytomegalovirus Strains

29

repressed, it was possible to reconstitute BAC-cloned viruses with wild-type RL13 sequences [23]. This new strain combines many of the advantages of BAC-cloned HCMV strains mentioned before such as genetic stability and purity with an isolate-like phenotype, that is, strictly cell-associated growth (see Note 2). Moreover, it allows investigators to test targeted mutations in short-term infection experiments in a system where the reconstituted Merlin viruses largely represent the genetic equivalents of an HCMV clinical isolate. The key here is the short term as after a few rounds of replication, RL13 will again be disrupted. One drawback of this system is that the virus can only be maintained in an immortalized cell line expressing the Tet repressor and high titers of virus progeny have not yet been obtained. 3.2 Choice of Strains for Particular Projects

As described in the previous section, many different HCMV strains are available, with quite distinctive features. There is no such thing as the ideal strain serving all purposes. The appropriate choice of an HCMV strain depends on the specific experimental design and requirements. The following paragraphs are meant as a guide for decision-making in the planning phase of a project.

3.2.1 Experiments Addressing Genetic Variability Between HCMV Isolates

Although herpesviral genomes are regarded as rather stable when compared to other viruses, genetic polymorphisms have been reported for many HCMV genes when different laboratory strains or isolates from different patients have been compared [7, 26]. For example, surface glycoproteins gN and gO show a high degree of polymorphism, probably reflecting the selective pressure of neutralizing antibodies [7, 27, 28]. Clinical HCMV strains are an important source for experiments in which naturally occurring polymorphisms within HCMV genes are to be considered. Examples are investigations into strain susceptibility to newly developed antiviral agents, and strain evolution in patients that have been unsuccessfully treated with experimental or prescribed pharmaceuticals and in which the emergence of drug-resistant strains is suspected. The drawback of using newly generated clinical isolates is that these viruses remain cell-associated and are not useful when infections with defined infection multiplicities using virus-containing supernatants are required. Protocols for focus expansion assays and plaque reduction assays have been developed to overcome this limitation and to allow for a certain degree of standardization when using clinical isolates. Moreover, cell-associated spread is mostly insensitive to neutralizing antibodies which prevents direct investigation of strain-specificity of antibodies with clinical isolates. To overcome this limitation, it may be convenient to use a “copy and paste” approach swapping sequences between HCMV strains [29]. For example, the glycoprotein of interest might be cloned

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into the genetic background of a laboratory strain to analyze its sensitivity to antibodies within the context of this cell-free virus. 3.2.2 Experiments Depending on High Infection Multiplicities

High virus titers are a prerequisite for all approaches where synchronous infection of a high percentage (>95%) of the cells is desired. For example, HCMV-induced downregulation of cellular factors may go undetected if too many uninfected cells are present in the sample. On the other hand, upregulation of cellular factors in an infected culture can be due to modifications in uninfected bystander cells, and again infection rates approaching 100% are desired to exclude this possibility and attribute the observed effect directly to infected cells. The fraction of infected cells (Finf) depends on the infection multiplicity (MOI, see Note 8) and is calculated according to the principles of Poisson distribution: Finf ¼ 1  (eMOI). This means that about 63% of cells are infected with an MOI of 1 infectious unit (IU)/cell and about 99% of cells are infected with an MOI of 5 IU/cell. High MOIs are desired for analyses of viral attachment and entry, especially when the number of virions per infected cell needs to be counted after infection with fluorescent-labeled virus or after immunofluorescence staining of virus particles. High MOIs are also desired when virion proteins or nucleic acids from input particles need to be detected by blotting techniques in infected cell lysates. Even more input virus is necessary for electron microscopic analyses of entry events: to detect only one virion per cell in an ultrathin section with conventional transmission EM, virus loads of >100 particles per cell are required. Cell culture-adapted HCMV strains like AD169, Towne, VHL/F, and TB40/F are particularly suitable for these purposes. Titers of >106 IU/ml can usually be achieved, allowing for experiments with MOIs of >10 with native virus preparations. Higher virus titers (>107 IU/ml) can be achieved by ultracentrifugation of large volumes of native virus preparations and resuspension of the virion pellet in small volumes of medium (see also Note 9 on the problem of particle–infectivity ratios and Note 10 on avoidance of dense bodies in virus preparations). Such enriched virus suspensions are often necessary for ultrastructural analyses of virus entry. If the cell type of interest cannot be infected by fibroblast-adapted strains, EC-adapted strains like VHL/E and TB40/E may be useful as they combine broad cellular tropism with relatively high titer.

3.2.3 Infection of Endothelial Cells, Epithelial Cells, or Professional Antigen Presenting Cells

For analyses of HCMV infection of endothelial cells, epithelial cells or professional antigen presenting cells, several options exist. Newly isolated clinical strains are a viable option as they have a naturally broad cellular tropism and are usually able to infect endothelial, epithelial, and myeloid cells. However, their strictly cell-associated growth prevents the generation of virus stocks containing high titers of cell-free progeny virus [3, 5]. This greatly limits the repertoire of possible experiments as one is limited to approaches based

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on coculture of infected and uninfected cells. Alternatively, cell culture-adapted RL13-/UL128+ derivatives of newly isolated clinical strains can be used if synchronous infection by cell-free supernatants is required. Still, virus titers produced by such strains (103 to 104 IU/ml) may be too low for a highly efficient infection of such cell cultures [5]. However, at least two well-established highly endotheliotropic HCMV strains (TB40/E and VHL/E) are available that produce high viral titers in cell-free supernatants and allow for high infection rates (Fig. 2). Alternatively, variants of AD169 and Towne are available in which the defective ORFs, UL131A and UL130, respectively, have been genetically repaired [30–32]. In addition to the choice of the appropriate strain, the choice of the producer cell type for generation of virus stocks is important. For example, progeny virus produced by infected fibroblast cultures was reported to be more endotheliotropic than progeny virus produced by infected endothelial cells [33]. Therefore, to achieve maximum infection rates in endothelial cells virus stocks should be produced in fibroblasts. However, one must be careful to use fibroblasts only to amplify a high-titer stock as repeated passage of endotheliotropic viruses in fibroblasts can lead to loss of UL128– UL131A, as discussed previously. A producer cell difference was also reported between fibroblasts and epithelial cells: virus propagated in the epithelial ARPE-19 cell line incorporated UL128UL131A proteins more efficiently into virions as compared to virus propagated in HFFs, associated with a respective shift in entry pathways and cell tropism [34, 35]. 3.2.4 Genetic Manipulation of HCMV Genomes

When a targeted mutation in viral genes in the context of replication-competent virus is desired, BAC-cloned viruses are the appropriate choice for most projects. Once the viral genome is available as a replicating bacterial artificial chromosome in E. coli, many tools for genetic engineering of plasmids can be applied in order to modify the viral genes contained in this vector. The tailored virus will then be recovered from fibroblasts that have been transfected with the mutated BACs. Importantly, the UL128 gene locus, which confers endothelial cell tropism, is stable during reconstitution in fibroblasts with most of the UL128+ BAC strains. This is surprising considering the selective pressure against this gene locus in recent clinical isolates and during reconstitution of UL128+ BAC-cloned Merlin variants. An explanation may be provided by strain-dependent expression levels in the UL128 locus [23, 36]. Obviously, a high pUL128 expression level—as in strain Merlin—renders the virus more prone to disrupting mutations in this gene during reconstitution or propagation. In this case, HCMV-BACs may also be reconstituted by transfection into cell types other than fibroblasts (e.g., epithelial cells) [23]. Many of the relevant HCMV strains are available as BAC-cloned viruses. They

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differ from each other regarding the viral genes that were replaced with the BAC cassette and they carry the genetic characteristics of their parental strain, which may influence the decision which of the BACs is suitable for the planned project. In order to have a valid wild-type control for a desired mutation, the chosen BAC clone should be intact regarding the gene of interest. Knockin mutants restoring the gene of interest are an alternative strategy. In addition to the integrity of the gene of interest, tropism and titer production may also influence the choice of a certain strain. 3.2.5 Virus–Host Defense Interactions

Virus genes that counteract antiviral actions of CD8 T cells, CD4 T cells, NK cells, and antibodies are usually nonessential for growth in cell culture and may therefore be counterselected during propagation of the virus in cell culture. For example, the NK evasion genes UL141 and UL142 are deleted in the genome of AD169varATCC (genetic defects of various strains are listed in Table 1). This has to be considered when studying virus–host defense interactions: clinical isolates can be assumed to encode the full set of HCMV genes, whereas “nonessential” genes may have been lost in cell cultureadapted strains. None of the available HCMV strains can guarantee genetic completeness. On the other hand, a certain gene usually is not disrupted in all strains (see Table 1). Therefore, inclusion of several strains reduces the risk of missing a viral immune evasion gene in a project addressing yet undefined immune evasive functions of HCMV. With regard to well-established immune evasive functions of HCMV (e.g., US2, 3, 6, and 11 for MHC class I mediated antigen presentation; UL16, 18, 40, 141, and 142 for NK cell function) [37], an appropriate strain can be chosen according to the available sequence information, in order to ensure that the viral genes of interest are intact.

3.2.6 Cell-Associated Spread

The fact that newly isolated clinical strains are able to grow in fibroblast cultures with almost no detectable virus in the culture supernatant suggests that cell-to-cell spread contributes to viral growth in vivo. The use of recent clinical isolates are appropriate for studying the underlying mechanisms of viral pathogenesis. Infected cells have to be cocultured with uninfected indicator cells before the extent of cell-to-cell transmissions can be analyzed by quantitative measurement of focus size or number [38]. The limits of such an approach are mentioned in Subheading 3.1.1. To analyze the effect of targeted mutations on cell-to-cell spread, cellassociated derivatives of BAC-cloned HCMV strains may under some circumstances serve as viable surrogates for clinical isolates. For example, deletion of UL99 or UL74 almost completely abrogates release of cell-free infectivity but still allows cell-associated focal spread of the virus [39, 40]. Such an experimental setup is advantageous in that the viral genome is known and can be further

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manipulated. The disadvantage of such an experimental setup, however, is that the mechanisms of virus transfer may differ from clinical isolates in which the UL99 and/or UL74 genes are intact. Therefore, results from such studies have to be interpreted carefully. The BAC-cloned version of the Merlin strain which conditionally represses RL13 in the presence of the Tet repressor expressed from stably transfected fibroblasts [23] (see Subheading 3.1.5) can help to overcome many limitations of clinical isolates, that is, allow for targeted mutation of genes of interest, ensure a high genetic stability, and allow for synchronous infection at the beginning of the experiment before derepressing the RL13 gene. 3.3

4

Conclusions

It has become evident that HCMV strains can differ genetically and phenotypically due either to natural interstrain polymorphisms or due to alterations occurring during extended propagation in cell culture. In particular, differences concerning the titers that can be achieved, the tropism for certain cell types, and the degree to which nonessential genes have been lost during propagation can be observed. As there is no ideal HCMV strain for all purposes, the choice of the most appropriate strain depends on the requirements of the particular experiment or project. Recent clinical isolates are ideal for correlative studies between viral genetic polymorphisms and clinical symptoms and may serve as a source for the identification of new viral genes or gene variants. The well-established fibroblast-adapted HCMV strains combine the advantage of high-titer production with a plethora of reference data available in the literature, however, the cost of using these strains is their restricted host cell tropism. For experiments in endothelial cells, epithelial cells, and leukocyte-derived cells, HCMV strains with unrestricted cell tropism are preferable. For genetic manipulation, BAC-cloned HCMV strains are the best choice as they can be easily engineered in E. coli. Many of the BAC-cloned viruses will, however, grow to lower titers than their parental strains and may have genetic defects due to insertion of the BAC cassette into the viral genome.

Notes 1. For isolation of HCMV from throat washing specimens directly start with step 2 (see Subheading 3.1.1) using 4 ml of the specimen. 2. “Clinical isolates” usually grow focally in a cell-associated manner and do not release virus progeny into the culture supernatant. As soon as the virus starts to release significant amounts of infectivity into the supernatant (usually within the first ten passages), the isolate becomes presumably adapted to growth

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in cell culture (including loss of RL13) and is therefore not “clinical isolate” in the true sense of the definition. This means that VHL/E, Toledo, TB40/E, VR1814, TR, and similar viruses are not “clinical isolates,” but cell culture-adapted “strains.” Sometimes, the term “clinical-like isolates/strains” is applied to indicate that these strains are genetically more complete than AD169 varATCC or Towne varS. To avoid confusion, we do not use the term “clinical strain” and we call a fresh patient isolate “clinical isolate” only as long as it grows cell-associated. 3. As demonstrated for strain TB40/E, plaque purification is no guarantee for genetic homogeneity [12, 20, 41]. The best way to achieve genetic homogeneity is by cloning the viral genome into a BAC and reconstituting replicating virus by transfection of the cloned genome into a permissive cell culture. 4. When a cytopathic effect (CPE) is mentioned in the protocols of this chapter, the detection of nuclear inclusions in phase contrast microscopy is meant, as this will indicate that the cell culture is producing virus progeny. Besides this CPE, which is indicative of the late phase of viral replication, HCMV can also cause other CPEs, such as cell rounding in the early phase of infection, syncytia formation in the late stage of infection, and cell destruction in the final stage of infection. While the time course and extent of the latter CPEs may vary depending on the virus strain, a nuclear inclusion will usually appear the latest on day 4 after infection in each productively infected HFF cell. 5. HCMV that has been isolated from patient material on fibroblasts can be transferred to endothelial cells by coculturing the infected fibroblast culture with uninfected endothelial cells. However, the uninfected fraction of the fibroblast culture may overgrow the endothelial cells due to their faster replication. To prevent this, mitosis in the fibroblast culture can be irreversibly inhibited with mitomycin C before coculturing the fibroblasts with endothelial cells. 6. Alternatively, the recent isolate can be transferred by cell-free transmission to endothelial cell monolayers directly after the appearance of cell-free virus, given that sufficient virus progeny is released into the supernatant. Isolation of HCMV directly from endothelial cell cultures is also possible. 7. When fibroblasts and endothelial cells are infected with the same preparation of an HCMV strain with extended tropism, the tropism of progeny virus differs to some degree depending on the producer cell type. Progeny from fibroblast cultures have a higher endothelial cell tropism than progeny from endothelial cells [33]. High-titer stocks of “EC-tropic” virus should therefore be produced in fibroblasts.

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8. MOI in this sense means the average number of infectious particles that have bound per cell in a given experiment. By definition, this MOI can only be determined retrospectively, for example, in the same cells infected in parallel with a dilution series of the same virus preparation. Often the MOI is estimated in advance as an extrapolation from previous experiments. This “estimated MOI” may differ from the “actual MOI” to some extent. 9. Particle–infectivity ratio is an important issue when a method basically detects the physical particle rather than the biologically active infectious unit. This concerns the detection of virions under the electron microscope, fluorescent virus particles in live microscopy, immunofluorescence detection of capsids, and also the determination of DNA copies with quantitative DNA detection methods. Usually, not all particles are infectious. Inevitably, a virus preparation will contain enveloped capsids lacking DNA or packed with damaged DNA. In addition, those particles that are infectious immediately after harvest have a biological half-life of about 1 day [42], which means that supernatant collected over a period of several days will contain a majority of inactive particles. Hence, the fraction of nonactive particles can be limited by using supernatants produced over a short time period, for example, overnight after a medium exchange. Unfortunately, procedures like freezing the virus and even ultracentrifugation will also affect the biological activity of HCMV virions. This has to be taken into account, when interpreting the localization of particles in EM or the behavior of particles in fluorescence assays. If necessary, the particle–infectivity ratio can be determined by counting the number of capsids per cell and comparing this to the MOI. A particular source of misunderstanding is the determination of a “particle–infectivity” ratio by comparison of the genome copies in a given supernatant with its ability to infect cell cultures. The genome copy number will reflect the total number of particles in the preparation, whereas only a small fraction of infectious particles in the inoculum is absorbed by a cell culture during the incubation. Such “particle–infectivity” ratio will reflect the absorption rate rather than the biological potency of the virions. Even if all virions were biologically active, only few will bind to and infect a cell, resulting in a very high “particle– infectivity” ratio. 10. If the presence of dense bodies needs to be avoided, virus preparations have to be gradient purified. However, the procedure of ultracentrifugation through a density gradient will increase the number of noninfectious particles in the purified virion fraction. Fresh preparations of pp65-deletion mutants are a good alternative in this situation as they do not produce

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dense bodies and can therefore be used directly without the need of gradient purification [43]. We apologize for not mentioning certain HCMV strains or not describing them in detail. We invite suggestions from colleagues who have experience with strains not commented here in order to improve future versions of this chapter. References 1. Rowe WP, Hartley JW, Waterman S et al (1956) Cytopathogenic agent resembling human salivary gland virus recovered from tissue cultures of human adenoids. Proc Soc Exp Biol Med 92:418–424 2. Prichard MN, Penfold ME, Duke GM et al (2001) A review of genetic differences between limited and extensively passaged human cytomegalovirus strains. Rev Med Virol 11:191–200 3. Sinzger C, Schmidt K, Knapp J et al (1999) Modification of human cytomegalovirus tropism through propagation in vitro is associated with changes in the viral genome. J Gen Virol 80(Pt 11):2867–2877. https://doi.org/10. 1099/0022-1317-80-11-2867 4. Yamane Y, Furukawa T, Plotkin SA (1983) Supernatant virus release as a differentiating marker between low passage and vaccine strains of human cytomegalovirus. Vaccine 1:23–25 5. Dargan DJ, Douglas E, Cunningham C et al (2010) Sequential mutations associated with adaptation of human cytomegalovirus to growth in cell culture. J Gen Virol 91:1535–1546. https://doi.org/10.1099/vir. 0.018994-0 6. Plotkin SA, Furukawa T, Zygraich N, Huygelen C (1975) Candidate cytomegalovirus strain for human vaccination. Infect Immun 12:521–527 7. Murphy E, Yu D, Grimwood J et al (2003) Coding potential of laboratory and clinical strains of human cytomegalovirus. Proc Natl Acad Sci U S A 100:14976–14981. https:// doi.org/10.1073/pnas.2136652100 8. Quinnan GV, Delery M, Rook AH et al (1984) Comparative virulence and immunogenicity of the Towne strain and a nonattenuated strain of cytomegalovirus. Ann Intern Med 101:478–483 9. Smith IL, Taskintuna I, Rahhal FM et al (1998) Clinical failure of CMV retinitis with intravitreal cidofovir is associated with antiviral resistance. Arch Ophthalmol 116:178–185. https://doi.org/10.1001/archopht.116.2. 178

10. Bradley AJ, Lurain NS, Ghazal P et al (2009) High-throughput sequence analysis of variants of human cytomegalovirus strains Towne and AD169. J Gen Virol 90:2375–2380. https:// doi.org/10.1099/vir.0.013250-0 11. Akter P, Cunningham C, McSharry BP et al (2003) Two novel spliced genes in human cytomegalovirus. J Gen Virol 84:1117–1122. https://doi.org/10.1099/vir.0.18952-0 12. Dolan A, Cunningham C, Hector RD et al (2004) Genetic content of wild-type human cytomegalovirus. J Gen Virol 85:1301–1312. https://doi.org/10.1099/vir.0.79888-0 13. Hahn G, Revello MG, Patrone M et al (2004) Human cytomegalovirus UL131-128 genes are indispensable for virus growth in endothelial cells and virus transfer to leukocytes. J Virol 78:10023–10033. https://doi.org/10.1128/ JVI.78.18.10023-10033.2004 14. Waldman WJ, Roberts WH, Davis DH et al (1991) Preservation of natural endothelial cytopathogenicity of cytomegalovirus by propagation in endothelial cells. Arch Virol 117:143–164 15. Sinzger C, Digel M, Jahn G (2008) Cytomegalovirus cell tropism. Curr Top Microbiol Immunol 325:63–83 16. Marchini A, Liu H, Zhu H (2001) Human cytomegalovirus with IE-2 (UL122) deleted fails to express early lytic genes. J Virol 75:1870–1878. https://doi.org/10.1128/ JVI.75.4.1870-1878.2001 17. Adler H, Messerle M, Koszinowski UH (2003) Cloning of herpesviral genomes as bacterial artificial chromosomes. Rev Med Virol 13:111–121. https://doi.org/10.1002/rmv. 380 18. Borst EM, Hahn G, Koszinowski UH, Messerle M (1999) Cloning of the human cytomegalovirus (HCMV) genome as an infectious bacterial artificial chromosome in Escherichia coli: a new approach for construction of HCMV mutants. J Virol 73:8320–8329 19. Hahn G, Khan H, Baldanti F et al (2002) The human cytomegalovirus ribonucleotide reductase homolog UL45 is dispensable for growth

Distinct Properties of Human Cytomegalovirus Strains in endothelial cells, as determined by a BAC-cloned clinical isolate of human cytomegalovirus with preserved wild-type characteristics. J Virol 76:9551–9555. https://doi.org/ 10.1128/jvi.76.18.9551-9555.2002 20. Sinzger C, Hahn G, Digel M et al (2008) Cloning and sequencing of a highly productive, endotheliotropic virus strain derived from human cytomegalovirus TB40/E. J Gen Virol 89:359–368. https://doi.org/10.1099/vir.0. 83286-0 21. Stegmann C, Rothemund F, Laib Sampaio K et al (2019) The N terminus of human cytomegalovirus glycoprotein O is important for binding to the cellular receptor PDGFRα. J Virol 93:e00138. https://doi.org/10.1128/ JVI.00138-19 22. Yu D, Smith GA, Enquist LW, Shenk T (2002) Construction of a self-excisable bacterial artificial chromosome containing the human cytomegalovirus genome and mutagenesis of the diploid TRL/IRL13 gene. J Virol 76:2316–2328. https://doi.org/10.1128/jvi. 76.5.2316-2328.2002 23. Stanton RJ, Baluchova K, Dargan DJ et al (2010) Reconstruction of the complete human cytomegalovirus genome in a BAC reveals RL13 to be a potent inhibitor of replication. J Clin Invest 120:3191–3208. https:// doi.org/10.1172/JCI42955 24. Laib Sampaio K, Weyell A, Subramanian N et al (2017) A TB40/E-derived human cytomegalovirus genome with an intact US-gene region and a self-excisable BAC cassette for immunological research. BioTechniques 63:205–214. https://doi.org/10.2144/000114606 25. Wilkinson GWG, Davison AJ, Tomasec P et al (2015) Human cytomegalovirus: taking the strain. Med Microbiol Immunol 204:273–284. https://doi.org/10.1007/ s00430-015-0411-4 26. Martı´-Carreras J, Maes P (2019) Human cytomegalovirus genomics and transcriptomics through the lens of next-generation sequencing: revision and future challenges. Virus Genes 55:138–164. https://doi.org/10. 1007/s11262-018-1627-3 27. Mattick C, Dewin D, Polley S et al (2004) Linkage of human cytomegalovirus glycoprotein gO variant groups identified from worldwide clinical isolates with gN genotypes, implications for disease associations and evidence for N-terminal sites of positive selection. Virology 318:582–597. https://doi.org/10. 1016/j.virol.2003.09.036 28. Pignatelli S, Dal Monte P, Landini MP (2001) gpUL73 (gN) genomic variants of human cytomegalovirus isolates are clustered into

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four distinct genotypes. J Gen Virol 82:2777–2784. https://doi.org/10.1099/ 0022-1317-82-11-2777 29. Zhang L, Zhou M, Stanton R et al (2018) Expression levels of glycoprotein O (gO) vary between strains of human cytomegalovirus, Influencing the Assembly of gH/gL Complexes and Virion Infectivity. J Virol 92: e00606. https://doi.org/10.1128/JVI. 00606-18 30. Adler B, Scrivano L, Ruzcics Z et al (2006) Role of human cytomegalovirus UL131A in cell type-specific virus entry and release. J Gen Virol 87:2451–2460. https://doi.org/10. 1099/vir.0.81921-0 31. Wang D, Shenk T (2005) Human cytomegalovirus UL131 open reading frame is required for epithelial cell tropism. J Virol 79:10330–10338. https://doi.org/10.1128/ JVI.79.16.10330-10338.2005 32. Lehmann C, Falk JJ, Bu¨scher N et al (2019) Dense bodies of a gH/gL/UL128/UL130/ UL131 pentamer-repaired towne strain of human cytomegalovirus induce an enhanced neutralizing antibody response. J Virol 93: e00931–e00919. https://doi.org/10.1128/ JVI.00931-19 33. Scrivano L, Sinzger C, Nitschko H et al (2011) HCMV spread and cell tropism are determined by distinct virus populations. PLoS Pathog 7: e1001256. https://doi.org/10.1371/journal. ppat.1001256 34. Wang D, Yu Q-C, Schro¨er J et al (2007) Human cytomegalovirus uses two distinct pathways to enter retinal pigmented epithelial cells. Proc Natl Acad Sci U S A 104:20037–20042. https://doi.org/10. 1073/pnas.0709704104 35. Wu K, Oberstein A, Wang W, Shenk T (2018) Role of PDGF receptor-α during human cytomegalovirus entry into fibroblasts. Proc Natl Acad Sci U S A 115(42):E9889. https://doi. org/10.1073/pnas.1806305115 36. Murrell I, Wilkie GS, Davison AJ et al (2016) Genetic stability of bacterial artificial chromosome-derived human cytomegalovirus during culture in vitro. J Virol 90:3929–3943. https://doi.org/10.1128/JVI.02858-15 37. Wilkinson GWG, Tomasec P, Stanton RJ et al (2008) Modulation of natural killer cells by human cytomegalovirus. J Clin Virol 41:206–212. https://doi.org/10.1016/j.jcv. 2007.10.027 38. Sinzger C, Knapp J, Plachter B et al (1997) Quantification of replication of clinical cytomegalovirus isolates in cultured endothelial

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cells and fibroblasts by a focus expansion assay. J Virol Methods 63:103–112 39. Jiang XJ, Adler B, Sampaio KL et al (2008) UL74 of human cytomegalovirus contributes to virus release by promoting secondary envelopment of virions. J Virol 82:2802–2812. https://doi.org/10.1128/JVI.01550-07 40. Silva MC, Yu Q-C, Enquist L, Shenk T (2003) Human cytomegalovirus UL99-encoded pp28 is required for the cytoplasmic envelopment of tegument-associated capsids. J Virol 77:10594–10605. https://doi.org/10.1128/ jvi.77.19.10594-10605.2003 41. Tomasec P, Wang ECY, Davison AJ et al (2005) Downregulation of natural killer cellactivating ligand CD155 by human

cytomegalovirus UL141. Nat Immunol 6:181–188. https://doi.org/10.1038/ni1156 42. Abdellatif MEA, Sinzger C, Walther P (2018) Investigating HCMV entry into host cells by STEM tomography. J Struct Biol 204:406–419. https://doi.org/10.1016/j. jsb.2018.10.007 43. Schmolke S, Kern HF, Drescher P et al (1995) The dominant phosphoprotein pp65 (UL83) of human cytomegalovirus is dispensable for growth in cell culture. J Virol 69:5959–5968 44. Dunn W, Chou C, Li H et al (2003) Functional profiling of a human cytomegalovirus genome. Proc Natl Acad Sci U S A 100:14223–14228. https://doi.org/10.1073/pnas.2334032100

Chapter 3 Using Diploid Human Fibroblasts as a Model System to Culture, Grow, and Study Human Cytomegalovirus Infection Elizabeth A. Fortunato Abstract Primary human diploid fibroblasts are used routinely to study host/pathogen interactions of human cytomegalovirus (HCMV). Fibroblasts’ ease of culture and tremendous permissiveness for infection allow the study of all facets of infection, an abbreviated list of which includes ligand–receptor interactions, activation of cell signaling responses, and dysregulation of the cell cycle and DNA repair processes. Another advantage to fibroblasts’ permissiveness for HCMV is the capability to grow high titer stocks of virus in them. This chapter will discuss the production of viral stocks of HCMV in primary human fibroblasts, commencing with culturing and infection of cells and continuing through harvest, titration (determining the infectious capacity of a particular virus preparation), and storage of viral stocks for use in downstream experiments. Key words Human cytomegalovirus, Culture of primary fibroblasts, Preparation and storage of virus stocks, Titrating viral stocks, Growth of virus in culture

1

Introduction HCMV has a wide range of permissiveness in vivo [1]. Although fibroblasts may not be the first cells that come to mind as clinically relevant to the drastic ramifications seen during congenital infection or transplant rejection, they provide a useful tool for the study of a fully permissive infection in the context of a tissue culture environment. Several compelling reasons to use fibroblasts include the following: (1) they are quite easy to culture and thrive for many passages; (2) they synchronize easily; (3) they can be grown in large quantities with relative ease; and (4) they display all the characteristics of a fully permissive infection, producing the full range of viral antigens and (with laboratory-adapted virus strains) producing high titer stocks of cell-free virus, which is readily harvested from the supernatant of infected cells. For a discussion of the use of other cell types to study distinct HCMV/host interactions see Chapters 2, 4–6.

Andrew D. Yurochko (ed.), Human Cytomegaloviruses: Methods and Protocols, Methods in Molecular Biology, vol. 2244, https://doi.org/10.1007/978-1-0716-1111-1_3, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Fibroblasts are not identical and different sources can be used. In general, primary cells (referred to as normal diploid fibroblasts) are utilized in most experiments. Cells can be obtained from cell banks such as ATCC or from local hospital sources and can be derived either from adult or fetal/newborn tissue. Cells from many different areas of the body, including lung, kidney and foreskin have been successfully used for HCMV studies. Fibroblasts have historically served as a “prototype” for lytic infection in vitro. An attractive characteristic of relatively high multiplicity of infection (MOI) HCMV infections in synchronized (via serum starvation or confluence arrest [2, 3] fibroblasts is a “synchronous” infection (see Note 7 for explanation of MOI). Following a high MOI infection, it is relatively easy to track the entire lytic life cycle in these cells. Initial interactions begin with binding to the cell surface receptors and continue through wholesale necrosis of the cells after 5–7 days (depending upon the virus strain used). As is discussed in Chapter 2, all HCMV strains possess the proteins necessary for interaction with, entry into and subsequent replication in fibroblasts (and [4]). Fibroblasts are also frequently used in virus titration (determining the infectious capacity of virions shed from infected cells). The latter is true even when clinical isolates are utilized or if an experiment is performed in another cell type. This is due to the ease with which all HCMV strains infect fibroblasts and produce assayable plaques. In essence, virus titration in fibroblasts can “level the playing field” for different virus strains when assessing functional virion output. It also allows for the relatively easy assessment of variation in plaque size when measuring the capacity of viral mutants to replicate in comparison to their wildtype parental counterparts. That all fibroblasts are permissive for all HCMV strains permits quick propagation of large-scale, high titer virus preparations within these cells. As is described in Chapter 2 (and [5]), growth in fibroblasts leads relatively quickly (usually within 10 or so passages) to tissue culture “adaptation” of clinical strains, first with loss of RL13, then mutation of the UL128–131A locus and eventual loss of the ULb0 cassette/suite of viral genes important for infection and growth in vivo (and in certain distinct tissue types). The laboratory-adapted strains Towne and AD169, which many laboratories utilize for the bulk of their studies, have undergone these adaptations and thus replicate to high levels in fibroblasts, allowing shedding of large quantities of cell-free virus. However, for this reason, long-term passage of clinical isolates within fibroblasts is not advisable. Clinical strains are highly cell-associated (when compared to the laboratory-adapted Towne and AD169 strains) and require different culturing and collection conditions in order to obtain high titer virus (see Notes 4, 15, and 22 for brief discussion of these differences). Therefore, the bulk of this chapter focuses on growth of the laboratory-adapted strains that are utilized by the

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large majority of laboratories working on HCMV for the bulk of their experiments. This chapter covers the steps needed to produce high titer viral stocks of HCMV for use in experiments. It will cover culturing fibroblasts (including the critical parameters important for maintaining healthy cells), the low multiplicity infection conditions necessary for the production of high titer stocks, harvesting viral supernatants, proper freezing and storage conditions and the titer assay for calculating the number of infectious virions/ml of supernatant. Also discussed will be other options for measuring antigen positivity and viral genomic replication in infected cultures when circumstances preclude the use of plaque assays during an experiment.

2

Materials All procedures described in this chapter will be carried out in a biological safety cabinet to maintain sterile conditions and biohazard abatement. HCMV is a biosafety level 2 pathogen and adequate precautions and disposal techniques must be utilized during all handling of the virus and virus-infected cultures. Unless they are presterilized by the manufacturer, all solutions should be either filter-sterilized or autoclaved before use.

2.1 Cells and Culture Media

1. Human diploid fibroblasts. As mentioned in the introduction, these cells can be obtained from several sources, the most common being American Type Culture Collection (ATCC). Many labs derive cells from tissue obtained from neonatal foreskins. 2. Culture media for fibroblasts: Eagle’s minimal essential medium (MEM, Gibco BRL) supplemented with final concentrations of 10% fetal bovine serum (FBS), penicillin (200 U/ ml), streptomycin (200 μg/ml), L-glutamine (2 mM), and amphotericin B, that is, Fungizone (1.5 μg/ml) (see Note 1). 3. Parameters for maintenance of cells in culture: In general, most tissue culture cells are maintained at 37  C in a humidified atmosphere containing 5% CO2.

2.2 Additional Solutions

1. Phosphate buffered saline (PBS): For 1 l add 8 g NaCl, 0.2 g KCl, 1.44 g Na2HPO4, and 0.24 g NaH2PO4 to 950 ml distilled H2O. Adjust the pH to 7.4, bring to volume, then autoclave. 2. 2.5% Trypsin: dilute tenfold in sterile PBS to a working 0.25% stock (store refrigerated at 4  C when not in use). Undiluted stock should be stored at 20  C.

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3. DMSO for frozen storage of virus stocks: add stock to final concentration of 1% in supernatant (see Note 2). 4. Agarose (2%) diluted in water for overlays. Agarose is added to water and then autoclaved to sterilize (see Note 3). 2.3

Plasticware

The following plasticware is needed (and should either be purchased sterile from the manufacturer or autoclaved): 1. Large format tissue culture flasks (~185 cm2 growth area). 2. 24-well tissue culture plates. 3. Conical tubes for spinning and freezing viral stocks. 4. Microcentrifuge tubes for serial dilutions. 5. Cotton-plugged barrier pipette tips for serial dilutions. 6. Serological pipettes supernatants.

2.4 Other Necessary Equipment

for

transferring

media

and

virus

The following equipment is also necessary to carry out these experiments: 1. A clinical centrifuge. 2. A hemocytometer for counting cells. 3. A tissue culture incubator for culturing cells and growing stocks. 4. A biological safety cabinet for all culturing of cells and handling of virus. 5. A 80  C freezer for storing stocks.

3

Methods

3.1 Growing Viral Stocks on Fibroblasts

1. Trypsinize and seed actively dividing, low passage human fibroblasts onto T185 flasks (~185 cm2 seeding area). Seed approximately 3.5  106 cells into each flask. In order to yield roughly 200 ml of supernatant per harvest, between 10 and 15 flasks should be seeded. Seed cells the night before they are to be infected. This allows the cells a chance to settle and adhere (see Notes 4–6). 2. The following morning infect the cells at a low MOI (0.02 is suggested). This low MOI allows ample time for cell to cell spread of the propagating virus (see Note 7). 3. Remove the inoculation media ~8–10 h postinfection (hpi). Each flask should be refed with 17–18 ml of fresh media (see Note 8). 4. Observe the monolayer each day, refeeding the cells every 2–3 days as necessary (see Note 9).

Diploid Fibroblasts as a Model System for HCMV Infection

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5. Watch for when the monolayer displays approximately 80% cytopathic effect (CPE). CPE is defined as enlarged/rounded cells containing clear virus replication centers visualized by light microscopy. At this point, refeed the cells again with 17 ml/flask in preparation for your first harvest. 6. After 3–4 days harvest the supernatants from the flasks and refeed your cells. If the monolayers have remained intact and there are only a small number of lysed cells in the flasks (see Note 10), another harvest should be possible in 2–3 days. Because a significant proportion of the cells are starting to die at this point, the titer from the second stock may be lower than the first stock. 7. For laboratory-adapted virus stocks, the supernatant collected from the infected cells can be cleared by centrifugation in a tabletop centrifuge. Supernatant can be dispersed equally in 50 ml conical tubes and spun at approximately 1500 rpm (500  g) for 5–10 min to pellet any cellular debris that might be present. If debris is still present after a first clearing spin, the supernatants can be transferred to new tubes and spun again for an additional 5 min. 8. Transfer the cleared supernatant to a T185 flask in order to pool the entire stock, being careful to avoid the cellular debris. Determine how many milliliters are in the flask. 9. Add 10% of a 10% DMSO stock made up in growth media to the supernatant, for example if you have 200 ml of supernatant, add 20 ml of your 10% DMSO stock. This yields a final 1% DMSO concentration in the virus stock (see Note 11). 10. Once the DMSO is added, cap the flask and mix well. Aliquot the supernatant into freezer-safe conical tubes (15 ml conical tubes are suggested) and store at 80  C (see Notes 12–15). 3.2 Titration of Virus Stocks

After virus stocks have been prepared and before they can be used in experiments the number of functional virions (or plaque-forming units—pfu) per ml must be determined (see Note 16). The following steps should be followed to determine the pfu/ml. 1. The day before beginning the titration process, seed a total of 1.5–1.8  106 fibroblasts into a 24-well plate. The cells should be resuspended in 24 mls of media and each well should receive 1 ml (see Note 17). 2. The following morning quickly thaw a 1 ml aliquot of frozen virus stock in a 37  C water bath. In complete media, dilute this stock serially, using ten-fold dilutions (see Note 18). 3. Aspirate the media from the wells of the 24-well plate seeded the previous day. Pipet 200–250 μl of each dilution into a separate well of the plate. Record and use a logical pattern

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when plating the dilutions (see Note 19). Rock the plate gently to ensure even coverage of the monolayers in each well. 4. After the dilutions are plated, replace the 24-well plate back into the incubator. Allow this inoculum to adsorb for 4–6 h (see Note 20). 5. Check the monolayers after 4–6 h of incubation; they should all be firmly attached. 6. Place an agarose overlay onto the wells. Final concentration of the agarose should be ~0.25% in media (diluted from a 2% stock made up in water; 3 ml agarose +21 mls media) (see Note 21). 7. Place 1 ml of overlay into each well (place directly over the supernatant already in the wells, DO NOT aspirate!). Allow the agarose to solidify (this usually takes 5–10 min at room temperature (RT) in the biosafety hood) and replace the plate into the 37  C incubator. 8. Monitor the plate for plaques every few days. Plaques should start appearing clearly on the lower dilutions between 4 and 5 days. Count the plaques twice between days 7 and 11 postplating (see Note 22). Ideally between 20 and 50 plaques per well will be counted in order to make accurate calculations. Calculate the titer based upon the number of plaques, the dilution and the amount of supernatant you plated (see Notes 23 and 24). 9. When are titer/plaque assays performed? Scenarios include determining the capability of mutant viruses to grow in culture versus their wild-type counterparts or to determine if the replication capacity of a given strain varies in different cell types. There are two accepted methods for performing growth curves, single round and multistep curves. In single round growth curves, all cells are infected at the outset of the experiment and then one of two methods of harvesting and assessing the output from the cells can be used. The first method determines “cumulative” yields (i.e., taking a small aliquot each day and leaving the media on for the entire experiment). The second method assesses how much virus the cells are releasing “each day.” In this method the entire supernatant is harvested each day and then total output for each day of the experiment is calculated. The second technique for performing growth curves is multistep curves. In these experiments the initial infection is done at a low MOI (i.e., between 0.01 and 0.1). This second method allows for the determination of whether the initial virus is capable of replicating and releasing additional virus that can infect another cell.

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4

45

Notes 1. Different laboratories use variations of this media including changes in base media (some use Dulbecco’s modified Eagle’s media), amount of serum added (anywhere from 5% to 20%), addition (or not) of antibiotics, and supplementation (or not) of extra glutamine. The use of heat-inactivated serum is recommended, since the complement present in serum can be detrimental to certain cell types in culture. Heat inactivated serum may also support increased viral growth in all cell types. It should be noted that some cell types do not grow well in the presence of Amphotericin B. Amphotericin B can be omitted, but cells must be monitored more closely for fungal contamination. 2. A stock of 10% DMSO solution in growth media can be made just prior to freezing. This solution can then be diluted tenfold (to a final concentration of 1%) into collected supernatants. 3. When not in use the 2% agarose stock can be allowed to solidify and stored at RT. Simply heat the stock in a microwave to liquefy and cool (in a 56  C water bath) a minimum of 1 h before use. Overlaying of a 24-well plate requires 24 ml of overlay medium (3 ml of melted 2% agarose mixed with 21 ml media for a ~0.25% final concentration of agarose). 4. The cell type in which to grow virus stocks should be carefully considered. Generally, HCMV laboratory-adapted virus strains are grown on primary fibroblasts. Other primary cells might be preferable for growing a clinical strain (i.e., endothelial cells to maintain the integrity of the RL13, UL128–131A, and ULb’ regions). Clinical isolates can be grown for a few passages on fibroblasts, but care must be used, as many clinical isolates very quickly lose/mutate these genes. In some strains this loss of integrity has been reported to occur as early as 1–2 passages after infection of fibroblasts. PCR assays should be performed to ensure that these genes are still intact within any clinical virus preparations. In addition, virus strains containing mutations in essential genes (e.g., pp71 and IE86) often must be grown on complementing cell lines in order to obtain high titer stocks [6, 7]. It is good practice when new cells are being used for experiments that they should be screened for the presence of viral gene expression and replication prior to use (either by immunofluorescence analysis and/or by PCR amplification of a region of the viral genome). Lastly, although this might seem obvious to the experienced herpes virologist, the cytomegaloviruses are highly species restricted and only human cells can be used for culturing HCMV.

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5. The condition of the cells in culture is extremely important to obtain high titer stocks. In essence, treat the cells delicately, affectionately and well! Maintaining cell cultures at sub-confluence is important to the vigor of the cells. DO NOT let cultures remain confluent for more than 1 or 2 days. Confluent cells are not exceptionally healthy and will yield suboptimal results. Cells should be refed 2–3/week and split 1:3 approximately once a week or when they are near confluence. 6. The number of cells seeded per flask should be on the high side (approximately 3.5  106/flask). In addition, anecdotal evidence has found that lower passage cells yield higher titer stocks. This is likely linked to the decreased growth of the primary cells as they age. Lower passage cells divide more regularly and are healthier in general. 7. What is an MOI? Multiplicity of infection refers to the plaqueforming units applied to each cell. In absolute terms, one can calculate the percent cells that will be infected at a given MOI using the formula [percent cells infected ¼ 1eMOI]. A synchronized infection is more likely to be produced by using a higher MOI (in general between 2 and 10, depending upon the cells). A high MOI ensures that all the cells are infected and produce viral antigens simultaneously. This is particularly the case if the cells have been previously synchronized in G0 versus an asynchronous infection [8, 9]. Producing high titer stocks requires starting the infection at a quite low MOI. A low MOI offers the virus a chance to “simmer” and replicate multiple times before the entire monolayer is lysed. If a high MOI were used to initiate the stock preparation (as might well be used during an actual experiment), only one round of infection will occur without any cell-to-cell spread of the virus. High MOI infections generally produce high particle to pfu ratios (see Note 16 for an explanation), which is not necessarily desirable in a stock preparation. 8. Although helpful when growing clinical isolates, leaving the inoculum of a lab-adapted virus strain on overnight can sometimes lead to the infection proceeding too quickly with subsequent lower virus yields. Maintaining a relatively low volume of media in flasks produces a more concentrated stock. 9. The infection should proceed relatively slowly at first, with the first harvest occurring between 7 and 10 days pi. Keep an eye on the monolayer. After a few days distinct “foci” of infected cells will be seen. As the infection proceeds these foci will coalesce and eventually the entire monolayer will exhibit strong cytopathic effect. When growing clinical isolates, it is often helpful to trypsinize and reseed the cells onto the same flasks

Diploid Fibroblasts as a Model System for HCMV Infection

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after large foci are observed. This seems to give the infection a “jumpstart” by spreading the infected cells across the entire flask. 10. Timing of the initial harvest is critical. If the supernatant is harvested too early the virus yield will be low. In addition, while waiting 3–4 days before the first harvest may seem like a long time, the virus in the supernatant will not suffer at 37  C. The best titer yields are produced when this first harvest is allowed to “simmer” on the cells. 11. Use care when adding the DMSO. Pipet slowly into the flask of supernatant while swirling the flask to avoid “flashing” the DMSO and creating “hotspots” of high concentrations, which could damage the virions. 12. Freeze 1 ml, 5 ml, and 10 ml aliquots so that appropriate quantity aliquots are available depending on requirements for future experiments. It is not acceptable to freeze and thaw virus stocks more than once. The virus loses significant viability following repeated freeze–thaw cycles rendering MOI calculations invalid. 13. “Mock-infected supernatant” for use as a control for infection can be created in parallel with virus stock preparation. Collect “spent” media that has been incubated for 2–4 days from an equivalent set of flasks of uninfected fibroblasts in culture. This “mock-infected supernatant” will have serum and a significant component of the chemokines/cytokines that are normally secreted by growing cells. This is referred to as “conditioned media” and is added in equivalent amounts to mock-infected cultures as virus is added to infected cultures. 14. Experiments that are sensitive to the effects of serum in virus stocks can benefit by creating virus stocks in serum free medium. In this case the cells are washed several times in warm PBS and then incubated in serum free media prior to collecting virions, however, this is a suboptimal condition for cell growth. An alternative approach is to pellet virions from the supernatant with a high-speed spin in an ultracentrifuge (23,000 rpm [90,000  g] for 75 min at 10  C). The pelleted virions can be washed in PBS to remove any residual serum and then respun under the same conditions. Some labs pellet these virions through a 25% sucrose “cushion” made in PBS to create serum/media free virus. Sorbitol can also be used as a cushion to purify virus. The theory is to cushion the envelope from the blows of multiple rounds of pelleting. The sucrose cushion may also serve to separate less dense cellular debris (which will float) from the denser pelleted particles. Once pelleted and washed, these particles can be resuspended either in an equivalent amount

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of serum-free media (or PBS) or, if more concentrated stocks are required, less volume can be used to resuspend. Note that this pelleting will not separate full/infectious virions from the other particles secreted by permissive cells, which include noninfectious enveloped particles (NIEPS) and dense bodies (DB). If pure virions are required, particles must be layered onto a gradient (e.g., glycerol tartrate gradients), which will isolate the three distinct bands of particles [10]. Purity of the fractions from these gradients should always be assessed before use. 15. Unlike laboratory-adapted strains, clinical isolates are very cellassociated. Additional steps must be taken when harvesting clinical virus stocks. Rather than harvesting only the supernatant, the cells must be harvested as well. First, the supernatant should be harvested and spun as described above. The cellular debris from this spin should be saved. In addition, trypsinize and harvest the cells on the flask. Spin these harvested cells and collect the pelleted cells. Add this cell pellet to the other cells derived from the supernatant clearing spin by resuspending in a small volume of media (5 ml or less). Sonicate all these cells (2 min on ice at an amplitude of 30% in a cuphorn sonicator). Alternatively, the resuspended cell pellet can be put through two rounds of freeze–thaw lysis (alternating between dry ice and 37  C baths for 5 min each). Spin out the debris in a tabletop centrifuge at ~1500 rpm (500  g) for 10 min. Pipet off the 5 ml of media and add it to the reserved supernatant, being careful to avoid the debris at the bottom. Respin if the supernatant still looks cloudy. When clear, pellet all the particles out of the supernatant in an ultracentrifuge (as described in Note 14) and resuspend in 1/10–1/20th the initial volume. This stock can now be titrated on fibroblasts. 16. A plaque forming unit (pfu) is defined as a functional virion that is capable of infecting a cell and producing enough virus within that cell to infect the cells that are surrounding it (i.e., capable of creating an infected plaque in a monolayer of uninfected cells). Note that the size of these plaques and the rapidity with which they are formed after infection of the monolayer can be used as a phenotypic gage when comparing virus mutants to their parental counterparts. Pfu is not a measurement of the number of viral particles released into the supernatant; the infectious virions are a small proportion of the total particles in the supernatant. There are many other particles interacting with and penetrating infected cells; these additional particles could elicit a response. These particles include NIEPs and DBs. The particle to pfu ratio for HCMV can be as high as 100:1 [11].

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17. Be aware that not all multiwell plates are the same. Occasionally, it appears that plates are unevenly coated or that there may be no coating on several wells. Corning plates seem to be the most consistent; this issue should be addressed prior to use and the best plates used for experimentation. Be careful when you are seeding cells on these small circular well plates. Gently rock the plates, do not swirl in a circular motion, as doing so will clump the cells into the middle of the well. The recommended “magic shake” consists of holding the plate in front of your body and gently moving it back and forth away from your body a few times followed by gently moving it back and forth across your body a few more times (sometimes a little dance does not hurt either!). 18. Make 101 to 106 dilutions in 1 ml each. It is very important to mix each dilution thoroughly before removing the next aliquot. Always change tips between tubes to avoid cross contamination. 19. As a general rule of thumb, when plating dilutions aspirate media from only half of the plate at a time in order to avoid the cells drying out. It is advisable to use at least duplicate (if not triplicate) wells for each dilution. This improves the accuracy, as the counts from the multiple wells of a given dilution will be averaged. 20. DO NOT allow the cells to dry out during this incubation. Since this is a relatively small amount of media, rock the plate every hour to ensure cells are continuously covered. 21. After completing the dilution series melt the agarose stock in a microwave and then place into a 56  C water bath to cool. This will ensure it will have cooled enough to add to the cells later without causing them harm. Work quickly making the agarose dilution in media. If it is too cool it will solidify in the pipette. Carboxymethyl cellulose is an alternate choice of overlay material. 22. Titrating clinical isolates may require a longer period of incubation. These strains often plaque slowly and only produce small plaques. 23. A sample calculation: Count plaques on the 105 dilution wells which were seeded with 250 μl of diluted stock each. Three wells have counts of 18, 22, and 20 plaques, respectively. The average number of plaques is therefore 20. The per ml titer is calculated as follows: 20 plaques  105  4 (because only 250 μl were plated) ¼ 8  106 pfu/ml. 24. One alternative to performing plaque assays to assess the titer of a virus stock is to assess the immediate early protein positivity (IE+) of cells after several hours of incubation. A second alternative is to compare the efficiency of a given virus strain

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(or mutant) to replicate its genome within a given cell type using quantitative PCR. It should be noted that neither of these assays establish whether the initiating virion that has entered a cell has the capacity to proceed through an entire life cycle and produce more infectious virions. Despite this caveat, if you know your virus replicates and makes appropriate new viral particles and you are just making stocks, this is a convenient and quick method to determine titer. These types of assays can also be utilized when a virus is incapable of actually producing plaques (e.g., mutant viruses that do not shed virus). References 1. Sinzger C, Jahn G (1996) Human cytomegalovirus cell tropism and pathogenesis. Intervirology 39:302–319 2. Casavant NC, Luo MH, Rosenke K, Winegardner T, Zurawska A, Fortunato EA (2006) Potential role for p53 in the permissive life cycle of human cytomegalovirus. J Virol 80:8390–8401 3. Fortunato EA, Spector DH (1998) p53 and RPA are sequestered in viral replication centers in the nuclei of cells infected with human cytomegalovirus. J Virol 72:2033–2039 4. Zhou M, Lanchy JM, Ryckman BJ (2015) Human cytomegalovirus gH/gL/gO promotes the fusion step of entry into all cell types, whereas gH/gL/UL128-131 broadens virus tropism through a distinct mechanism. J Virol 89:8999–9009 5. Wilkinson GW, Davison AJ, Tomasec P, Fielding CA, Aicheler R, Murrell I, Seirafian S, Wang EC, Weekes M, Lehner PJ, Wilkie GS, Stanton RJ (2015) Human cytomegalovirus: taking the strain. Med Microbiol Immunol 204:273–284 6. Bresnahan WA, Hultman GE, Shenk T (2000) Replication of wild-type and mutant human cytomegalovirus in life-extended human diploid fibroblasts. J Virol 74:10816–10818

7. Sanders RL, Clark CL, Morello CS, Spector DH (2008) Development of cell lines that provide tightly controlled temporal translation of the human cytomegalovirus IE2 proteins for complementation and functional analyses of growth-impaired and nonviable IE2 mutant viruses. J Virol 82:7059–7077 8. Fortunato EA, Sanchez V, Yen JY, Spector DH (2002) Infection of cells with human cytomegalovirus during S phase results in a blockade to immediate early gene expression that can be overcome by inhibition of the proteasome. J Virol 76:5369–5379 9. Salvant BS, Fortunato EA, Spector DH (1998) Cell cycle dysregulation by human cytomegalovirus: influence of the cell cycle phase at the time of infection and effects on cyclin transcription. J Virol 72:3729–3741 10. Irmiere A, Gibson W (1983) Isolation and characterization of a noninfectious virion-like particle released from cells infected with human strains of cytomegalovirus. Virology 130:118–133 11. Benyesh-Melnick M, Probstmeyer F, McCombs R, Brunschwig JP, Vonka V (1966) Correlation between infectivity and physical virus particles in human cytomegalovirus. J Bacteriol 92:1555–1561

Chapter 4 Using Primary Human Cells to Analyze Human Cytomegalovirus Biology Emma Poole, Ian Groves, Sarah Jackson, Mark Wills, and John Sinclair Abstract The extensive tropism of human cytomegalovirus (HCMV) results in the productive infection of multiple cell types within the human host. However, infection of other cell types, such as undifferentiated cells of the myeloid lineage, give rise to nonpermissive infections. This aspect has been used experimentally to model latent infection, which is known to be established in the pluripotent CD34+ hematopoietic progenitor cell population resident in the bone marrow in vivo. The absence of a tractable animal model for studies of HCMV has resulted in a number of laboratories employing experimental infection of cells in vitro to simulate both HCMV lytic and latent infection. Herein, we will focus on the techniques used in our laboratory for the isolation and use of primary cells to study aspects of HCMV latency, reactivation, and lytic infection. Key words Fibroblasts, Endothelial cells, Epithelial cells, Monocytes, CD34+ cells, HCMV latency, Myeloid differentiation, Primary cell Isolation

1

Introduction HCMV is a species-specific pathogen, which has precluded extensive analyses in animal models [1]. Consequently, many studies have employed the use of cell lines to provide valuable insights into various aspects of HCMV biology. However, the usefulness of cell lines can be compromised by their biological relevance to HCMV as a human pathogen and, as such, much current research has focused on the ex vivo use of primary human cells and, more recently, the development of humanized mouse models [2] . In the human host, latency can be established in the CD34+ hematopoietic progenitor cells. Despite the pluripotency of this population, the carriage of viral genomes appears to be restricted to the cells of the myeloid lineage. Viral genome carriage occurs in the absence of the normal lytic transcription program/virus reactivation and virus production is observed upon terminal

Andrew D. Yurochko (ed.), Human Cytomegaloviruses: Methods and Protocols, Methods in Molecular Biology, vol. 2244, https://doi.org/10.1007/978-1-0716-1111-1_4, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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differentiation of these myeloid progenitors to a macrophage or dendritic cell (DC) phenotype [3]. In this chapter, we describe methods routinely used in HCMV research to elucidate the mechanisms that regulate HCMV latency and reactivation. In general, we use two primary myeloid cell types. One is CD34+ cells isolated from G-CSF mobilized patients after leukapheresis and the other is the CD14+ compartment of peripheral blood; both are sites of natural HCMV latency and both can also be infected in culture to establish experimental latency. Additionally, CD34+ cells and monocytes can be differentiated into Langerhans DCs or interstitial DCs/macrophages, respectively, to reactivate latent HCMV; both from natural latency or experimental latency. The advantage of CD14+ monocytes is their relative ease of isolation from venous blood and their ubiquitous availability. In order to allow us to understand the role of the immune system in the context of both lytic and latent infection, we also isolate autologous dermal fibroblasts from HLA-typed donors to preclude issues arising from class restriction, as experiments involving different cell types from nonmatched donors may lead to nonspecific immune responses. Thus, for example, fibroblasts isolated from a specific donor can be infected with HCMV and then cocultured with immune cells from that donor to analyze HCMVspecific immune responses [4]. Infection of CD34+ and CD14+ cells with HCMV results in a latent infection which is marked by the carriage of viral genome in the presence of expression of the so-called latency associated transcripts, such as UL81-82ast and UL138, but the relative absence of the immediate early (IE) transcripts [5–8] and, importantly, the lack of production of infectious virions. This contrasts with infection in differentiated cells and fibroblasts where the IE RNA and protein are readily detectable to high levels and where infectious virus is produced. For the study of natural latency, HCMV genome carriage can be detected directly ex vivo from myeloid cells or progenitors from seropositive individuals using more sensitive methods such as droplet PCR [9, 10]. The techniques described in this report are used routinely in our laboratory to underpin our research programmes analyzing the precise cellular and molecular mechanisms that govern the establishment, maintenance, and, ultimately, reactivation of HCMV.

2

Materials Ensure that all work with blood products is carried out in accordance with local safety guidelines and with appropriate ethical consideration and permission/regulatory approval. Isolation of blood cell populations are routinely carried out in a Class II

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Microbiological Safety Cabinet (Class II MSC) for user protection and sterility, usually located in a Containment Level 2/3 (CL2/3) laboratory (UK) or Biosafety Level 2 (BSL2, US). 2.1 Mononuclear Cell Isolation

1. Appropriate venipuncture equipment (see Note 1). 2. Sodium Heparin 5 ml vials (1000 U/ml) (see Note 2). 3. Density gradient media—1.077 g/ml (e.g., Histopaque, Lymphoprep, Ficoll-Paque). 4. Endotoxin-free DPBS (Dulbecco’s phosphate buffered saline) without calcium and magnesium. 5. Centrifuge with swinging bucket rotor and no-brake option to allow slow reduction of speed and prevent cell resuspension when the brake is applied (50 ml tube capacity). 6. 50 ml polypropylene conical nonskirted tubes. 7. Sterile plasticware (10 ml and 25 ml serological pipettes and 3 ml Pasteur pipettes). 8. Apheresis cone (Blood transfusion product) [11]. 9. Endotoxin-free PBS with calcium and magnesium.

2.2 Magnetic Isolation of CD34+ and CD14+ Cells

1. Magnetic-activated cell sorting (MACS) magnet (Miltenyi). 2. 30 μm preseparation filter (Miltenyi). 3. LS columns (all Miltenyi Biotec) or an AutoMACS Pro (located in a Class II MSC for sterility) (Miltenyi). 4. MACS chill 15 or 50 racks (Miltenyi). 5. AutoMACS column (Miltenyi). 6. AutoMACS washing solution (Miltenyi). 7. Storage solution (Miltenyi). 8. MACS running buffer (see Note 3). 9. DPBS without calcium and magnesium. 10. 50 ml polypropylene conical tubes (Falcon). 11. 14 ml snap-cap polypropylene tubes (Falcon) or 15 ml screwtop conical polypropylene tubes. 12. Sterile plasticware (10 ml and 25 ml serological pipettes; 3 ml Pasteur pipettes; and 20 μl, 200 μl, and 1 ml pipette tips). 13. Freezing medium (Bambanker™). Alternative freezing media—10% DMSO (sterile tissue culture grade, Sigma) with 90% FBS or 60% IMDM (Iscove’s Modified Dulbecco’s Medium, Sigma), 10% DMSO, and 30% Serum replacement (Panexin, Pan Biotech). 14. For isolation of CD14+ monocytes: CD14+ Microbeads (Miltenyi Biotec)—contains 2 ml CD14 MicroBeads for 1
109 total cells.

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15. For isolation of CD34+ cells: CD34+ Microbeads UltraPure Kit (Miltenyi Biotec)—contains 2 ml CD34 MicroBeads UltraPure and 2 ml FcR Blocking Reagent, for 2
109 total cells. 2.3 Dermal Fibroblast Isolation

1. Sterile disposable curved blade scalpel. 2. Sterile fine and blunt-nosed forceps. 3. Sterile straight needle (e.g., 21G sharp and blunt). 4. Glass coverslips (sterilized). 5. Sterile plastic petri dish (10 cm diameter). 6. Tissue culture grade plasticware: 6-well plates. 7. Sterile plasticware (3 ml Pasteur pipettes and 10 ml and 25 ml serological pipettes). 8. Sterile polypropylene universal tube (25 ml) with 20 ml media. 9. DMEM-high glucose (Dulbecco’s Modified Essential Medium) supplemented with 10% FBS (fetal bovine serum— see Note 4) and 100 U/ml Penicillin and 100 μg/ml Streptomycin. 10. Skin punch biopsy (see Note 5).

2.4 Infection and Differentiation

1. Prepared cells (see Subheading 3). 2. Iscoves medium: (500 ml Iscoves Modified Eagle’s Medium (IMEM) supplemented with 50 ml FCS and 75 ml Horse serum as well as 5 ml pen/strep. Medium will look cloudy). 3. Phorbol 12-myistate 13-acetate (PMA). 4. Hydrocortisone. 5. X-Vivo15 medium (Lonza Biowhittaker™) supplemented with 2.5 mM L-glutamine. 6. Cytokines for Langerhans cell differentiation: TGF-beta, TNF-alpha, SCF, Flt-3 L, and GM-CSF (all high purity and endotoxin levels 95% purity). Visual representations of these gradients including cellular distribution within both the Ficoll-Hypaque density gradient and the Percoll density gradient are available in Figs. 1, 2 and 3.

3.1 Donation Preparation and Venipuncture Blood Donation

1. Add 2 mL of sterile EDTA into the tip of each 4
60 mL syringes using a repeater pipet (see Note 5). 2. Add 10 mL RPMI to 8
50 mL conical tubes. 3. Warm Ficoll-Hypaque temperature (RT).

and

1%

HSRPMI

to

room

4. Using materials outlined in Subheading 2.1 and in accordance with laboratory and IRB specifications and requirements, perform venipuncture for blood donation. 3.2 Ficoll-Hypaque Density Gradient 1 (See Figs. 1 and 2)

1. Carefully pour 30 mL of the whole blood into each of the 8
50 mL conical tubes prepared with 10 mL RPMI in Subheading 3.1. 2. Carefully and slowly underlay 12 mL of Ficoll-Hypaque into each conical tube containing 30 mL whole blood and 10 mL RPMI. 3. Centrifuge the gradients at 1320 rpm (370
g) for 30 min at RT. 4. Collect and discard the top 10 mL (plasma) from each tube. 5. Collect each fraction containing total mononuclear leukocytes (buffy coat; the remaining 20 mL), taking care to avoid collection of erythrocytes and the polymorphonuclear leukocytes that sit on top of the pelleted erythrocytes.

Fig. 1 Final Products of Ficoll-Hypaque and Percoll Density Gradients. (a) Representative image of Ficoll-Hypaque centrifugation result. After whole blood is run through the Ficoll density gradient, the least dense fraction (plasma) is displaced to approximately the top 10 mL of the 50 mL conical tube. The cell fraction that spins into the mid-range density is the mononuclear cell fraction, comprised of approximately the middle 20 mL of the conical. Mononuclear cells are typically found within the middle of this fraction and may stick to the wall of the conical tube. The denser polymorphic cells are found just above the erythrocyte fraction. The cells with greatest density (erythrocytes) make up the last 20 mL of volume within the conical (Note: platelets spin down with the mononuclear cell fraction). (b) Representative image of Percoll density gradient result. After the isolated mononuclear cells are spun through the Percoll gradient, monocytes will be visible at the junction of the 45% and 52.5% layers (at approximately the 4.5 mL mark). The denser mononuclear cells (lymphocytes) and any remaining erythrocytes will be located in the cell pellet at the bottom of the tube

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Fig. 2 Ficoll-Hypaque Gradient. A 12 mL Ficoll-Hypaque underlay is carefully and slowly dispensed beneath 40 mL of whole blood and RPMI to create the density gradient that will allow the isolation of the mononuclear cell fraction. This gradient is then centrifuged, and the distribution of fractions occurs as described in Fig. 1

Fig. 3 Percoll Density Gradient. After the 52.5% density Percoll layer is placed at the bottom of the 15 mL conical, a 45% density layer is slowly and carefully layered on top of the 52.5% bottom later. Mononuclear cells isolated from the Ficoll-Hypaque density gradient are then layered slowly and carefully on top of the 45% density Percoll layer. All gradients are then centrifuged, and the distribution of the monocytes and lymphocytes are shown in the figure

6. Deposit each isolated mononuclear leukocyte fraction into new 8
50 mL conical tubes. 7. Fill each conical tube with saline (0.9%, Sodium chloride irrigation solution). Centrifuge at 1050 rpm (250
g) for 10 min at RT to begin removing platelets. 8. Remove and discard the supernatant. Resuspend one cell pellet with approximately 10 mL saline and combine all tubes. 9. Fill the remaining tubes with saline and centrifuge at 1050 rpm (250
g) for 10 min at RT.

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10. Repeat step 9 until supernatant is clear, and thus all (most) platelets are removed (see Note 6). 11. Resuspend final cell pellet in 12 mL 1% HSRPMI. 12. Move on to the next step (combined mononuclear leukocyte fraction should be in a single 50 mL conical tube). 3.3 Percoll Density Gradient 2: (See Note 7)

1. Vortex stock solution of Percoll. 2. Add 2 mL 10
dPBS to 18 mL Percoll to make what we call 100% Percoll. 3. Make 52.5% fraction (bottom): 10.7 mL 100% Percoll, 9.3 mL 1% HSRPMI. Vortex. 4. Make 45% fraction (top): 9.3 mL 100% Percoll, 11.5 mL 1% HSRPMI. Vortex. 5. Add 4.5 mL of 52.5% fraction to into 4
15 mL conical tubes. 6. Slowly layer 4 mL of the 45% fraction on top of the 52.5% fraction in each conical. 7. Slowly layer 3 mL of final cell suspension from Subheading 3.2 to the top of each 15 mL conical. 8. Centrifuge at 1700 rpm (610
g) for 30 min at RT. 9. Proceed to Subheading 3.4.

3.4 Monocyte Collection and Cell Counting

1. Collect and discard the first 5.5 mL of liquid from the Percoll gradients. 2. Collect the next 4 mL—the monocyte fraction—and dispense into a new 50 mL conical. 3. Once each fraction is collected, fill the 50 mL conical with saline. Centrifuge at 1050 rpm (250
g) for 10 min at RT (Size of cell pellet will vary from donor to donor). Repeat for a total of 2 saline washes. 4. Collect and discard supernatant. Resuspend the monocyte pellet in 10 mL in working media for planned experiments (Typically 1–10% HSRPMI). 5. Count cells by making a 1:10 dilution (i.e., 10 μL cell suspension into 90 μL 1%HSRPMI) in an Eppendorf tube. Load dilution onto a hemocytometer and then via a light microscope count the isolated monocytes, which unlike lymphocytes, turn dark as the cells settle, flatten and stick to the glass hemocytometer. Additionally, trypan blue may be added to the cell dilution at this step in order to visually determine cell viability. 6. After assessing cell numbers, the isolated monocytes are ready to use for experiments. It is optional to store isolated monocytes in a 5% CO2 37  C incubator overnight. However, it is not advisable to let the isolated monocytes sit for more than 24 h (see Note 8).

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Conclusion

HCMV exhibits a unique and complex biology within peripheral blood monocytes that is directly linked to HCMV pathogenesis [8, 12, 19, 22–26]. To study this host–pathogen interaction in our laboratory, we utilize primary human peripheral blood monocytes as an ex vivo model. As discussed above, monocytic leukemic cell lines exist and can be used as a model to study of aspects pathogenmonocyte biology or to study monocyte immunology. Nevertheless, primary cells are the most biologically relevant model system and thus might be the right cell choice for your experimentation. There are disadvantages to primary monocyte usage. For example, genetic manipulation is often limited in primary cells and there is the inherent variability that occurs between different donors. While these challenges can be overcome with experimental design, additional hurdles can present themselves during use of peripheral blood monocytes. Monocytes exist as a dynamic population of cells within the blood and are biologically programmed to differentiate or undergo apoptosis in reaction to external stimuli. Because of these characteristics, primary monocyte isolation requires careful and stringent methodology. While other isolation methods are available, different techniques must be chosen based on the specific question being asked. The dual density gradient centrifugation procedure as outlined in this chapter is a method both sensitive and stringent enough to allow isolation of naı¨ve monocytes from whole blood that have minimal activation and a high purity.

Notes 1. Monocytes are innate immune cells and thereby are highly sensitive to LPS. Amounts of LPS approaching the picogram (pg) level are detectable by monocytes, causing activation and/or cell death [27]. Thus it is very important to use only products that are free of LPS (pyrogen-free). This point includes autoclaved glassware, which is likely sterile, but not LPS free. Glassware needs to be appropriately baked to destroy residual LPS. One can use polymyxin B to minimize the potential impact of LPS [26]. 2. Monocytes stick to polystyrene, but not polypropylene. Therefore, using plastics during isolation composed of polystyrene will increase cell loss and is thus not advisable for use during isolation and manipulation of primary monocytes. 3. This protocol is set for collection of 240 mL of whole blood. Aspects of the protocol can to be altered (scaled) to accommodate different volumes of whole blood. 4. Product name may vary from company to company. The reagent used should contain polysucrose (Ficoll) and sodium diatrizoate, adjusted to a density of 1.077 g/mL in order to

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isolate mononuclear cells from whole blood. The reagent should be tested for endotoxin ( 0.5) dilution on an LB agar plate with 34 μg/mL chloramphenicol and 1% Larabinose. 7. Incubate at 32  C for 1–2 days until average size bacterial colonies are grown. 8. Optional: Pick bacterial colonies from the arabinosecontaining agar plates and replicate plate on LB agar plates with 34 μg/mL chloramphenicol and to plates with 34 μg/ mL chloramphenicol plus 30 μg/mL kanamycin (or 20 μg/mL zeocin when using it as selection marker). The colonies that are resistant to chloramphenicol, but not to kanamycin (or zeocin) contain the BACs with resolved cointegrates. 9. Confirm positive clones with restriction fragment analysis, colony PCR, sequencing, or any other appropriate technique. 3.3 Transfection of HCMV BAC into Human Cells and HCMV Reconstitution

Once the recombinant viral clones have been generated and their identity verified, one may proceed to reconstitute infectious virus out of the viral DNA. Viral DNA is best isolated from approximately 200 mL of bacteria by means of a commercial kit for column purification of BAC DNA (several kits may be used here, for instance, the EndoFree Plasmid Maxi Kit from QIAGEN). This results in a typical yield of 50 μL of DNA preparation with a concentration of DNA at 200–500 ng/μL. The BAC DNA may be then used for transfection of human cells, upon which the viral genes start to get expressed and virus restarts its infectious cycle. There are several commercial transfection reagents available, including Metafectene Pro (Biontex, Germany), Lipofectamine2000 (Invitrogen) or FuGENE® HD (Promega, Germany). Essentially all of these reagents allow for the reconstitution of the infectious process, provided that DNA preparations are of high

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purity and concentration. We will describe here a protocol adapted for Fugene® HD mediated transfection of BAC DNA into MRC-5 cells, a commonly used fibroblast cell line. 1. Grow MRC-5 cells in tissue culture flasks and transfer them to 6-well plates one day prior to transfection. Distribute cells at 50% of maximum confluency (approximately 105 cells per well). 2. On the day of transfection, inspect cells to make sure that cells are spread uniformly across the plate. Allow FuGENE® HD Reagent, DNA, and Opti-MEM to adjust to room temperature and vortex all reagents except BAC DNA prior to use. 3. Mix 2 μg BAC DNA with Opti-MEM in a polystyrene tube (see Note 2). Adjust total volume to 100 μL. 4. Add transfection reagent to the diluted DNA without touching the plastic of the tube at 4:1 FuGENE® HD (μL)–DNA (μg). 5. Mix contents thoroughly, by pipetting up and down ten times. 6. Incubate mixture for 15 min at room temperature. 7. Meanwhile replace the cell culture medium with fresh medium without supplement (FCS or antibiotics). 8. Add transfection complex to cells in a dropwise manner and swirl plate to allow for distribution over the entire plate surface. 9. At 3–8 h (6 h is optimally) after transfection, replace medium with fully supplemented cell culture medium (see Note 4). 10. Following transfection, incubate cells for 48–72 h. 11. Split cells of each transfected well to two T25 flasks. Incubate cells further and monitor by microscopy for areas of viral CPE starting at 7 days post transfection. 12. Once cells become confluent, combine the cells from the T25 flasks into a T175 flask. You will likely need to split the cells of the flask again a few days later (see Note 29). 13. Supernatant of completely infected and lysed cells can be used for virus passaging and virus stock generation. Supernatant that is not used immediately can be stored at
80  C.

4

Notes 1. pEB1097 allows for a ready-to-go approach for the generation of a novel HCMV BAC, because it already flanks the BAC vector with sites of HCMV homology to ORFs US2 and US6 respectively. However, this vector results in recombinants that omit the US2-US6 ORFs, which may be suboptimal for some applications. Therefore, the insertion of target HCMV sequences in a novel BAC vector, like the pBeloBAC11, allows

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for the generation of CMV recombinants that contain the US2-US6 region. In generating such mutants, it is important to consider that the replacement of some nonessential genes of the CMV genomes by the BAC vector sequences is favorable in order to prevent overlength of the resulting recombinant CMV genome. Once cloned in E. coli the missing sequences could be easily readded to the BAC. One integration site commonly used was between the ORFs US2 and US6 [8, 9]. Other chosen insertion sites were between US28 and US29 [25] or between US28 and US34 [14]. 2. Optimum FuGENE® HD–DNA ratio depends on the cell lines that is being transfected. We use 4:1 ratio regularly with multiple cell lines (MRC-5, NIH3T3, HEK293, etc.) in our lab. Also, the amount of the mix should be scaled up from 100 μL, if larger cell culture vessel is used for transfection. 3. We recommend performing the protocol at least in triplicate, using different amounts of the digested recombination plasmid. It is hard to predict if recombination will take place, with which frequency, and which conditions will be optimal. There is a lot of randomness in this approach. Consequently, one should not rely on just a single attempt. 4. FuGENE® HD is relatively mild transfection reagent and is well tolerated by cells. Most cell lines can be incubated overnight with FuGENE®HD. However, in most cases a 5–8 h incubation with the transfection mix is sufficient to obtain high transfection rates. 5. Please note that the additives will lead to a growth arrest of the cells. It is advisable to first establish suitable conditions to determine how fast the growth arrest occurs with the particular passage of fibroblasts that the lab is using. Moreover, the presence of a sufficient amount of cells is required to allow for the spread of the viruses to neighboring cells in order to generate a sufficient amount of progeny virus. 6. Since it is hard to predict which transfection experiment was successful and how much of the recombinant virus may be present in the supernatant, we recommend performing step 2 with supernatant from the independent attempts. Furthermore, it is advisable to add different amounts of the supernatants. As a rule of thumb, we recommend using substantial amounts for the second step (1–5 mL). 7. This time point does not seem to be critical; however, we recommend waiting until the late phase of the infection cycle to guarantee that sufficient amounts of circular genomes are present.

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8. Depending on how much virus is present in the inoculum, it can take 2–3 weeks until complete CPE occurs. 9. It is critical to use highly competent E. coli preparations. One useful strain is DH10B. Transformation of chemically competent bacteria by heat shock treatment was not successful in our hands, as it does not yield sufficient transformation efficiency. Alternatively, commercial chemically competent cells can be used. We recommend using electrocompetent cells either made in the lab or bought from a commercial vendor. 10. We recommend using SnapGene® for primer designing and in silico analysis of the sequences. SnapGene® viewer is a free version of the full software package with certain features missing. It has robust graphical interface that simplify the in silico analysis and manipulation of large sequences such as BACs and whole genome. Online tools such as NEBuilder® can also be used to design primers for Gibson cloning. These tools are also helpful in identifying the annealing temperature for primer and the thermocycler program for fragment amplification. 11. The overlapping sequence between fragments should be 15–25 bp. We recommend using 20 bp overlap to achieve high efficiency of assembly. The annealing temperature of the overlap sequences for all fragments should be greater than 48  C. 12. The vector backbone can also be generated by PCR amplification. This approach allows for the addition of overlapping sequence on the vector backbone. However, we recommend digestion of plasmid to generate linear vectors, as in some cases it might be difficult to reliably amplify the whole vector sequence with PCR. 13. Gel extraction is associated with a low yield of DNA. While small amounts of DNA (Nano grams) are required for Gibson cloning reaction, the eluted amount may not be sufficient. If the yield is low (e.g., 10 ng/mL), upscale the enzyme digestion reaction with larger amounts of plasmid DNA. Furthermore, the yield can be improved by eluting the DNA in a smaller volume than recommended by the manufacturer. We use a 30 μL volume rather than a 50 μL for final DNA elution from QIAquick Spin Columns. 14. The PCR product can be directly purified if the template plasmid do not express the kanamycin resistance gene or other selection markers that will be used to select the final assembled plasmid. If the template plasmid and the final assembled plasmid have the same resistance genes, it is recommended to isolate the required band by gel extraction.

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15. The molar concentration of dsDNA can be calculated with the given formula. Molarity ðnMÞ ¼ ½DNA concentration in ng=μL  ð660 g=mol  fragment size in bpÞ  106 Alternatively, online tools like NEBioCalculator can be used to calculate molarity of dsDNA. If assembly of one fragment with the vector is required, use 1:2 (vector–insert) molar ratio of DNA. However, when assembling multiple fragments in a vector, use equimolar amounts of all fragments. 16. It is recommended that the size of the homology region be 50 bp, but we have used as little as 40 bp homology for recombination. Primers with >100 nucleotides can be ordered commercially. It needs to be considered that these large primers tend to have high annealing temperature that is even greater than the polymerase extension temperature. In the initial cycle of the PCR program, the primers are binding to the template with as little as 20 bp and the annealing temperature is considerably lower for this reaction as compared to the annealing temperature reaction when the whole primer can bind to the template later after some amplification cycles. Due to this reason, we modify our thermocycler program to accommodate for this change in annealing temperature. For instance, primers used to generate the cassette for insertion of stop codons in UL36 gene are shown. Forward Primer: TCAACGACGTGGGGCTTACCTTGCGA ACAGACGGTGCCTCACTTGCCCACGAAGGGCCC CTCGAGCTCTCCCGGGAATTC. Reverse Primer: CTCCGTTCGCGCAGCGCCCTGGGGCC CTTCGTGGGCAAGTGAGGCACCGTCTGTTCGCA ATAGCAGCCAGTGTTACAACCA. The pGP704-I-SceI-kan plasmid was used as a template. In both primers, the nucleotides highlighed in bold and underlined anneal to the template plasmid, while the rest of the sequence carries the target sequence for homologous recombination. Thus, the annealing temparature of the primers to the template is 57  C for HiFi Platinum™ Taq. Therefore, the PCR cycling program used to amplify the construct proceeds as follows: 30 s 95  C/30 s 55–65  C (increase 1  C per cycle)/2 min 68  C for 10 cycles 30 s 95  C/2 min 68  C for 30 cycles 17. Generally, it is recommended to avoid gel purification as it reduces the DNA yield drastically, especially in inexperienced hands. If the gel extration is necessary (e.g., due to mutiple

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PCR bands), upscale the PCR reaction volume and purify using one extraction column and elute the product in the minimum recommended volume (30 μL for QIAquick Spin Columns). 18. Electrocompetent and recombination-proficient E. coli preparations can be stored for several weeks at
80  C (up to 3 months in our hands) without losing the ability for recombination. 19. Time until completion may vary. To catch the cells in the growth phase, start to measure OD after 2 h with 20 min intervals. 20. It is necessary to grow GS1783 bacteria at 32  C, in order to maintain activity of the λ CI857 repressor. Conversely, growing them at 42  C for 15 min activates the promoter and leads to Red recombinase expression, making the bacteria ready to recombine any homologous DNA sequences. This induction should be brief to minimize off-target recombination events. 21. E. coli strains have high competency for transformation at 0.4–0.6 OD600nm. The efficiency drops above 0.6 OD600nm. 22. During log phase, E. coli is present as a spheroplast with a relatively weaker cell wall that makes the bacterial cell sensitive to mechanical damage. It is recommended to handle the cultures gently. Furthermore, it is better to centrifuge the culture with lower g forces for longer period rather than high g forces for shorter times. Keep pipetting to a minimum and centrifuge cells at low g forces, so that tight pellets are not formed. 23. Although, more concentrated electrocompetent cells are prone to mechanical damage, generally they have higher competency. 24. We recommend incubating the transformed bacteria for up to 3 h in the absence of antibiotic selection, in order to allow for segregation of the recombined and nonrecombined copy of the BAC molecules to daughter cells. 25. Incubation in 5 or 10 mL bacterial culture tubes results in optimal air exchange during shaking as compared to the microcentrifuge tubes. This gives better recovery of the bacteria after electroporation. 26. Colony PCR allows for the identification of clones carrying cointegrates in the BAC. This is achieved with diagnostic primers, which generate PCR products of different lengths in the recombined and in the nonrecombined BAC (see Fig. 4). We suggest the use of MangoMix PCR reaction Kit and 20 μL volume per reaction. Before setting the PCR reaction, prepare 3 mL aliquots of LB medium with an adequate amount of selective antibiotic for each tested colony. Pick single colonies with a clean toothpick or yellow pipette tip and resuspend them in the PCR reaction mix. Then put some material from the

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Fig. 4 Presence of recombined and nonrecombined BAC after first step of en passant recombination. Single colonies were picked to perform colony PCR after insertion of GFP gene into the HCMV genome. The lower band at 200 bp represents the WT BAC and the upper band at 2000 bp show the recombined BAC with the en passant cassette inserted

same colonies in the LB medium. Run the PCR reaction with an appropriate program. Incubate liquid cultures overnight at 32  C with 220 rpm shaking. 27. It is recommended to use colony PCR primers that will anneal in the HCMV BAC flanking the recombined sequence and not within the recombined sequence. We have observed that the use of colony primers, which anneal in the inserted sequence show increased false-positive results. Since lambda red recombination occurs through a single-stranded intermediate with a preference for the lagging strand, it is likely that one bacterium will have two BAC copies after recombination, one with recombined and the other with nonrecombined sequence. If the culture is not given enough time during the recovery phase in SOC medium, colony PCR on single colonies will give two bands, one for the WT and another for the recombined BAC. We observe this phenomenon regularly (see Fig. 4). It is recommended to passage the bacteria once in media supplemented with 34 μg/mL chloramphenicol and 30 μg/mL kanamycin. This should result in segregation of recombined and nonrecombined BAC and selection of BAC with kanamycin resistance gene (recombined BAC). 28. Use the incubation time to prepare the L-arabinose solutions in LB medium. It should always be freshly prepared, sterile filtered and not autoclaved. 29. It will likely take 2–3 weeks before you may be able to observe CPE. It is not uncommon for this amount of time to pass before significant CPE is observed, so it is imperative to be patient and continue passaging confluent cell cultures until the CPE is evident.

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References 1. Spaete RR, Mocarski ES (1987) Insertion and deletion mutagenesis of the human cytomegalovirus genome. Proc Natl Acad Sci U S A 84:7213–7217 2. Chee MS, Bankier AT, Beck S, Bohni R, Brown CM, Cerny R, Horsnell T, Hutchison CA 3rd, Kouzarides T, Martignetti JA et al (1990) Analysis of the protein-coding content of the sequence of human cytomegalovirus strain AD169. Curr Top Microbiol Immunol 154:125–169 3. Murphy E, Rigoutsos I, Shibuya T, Shenk TE (2003) Reevaluation of human cytomegalovirus coding potential. Proc Natl Acad Sci U S A 100:13585–13590 4. Davison AJ, Dolan A, Akter P, Addison C, Dargan DJ, Alcendor DJ, McGeoch DJ, Hayward GS (2003) The human cytomegalovirus genome revisited: comparison with the chimpanzee cytomegalovirus genome. J Gen Virol 84:17–28 5. Dargan DJ, Douglas E, Cunningham C, Jamieson F, Stanton RJ, Baluchova K, McSharry BP, Tomasec P, Emery VC, Percivalle E, Sarasini A, Gerna G, Wilkinson GW, Davison AJ (2010) Sequential mutations associated with adaptation of human cytomegalovirus to growth in cell culture. J Gen Virol 91:1535–1546 6. Sinzger C, Schmidt K, Knapp J, Kahl M, Beck R, Waldman J, Hebart H, Einsele H, Jahn G (1999) Modification of human cytomegalovirus tropism through propagation in vitro is associated with changes in the viral genome. J Gen Virol 80(Pt 11):2867–2877 7. Murrell I, Wilkie GS, Davison AJ, Statkute E, Fielding CA, Tomasec P, Wilkinson GW, Stanton RJ (2016) Genetic stability of bacterial artificial chromosome-derived human cytomegalovirus during culture in vitro. J Virol 90:3929–3943 8. Borst EM, Hahn G, Koszinowski UH, Messerle M (1999) Cloning of the human cytomegalovirus (HCMV) genome as an infectious bacterial artificial chromosome in Escherichia coli: a new approach for construction of HCMV mutants. J Virol 73:8320–8329 9. Hahn G, Rose D, Wagner M, Rhiel S, McVoy MA (2003) Cloning of the genomes of human cytomegalovirus strains Toledo, TownevarRIT3, and Towne long as BACs and sitedirected mutagenesis using a PCR-based technique. Virology 307:164–177 10. Marchini A, Liu H, Zhu H (2001) Human cytomegalovirus with IE-2 (UL122) deleted

fails to express early lytic genes. J Virol 75:1870–1878 11. Hahn G, Khan H, Baldanti F, Koszinowski UH, Revello MG, Gerna G (2002) The human cytomegalovirus ribonucleotide reductase homolog UL45 is dispensable for growth in endothelial cells, as determined by a BAC-cloned clinical isolate of human cytomegalovirus with preserved wild-type characteristics. J Virol 76:9551–9555 12. Sinzger C, Hahn G, Digel M, Katona R, Sampaio KL, Messerle M, Hengel H, Koszinowski U, Brune W, Adler B (2008) Cloning and sequencing of a highly productive, endotheliotropic virus strain derived from human cytomegalovirus TB40/E. J Gen Virol 89:359–368 13. Murphy E, Yu D, Grimwood J, Schmutz J, Dickson M, Jarvis MA, Hahn G, Nelson JA, Myers RM, Shenk TE (2003) Coding potential of laboratory and clinical strains of human cytomegalovirus. Proc Natl Acad Sci U S A 100:14976–14981 14. Stanton RJ, Baluchova K, Dargan DJ, Cunningham C, Sheehy O, Seirafian S, McSharry BP, Neale ML, Davies JA, Tomasec P, Davison AJ, Wilkinson GW (2010) Reconstruction of the complete human cytomegalovirus genome in a BAC reveals RL13 to be a potent inhibitor of replication. J Clin Invest 120:3191–3208 15. Tischer BK, Smith GA, Osterrieder N (2010) En passant mutagenesis: a two step markerless red recombination system. Methods Mol Biol 634:421–430 16. Stone NE, Fan JB, Willour V, Pennacchio LA, Warrington JA, Hu A, de la Chapelle A, Lehesjoki AE, Cox DR, Myers RM (1996) Construction of a 750-kb bacterial clone contig and restriction map in the region of human chromosome 21 containing the progressive myoclonus epilepsy gene. Genome Res 6:218–225 17. Sampaio KL, Weyell A, Subramanian N, Wu Z, Sinzger C (2017) A TB40/E-derived human cytomegalovirus genome with an intact US-gene region and a self-excisable BAC cassette for immunological research. BioTechniques 63:205–214 18. Hammer Q, Ruckert T, Borst EM, Dunst J, Haubner A, Durek P, Heinrich F, Gasparoni G, Babic M, Tomic A, Pietra G, Nienen M, Blau IW, Hofmann J, Na IK, Prinz I, Koenecke C, Hemmati P, Babel N, Arnold R, Walter J, Thurley K, Mashreghi MF, Messerle M, Romagnani C (2018) Peptide-specific recognition of human

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cytomegalovirus strains controls adaptive natural killer cells. Nat Immunol 19:453–463 19. Sambrook JF, Russel DW (2000) Molecular cloning: a laboratory manual. Cold Spring Harbor laboratory press. Cold Spring Harbor, New York 20. Borst EMC-M, I.; Messerle, M. (2004) Cloning of β-herpesvirus genomes as bacterial artificial chromosomes. In: Zhao SS, M. (eds) Bacterial artificial chromosomes: volume 2: functional studies vol 256. Humana Press Inc., Totowa, NJ, pp 221–240 21. Tischer BK, von Einem J, Kaufer B, Osterrieder N (2006) Two-step red-mediated recombination for versatile high-efficiency markerless DNA manipulation in Escherichia coli. BioTechniques 40:191–197 22. Mosberg JA, Lajoie MJ, Church GM (2010) Lambda red recombineering in Escherichia coli

occurs through a fully single-stranded intermediate. Genetics 186:791–799 23. Thomason LC, Sawitzke JA, Li X, Costantino N, Court DL (2014) Recombineering: genetic engineering in bacteria using homologous recombination. Curr Protoc Mol Biol 106:1 16 1–1 1639 24. Gibson DG, Young L, Chuang RY, Venter JC, Hutchison CA 3rd, Smith HO (2009) Enzymatic assembly of DNA molecules up to several hundred kilobases. Nat Methods 6:343–345 25. Yu D, Smith GA, Enquist LW, Shenk T (2002) Construction of a self-excisable bacterial artificial chromosome containing the human cytomegalovirus genome and mutagenesis of the diploid TRL/IRL13 gene. J Virol 76:2316–2328

Chapter 9 Methods for Studying the Function of Cytomegalovirus GPCRs Christine M. O’Connor and William E. Miller Abstract All of the cytomegaloviruses discovered to date encode two or more genes with significant homology to G protein–coupled receptors (GPCRs). The functions of these cytomegalovirus GPCRs continue to be actively studied and it is clear that they exhibit numerous interesting functions in vitro and in vivo. In this chapter, we review the various methodologies that can be used to examine biochemical aspects of viral GPCR signaling in vitro, as well as examine the biological activity of these viral GPCRs in vitro and in vivo in virus infected cells using recombinant cytomegaloviruses. Key words G protein–coupled receptors, Human cytomegalovirus, Murine cytomegalovirus, Virus genetics, Signal transduction, Virological methods

1

Introduction Human cytomegalovirus (HCMV) is a beta herpesvirus that infects a large majority of the world’s population. Infection with HCMV in utero is the leading cause of infectious congenital birth defects in developed countries, resulting in developmental disabilities. Although infection with the virus remains, for the most part, asymptomatic in healthy individuals, immunocompromised individuals who undergo viral reactivation or receive a primary infection suffer from severe morbidity and often mortality as a direct consequence of HCMV-associated disease [1]. Sequence analysis of the HCMV genome reveals that this virus encodes at least four G protein-coupled receptors (GPCR), including UL33, UL78, US27, and US28 [2, 3]. GPCRs are cell surface molecules that contain seven transmembrane domains and function in signal transduction [4]. The binding of an appropriate ligand to its cognate GPCR activates the receptor (agonist-dependent signaling), while dissociation of the ligand converts the GPCR to its inactive state. Some GPCRs exhibit significant signaling activity in the absence of a bound ligand and in this case the signaling is

Andrew D. Yurochko (ed.), Human Cytomegaloviruses: Methods and Protocols, Methods in Molecular Biology, vol. 2244, https://doi.org/10.1007/978-1-0716-1111-1_9, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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termed constitutive. When in their active state, GPCRs induce a variety of signal transduction pathways that alter the cellular environment by activating molecules involved in adhesion, migration, proliferation, differentiation, cytoskeletal dynamics, contractility, and so on [5]. Both the primate (i.e., human, rhesus) and nonprimate (i.e., murine, rat) CMVs encode members of the UL33 and UL78 family, whereas only the primate CMVs additionally encode US27 and US28 [6]. Cytomegalovirus encoded GPCRs have been demonstrated to respond to external ligands and/or signal constitutively; some examples are provided here with references—HCMV US28 [7– 11], HCMV UL33 [12], HCMV US27 [13], Rat CMV (RCMV) R33 [12, 14], and Murine CMV (MCMV) M33 [9, 15]. The murine UL33 ortholog M33 contributes to pathogenesis in vivo, as assessed by a requirement of the GPCR and its constitutive signaling for viral replication within the host’s salivary glands [16–22]. Similar results were demonstrated with the RCMV UL33 ortholog, R33 [23]. Although members of the UL33 gene family are required for pathogenesis in vivo; UL33, R33, and M33 are dispensable for replication in fibroblasts [24, 25]. Similar to UL33, HCMV UL78 has orthologs across the betaherpesvirus family. MCMV M78 is a virion constituent, and upon infection of host cells, promotes immediate early (IE) viral mRNA accumulation [26]. Infection of the respective host with a virus harboring a deletion of M78 or R78 decreases viral titers in the spleen, liver, and salivary glands, while increasing the survival rates in these animals [26–28], suggesting a role for these GPCRs in viral pathogenesis in both the mouse and rat CMV models. HCMV UL78 is assembled into the mature viral particle [29], and although not essential for efficient viral replication in fibroblasts [29, 30] or in a renal artery tissue culture model [30], it is critical for replication in epithelial and endothelial cells [29]. In epithelial cells, UL78 is necessary for appropriate delivery of the viral particle to the nucleus [29]. HCMV US27 is important for efficient spread of HCMV via the extracellular route in both endothelial cells and fibroblasts [31]. Recent work has revealed US27 has signaling capabilities. US27 constitutively activates nuclear respiratory factor-1 (NRF-1)/antioxidant response element (ARE) via Gβγ and phosphoinositide 3-kinase (PI3K) [32]. Furthermore, US27 expression during lytic infection modulates CXCR4 signaling [33] and internalization [34], as well as enhances the CXCL12/CXCR4 signaling axis [13], suggesting this viral GPCR (vGPCR) impacts cellular GPCR signaling. Additionally, US27’s canonical ‘DRY’ box, to which G-proteins couple, functions to promote HEK293 cell growth and survival in transient transfection assays [35], suggesting that US27’s signaling abilities may impact these cellular processes. Currently UL78 and US27 chemokine interaction(s) and natural ligand(s) remain unknown, and UL78 signaling (constitutive and/or ligand-induced) remains elusive.

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Arguably the best-studied HCMV GPCR is encoded by US28. US28 exhibits both constitutive and agonist-dependent signaling and has been demonstrated to bind numerous C-C chemokines (RANTES, MIP-1α, MIP-1β, and MCP-1) and the CX3Cchemokine fractalkine [7, 8, 12, 36–40]. US28 constitutive signaling is exemplified by its ability to activate phospholipase C-β in the absence of ligand leading to increased adhesion of monocytes, while its agonist-dependent signaling is exemplified by its ability to modulate Ca2+ flux, direct vascular smooth muscle cell migration [7, 10, 11, 39–45]. HCMV US28 appears to be somewhat “promiscuous” in its G-protein coupling as it appears to be able to activate both Gq/11 and G12/13 G-proteins [7, 43, 46, 47]. As a consequence of this G-protein signaling, US28 influences the activity of a variety of downstream effectors such as nuclear factor kappalight-chain-enhancer of activated B cells (NFκB), nuclear factor of activated T cells (NFAT), cyclic adenosine monophosphate (cAMP) response element binding protein (CREB), mitogen-activated protein kinase (MAPK), Rho, activator protein (AP-1), and signal transducer and activator of transcription (STAT)/interleukin 6 (IL6) [9, 46, 48–52]. In addition to its involvement in cell adhesion and migration, HCMV US28 has been shown to have oncogenic potential. Expression of US28 enhances cell growth and cell cycle progression, and induces a proangiogenic and transformed phenotype in vitro [53, 54]. In vivo, injection of NIH3T3 fibroblasts expressing US28 into nude mice does indeed promote tumorigenesis [53, 55], possibly via cyclooxygenase2 (COX-2) up-regulation [54]. Additionally, investigators have shown the HCMV US28 RNA is found in both glioblastomas [49] and medulloblastomas [56], and in the former, promotes an invasive and angiogenic phenotype [49]. Indeed, constitutive US28 signaling accelerates tumor growth in murine models of glioblastoma [57]. Finally, US28 has been linked to other cancers, including the progression of intestinal neoplasia in transgenic mice [58] and colorectal cancer [59]. Taken together, these results argue for an oncomodulatory role for US28. US28, UL33, and UL78 are all detected during both natural and experimental latency [60–63], although the functional ramifications of UL33 and UL78 expression remain undefined. US28, however, is required to establish [52, 60] and maintain viral latency in hematopoietic progenitor cells [52, 60, 64]. Furthermore, US28 signaling impacts viral latency, and several pathways, including MAPK, NFκB, and AP-1 are attenuated, while STAT3-inducible nitric oxide synthase (iNOS) signaling is activated in response to the expression of this vGPCR [52, 64, 65]. While our understanding of US28’s role during this phase of infection has significantly broadened over the recent years, a complete understanding of this vGPCR’s function(s) during latency and reactivation are likely far from complete.

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This chapter focuses on recent advancements in methodologies used for studying the function of CMV GPCRs, emphasizing protocols that can be performed in classical cellular signaling models as well as in the context of infection. In particular, we will address methods for examining viral GPCR signaling by transient assays and focus on the recent breakthroughs in generating viral mutants making possible the investigation of GPCR function in the context of infection. Finally, we will discuss the importance of animal models to our understanding of the CMV GPCRs in viral pathogenesis.

2 2.1

Materials Cell Culture

1. Primary human fibroblasts, such as HS68 cells (ATCC, CRL-1635) used between passages 10 and 20 (see Note 1). 2. Human embryonic kidney cell line HEK-293 (ATCC, CRL-1573). 3. Human embryonic kidney cell line with SV40 T-antigen HEK-293T (ATCC, CRL-3216). 4. Primary human retinal pigment epithelial cells (ARPE19 [ATCC, CRL-2302]) used between passages 22 and 35. 5. Human umbilical vascular endothelial cells (HUVECs) isolated from umbilical cords by collagenase digestion (or purchased from Lonza) maintained on either Primaria tissue culture plates (BD Falcon) or plates precoated with 0.1% pig gelatin (MilliporeSigma) in 1
PBS; used between passages 2 and 8. 6. Human THP-1 monocytic cell line (ATCC, TIB-202), maintained between 2
105 and 8
105 cells/ml. 7. Human Kasumi-3 cell line (ATCC, CRL-2725), maintained between 3
105 and 1
106 cells/ml. 8. Primary human cord blood- or bone marrow–derived CD34+ hematopoietic progenitor cells (see Note 2). 9. Dulbecco’s modified eagle medium (DMEM) containing 10% Fetal Clone III serum (Hyclone), supplemented with 100 U/ ml each of penicillin and streptomycin is used to culture primary human fibroblasts and HEK-293T cells. 10. Dulbecco’s modified eagle medium (DMEM) containing 10% fetal bovine serum (Hyclone), supplemented with 100 U/ml each of penicillin and streptomycin is used to culture HEK-293T cells. 11. Minimal essential medium MEM containing 10% fetal bovine serum (FBS; Hyclone), supplemented with 100 U/ml each of penicillin and streptomycin is used to culture HEK-293 cells.

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12. DMEM-HAM’s F12 containing 10% FBS, 2.5 mM L-glutamine, 0.5 mM sodium pyruvate, 15 mM HEPES, 1.2 g/L NaHCO3 and 100 U/ml each of penicillin and streptomycin is used to culture ARPE19 cells. 13. EGM-2 medium supplemented with the EGM-2 additives (Lonza) is used to culture HUVECs. 14. Roswell Park Memorial Institute 1640 (RPMI 1640) supplemented with 10% FBS (MilliporeSigma) and 100 U/ml each of penicillin and streptomycin is used to culture human THP-1 cells. 15. RPMI 1640 (ATCC, 30–2001) containing 20% FBS (MilliporeSigma), 100 μg/ml gentamycin, and 100 U/ml each of penicillin and streptomycin is used to culture Kasumi-3 cells. 16. X-VIVO 15 Serum-free Hematopoietic Cell Medium (X-VIVO 15; Lonza) containing 100 U/ml each of penicillin and streptomycin is used for the infection of Kasumi-3 cells. 17. Trypsin–EDTA: 0.05% Trypsin, 0.53 mM EDTA. 18. Transfection reagent (e.g., Mirus TransIT® LT1 or Invitrogen Lipofectamine 2000). 2.2 Assessing PLC-β Activity by Measuring IP3 Accumulation

1. Wash dowex in formate phase (AG1-X8, Bio-Rad 140-1444) with 20 L dH2O. Store as 50% slurry in dH2O at 4  C. Add 1 ml of slurry to column prior to use. A variety of reusable columns can be used. Bio-Rad Poly-Prep® chromatography columns (part no. 731-1550) work well. 2. [2-3H(N)]-myoinositol (PerkinElmer, NET-114A).

2.3

Cell Lysis

1. Standard lysis buffer: 50 mM Hepes, pH 7.4, 0.5% NP-40, 250 mM NaCl, 10% glycerol, 2 mM EDTA, 1 mM PMSF, 2.5 μg/ml aprotinin, 5.0 μg/ml leupeptin, 200 μM activated sodium orthovanadate, 1 mM sodium fluoride (see Note 3). 2. RIPA lysis buffer: 10 mM Tris pH 7.5, 0.1% SDS, 1.0% Triton X-100, 1.0% deoxycholate, 150 mM NaCl, 5 mM EDTA, 1 mM PMSF, 2.5 μg/ml Aprotinin, 5.0 μg/ml Leupeptin, 200 μM activated sodium orthovanadate, and 1 mM sodium fluoride. 3. Laemmli Sample Buffer: 14.0 ml 4
Tris Stacking buffer pH 6.8, 14.4 ml 50% Glycerol, 2.0 g SDS, 240 μl betamercaptoethanol, and 9.4 ml dH2O. 4. 10
Red Blood Cell (RBC) Lysis Buffer: 40.15 g NH4CL, 5.0 g NaHCO3, 0.186 g EDTA in 200 ml dH2O. Dilute RBC Lysis buffer to 1
prior to use.

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Western Blotting

1. Tris buffered saline containing Tween 20 (TBST): 50 mM Tris–HCl, pH 7.4, 150 mM NaCl, 0.1% Tween 20. 2. Blocking Buffer: TBST containing 5% nonfat dried milk or phosphate buffered saline containing Tween 20 (PBST) containing 5% BSA. 3. PBST: 10 mM Na2HPO4, 1.8 mM KH2PO4, 2.7 mM KCL, 137 mM NaCl, 0.05% Tween 20. 4. Blocking Buffer: PBST containing 5% bovine serum albumin (BSA). 5. Phospho-p38 MAPK (Thr180/Tyr182) Antibody, Cell Signaling Technology, Cat#9211. 6. Monoclonal Cat#F1804.

anti-FLAG

M2

antibody,

MilliporeSigma,

7. Diluent for anti-FLAG M2 antibody: PBST containing 2% BSA. 8. Supported nitrocellulose (e.g., Schleicher and Schuell). 9. Chemiluminescence Detection Kit (e.g., Amersham ECL or Pierce SuperSignal western blotting kits). 10. Infrared Detection Kit (e.g., Li-Cor Odyssey western blotting kits). 2.5 Luciferase Reporter Assays

1. Dual-Luciferase Cat#E1910.

Reporter

Assay

System,

2.6 BAC Recombineering

1. 0.2 mg/ml D-biotin: sterile filtered, made fresh.

Promega,

2. 20% galactose: autoclaved, stored at 4  C. 3. 20% 2-deoxy-galactose: autoclaved, made fresh. 4. 20% glycerol: autoclaved, stored at RT. 5. 10 mg/ml L-leucine: Heat to get into solution but do not let boil. Sterile filtered, stored at 4  C. 6. 12.5 mg/ml chloramphenicol in EtOH, stored at 20  C. 7. 1 M MgSO4·7H2O stored at RT. 8. 1
M9 medium (1 L, autoclaved, stored at room temperature): 42.3 mM Na2HPO4 (6 g per liter), 22 mM KH2PO4 (3 g per liter), 18.7 mM NH4Cl (1 g per liter), 8.6 mM NaCl (500 mg per liter). 9. 5
M63 (1 L, autoclaved, stored at room temperature): 75.5 mM (NH4)2SO4 (10 g per liter), 0.5 M KH2PO4 (68 g per liter), 9.0 μM FeSO4·7H2O (2.5 mg per liter). Adjust to pH 7 with KOH. 10. M63 minimal plates (500 ml makes 20–25 plates): 7.5 g agar in 400 ml ddH2O in a 500 ml bottle with a stir bar and autoclaved, cooled to a “touchable” temperature of ~50–55  C,

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100 ml 5
M63 medium, 500 μl 1 M MgSO4·7H2O (1 μM), 500 μl chloramphenicol (12.5 μg/ml), 2.5 ml biotin (0.5 mg) (see Note 4), 5 ml galactose (0.2%), 2.25 ml leucine (45 mg). 11. 2-DOG plates (500 ml makes 20–25 plates): 7.5 g agar in 400 ml H2O in a 500 ml bottle with a stir bar and autoclaved, cooled to a “touchable” temperature of 50–55  C, 100 ml 5
M63 medium, 500 μl 1 M MgSO4·7H2O (1 μM), 500 μl chloramphenicol (12.5 μg/ml), 2.5 ml biotin (0.5 mg) (see Note 4), 5 ml 2-deoxy-D-galactose (0.2%), 2.25 ml leucine (45 mg), and 5 ml glycerol (0.2%). 12. MacConkey indicator plates: Prepare MacConkey agar plus galactose according to manufacturer’s instructions (e.g., BD, Cat#281810), with the addition of 12.5 μg/ml chloramphenicol. 13. Primers for galactokinase (galK) gene amplification. Primers must contain a minimum of 50 bp of homologous sequence to the intended site of mutagenesis within the bacterial artificial chromosome (BAC). The underlined sequences below are complimentary to the pGalK cassette: Forward Primer: 50 -50 bp homology-CCTGTTGACAATTAATCATCGGCA-30 . Reverse Primer: 50 -50 bp complimentary strand homologyTCAGCACTGTCCTGCTCCTT-30 . 14. PCR cleanup columns (e.g., GE Healthcare GFX columns). 2.7 Purification of BAC DNA

1. CMPS1 [Similar to Qiagen P1 buffer, +RNAse]: 50 mM Tris– HCl, pH 8.0, 10 mM EDTA, pH 8.0, 200 μg/ml RNAse A added just prior to use (20λ of 10 mg/ml stock, per 1.0 ml CMPS1). 2. Alkaline SDS Solution [Similar to Qiagen P2 buffer]: Final concentrations: 0.2 N NaOH, 1% SDS. Make stock solutions at 2
concentrations, so mix equal parts just prior to use. 3. TEN Solution: 10 mM Tris–HCl, pH 7.4 or pH 8.0, 1 mM EDTA, pH 8.0, 150 mM NaCl. 4. 10.1 TE Solution: 10 mM Tris–HCl, pH 7.4 or pH 8.0, 0.1 mM EDTA, pH 8.0. 5. Endotoxin Removal Kit: MilliporeSigma, Cat#E4274. 6. Column BAC purification kit (e.g., Macherey-Nagel Nucleobond BAC purification kit).

2.8

Plasmids

1. pcDNA3 (or similar) vector for viral GPCR of choice (HCMVUS27, HCMV-US28, MCMV-M33, etc.). 2. pcDNA3 (or similar) vector for pp71. 3. pGL3 3
MHC-Luc (or similar) to assess NFκB activity.

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4. pGL3 9
NFAT-LUC (or similar) to assess NFAT activity. 5. pFRLUC and (Stratagene).

pFA2CREB

to

assess

CREB

activity

6. pHRG-TK to control for transfection variation and generalized effects of viral GPCRs on basal transcriptional activity (Promega). 7. pSLIK-Venus lentiviral vector for viral GPCR of choice (HCMV-US27, HCMV-US28, MCMV-M33, etc.) (Addgene). 8. psPAX2 lentiviral packaging vector (Addgene). 9. pCMV-VSV-G envelope expression vector (Addgene). 2.9

Miscellaneous

1. Antifade mounting medium (e.g., Vector Labs Vectashield or Molecular Probes SlowFade). 2. Biotinylated anti-FLAG antibody (e.g., MilliporeSigma M2 biotinylated anti-FLAG). 3. Cell Surface Protein Isolation kit (Thermo Scientific). 4. High Pure RNA Isolation Kit (Roche). 5. TaqMan Reverse Transcription Reagents (ThermoFisher). 6. SYBR Green (ThermoFisher).

3

Methods While it is not possible to generalize the signaling activities of the cytomegalovirus GPCRs into a single pathway, it is clear that several of these receptors (US28/M33/R33) signal via G-proteins such as Gq/11 and drive a number of downstream signals including accumulation of the second messenger IP3, activation of protein kinases, and stimulation of transcription factor activity. In this section, we will describe basic methodology that can be used to assess these particular signaling activities.

3.1 Measuring Viral GPCR-Stimulated Inositol Triphosphate (IP3) Accumulation

The following protocol was designed for the study of HCMV US28 stimulated IP3 accumulation (a.k.a. PIP2 hydrolysis, PLC activity, inositol triphosphate accumulation) in HCMV-infected fibroblasts, transiently transfected HEK-293 cells, or in lentivirus transduced THP-1 cells. These protocols can easily be adapted for use in a number of different cell types and can be modified to study other cytomegalovirus GPCRs [7, 9, 10, 12, 39, 40, 43, 66]. In the case of HCMV infection, the methodology described uses the primary human fibroblast cell line, HS68 (ATCC, CRL-1635), in the case of transient transfection, the methodology described uses the embryonic kidney cell line HEK-293 (ATCC, CRL-1573), and in the case of lentiviral transduction, the methodology described uses

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the human monocytic cell line THP-1 (ATCC, TIB-202). Gq/11 stimulated PLC-β activity cleaves phosphatidylinositol 4,5-bisphosphate (PIP2) into inositol 1,4,5-trisphosphate (IP3) and diacylglycerol (DAG). Accumulation of IP3 is easily measured in the lab with standard equipment and reagents. 1. Plate cells into 12-well culture plates so that they will be ~75% confluent at time of plating (adherent cells) or into 6-well plates at approximately 400,000 cells/ml (suspension cells). Incubate in humidified incubators at 37  C and 5% CO2. For infection of HS68 fibroblasts with HCMV, the suggested cell number is ~100,000 cells per well in a total volume of 1 ml of medium. For transient transfection of HEK-293 cells, the suggested cell number is ~250,000 cells per well in a total volume of 1 ml medium. HEK-293 cells and their derivatives are not tightly adherent and care should be taken to facilitate adherence such as coating the culture wells with 5 mg/ml collagen prior to plating cells. For lentiviral transduction of THP-1 cells, the suggested cell number is ~400,000 cells per well in a total volume of 500 μl of medium. 2. Let cells such as HS68 or HEK-293 adhere overnight. In the case of the suspension cells such as THP-1, immediately proceed to step 3. 3. Infect with virus (HCMV or lentivirus) or transfect with appropriate viral GPCR expression construct. The length of time to let the infection or transfection proceed prior to harvesting should be determined empirically depending on timing of viral GPCR expression, and so on. In the case of HCMV infection experiments, US28 reaches maximal expression at approximately 48 h postinfection (hpi); therefore, 48 h would be an appropriate time to analyze US28 dependent IP3 accumulation. Similarly, 48 hpi is a typical time at which to analyze transient transfection-based experiments as this is the time at which most transient gene expression peaks. For lentiviral transductions, since the pSLIK virus is DOX inducible, DOX should be added at 48 h posttransduction and US28 dependent IP3 accumulation analyzed at 96 h posttransduction. For infection-based experiments proceed to step 4, for transient transfection-based experiments proceed to step 5, for lentiviral transduction-based experiments, proceed to step 6. 4. Infection of HS68 fibroblasts with HCMV. Adsorb virus to cells at appropriate multiplicity of infection (MOI) for 3–6 h. Both wild-type and ΔUS28 strains should be used to ascertain the specific effects of US28 on driving the activation of this signaling pathway. To achieve roughly 95–99% infection, an MOI of 3–5 should be chosen. At the end of the adsorption period remove the medium containing virus and feed with fresh medium.

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5. Transient transfection of HEK-293 cells. The following describes the amount of DNA and lipid required for each well of a 12-well plate, although the amounts can be scaled up or down depending on the scale of transfection required. 250 ng of plasmid DNA is diluted in 50 μl serum free medium, supplemented with 1 μl of Mirus TransIT® LT1 transfection reagent and incubated for 15 min at room temperature. The 50 μl transfection reaction is then transferred to the appropriate wells of a 12-well plate and the transfection is allowed to proceed for 6 h. At the end of the 6 h incubation, remove the medium containing DNA/transfection reagent and feed with fresh medium. The TransIT® LT1 transfection reagent is highly efficient and exhibits low toxicity and therefore can be left on the cells overnight if desired. The viral GPCRs themselves are somewhat toxic in nature and thus should be tested at various concentrations (i.e., 10 ng, 50 ng, and 250 ng of DNA per well). All transfections should contain a total of 250 ng plasmid DNA, so in cases were less than 250 ng of viral GPCR DNA is used, the transfection cocktail should be supplemented with empty vector. It is recommended that the experiments be performed in duplicate or triplicate. 6. Lentiviral transduction of THP-1 cells. Lentiviral particles containing the pSLIK-US28 gene should be packaged as described below in Subheading 3.2. Two ml of lentivirus containing media should be added to each well of the THP-1 cells and “spinfected” by centrifuging the THP-1 cells and lentiviral media at 1000
g for 90 min at 37  C. After spinfection, THP-1 cells are cultured overnight, and media is replaced with fresh media in the morning. At day 2 after transduction, THP-1 cells are typically analyzed by flow cytometry for Venus expression to examine transduction efficiencies. Transduced cells are then transferred to 12-well plates at a concentration of 200,000 cells per ml and either left untreated or treated with 1 μg/ml doxycycline to induce US28 expression from the pSLIK lentivirus construct. 7. At 24 h postinfection/transfection or 72 h posttransduction, aspirate medium and add 1 ml per well of fresh medium containing 1.0 μCi/ml (HS68/HEK-293 cells) or 2.0 μCi/ml (THP-1 cells) [2-3H(N)]-myoinositol. Doxycycline should be readded to the media of pSLIK transduced cells to maintain US28 expression. The concentration of myo-inositol can be increased if necessary and the cells can be labeled in either serum free medium or serum containing medium (see Note 5). 8. The following day, wash cells 1
with 1 ml serum free medium. 9. Feed cells with 1 ml serum free medium containing 20 mM LiCl. If using chemokines or other potential agonists, add simultaneously with serum free medium containing LiCl. The

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LiCl inhibits endogenous inositol phosphatase activity and enables newly produced, receptor stimulated IP3 to accumulate. 10. Let inositol phosphates accumulate for 2–3 h. 11. Stop the reaction by aspirating medium, adding 1 ml of 0.4 M perchloric acid per well, and incubating for 15 min in the cold room. The perchloric acid will not cause the cells to lift off, but the perchloric acid at this point will contain the accumulated IPs. 12. Transfer 800 μl of perchloric acid from each well to a microfuge tube containing 400 μl of 0.72 M KOH/0.6 M KHCO3. This will form a white fluffy precipitate. 13. Vortex and centrifuge for 1 min at 15,000
g. 14. Transfer 50 μl of supernatant to scintillation vials, add 10 ml scintillation fluid, and count. (This step is optional and may be used to internally control for the relative labeling and cell number used). 15. Transfer 1 ml of the supernatant to Fisherbrand 12
75 mm glass tubes containing 3 ml dH2O. 16. Prepare dowex columns by adding 1 ml of dowex slurry (described in materials section) and let settle. A variety of reusable columns can be used for this step. 17. Pour sample from step 12 over column. Let sample flow through. 18. Wash columns with 2 bed volumes (~25 ml) of dH2O. 19. Wash columns with 1 bed volume of 60 mM sodium formate/ 5 mM disodium tetraborate. After washing the columns the inositol phosphates should be eluted using one of the following elution steps. Be sure to transfer the chromatography columns into a fresh scintillation vial prior to elution. Alternatively total IPs can be eluted if it is not necessary to differentiate between the exact form of the IP being generated (see Note 6). 20. Elute IP1 with 4 ml of 0.2 M ammonium formate/0.1 M formic acid. Wash the column with 1 bed volume of the same buffer. 21. Elute IP2 with 4 ml of 0.4 M ammonium formate/0.1 M formic acid. Wash the column with 1 bed volume of the same buffer. 22. Elute IP3 with 4 ml of 0.8 M ammonium formate/0.1 M formic acid. 23. Add 10 ml scintillation fluid to each eluted sample. Count in scintillation counter.

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3.2 Packaging of pSLIK-Based Lentiviral Constructs

The following protocol is specifically designed to package pSLIKVenus based lentiviral constructs, but it can easily be adapted for other lentiviral constructs. We prefer the pSLIK-Venus lentiviruses as gene expression is tightly regulated and inducible by doxycycline. Moreover, Venus expression is easy to monitor enabling flow cytometric analyses of transduction efficiencies and cell sorting when necessary. pSLIK-Neo and pSLIK-Hyg constructs are also useful as they facilitate selection of transduced cells in G418 or Hyg containing media. 1. Plate HEK-293T cells at 2.5
106 in standard 100-mm cell culture treated dishes. 2. Let cells adhere overnight. 3. Dilute 3 μg of pSLIK-Venus-US28FL, 1.5 μg psPAX2, and 0.5 μg pCMV-VSV-G plasmids into 1 ml serum free DMEM. 4. Slowly add 20 μl of Mirus LT-1 transfection reagent in a dropwise manner (4:1 ratio of Mirus LT-1 to DNA) and incubate at room temperature for 30 min. 5. Transfer the transfection mix (DNA, Mirus LT-1, and 1 ml DMEM) to a 100-mm dish of HEK-293T cells as described above in step 1. Incubate cells overnight in a humidified incubator at 37  C and 5% CO2. 6. Twenty-four hours after transfection, aspirate media and feed with 8 ml of fresh media supplemented with serum and antibiotics. 7. Forty-eight hour after transfection, reduce the volume of media from 8 ml to 4 ml. This enables the generation of lentiviral stocks with more concentrated viral particles. 8. Culture media containing lentivirus is harvested at 72 h posttransfection, centrifuged for 15 min at 500
g, and filtered through a 0.45 μM polyethersulfone (PES) membrane filer to ensure removal of residual cells. 9. Lentiviruses are then concentrated by ultracentrifugation at 25,000 rpm for 90 min at 4  C and viral pellets are resuspended in 1/10 volume of media of choice; for example, for transduction of THP-1 cells, viral pellets are resuspended in RPMI1640.

3.3 Measuring Viral GPCR-Stimulated Protein Kinase Activation

The following protocol is specifically designed for the detection of US28 or M33 stimulated p38-MAPK kinase activation in transiently transfected HEK-293 cells, but it can easily be adapted for use in a number of different cell types or for different protein kinases [51, 66, 67]. Moreover, the protocol can be modified to study other cytomegalovirus GPCRs in conditions of HCMV/ MCMV infection or lentivirus transduction. The protocol takes

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advantage of phospho-specific antibodies (which recognize activated forms of protein kinases) to assess vGPCR mediated activation of the protein kinase in question. 1. Plate HEK-293 cells into 12-well culture plate so that they will be ~75% confluent at time of plating. Incubate in humidified incubators at 37  C and 5% CO2. The suggested cell number is ~250,000 cells per well in a total volume of 1 ml of medium. HEK-293 cells and their derivatives are not tightly adherent and care should be taken to facilitate adherence, such as coating the culture wells with 5 mg/ml of collagen prior to plating cells. 2. Let cells adhere overnight. 3. Transfect with an appropriate viral GPCR expression construct as described above in Subheading 3.1, step 5. 4. Forty-eight hours posttransfection, lyse cells and prepare protein extracts for gel electrophoresis. Protein extracts can be prepared using several different lysis buffers, depending on the preference of the investigator (see Subheading 2). Extracts prepared directly in Laemmli sample buffer (step 5) maintains the phosphorylation status of most kinases. However, the use of this buffer eliminates the possibility of quantifying protein concentrations and therefore requires accurate cell counts prior to preparation of the extracts. When extracts are prepared in standard lysis buffer (step 6), one must ensure that phosphatase activity does not affect the results of the experiments. In particular it is important to use NaF and activated Na3VO4 in lysis buffers to inhibit serine/threonine and tyrosine phosphatases respectively (see Note 3). 5. To prepare whole cell extracts directly in Laemmli sample buffer, medium is aspirated from the 12-well plates, 250 μl of Laemmli sample is added directly to the wells, wells are scraped briefly with a cell scraper, and the extracts are transferred to microcentrifuge tubes. Extracts are sonicated briefly to disrupt chromosomal DNA. 6. To prepare whole cell extracts in standard lysis buffer, medium is aspirated from the 12-well plates, wells are washed 1
with 1
PBS, and 250 μl of standard lysis buffer is added directly to the wells. Wells are scraped briefly with a cell scraper, extracts are transferred to microcentrifuge tubes, and incubated on ice for 15–30 min. Extracts are clarified by centrifugation at 12,000
g and supernatant is transferred to a fresh tube. Protein concentration is then quantified by standard protein assays (Bradford, Bio-Rad Protein Assay, etc.).

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7. Extracts prepared by the procedure in steps 5 or 6 are then subjected to SDS-PAGE using standard protocols for gel electrophoresis. 8. Transfer resolved proteins to supported nitrocellulose membranes and block nonspecific reactivity with Tris-buffered saline containing 0.1% Tween 20 (TBST) and 5% nonfat dried milk. In some cases, blocking with 5% nonfat dried milk can increase nonspecific reactivity due to the presence of phosphoproteins present in the milk. In this case, 1% bovine serum albumin (BSA) can be used as a substitute for the milk. 9. Antibody directed against the phosphorylated/activated form of the protein kinase of interest is then used to probe western blots. In the case of p38-MAPK, the anti-phosphospecific p38 antibody is diluted 1:1500 in TBST. Bound primary antibody is then detected using the appropriate secondary antibodies using enhanced chemiluminescence or infrared fluorescence systems. 3.4 Measuring vGPCR-Stimulated Transcription Factor Activity

The cytomegalovirus GPCRs activate a number of transcription factors including NFκB, CREB, and NFAT [7, 12, 17, 19, 51]. The following protocol is specifically designed for the detection of US28 or M33 stimulated transcriptional reporter activity in transiently transfected HEK-293 cells, but it can easily be adapted for use in a number of different cell types or transcription factors and can be modified to study other cytomegalovirus GPCRs in conditions of HCMV/MCMV infection or lentivirus transduction. Two important considerations should be taken into account when assessing vGPCR-stimulated transcription factor activity in infected cells. First, it is important to use wild-type and vGPCR null mutants (i.e., ΔUS28 mutants) to differentiate between vGPCR effects and those due to either the virion itself or to other cytomegalovirus proteins. Many of the transcription factors stimulated by the vGPCRs are in fact activated during cytomegalovirus infection, but it is clear that the viruses use multiple mechanisms to activate transcription factors at different stages of infection. Such is the case for NFκB, which is activated within minutes after virion binding, due to virus engagement of NFκB linked cell surface receptors and also during the IE and E phases of infection [68]. Second it is also important to use internal controls such as the pHRG-TK Renilla luciferase control reporter. This will allow the investigator to control for generalized effects of cytomegalovirus infection on the basal transcription machinery itself, which can lead to artifactual conclusions regarding specific changes in transcription factor activity. 1. Plate HEK-293 cells into 12-well culture plate so that they will be ~75% confluent at the time of plating. Incubate in humidified incubators at 37  C and 5% CO2. The suggested cell number is ~250,000 cells per well in a total volume of 1 ml

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medium. HEK-293 cells and their derivatives are not tightly adherent and care should be taken to facilitate adherence such as coating the culture wells with 5 mg/ml collagen prior to plating cells. 2. Let cells adhere overnight. 3. Transfect with appropriate viral GPCR expression constructs and reporter genes. For transient transfection of HEK-293 cells in one well of a 12-well culture plates, 250 ng of plasmid DNA is diluted in 50 μl serum free medium, supplemented with 1 μl of Mirus TransIT® LT1 transfection reagent and incubated for 15 min at room temperature. The 50 μl transfection reaction is then transferred to cells plated as described in step 1 above and the transfection is allowed to proceed for 6 h. At the end of the 6 h incubation, remove the medium containing DNA/transfection reagent and feed with fresh medium. The TransIT® LT1 transfection reagent is highly efficient and exhibits low toxicity and therefore can be left on the cells overnight if desired. The vGPCRs themselves are somewhat toxic in nature and thus should be tested at various concentrations (i.e., 10 ng, 50 ng, and 250 ng of DNA per well). The concentration of the reporter gene DNA per well is as follows: for assessing NFκB activity (15 ng of pGL3-3
MHC-Luc), for assessing NFAT activity (15 ng of pGL3-9
NFAT-Luc) and for assessing CREB activity (30 ng pFR-LUC/10 ng pFA2-CREB) (see Note 7). The control pHRG-TK renilla luciferase plasmid should be included in all transfections at a concentration of 15 ng per well. All transfections should contain a total of 250 ng plasmid DNA, so in cases where less than 250 ng of viral GPCR DNA is used, the transfection cocktail should be supplemented with empty vector. To control for transfection variability, it is recommended that the experiments be performed in duplicate or triplicate. 4. Forty-eight hours posttransfection, aspirate medium and wash wells with PBS. 5. Add 200 μl of 1
Passive Lysis Buffer (PLB) per well (see Note 8). Incubate for 15–30 min at room temperature. Transfer lysate to microcentrifuge tubes. The samples can be stored at 80  C at this point. 6. Experimental luciferase (firefly) and control luciferase (renilla) can be examined on a luminometer using Luciferase Assay Reagent II (LAR II) and Stop&Glo Reagent according to the manufacturer’s instructions (see Note 8). These techniques provide the basis from which to examine proximal (IP3 accumulation), intermediate (p38-MAP kinase) and distal (transcription factor) signaling activity emanating from cytomegalovirus encoded GPCRs. The number of antibodies that

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recognize phosphorylated and thus activated protein kinases is increasing at a rapid pace, thus enabling researchers to continue to explore a variety of signaling pathways that lie downstream of the viral GPCRs. There are numerous other methodologies that have been used to examine viral GPCR signaling, however space constraints simply prevent us from covering each of these techniques in detail. For example, Smit and colleagues have used limited microarray analyses to uncover genes upregulated in response to US28 expression, and data mining approaches could easily be combined with large scale gene expression studies to identify networks of signaling pathways downstream of the vGPCRs [57, 69]. Finally, several investigators have used pharmacological inhibitors of signaling proteins such as phospholipase C-beta (PLC-β), protein kinase C (PKC), and PI3K to identify additional signaling proteins downstream of the vGPCRs [9, 19, 40, 51, 53–55, 57]. It is important to note that most of the signaling information that we currently have regarding the vGPCRs has been generated in in vitro transfection/ overexpression systems and it will be essential to use this current knowledge and extend these studies to identify what signals are truly generated in cytomegalovirus infected cells. 3.5 Generating and Analyzing Recombinant Cytomegaloviruses with Mutant GPCRs

Although significant advancements in our understanding of cytomegalovirus GPCRs have been made using the transient systems described above, recombineering methodologies have made possible the ability to investigate the CMV GPCRs in the context of viral infection.

3.5.1 BAC Recombineering of GPCR Genes

Early work aimed at generating recombinant viruses with mutations in viral GPCR genes took advantage of homologous recombination in mammalian cells, whereby a selectable marker was introduced by site-directed mutagenesis (Table 1). This method proved laborious and inefficient, and for some genes, impossible. Open reading frames (ORFs) that were essential for growth or those that conferred a severe growth defect when compared to wild-type, could only be mutated by this method if they were generated on complementing cell lines, which expressed the ORF of interest in trans [71]. Fortunately RCMV, RhCMV, MCMV, guinea pig CMV (GPCMV), and a variety of laboratory and clinical HCMV strains have been cloned into bacterial artificial chromosomes (BACs). Original protocols for BAC recombineering generated recombinant viral BACs that remained “marked” with either an insertion cassette or partial sequence from the shuttle plasmids used for RecE/T-mediated recombineering techniques [72]. Thus, this procedure does not yield seamless recombinants, which complicates the generation of viral BACs with multiple mutations and/or tags as well as revertants.

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Table 1 Cytomegalovirus GPCR recombinant virus constructs Viral GPCR

Host

M33

Murine K181

M33

K181

M78

Murine K181

R33

Rat

Maastricht Homologous recombination in mammalian cells [23]

R78

Rat

Maastricht Homologous recombination in mammalian cells [27, 28]

UL33

Human AD169

UL78

US27

US28

US27:US28 Multiple deletion mutant

Strain

Recombination Method Homologous recombination in mammalian cells [16–18, 70] Kan-frt BAC [19, 20] Homologous recombination in mammalian cells [26]

Kan-frt BAC [12, 25], homologous recombination in mammalian cells [24], galK BAC [25]

FIX

Kan-frt BAC [25], galK BAC [25]

TB40/E

Kan-frt BAC [25], galK BAC [25]

Human AD169

Kan-frt BAC [25], galK BAC[25], pST shuttle vector BAC [30]

FIX

Kan-frt BAC [29], galK BAC [29]

TB40/E

Kan-frt BAC [29], galK BAC [29]

Human AD169

Homologous recombination in mammalian cells [37]

AD169

Kan-frt BAC [25], galK BAC [25]

FIX

Kan-frt BAC [31], galK BAC [31]

TB40/E

Kan-frt BAC [31], galK BAC [31]

Human AD169

Homologous recombination in mammalian cells [37]

Toledo

Homologous recombination in mammalian cells [42]

Towne

Homologous recombination in mammalian cells [38]

AD169

Kan-frt BAC, galK BAC [25], pST shuttle vector BAC [10, 66]

FIX

Kan-frt BAC [39, 43, 44], galK BAC [25]

TB40/E

Kan-frt BAC [25], galK BAC [44, 45, 52]

Titan

Kan-frt BAC [48, 53, 54]

Human AD169

Homologous recombination in mammalian cells [37] (continued)

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Table 1 (continued) Viral GPCR

Host

Strain

UL33:UL78:US27:US28 multiple deletion mutant

Human AD169

Recombination Method I-SceI BAC [55]

FIX

galK BAC [25]

TB40/E

galK BAC [44]

UL33:UL78:US27 Multiple deletion mutant

Human TB40/E

galK BAC [60]

UL33:UL78:US28 Multiple deletion mutant

Human TB40/E

galK BAC [32]

The field of bacterial recombineering has advanced greatly over the recent years, and researchers have adapted these methods for generating recombinant CMV BACs to study the function of the CMV GPCRs (Table 1). In particular, two methods including galK [73] and I-SceI [74] recombineering have proved extremely useful, as each of these protocols results in recombinant viral BAC DNA that is seamless at the site of recombination. The advantage of seamless recombineering is such that one can generate multiple site-specific mutations, epitope tags, fusion proteins, gene insertions or whole ORF deletions within a single background. Additionally, these recombineering protocols are more efficient than previous BAC-mediated methods or site-directed mutagenesis in mammalian cells, have lower rates of off-site spontaneous recombination, require less time to generate mutants, permit reversion of the mutation, and unlike homologous recombination in mammalian cells, support the mutagenesis of essential ORFs. GalK and I-SceI recombination each take advantage of the Red recombinase system [73, 74]. The I-SceI method has previously been described for the study of HCMV GPCRs, and thus this chapter will focus on the utilization of the galK recombineering system [55]. Recombineering by galK uses a straightforward methodology, which involves introduction of the galK insertion cassette into the locus of interest, followed by homologous recombination with either a double-stranded oligonucleotide or a purified PCR containing the intended final change to the locus of interest, product by counterselection against the galK cassette. 1. PCR amplify galK gene using no more than 2 ng of pGalK plasmid as the template with primers (Subheading 2) that contain a minimum of 50 bp of homologous sequence to the intended site of mutagenesis within the BAC.

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2. To remove the pGalK template, digest the PCR product by adding 1 μl of DpnI directly into the PCR reaction and incubate at 37  C for 1 h. 3. Use PCR clean-up columns to purify the PCR product and elute the PCR product in 25 μl of dH2O. This cassette is then inserted into the CMV BAC genome by homologous recombination mediated by heat shock induced Red recombinase enzymes. It is critical to maintain the BAC in a recombination competent bacterial strain, such as SW102, SW105, or SW106 all of which are galK. Bacteria should be grown at 32  C to maximize stability of the CMV BAC genome. 4. Following overnight culture of the BAC-containing bacteria at 32  C in 5–10 ml of medium containing 12.5 μg/ml chloramphenicol, inoculate 25 ml of medium containing 12.5 μg/ml chloramphenicol with a 1:50 dilution of the overnight culture. Grow the bacteria to an OD600 of 0.5–0.6. Heat shock the cells at 42  C for 15 min in a shaking water bath. 5. Quickly cool the bacteria in an ice bath slurry, with shaking, then transfer 10 ml of the culture to a prechilled conical and pellet the bacteria at 4  C. Gently resuspend the pellet in 1 ml ice-cold ddH2O and transfer to a microcentrifuge tube. Wash the cell pellet 3 additional times in ice-cold ddH2O, and resuspend the pellet after the final wash in 100 μl of ice-cold ddH2O. 6. Use 2.5 μl of the PCR product to transform 50 μl of the bacteria by electroporation in a prechilled 2 mm gap electroporation cuvette. Recover for 1 h in 1 ml of medium without antibiotic at 32  C, and then wash the pellet three times in M9 salts taking care not to vortex or pipette too harshly. Resuspend the final pellet in 1 ml M9 salts. 7. Plate 100 μl of undiluted cells and 100 μl of a 1:10 dilution onto M63 minimal medium plates and incubate at least 3 days at 32  C. BACs that recombine to express galK are chosen by positive screen for growth on minimal medium containing galactose as the sole carbon source. Although all colonies that grow on these plates should ideally contain an integrated galK cassette, these plates screen for the preferential growth of potentially successful recombinants. 8. Successful galK recombinants are further selected using MacConkey’s indicator plates containing galactose, on which galKpositive clones will grow as single red colonies, while galKnegative clones will grow as white colonies. This step is critical to ensuring that the clones used in the counter-selection step do indeed contain the galK cassette.

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9. Patch single red colonies from step 8 onto LB/chloramphenicol plates and confirm the insertion of the galK cassette at the proper location of interest by PCR using flanking primer sets. Next, counterselect against galK, by substituting either a PCR product or a double stranded oligo that contains the mutation or epitope tag of one’s choice, all of which also contain flanking arms to the region in the BAC DNA being mutated. In this step the PCR product or double stranded oligo is inserted into the CMV BAC genome by homologous recombination mediated by red recombinase as described in step 2, with slight modification. 10. Prepare competent bacteria as described above (see steps 4 and 5). 11. Mix 10 mg of each oligo in a volume of 100 ml 1
PCR buffer. Boil for 5 min and cool slowly to room temperature. EtOH precipitate the annealed oligos and resuspend the final pellet in 100 μl ddH2O to yield a final concentration of 200 ng/ml. Use 1 μl per transformation. For reversion using PCR products, PCR amplify the desired insert with at least 50 bp of flanking sequence to the site of recombination, generating a PCR product of ~1000 bp. If the product is larger, increase the size of the flanking sequence. For example, use 500 bp for products >2 kb. Use 2.5 μl per transformation. 12. Following transformation, recover the bacteria in 10 ml medium without antibiotic in a 100 ml baffled flask for 4.5 h in a 32  C shaking incubator. Remove 1 ml of the culture and wash three times with M9 salts. Resuspend the final pellet in 1 ml M9 salts. The cells are diluted and plated (as described in step 7) on 2-deoxy-D-galactose (2-DOG) plates. 13. Selection against galK involves resistance to 2-DOG on minimal plates with glycerol as the carbon source. 2-DOG is harmless to bacteria, unless phosphorylated by functional galK. As a result, 2-DOG becomes 2-deoxy-galactose-1-phosphate, which bacteria cannot metabolize, and thus it is a toxic intermediate to those clones that still harbor the galK cassette. The resulting 2-DOG-resistant colonies are recombinant clones that have no residual foreign DNA sequences as a result of the recombineering protocol. 14. Patch colonies on both 2-DOG and M63 minimal plates to ensure for the excision of galK. Verify that the clones lack galK by PCR and finally sequence the recombinants to ensure genomic integrity at the site of the recombination. One can now use this clone to generate additional recombinants within the same background, or alternatively, reconstitute infectious virus (see Subheading 3.4, step 2).

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Following the successful generation of recombinant BACs for the GPCR(s) of interest, one can easily reconstitute infectious virus. The HCMV clinical strain TB40/E [75], for example, yields high titers following reconstitution, providing ample virus with which to perform a multitude of experiments. Additionally, as none of the GPCR mutants, including those recombinants that harbor multiple GPCR deletions, show a particle to PFU defect, each recombinant will indeed yield a stock with a titer that is sufficient [25]. The first step in reconstituting recombinant viruses is purifying the BAC DNA by either alkaline lysis/precipitation or column purification kit. The protocol described here involves purification of BAC DNA by alkaline lysis and we have modified the protocol to also include an additional step to remove endotoxins. Compositions of the buffers used in this protocol are given in the Materials section of this chapter. 1. Grow 10 ml overnight (~16–18 h) culture of bacteria containing the BAC of interest. 2. Pellet bacteria, resuspend in 200 μl CMPS1 w/RNaseA solution, lyse with 400 μl alkaline SDS solution and neutralize with 300 μl KAc. Pellet debris and treat clarified supernatant with the Endotoxin Removal solution (MilliporeSigma) according to the manufacturer’s instructions (see Note 9). Precipitate DNA with 0.7 volumes isopropanol to pellet DNA. 3. Dissolve the resulting pellet in 500 μl TEN buffer at room temperature. Once the pellet has dissolved (roughly 10 min) centrifuge briefly to remove any remaining cellular debris, and precipitate the DNA from the supernatant with 2 volumes ethanol. The resulting BAC DNA is resuspended in 10.1 TE buffer, and should be used within 24 h for transfection. Importantly, one should refrain from freezing BAC DNA that is slated for transfection, as this greatly reduces the efficiency. Additionally, when manipulating BAC DNA, one should take care not to shear the DNA by rapid pipetting or using standard pipette tips (wide-bore tips are optimal). 4. To reconstitute virus, transfect low-passage, primary fibroblasts (1.7
106 cells) in a 4 mm cuvette by electroporation (960 μF, 0.26 V) in 500 μl of Opti-MEM with 1 μg of pCGN-pp71 (or equivalent pp71-expressing plasmid) and the BAC DNA of interest. Plate transfected cells in either T75 flasks or 100 mm dishes. 5. Feed transfected cells every 2–3 days until significant cytopathic effect (CPE) is observed. 6. To generate TB40/E stocks proceed to step 7, to generate FIX stocks proceed to step 10.

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7. When fibroblasts have reached 100% CPE, infectious supernatant and cells are collected by low-speed centrifugation. 8. Reserve the supernatant, and bath sonicate the cells to release the cell-associated virus, spin the cell debris as before, and combine the supernatants. 9. This combined supernatant can now be used to generate high titer stocks by further expansion. To generate TB40/E stocks, only 1/10th of the infectious supernatant from step 8 is necessary to infect at least 5
107 cells. The remaining supernatant can be stored at 80  C in 1–2 ml aliquots and used to generate additional future stocks at another time. 10. For generating FIX stocks, when fibroblasts have reached 100% CPE, remove the medium and trypsinize the cells. Seed the infected cells onto approximately 4
107 to 5
107 cells. FIX is highly cell-associated, and thus seeding the cells rather than the medium from the transfection plate is critical. 11. For both FIX and TB40/E, generating a high titer usable stock may require one to concentrate the viral stock. To concentrate virus, harvest cells and medium from the expanded stock as described above. Pipette the cleared medium into ultracentrifugation tubes, and underlay with 20% D-sorbitol, containing 50 mM Tris–HCl, pH 7.2 and 1 mM MgCl2. Concentrate infectious virus by ultracentrifugation at 72,128
g for 90 min at 25  C. Virus stocks can be stored at 80  C in complete medium containing 1.5% BSA for long-term storage. 3.5.3 Assessment of Viral Growth Properties

Many of the early assessments of the HCMV ORFs’ necessities for viral replication in tissue culture were performed using fibroblasts [76, 77]. Although fibroblasts are invaluable to the study of HCMV lytic replication, they do not afford the ability to uncover functions of HCMV genes that are essential for growth in other clinically relevant cell types and tissues. Such is the case for the viral GPCRs that are not required for HCMV replication in fibroblasts. Thus, many investigators have taken advantage of clinical strains of HCMV that exhibit a broader cell tropism. The use of these clinical strains (i.e., TB40/E, TR, or FIX) allows for studies in an expanded repertoire of cell types including, but not limited to fibroblasts, hematopoietic progenitor cells, monocytes, macrophages, epithelial and endothelial cells. Assessing the growth properties of a mutant virus in range of cell types is critical, as CMV pathogenesis in vivo is complicated and involves a plethora of different cells and tissues. The following seven steps (steps 1–7) are used for assessing production and/or spread of virus occurring via the extracellular route.

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1. Plate cells in 6-well plates (~5
105 to 1
106 cells, depending on cell type), designating two wells for each virus being tested. These two wells will serve as duplicate infections. Note that infection of ARPE19 cells does not result in efficient extracellular spread of HCMV (e.g., FIX and TB40/E) therefore quantifying viral replication is performed by cell-associated viral assays (see below). 2. For multistep growth curves in fibroblasts or endothelial cells use a low MOI (e.g., 0.01–0.1 PFU/cell). For single-step growth curves, a high MOI between 1.0 and 3.0 PFU/cell is recommended. 3. Dilute the viral stock in the medium specific to the cell type being used. Ensure that enough inoculum is prepared to cover each well in addition to some that is reserved to assess the input titer. This is important when comparing viral growth between wild-type and recombinants, as one needs to ensure an equal amount of virus was used in the initial infection. Thus, reserving some inoculum that was not used to infect cells is important when titering the growth curve. 4. Remove the medium and wash one time with 1
PBS. Add the inoculum to the cells in low volume (750 μl for a well of a 6-well plate) to ensure sufficient contact of the virus with the cells. Incubate at 37  C/5% CO2 for 1 h, rocking the plate every 15 min. 5. Remove the inoculum, wash three times with 1
PBS, add fresh medium to the cells, and return to the incubator. 6. For low MOI infections, suggested time points include 0, 4, 8, 12, and 15 days postinfection (dpi). For high MOI infections, suggested time points include 0, 24, 48, 72, 96, and 120 hpi. The investigator should adjust these, as necessary. At each time point, remove a portion of the supernatant. This will vary depending on the assay being used to titer viral growth. One should reserve enough supernatant from the cells such that the titering assay can be performed in triplicate. Replenish the cultures with the same volume of fresh medium that was removed for the time point. Store all of the collected samples at 80  C until the time course is completed. 7. Once all of the time points have been collected, thaw the samples in a 37  C water bath, and assess the titers by plaque assay, TCID50 analyses, or modified IFA for IE protein expression. Each time point for each virus should be measured in triplicate. The following nine steps (steps 8–17) are used for assessing production and/or spread of virus occurring via the cellassociated route.

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8. Plate cells in 6-well plates (~5
105 to 1
106 cells, respectively). Dedicate one well for each time point for every virus being assessed. 9. Both endothelial cells and fibroblasts support infection with low MOIs of 0.01–0.1 PFU/cell or high MOI infections of at least 1.0 PFU/cell. Viral infection of ARPE19 cells spreads exclusively by cell-to-cell contact following either FIX or TB40/E infection. For ARPE19 cells, it is advisable to use an MOI of approximately 0.1 PFU/cell for multistep growth curves and an MOI of at least 1.0 PFU/cell for single-step growth curves (see Note 10). 10. Prepare the inoculum as above, diluting virus in the appropriate medium as described above in step 3. 11. Remove the medium from the cells and wash the cells with 1
PBS. 12. Add the inoculum to the appropriate wells as above reserving an aliquot of the inoculum, and infect the cells for 1 h at 37  C/5% CO2, rocking the plates every 15 min. Although not required, infection of ARPE19 cells is increased by centrifugal enhancement at 1000
g for 30 min at room temperature. If this step is performed, the cells should next be incubated at 37  C/5% CO2 for an additional 1 h with rocking every 15 min. 13. Remove the inoculum and wash the cells three times with 1
PBS to remove any residual virus that had not entered the cells. Replenish the cultures with fresh medium and return to the incubator. 14. For either fibroblasts or endothelial cells, collect the cellassociated virus at the times described above in Subheading 4.3.1, step 3 for low and high MOIs. For ARPE19 cells, infection progresses at a slower rate, and thus, low MOI time points include 0, 10, 20, and 30 dpi. Additionally, the medium on ARPE19 cultures should be changed every 5 dpi to ensure cell health over the time course of infection. For single-step growth analyses at high MOI, suggested time points include 0, 4, 8, and 12 dpi. 15. To collect cell-associated virus at each time point, remove the medium from the cultures, and wash 2–3 times with 1
PBS. Add back at least 1 ml of fresh medium and scrape the cells into the medium. Samples should be stored at 80  C until the time course is completed. 16. Evaluating the titer of the cell-associated virus requires three freeze–thaw cycles to disrupt the cells, thereby releasing the virus. Thaw samples in a 37  C water bath, ensuring that the samples completely thaw, and then quickly refreeze in liquid

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nitrogen. Following the third thaw, spin down cellular debris and transfer the medium to a fresh tube for use in a titering assay as described above. Undoubtedly, the types of approaches described in this section will generate important information on the roles of viral GPCRs in viral replication and spread in clinically relevant cell types in vitro, and when combined with in vivo viral replication experiments in animal models, will provide clues as to how these proteins function to facilitate replication and pathogenesis during the natural course of cytomegalovirus infection. 3.6 Detecting Viral GPCR Proteins in Infected Cells

As mentioned above, bacterial recombineering techniques have afforded investigators the ability to epitope tag viral proteins, in particular the viral GPCRs. Previous studies assessing the expression and subcellular localization of the CMV GPCRs generally included the overexpression of individual GPCRs in cell types in which CMV infection is not supported. Although these studies yield important information about potential function, these were not performed in the context of viral infection. Moreover, the expression level of viral GPCRs in infected cells may be very different than that observed in transient assays and may result in qualitative and quantitative differences in signaling. Antibodies directed against several of the CMV GPCRs including US27 [78], US28 [79], UL33 [24, 80], and MCMV M78 [26] have been generated. However, construction of viral recombinants expressing epitopetagged GPCRs allows investigators to utilize commercially available validated antibodies that work across a variety of techniques including immunofluorescence assay (IFA), immunoprecipitation/western blot, immuno-electron microscopy, and fluorescence activated cell sorting (FACS).

3.6.1 Detection and Localization of Viral GPCRs by Immunofluorescence Assay (IFA)

IFA provides a useful platform for determining the cellular localization of a given GPCR and provides a convenient tool for determining the percentage of cells expressing the GPCR in question. 1. Grow cells on glass coverslips and infect at an MOI of at least 0.5 PFU/cell. Using an MOI of 3 PFU/cell will typically guarantee that >95% of the cells are infected. 2. At the desired time postinfection, wash cells with 1
PBS, and fix with 2% paraformaldehyde at 37  C for 15 min. Alternatively, one can use cold 100% EtOH to fix the infected cells, although it is important to note that this will destroy any color marker (e.g., eGFP or mCherry) that is expressed from the viral genome. 3. Following fixation, wash cells three times with 1
PBS at room temperature, and then permeabilize with 0.1% Triton X-100 for 15 min at room temperature.

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4. Wash cells with PBS containing 0.2% Tween 20, and then block for at least 1 h at room temperature in 2% BSA/0.2% Tween 20 in 1
PBS. Alternatively, one can block the coverslips overnight at 4  C. If multiple time points are necessary, the blocking step is an excellent step at which to stop until the remaining slides are harvested. 5. After blocking all of the coverslips, stain with primary antibody in blocking buffer for at least 1 h at room temperature. 6. After staining with primary antibody, wash the coverslips with 1
PBS containing 0.2% Tween 20 at least three times at room temperature. 7. Stain the cells/coverslips with secondary antibody containing the appropriate conjugated fluorophore for at least 1 h in the dark at room temperature. A nuclear dye, such as 40 ,6-diamidino-2-phynelindole (DAPI) or Hoechst should also be included, as this serves as an excellent control. 8. Wash the cells/coverslips three times in 1
PBS containing 0.2% Tween 20. 9. Mount and seal the coverslips onto slides using an appropriate antifade mounting medium. 10. View the cells using standard fluorescent or confocal microscopy techniques. We have taken advantage of recombineering techniques to generate FLAG-tagged GPCR recombinants in the AD169, FIX, and TB40/E backgrounds. Using these recombinants coupled with IFA, we have shown cellular localization for each of the GPCRs, and have determined the presence of each in the mature HCMV virion [25, 29, 31]. 3.6.2 Detection and Localization of Viral GPCRs (and Interacting Partners) by FLAG Immunoprecipitation/ Western Blot

Immunoprecipitation followed by western blotting is a very sensitive technique that can be used to detect viral GPCRs. This sensitivity is essential as future investigations designed to analyze vial GPCR expression function are likely to be performed in clinically relevant cell types that may not exhibit lytic expression patterns comparable to that observed in standard HCMV-infected fibroblasts. 1. Plate cells in 100 mm dishes at 50–75% confluent. Infect with viruses or expression constructs expressing FLAG-tagged GPCRs as described elsewhere in this report. 2. At the appropriate times postinfection or posttransfection, remove medium and wash cells 1
with PBS. 3. Add 1.0 ml of RIPA buffer containing protease and phosphatase inhibitors.

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4. Transfer the lysate to microcentrifuge tubes and shear the DNA by passing lysates through a 22-gauge needle and syringe 15–20 times. 5. Clarify the supernatant by centrifugation at 15,000
g for 15 min at 4  C. 6. Transfer supernatant to a clean microcentrifuge tube and preclear lysates by adding 50 μl Sepharose 4B and rotating for 30–60 min at 4  C. 7. Pellet Sepharose 4B by centrifugation at 15,000
g for 15 min at 4  C. 8. Transfer the supernatant to fresh tube and add 20 μl of antiFLAG M2 beads. Rotate for 4 h at 4  C. 9. Pellet M2 beads by centrifugation at 15,000
g for 15 min at 4  C. Wash beads four times with 1 ml RIPA buffer. 10. Resuspend washed beads in 50 μl of 3
sample buffer and incubate for 30 min at room temperature or 10 min at 42  C. It is important to avoid boiling the immunoprecipitated samples as GPCRs have a tendency to aggregate and can form altered species that do not migrate at the predicted molecular weight on SDS-PAGE gels. 11. Separate immunoprecipitates by SDS-PAGE and analyze by western blot as described above in Subheading 3.2. 12. For western blot analyses of immunoprecipitated viral GPCRs and interacting proteins, it is important to use antibody reagents that are derived from a species different from that used in the immunoprecipitation step (in this case, the immunoprecipitating M2 antibody is mouse) to prevent cross reactivity between the immunoprecipitating and primary western antibody. 3.6.3 Detection and Localization of Viral GPCRs by Fluorescence Activated Cell Sorting (FACS)

FACS analysis is a powerful and rapid tool for assessing the expression of a given viral GPCR. Investigators have successfully used this method to demonstrate cell surface expression of epitope-tagged CMV GPCRs [39, 43, 79, 81–86]. For example, Stropes and Miller generated a variety of FLAG-tagged US28 recombinants to study US28 signaling in infected cells and demonstrated that while wildtype and a N-terminal truncation mutant exhibited similar constitutive signaling activities, the N-terminal truncation mutant exhibited decreased cell surface accumulation in comparison to wild-type US28 [39]. 1. Using as few as 1
105 cells, infect cells with a recombinant virus of choice as described throughout this chapter. Cells can be infected with a wide range of MOIs as FACS can accurately detect a positive cell population as small as 2–3%.

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2. Harvest the infected cells by trypsinization Trypsin-EDTA (0.05% Trypsin, 0.53 mM EDTA) and neutralize the trypsinized population by resuspending cells in complete medium containing serum. 3. Wash cells two times with 1
PBS. 4. Stain cells with a primary antibody directed at the epitope tag for 1 h at 4  C. The antibody should be diluted in 1
PBS containing 0.5% BSA. When using FLAG-tagged viral GPCRs, it may be beneficial to use biotinylated anti-FLAG antibody as this coupled with fluorophore conjugated streptavidin can enhance the signal significantly. 5. If the primary antibody used is not preconjugated with a fluorophore, wash the cells as above in PBS, and then stain the cells with the appropriate secondary antibody in the aforementioned buffer for 1 h at 4  C. 6. After a final series of washes in PBS, analyze cells by FACS. 3.6.4 Detecting vGPCRs During Latency by RT-qPCR

UL33, UL78, and US28 are expressed during latency, but these genes, along with US27, are also transcribed during lytic HCMV infection. To detect their transcript expression during latency, one must assess their expression as a ratio relative to that of a gene that is highly expressed during lytic infection, yet is significantly repressed during latency, described in detail below. 1. Plate Kasumi-3 cells in 6-well plates at a density of 5
105 cells/ml in serum-low X-VIVO 15 medium (Lonza) for 48 h. 2. Recount the cells and infect cells at a density of 5
105 cells/ ml with viruses expressing 3
FLAG-tagged vGPCRs at an MOI of 1.0 by centrifugal enhancement at room temperature for 30 min at 450
g. Return the infected cells to the incubator overnight. 3. Remove the cells and infectious media and pellet the cells at room temperature for 4 min at 500
g. Wash cells 3
with PBS, then replate at 5
105 cells/ml in X-VIVO 15 medium (Lonza). 4. At the desired time point postinfection, remove medium and the cells, and gently pellet the cells at room temperature for 4 min at 500
g. Wash cells 3
with PBS. Freeze pellets or move directly to step 5. 5. RNA is isolated using the High Pure RNA Isolation Kit (Roche) following the manufacturer’s protocol. 6. 0.5–1.0 μg RNA is used for reverse transcription in the following reaction (final volume, 50 μl), using TaqMan Reverse Transcription Reagents (ThermoFisher): 5 μl 10
RT Buffer, 10 μl dNTPs, 11 μl MgCl2, 2.5 μl Random Hexamers, 1 μl RNAsin, 1.25 μl RT.

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7. Equal volume (1 μl/well) of each cDNA is then used for the qPCR reaction. Each sample is analyzed in triplicate. We prefer to use SYBR Green (ThermoFisher), though this is not required for the success of this part of the protocol. 8. Use primers for the vGPCR of choice, as well as primers for a cellular control (e.g., GAPDH ) and a highly expressed lytic transcript (e.g., UL122 or UL123). To determine the latent expression of the vGPCR of interest, normalize the vGPCR and lytic transcript to GAPDH , and then take the ratio of vGPCR to lytic transcript. A high ratio denotes latent expression. 3.6.5 Detecting vGPCRs During Latency by Immunoblot

Detection of viral proteins during HCMV latent infection is not trivial. Nonetheless, we have taken advantage of C-terminal triple FLAG epitope tags (3
F) to detect the expression of the vGPCRs during latency. 1. Plate Kasumi-3 cells in 6-well plates at a density of 5
105 cells/ml in serum-low X-VIVO 15 medium (Lonza) for 48 h. 2. Recount the cells and infect at a density of 5
105 cells/ml with viruses expressing 3
FLAG-tagged vGPCRs at an MOI of 1.0 by centrifugal enhancement at room temperature for 30 min at 450
g. Return the infected cells to the incubator overnight. 3. Remove the cells and infectious media and pellet the cells at room temperature for 4 min at 500
g. Wash cells 3
with PBS, then replate at 5
105 cells/ml in X-VIVO 15 medium (Lonza). 4. At 7 days postinfection, remove medium and the cells, and gently pellet the cells at room temperature for 4 min at 500
g. Wash cells 3
with PBS. Freeze pellets or move directly to step 3. 5. Add 100 μl of RIPA buffer containing protease and phosphatase inhibitors. 6. Transfer the lysate to microcentrifuge tubes and incubate on ice for 1 h, vortexing the tubes every 15 min. 7. Clarify the supernatant by centrifugation at 15,000
g for 10 min at 4  C. 8. Transfer supernatant to a clean microcentrifuge tube and determine the protein concentration as described in Subheading 3.2 (Bio-Rad Protein Assay, Bradford, etc.). 9. Denature 100 μg protein in 6
sample buffer and incubate for 30 min at room temperature or 10 min at 42  C. It is important to avoid boiling the samples as GPCRs have a tendency to aggregate and can form altered species that do not migrate at the predicted molecular weight on SDS-PAGE gels.

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10. Separate proteins by SDS-PAGE and analyze by western blot as described above in Subheading 3.2. Here, membranes are blocked using the PBST-based blocking buffer overnight at 4  C with rocking. The primary anti-M2 FLAG antibody is diluted 1:7500 in PBST/2% BSA. 3.6.6 Other Potential Methodologies for the Detection and Localization of Viral GPCRs

Investigators have also utilized a variety of additional techniques, including enzyme-linked immunosorbent assay (ELISA) and immune-electron microscopy (immuno-EM) to assess the intracellular and/or surface expression of vGPCRs [9, 24, 26, 29, 31, 66, 78, 80, 82, 87–89]. The finding that US27 also localizes to the membranes of the cells was demonstrated by immunoprecipitation of cell surface proteins following infection with a US27 FLAGtagged, yet localizes as well to the perinuclear region as shown by IFA [31]. Additionally, Tschische et al. described the heteromerization of HCMV UL33, UL78, and US27 each with US28 in transient transfection assays, and provided evidence of their colocalization using a combination of IFA, immunoprecipitation, and bioluminescence resonance energy transfer (BRET) analyses [89]. Also, Fraile-Ramos and colleagues utilized immuno-EM to discern the intracellular localization of HCMV UL33 and US27 [80]. Proximity ligation assay (PLA) coupled with immunofluorescence microscopy, a powerful approach to determine protein interactions and modifications, was used to show US27’s interaction with cellular CXCR4 [13]. Taken together, these methodologies provide useful tools in examining the expression and localization of the CMV GPCRs within infected cells.

3.7 Methods for Studying vGPCR Function in Animal Models

The experimental approaches and methodology described thus far enable a thorough examination of the biochemical and molecular signaling activities of the vGPCRs and can be used to study the in vitro function of these interesting and conserved cytomegalovirus proteins. However, they fall short of addressing perhaps the most important fundamental questions regarding the CMV GPCRs: (1) What are the primary biological functions of these CMV GPCRs in vivo? (2) How do these functions affect pathogenesis? and (3) How does the signaling activity of the CMV GPCRs mediate their roles in pathogenesis? Therefore, it is essential to extend the biochemical and molecular genetic experiments described thus far with pathogenesis experiments performed in animal models. The results obtained from in vivo studies will provide the genesis for the rational design of experiments aimed at exploring the molecular functions of CMV GPCRs in biologically relevant cellular models. Of the models available for CMV research, the mouse model appears to be the best suited for studies on the role that the GPCRs play in pathogenesis in vivo. Both the M33 and M78 genes exhibit profound growth defects in organs important for viral persistence such as the salivary gland [16–19, 26,

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70]. Moreover, the mouse is easily amenable to genetic manipulation such as transgenesis and gene knock-out, thus allowing investigators to extend pathogenesis studies and potentially explore detailed mechanisms underlying pathogenic processes. Finally, the mouse is a cost-effective model in which one can functionally and mechanistically examine these CMV-encoded GPCRs before moving on to more complex primate models, if warranted. In this section, we will describe basic methodology to assess cytomegalovirus replication/dissemination in the mouse using wild-type and M33 null MCMVs as an example. BACs containing the Smith and K181 strains have been generated and can be manipulated to delete entire GPCR ORFs or induce subtler mutations in signaling motifs, and so on using recombineering methodologies similar to that discussed above [90, 91]. There are a multitude of different strains of mice that have been used to study cytomegalovirus pathogenesis, many offering unique attributes that can be exploited to gain additional insight into the mechanisms of cytomegalovirus replication and spread in vivo. Briefly, strains such as Balb/C are relatively sensitive to MCMV infection, while other strains, such as C57BL/6 are much more resistant to MCMV infection [92, 93]. That nature of this difference lies in the cmv1 locus which encodes the activating NK receptor LY49H in the resistant, but not sensitive strains. Severely immunodeficient mouse strains such as CB17SCID and NOD-SCID-gammaCnull (NSG) mice have emerged as useful models to explore CMV replication and trafficking in the absence of adaptive (CB17SCID) or adaptive/innate NK (NSG) immune function [94– 97]. Particular care must be given to dosage and duration of infection when using the immunodeficient animals as these animals are particularly sensitive to CMV and quickly succumb to the infection. The following five steps (steps 1–5) specifically describe the examination of MCMV replication and spread in the mouse using wildtype and M33 vGPCR null viruses, but can easily be adapted to analyze viral recombinants with deletions/mutation in any vGPCR gene of interest. 1. Five- to six-week-old female mice are obtained from the appropriate vendor and housed under pathogen-free conditions in barrier-filtered SMI cages according to Association for Assessment and Accreditation of Laboratory Animal Care (AALAC) approved guidelines. Mice are given water and chow ad libitum for the duration of the experiment. 2. Six- to twelve-week-old mice are infected with 1
105 to 1
106 PFU/animal of tissue culture derived wild-type or M33 null viruses. Salivary gland derived stocks of many MCMV strains can alternatively be used. However, mutants such the M33 null viruses do not exhibit strong salivary gland

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tropism and thus do not allow for the generation of salivary gland stocks. Thus, in the case of M33 null viruses or other mutants that do not grow in the salivary gland, one is limited to tissue culture derived virus. 3. Virus is injected into animals via one of several routes including intraperitoneal (i.p.), intravenous (i.v.), or subcutaneous (s.c.) into the rear footpad. The i.p. route is the most convenient for routine assessment of MCMV growth in various tissues and in this case a 28-gauge insulin syringe containing up to 300 μl of virus diluted in PBS is used for the infection. 4. At appropriate times postinfection (see Note 11) animals are sacrificed by CO2 asphyxiation and blood is immediately obtained by cardiac puncture and placed into EDTA-treated blood collection tubes. The blood can be used to assess the number of MCMV-infected blood leukocytes as described in step 6. 5. Internal organs and/or tissues of interest (such as spleen, liver, and salivary gland) are removed via dissection and placed into 1 ml DMEM, flash-frozen, and stored at 80  C until use. To assess virus titers in organs and/or tissues proceed to step 9. The following three steps (steps 6–8) are used to assess the number of infected leukocytes in blood by infectious center assay. 6. Dilute 500 μl of blood in 5 ml RBC lysis buffer. Mix and incubate at RT until RBCs lyse (the solution will change from opaque to clear; but remain deep red). Pellet WBCs for 10 min at 400
g. Wash the WBCs three times with sterile 1
PBS to remove hemoglobin and platelets. Resuspend washed WBCs in 1 ml of 1
PBS. 7. Transfer 1
104 to 1
105 WBC to MEF monolayers (see Note 12), incubate for 3–4 h to allow WBCs to settle to bottom of well and come in contact with MEFs. Carefully remove medium without disturbing settled WBCs and overlay with DMEM containing 0.75% CMC. Incubate undisturbed for 6–7 days. 8. Remove medium, fix monolayers with methanol, and stain with Giemsa diluted 1:5. Count plaques. Plaques that develop arise as a consequence of a single infected WBC that is productively shedding virus, hence the term infectious center. The following four steps (steps 9–12) are used to assess virus titers in organs/tissues by plaque assay. 9. Thaw tissue that was suspended in 1 ml DMEM and flashfrozen as described in step 5. Transfer to Dounce and homogenize organ using 20–30 passes with the tightfitting glass pestle. Ensure visually that the tissue is completely homogenized—if not, proceed with additional passes until the tissue is completely disrupted.

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10. Centrifuge for 5 min in a microcentrifuge at 5000
g to pellet cellular and tissue debris. Transfer supernatant to fresh tube. 11. Transfer dilutions of tissue supernatant to MEF monolayers (see Note 12) and incubate for 3–4 h to allow virus adsorption. Carefully remove medium and overlay with DMEM containing 0.75% CMC. Incubate undisturbed for 4–5 days. Virus titers in organs vary dramatically depending on initial virus dose and dpi, so care should be taken to ensure that the dilutions of tissue supernatant used in the assay will allow for quantitation of plaques in each well. 12. Remove medium, fix monolayers with methanol, and stain with Giemsa diluted 1:5. Count plaques. Using in vivo assays like the one just described, it is evident that the CMV GPCRs confer important activities that facilitate viral replication in the whole organism. It is important to ensure that the observed phenotype is due to deletion/alteration of the targeted gene, and this can be accomplished by using “rescue” viruses in which the mutated region is reverted to wild-type. While it is clear that the GPCRs and their ability to signal through G-proteins are essential for replication in vivo, it is not clear what specific signaling pathways are involved or how activation of these signaling pathways facilitate replication. The power of mouse genetics combined with in vivo growth assessment of viruses with GPCR mutations should provide important answers to these questions. It is the answers to these questions that should be at the forefront of future investigations aimed at exploring molecular and biochemical properties of the vGPCRs. 3.8 Conclusions and Discussion of Current State of the Art Techniques Useful for Studying vGPCR Signaling/Function

Techniques such as transient transfections and related gene delivery methodologies have proved to be invaluable in providing a basic understanding of the CMV GPCRs and how they function in vitro. However, continued vertical advancement of our understanding of the CMV GPCRs requires us as investigators to distance ourselves from standard in vitro techniques and begin to perform studies in the context of virus-infected cells using clinical strains of HCMV and cell types important for in vivo pathogenesis. Taking advantage of CMV GPCR mutants constructed by recombineering techniques is critical for the successful transition to these more sophisticated types of experiments. The tools and resources, including bacterial recombineering techniques, now exist for the cytomegaloviruses, therefore making such studies possible. BAC recombineering protocols provide efficient means to derive viral recombinants for use in both in vitro and in vivo studies. The benefit to generating mutants in BAC viral DNA as opposed to utilizing expression plasmids is that one can investigate the function of the GPCRs in the milieu of the remaining viral ORFs and at

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physiologically relevant expression levels. Finally, the tools now available for studying CMV GPCRs affords us as investigators the ability to perform high-throughput screens to search for novel viral GPCR therapeutics that may influence HCMV infection and/or replication. Over 1/3rd of marketed drugs target cellular GPCRs, and thus the CMV GPCRs are attractive targets. Both the gammaand beta-herpesvirus subfamilies encode GPCRs, and in animal models, these proteins have been shown to aid in viral pathogenesis. Thus, it seems likely that herpesviruses have hijacked cellular GPCRs to promote viral replication and dissemination in the host. Utilizing the current resources and technologies, we will undoubtedly uncover the function of these proteins, and perhaps exploit their activities in an effort to develop novel antiviral therapies to combat HCMV infections.

4

Notes 1. Typical doubling times are 48–72 h for HS68 fibroblasts and 18–24 h for HEK-293 cells, HEK-293T cells, ARPE-19 cells, and THP-1 cells. 2. Primary human CD34+ HPC isolation, culture methods, media requirements, and infection methods are described in detail in Chapter 8 of this book. 3. To activate the sodium orthovanadate, prepare a 200 mM stock solution, adjust the pH to 10 using NaOH or HCl and boil until colorless. Cool to room temperature, readjust to pH 10 and repeat until the solution stabilizes at pH 10 and remains colorless. Store the activated sodium orthovanadate in aliquots in the 20  C freezer. The protease inhibitors aprotinin, leupeptin, and PMSF can be substituted for Complete Mini Protease tabs (Roche). 4. Biotin is degraded by light—make fresh and use plates 1 month. 5. The different medium formulations contain various amounts of unlabeled myoinositol and therefore, it may be beneficial to use MEM as the inositol concentration is lower and gives better labeling. In the case of THP-1 cells, cells should be labeled in RPMI-160 media lacking inositol. 6. In many cases, simply eluting the total inositol phosphates will provide an extremely accurate measurement of receptor signaling. In this case after Subheading 3.1, step 18, simply transfer the columns to scintillation vials and elute the total inositol phosphates with 4 ml 0.1 M formic acid/1.0 M ammonium formate.

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7. The pFR-LUC and pFA2 plasmids are part of the PathDetect in vivo Signal Transduction trans-reporting system available from Agilent Technologies™. More information about these plasmids and the trans-reporting system can be found at: http://www.genomics.agilent.com/files/Manual/219000. pdf 8. Passive Lysis Buffer (PLB), Luciferase Assay reagent II (LAR II), and Stop&Glo are components of the Promega DualLuciferase® Reporter Assay System. More information about this system can be found at http://www.promega.com/ products/reporter-assays-and-transfection/reporter-assays 9. The bacterial strains SW102, SW105, and SW106 produce endotoxins, which are co-purified with the BAC DNA and thus can be introduced into mammalian cells during the transfection process. Endotoxins stimulate components of the innate immune response in mammalian cells, and therefore are toxic to cells in tissue culture. Thus, adding a step within the alkaline lysis protocol to remove endotoxins will greatly improve the health of the mammalian cells posttransfection of the BAC DNA and greatly improve the overall transfection efficiency. 10. Infections below an MOI of 0.1 PFU/cell in ARPE19 cells result in insufficient viral output for assessment by plaque assay and are therefore not recommended. 11. MCMV replicates in a large number of cell types and tissue. In particular, the virus can be found at high levels during the acute phase in organs such as spleen and liver (3–5 dpi) and during the persistent phase in tissues such as the salivary gland (12–21 dpi). 12. One day prior to using for plaque or infectious center assays, plate 100,000 primary MEFs (passage 2–8) into each well of a 12-well plate. References 1. Khanna R, Diamond DJ (2006) Human cytomegalovirus vaccine: time to look for alternative options. Trends Mol Med 12:26–33 2. Chee MS et al (1990) Analysis of the proteincoding content of the sequence of human cytomegalovirus strain AD169. Curr Top Microbiol Immunol 154:125–169 3. Chee MS, Satchwell SC, Preddie E, Weston KM, Barrell BG (1990) Human cytomegalovirus encodes three G protein-coupled receptor homologues. Nature 344:774–777 4. Dorsam RT, Gutkind JS (2007) G-proteincoupled receptors and cancer. Nat Rev Cancer 7:79–94

5. Sodhi A, Montaner S, Gutkind JS (2004) Does dysregulated expression of a deregulated viral GPCR trigger Kaposi’s sarcomagenesis? FASEB J 18:422–427 6. Miller-Kittrell M, Sparer TE (2009) Feeling manipulated: cytomegalovirus immune manipulation. Virol J 6:4 7. Casarosa P et al (2001) Constitutive signaling of the human cytomegalovirus-encoded chemokine receptor US28. J Biol Chem 276:1133–1137 8. Kuhn DE, Beall CJ, Kolattukudy PE (1995) The cytomegalovirus US28 protein binds

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multiple CC chemokines with high affinity. Biochem Biophys Res Commun 211:325–330 9. Waldhoer M, Kledal TN, Farrell H, Schwartz TW (2002) Murine cytomegalovirus (CMV) M33 and human CMV US28 receptors exhibit similar constitutive signaling activities. J Virol 76:8161–8168 10. Minisini R et al (2003) Constitutive inositol phosphate formation in cytomegalovirusinfected human fibroblasts is due to expression of the chemokine receptor homologue pUS28. J Virol 77:4489–4501 11. Gao JL, Murphy PM (1994) Human cytomegalovirus open reading frame US28 encodes a functional beta chemokine receptor. J Biol Chem 269:28539–28542 12. Casarosa P et al (2003) Constitutive signaling of the human cytomegalovirus-encoded receptor UL33 differs from that of its rat cytomegalovirus homolog R33 by promiscuous activation of G proteins of the Gq, Gi, and Gs classes. J Biol Chem 278:50010–50023 13. Tu CC, Arnolds KL, O’Connor CM, Spencer JV (2018) Human cytomegalovirus UL111A and US27 gene products enhance the CXCL12/CXCR4 signaling axis via distinct mechanisms. J Virol 92:e01981–e01917 14. Gruijthuijsen YK et al (2002) The rat cytomegalovirus R33-encoded G protein-coupled receptor signals in a constitutive fashion. J Virol 76:1328–1338 15. Sherrill JD, Miller WE (2006) G proteincoupled receptor (GPCR) kinase 2 regulates agonist-independent Gq/11 signaling from the mouse cytomegalovirus GPCR M33. J Biol Chem 281:39796–39805 16. Davis-Poynter NJ et al (1997) Identification and characterization of a G protein-coupled receptor homolog encoded by murine cytomegalovirus. J Virol 71:1521–1529 17. Case R et al (2008) Functional analysis of the murine cytomegalovirus chemokine receptor homologue M33: ablation of constitutive signaling is associated with an attenuated phenotype in vivo. J Virol 82:1884–1898 18. Cardin RD, Schaefer GC, Allen JR, DavisPoynter NJ, Farrell HE (2009) The M33 chemokine receptor homolog of murine cytomegalovirus exhibits a differential tissue-specific role during in vivo replication and latency. J Virol 83:7590–7601 19. Sherrill JD et al (2009) Activation of intracellular signaling pathways by the murine cytomegalovirus G protein-coupled receptor M33 occurs via PLC-{beta}/PKC-dependent and -independent mechanisms. J Virol 83:8141–8152

20. Bittencourt FM, Wu SE, Bridges JP, Miller WE (2014) The M33 G protein-coupled receptor encoded by murine cytomegalovirus is dispensable for hematogenous dissemination but is required for growth within the salivary gland. J Virol 88:11811–11824 21. Farrell HE, Bruce K, Ma J, Davis-Poynter N, Stevenson PG (2018) Human cytomegalovirus US28 allows dendritic cell exit from lymph nodes. J Gen Virol 99:1509–1514 22. Farrell HE et al (2017) Murine Cytomegalovirus Spreads by Dendritic Cell Recirculation. mBio 8:e01264–e01217 23. Beisser PS et al (1998) The R33 G proteincoupled receptor gene of rat cytomegalovirus plays an essential role in the pathogenesis of viral infection. J Virol 72:2352–2363 24. Margulies BJ, Browne H, Gibson W (1996) Identification of the human cytomegalovirus G protein-coupled receptor homologue encoded by UL33 in infected cells and enveloped virus particles. Virology 225:111–125 25. O’Connor CM, Shenk T. Unpublished observations. In 26. Oliveira SA, Shenk TE (2001) Murine cytomegalovirus M78 protein, a G protein-coupled receptor homologue, is a constituent of the virion and facilitates accumulation of immediate-early viral mRNA. Proc Natl Acad Sci U S A 98:3237–3242 27. Beisser PS, Grauls G, Bruggeman CA, Vink C (1999) Deletion of the R78 G protein-coupled receptor gene from rat cytomegalovirus results in an attenuated, syncytium-inducing mutant strain. J Virol 73:7218–7230 28. Kaptein SJ et al (2003) The rat cytomegalovirus R78 G protein-coupled receptor gene is required for production of infectious virus in the spleen. J Gen Virol 84:2517–2530 29. O’Connor CM, Shenk T (2012) Human cytomegalovirus pUL78 G protein-coupled receptor homologue is required for timely nuclear accumulation of virion constituents in epithelial cells but not fibroblasts. J Virol 86:11425–11433 30. Michel D et al (2005) The human cytomegalovirus UL78 gene is highly conserved among clinical isolates, but is dispensable for replication in fibroblasts and a renal artery organculture system. J Gen Virol 86:297–306 31. O’Connor CM, Shenk T (2011) Human cytomegalovirus pUS27 G protein-coupled receptor homologue is required for efficient spread by the extracellular route but not for direct cellto-cell spread. J Virol 85:3700–3707 32. Boeck JM, Stowell GA, O’Connor CM, Spencer JV (2018) The human cytomegalovirus

vGPCR Methods US27 gene product constitutively activates antioxidant response element-mediated transcription through Gbetagamma, phosphoinositide 3-kinase, and nuclear respiratory factor 1. J Virol 92(23):e00644–e00618 33. Arnolds KL, Lares AP, Spencer JV (2013) The US27 gene product of human cytomegalovirus enhances signaling of host chemokine receptor CXCR4. Virology 439:122–131 34. Boeck JM, Spencer JV (2017) Effect of human cytomegalovirus (HCMV) US27 on CXCR4 receptor internalization measured by fluorogen-activating protein (FAP) biosensors. PLoS One 12:e0172042 35. Tu CC, Spencer JV (2014) The DRY box and C-terminal domain of the human cytomegalovirus US27 gene product play a role in promoting cell growth and survival. PLoS One 9: e113427 36. Kledal TN, Rosenkilde MM, Schwartz TW (1998) Selective recognition of the membrane-bound CX3C chemokine, fractalkine, by the human cytomegalovirus-encoded broad-spectrum receptor US28. FEBS Lett 441:209–214 37. Bodaghi B et al (1998) Chemokine sequestration by viral chemoreceptors as a novel viral escape strategy: withdrawal of chemokines from the environment of cytomegalovirusinfected cells. J Exp Med 188:855–866 38. Vieira J, Schall TJ, Corey L, Geballe AP (1998) Functional analysis of the human cytomegalovirus US28 gene by insertion mutagenesis with the green fluorescent protein gene. J Virol 72:8158–8165 39. Stropes MP, Miller WE (2008) Functional analysis of human cytomegalovirus pUS28 mutants in infected cells. J Gen Virol 89:97–105 40. Wu SE, Miller WE (2016) The HCMV US28 vGPCR induces potent Galphaq/PLC-beta signaling in monocytes leading to increased adhesion to endothelial cells. Virology 497:233–243 41. Billstrom MA, Johnson GL, Avdi NJ, Worthen GS (1998) Intracellular signaling by the chemokine receptor US28 during human cytomegalovirus infection. J Virol 72:5535–5544 42. Streblow DN et al (1999) The human cytomegalovirus chemokine receptor US28 mediates vascular smooth muscle cell migration. Cell 99:511–520 43. Stropes MP, Schneider OD, Zagorski WA, Miller JL, Miller WE (2009) The carboxyterminal tail of human cytomegalovirus (HCMV) US28 regulates both chemokineindependent and chemokine-dependent

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signaling in HCMV-infected cells. J Virol 83:10016–10027 44. Miller WE et al (2012) US28 is a potent activator of phospholipase C during HCMV infection of clinically relevant target cells. PLoS One 7:e50524 45. Noriega VM et al (2014) Human cytomegalovirus US28 facilitates cell-to-cell viral dissemination. Viruses 6:1202–1218 46. Melnychuk RM et al (2004) Human cytomegalovirus-encoded G protein-coupled receptor US28 mediates smooth muscle cell migration through Galpha12. J Virol 78:8382–8391 47. Moepps B et al (2008) Constitutive serum response factor activation by the viral chemokine receptor homologue pUS28 is differentially regulated by Galpha(q/11) and Galpha (16). Cell Signal 20:1528–1537 48. Slinger E et al (2010) HCMV-encoded chemokine receptor US28 mediates proliferative signaling through the IL-6-STAT3 axis. Sci Signal 3:ra58 49. Soroceanu L et al (2011) Human cytomegalovirus US28 found in glioblastoma promotes an invasive and Angiogenic phenotype. Cancer Res 71:6643–6653 50. Boomker JM, The TH, de Leij LF, Harmsen MC (2006) The human cytomegalovirusencoded receptor US28 increases the activity of the major immediate-early promoter/ enhancer. Virus Res 118:196–200 51. McLean KA, Holst PJ, Martini L, Schwartz TW, Rosenkilde MM (2004) Similar activation of signal transduction pathways by the herpesvirus-encoded chemokine receptors US28 and ORF74. Virology 325:241–251 52. Krishna BA, Humby MS, Miller WE, O’Connor CM (2019) Human cytomegalovirus G protein-coupled receptor US28 promotes latency by attenuating c-fos. Proc Natl Acad Sci U S A 116:1755–1764 53. Maussang D et al (2006) Human cytomegalovirus-encoded chemokine receptor US28 promotes tumorigenesis. Proc Natl Acad Sci U S A 103:13068–13073 54. Maussang D et al (2009) The human cytomegalovirus-encoded chemokine receptor US28 promotes angiogenesis and tumor formation via cyclooxygenase-2. Cancer Res 69:2861–2869 55. Maussang D, Vischer HF, Schreiber A, Michel D, Smit MJ (2009) Pharmacological and biochemical characterization of human cytomegalovirus-encoded G protein-coupled receptors. Methods Enzymol 460:151–171

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56. Baryawno N et al (2011) Detection of human cytomegalovirus in medulloblastomas reveals a potential therapeutic target. J Clin Invest 121:4043–4055 57. Heukers R et al (2018) The constitutive activity of the virally encoded chemokine receptor US28 accelerates glioblastoma growth. Oncogene 37:4110–4121 58. Bongers G et al (2010) The cytomegalovirusencoded chemokine receptor US28 promotes intestinal neoplasia in transgenic mice. J Clin Invest 120:3969–3978 59. Cai ZZ et al (2016) Human cytomegalovirusencoded US28 may act as a tumor promoter in colorectal cancer. World J Gastroenterol 22:2789–2798 60. Humby MS, O’Connor CM (2015) Human cytomegalovirus US28 is important for latent infection of hematopoietic progenitor cells. J Virol 90:2959–2970 61. Goodrum F, Reeves M, Sinclair J, High K, Shenk T (2007) Human cytomegalovirus sequences expressed in latently infected individuals promote a latent infection in vitro. Blood 110:937–945 62. Cheng S et al (2017) Transcriptome-wide characterization of human cytomegalovirus in natural infection and experimental latency. Proc Natl Acad Sci U S A 114:E10586–E10595 63. Cheung AK, Abendroth A, Cunningham AL, Slobedman B (2006) Viral gene expression during the establishment of human cytomegalovirus latent infection in myeloid progenitor cells. Blood 108:3691–3699 64. Krishna BA et al (2017) Latency-associated expression of human cytomegalovirus US28 attenuates cell signaling pathways to maintain latent infection. mBio 8:e01754–e01717 65. Zhu D et al (2018) Human cytomegalovirus reprogrammes haematopoietic progenitor cells into immunosuppressive monocytes to achieve latency. Nat Microbiol 3:503–513 66. Waldhoer M et al (2003) The carboxyl terminus of human cytomegalovirus-encoded 7 transmembrane receptor US28 camouflages agonism by mediating constitutive endocytosis. J Biol Chem 278:19473–19482 67. Miller WE et al (2001) Identification of a motif in the carboxyl terminus of beta -arrestin2 responsible for activation of JNK3. J Biol Chem 276:27770–27777 68. Goodwin CM, Ciesla JH, Munger J (2018) Who’s driving? Human cytomegalovirus, interferon, and NFkappaB signaling. Viruses 10:447 69. de Wit RH et al (2016) Human cytomegalovirus encoded chemokine receptor US28

activates the HIF-1alpha/PKM2 axis in glioblastoma cells. Oncotarget 7:67966–67985 70. Farrell HE et al (2011) Partial functional complementation between human and mouse cytomegalovirus chemokine receptor homologues. J Virol 85:6091–6095 71. Brune W, Messerle M, Koszinowski UH (2000) Forward with BACs: new tools for herpesvirus genomics. Trends Genet 16:254–259 72. Datsenko KA, Wanner BL (2000) One-step inactivation of chromosomal genes in Escherichia coli K-12 using PCR products. Proc Natl Acad Sci U S A 97:6640–6645 73. Warming S, Costantino N, Court DL, Jenkins NA, Copeland NG (2005) Simple and highly efficient BAC recombineering using galK selection. Nucleic Acids Res 33:e36 74. Tischer BK, von Einem J, Kaufer B, Osterrieder N (2006) Two-step red-mediated recombination for versatile high-efficiency markerless DNA manipulation in Escherichia coli. BioTechniques 40:191–197 75. Sinzger C et al (2008) Cloning and sequencing of a highly productive, endotheliotropic virus strain derived from human cytomegalovirus TB40/E. J Gen Virol 89:359–368 76. Dunn W et al (2003) Functional profiling of a human cytomegalovirus genome. Proc Natl Acad Sci U S A 100:14223–14228 77. Yu D, Silva MC, Shenk T (2003) Functional map of human cytomegalovirus AD169 defined by global mutational analysis. Proc Natl Acad Sci U S A 100:12396–12401 78. Margulies BJ, Gibson W (2007) The chemokine receptor homologue encoded by US27 of human cytomegalovirus is heavily glycosylated and is present in infected human foreskin fibroblasts and enveloped virus particles. Virus Res 123:57–71 79. Mokros T et al (2002) Surface expression and endocytosis of the human cytomegalovirusencoded chemokine receptor US28 is regulated by agonist-independent phosphorylation. J Biol Chem 277:45122–45128 80. Fraile-Ramos A et al (2002) Localization of HCMV UL33 and US27 in endocytic compartments and viral membranes. Traffic 3:218–232 81. Droese J et al (2004) HCMV-encoded chemokine receptor US28 employs multiple routes for internalization. Biochem Biophys Res Commun 322:42–49 82. Penfold ME, Schmidt TL, Dairaghi DJ, Barry PA, Schall TJ (2003) Characterization of the rhesus cytomegalovirus US28 locus. J Virol 77:10404–10413

vGPCR Methods 83. Pleskoff O et al (2005) The human cytomegalovirus-encoded chemokine receptor US28 induces caspase-dependent apoptosis. FEBS J 272:4163–4177 84. Pleskoff O, Treboute C, Alizon M (1998) The cytomegalovirus-encoded chemokine receptor US28 can enhance cell-cell fusion mediated by different viral proteins. J Virol 72:6389–6397 85. Pleskoff O et al (1997) Identification of a chemokine receptor encoded by human cytomegalovirus as a cofactor for HIV-1 entry. Science 276:1874–1878 86. Vomaske J et al (2009) Differential ligand binding to a human cytomegalovirus chemokine receptor determines cell type-specific motility. PLoS Pathog 5:e1000304 87. Casarosa P et al (2005) CC and CX3C chemokines differentially interact with the N terminus of the human cytomegalovirus-encoded US28 receptor. J Biol Chem 280:3275–3285 88. Casarosa P et al (2003) Identification of the first nonpeptidergic inverse agonist for a constitutively active viral-encoded G proteincoupled receptor. J Biol Chem 278:5172–5178 89. Tschische P, Tadagaki K, Kamal M, Jockers R, Waldhoer M (2011) Heteromerization of human cytomegalovirus encoded chemokine receptors. Biochem Pharmacol 82:610–619 90. Redwood AJ et al (2005) Use of a murine cytomegalovirus K181-derived bacterial

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artificial chromosome as a vaccine vector for immunocontraception. J Virol 79:2998–3008 91. Messerle M, Crnkovic I, Hammerschmidt W, Ziegler H, Koszinowski UH (1997) Cloning and mutagenesis of a herpesvirus genome as an infectious bacterial artificial chromosome. Proc Natl Acad Sci U S A 94:14759–14763 92. Lee SH et al (2001) Susceptibility to mouse cytomegalovirus is associated with deletion of an activating natural killer cell receptor of the C-type lectin superfamily. Nat Genet 28:42–45 93. Webb JR, Lee SH, Vidal SM (2002) Genetic control of innate immune responses against cytomegalovirus: MCMV meets its match. Genes Immun 3:250–262 94. Upton JW, Kaiser WJ, Mocarski ES (2010) Virus inhibition of RIP3-dependent necrosis. Cell Host Microbe 7:302–313 95. Ghazal P, Messerle M, Osborn K, Angulo A (2003) An essential role of the enhancer for murine cytomegalovirus in vivo growth and pathogenesis. J Virol 77:3217–3228 96. Pollock JL, Virgin HW (1995) Latency, without persistence, of murine cytomegalovirus in the spleen and kidney. J Virol 69:1762–1768 97. Okada M, Minamishima Y (1987) The efficacy of biological response modifiers against murine cytomegalovirus infection in normal and immunodeficient mice. Microbiol Immunol 31:45–57

Chapter 10 Analysis of Cytomegalovirus Glycoprotein and Cellular Receptor Interactions Jamil Mahmud and Gary C. Chan Abstract Human cytomegalovirus (HCMV) entry into host cells is a complex process involving interactions between an array of viral glycoproteins with multiple host cell surface receptors. A significant amount of research has been devoted toward identifying these glycoprotein and cellular receptor interactions as the broad cellular tropism of HCMV suggests a highly regulated yet adaptable process that controls viral binding and penetration. However, deciphering the initial binding and cellular receptor activation events by viral glycoproteins remains challenging. The relatively low abundance of receptors and/or interactions with glycoproteins during viral entry, the hydrophobicity of membrane receptors, and the rapid degradation and recycling of activated receptors have complicated the analysis of HCMV entry and the cellular signaling pathways initiated by HCMV engagement to the host membrane. Here, we describe the different methodologies used in our laboratory and others to analyze the interactions between HCMV glycoproteins and host cellular receptors during the entry stage of the viral life cycle. Key words Gradient purification, Membrane receptor, Soluble glycoprotein, Retroviral vector, Western blot, Coimmunoprecipitation

1

Introduction Human cytomegalovirus (HCMV) encodes an array of glycoproteins allowing for a broad infection tropism. During the entry process HCMV glycoproteins mediate viral attachment and entry into the host cells in an orderly fashion [1, 2]. Initial tethering of HCMV to the cell surface is facilitated by glycoprotein gM/gN and/or gB binding to heparin sulfated proteoglycans (HSPGs) [3, 4]. This reversible binding is rapidly replaced by an irreversible attachment mediated by specific glycoprotein and cellular receptor interactions, which appear to be dependent on virus strain and cell type. The key glycoproteins involved during viral entry include gB, gH, gL, and gO, as well as HCMV proteins UL128-131 [5– 10]. Glycoprotein gH is found in three complexes: a gH/gL

Andrew D. Yurochko (ed.), Human Cytomegaloviruses: Methods and Protocols, Methods in Molecular Biology, vol. 2244, https://doi.org/10.1007/978-1-0716-1111-1_10, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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dimer, a gH/gL/gO trimer, and a gH/gL/UL128-131 pentamer [11–13]. Following initial tethering of the virion to the cell surface, gH complexes bind integrins to induce cell signaling needed for viral entry into target cells. Specifically, the trimeric gH complex mediates entry into fibroblasts, while the pentameric gH complex is required for entry into epithelial, endothelial, and myeloid cells [14–17]. Glycoprotein gB forms a homotrimer through disulfide bonds [18] and binds epidermal growth factor receptor (EGFR) or platelet-derived growth factor receptor (PDGFR) α to trigger host signaling required for viral entry [8, 19]. However, there are conflicting reports about whether EGFR and/or PDGFRα act as bona fide entry receptors [8, 10, 19–24]. Although the discrepancies between studies are likely in part due to differences in the HCMV strains and cell types used, the inherent difficulty in assessing direct glycoprotein binding and activation of EGFR and PDGFRα has also contributed to the lack of clarity surrounding the involvement of these receptors during viral entry. For example, EGFR forms aggregation during normal sample preparation for coimmunoprecipitation (co-IP) and western blotting due to its highly hydrophobic nature (contains >60% hydrophobic amino acids) and long cytoplasmic region (46% of total protein) [25]. Aggregation of EGFR often results in failure to co-IP and/or detect protein expression. Modifications to traditional protocols are needed to detect EGFR and are particularly important when working with cell types expressing low levels of EGFR such as monocytes (Fig. 1). HCMV binding and activation of cellular receptors during viral entry disrupts normal cell signaling to induce profound changes in the host cell microenvironment, which are required for almost every aspects of the viral lifecycle, including entry, replication, and persistence [26–28]. As expected by the broad cellular tropism, HCMV binds and activates a number of different receptors during HCMV entry, including EGFR, PDGFRα, integrins, TLR2, Nrp1, CD147, CD90, and BST2 [8, 9, 19, 23, 24, 29–34]. However, identifying interactions between viral glycoproteins and cell surface receptors has been difficult due to the low abundance of receptors and/or interactions, the hydrophobicity of membrane receptors, and the rapid degradation and recycling of activated receptors. Consequently, much consideration must be taken when attempting to identify and/or confirm glycoprotein and cellular receptor interactions during viral entry. This chapter details the different techniques used routinely in our laboratory to study binding and activation of cellular receptors by HCMV glycoproteins.

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Fig. 1 Detection of EGFR versus integrin β1 by western blotting requires different sample preparation techniques. EGFR has a longer cytoplasmic domain (~46% of total protein) than integrin β1 (~6%) and has overall higher abundance of hydrophobic amino acids residues (~60% vs. ~30%). Due it its hydrophobic nature, EGFR forms aggregates at higher temperature, and thus resolves best with lower temperature sample preparation [25]. Contrarily, integrin β1 resolves best with typical sample preparation at 95
C in the presence of a reducing agent. p-EGFR phospho-EGFR, p integrin β1 phospho-integrin β1, SDS Sodium dodecyl sulfate, BME 2-mercaptoethanol, RT room temperature

2

Materials

2.1 Purification of Virions

1. Human embryotic lung cells (HEL) (see Note 1). 2. Dulbecco’s minimal essential media (DMEM) supplemented with 4% fetal bovine serum (FBS) (4% DMEM). 3. TB40E or other clinical/laboratory isolates of HCMV (see Note 2). 4. 10 mM Tris–HCl, 150 mM NaCl, pH 7.5 Tris-buffered saline (TBS). 5. Sorbitol gradient solutions: 20%, 30%, 40%, and 60% sorbitol solutions (w/v) in TBS.

2.2 Purification of Soluble Glycoproteins gB and gH (sgB and sgH) 2.2.1 Isolation of HCMV Bacterial Artificial Chromosome (BAC)

1. Ribonuclease A. 2. Buffer S1: 50 mM pH 8.0 Tris, 10 mM pH 8.0 EDTA, and 100 μg/ml RNase A. 3. Buffer S2: Part 1 (400 mM NaOH), Part 2 (2% SDS). Mix at 1:1 ratio before use for a final concentration of 200 mM NaOH and 1% SDS. 4. Buffer S3: 2.8 M pH 5.1 KAc. 5. Buffer TEN: 10 mM pH 7.4–8.0 Tris, 1 mM pH 8.0 EDTA, and 150 mM NaCl.

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2.2.2 Cloning HCMV sgB and sgH

1. Retroviral vector pQCXIN (Clontech). 2. Platinum™ PCR SuperMix High Fidelity (ThermoFisher Scientific). 3. E.Z.N.A.® Gel Extraction Kit (V-spin) (Omega). 4. Restriction enzymes. 5. T4 DNA ligase. 6. Competent cells for cloning, 5-alpha Competent E. coli (High Efficiency) (New England Biolabs). 7. Endotoxin free plasmid extraction kit (Qiagen). 8. Forward and reverse primers with AgeI and BamHI restrictions sites (see Note 3): sgB-AgeI forward primer, 50 -CCCACCGGT GACGAACATGGAATCCAGGAT-30 ; sgB-BamHI reverse primer, 50 -GCGGGATCCCTAATGGTGATGGTGATGA TGCTGCTTGTACGAGTTGAATTC-30 ; sgH-AgeI forward primer, 50 -CCCACCGGTCCGCGCTATGCGGCCCGG 0 CCT-3 ; sgH-BamHI reverse primer, 50 - GCGGGATCCC TAATGGTGATGGTGATGATGGTCGGTGGCGTCCACG ACGAC-30 .

2.2.3 Generating Stably sgB and sgH Expressing Expi293F Cells

1. Expi293™ Expression System Kit (ThermoFisher Scientific). 2. GP2-293 retroviral packaging system (Clontech). 3. G418, geneticin. 4. Baffled polycarbonate Erlenmeyer flask with vent cap.

2.2.4 Purifying sgB and sgH

1. Lysis buffer: 50 mM pH 8.0 sodium phosphate, 200 mM NaCl, 10% glycerol, 25 mM imidazole. Prepare 1 l of buffer by mixing 7.1 g sodium phosphate dibasic, 11.69 g NaCl, 100 ml glycerol, and 12.5 μl of imidazole from 2 M stock with H2O to a volume of 980 ml. Adjust the pH to 8 using concentrated hydrochloric acid. Adjust the volume to 1000 ml with water. Check the pH and adjust if necessary. Filter through a 0.2 μm polyethersulfone membrane and store at 4
C. 2. Phosphate buffer saline, PBS (10 mM phosphate buffer and 140 mM NaCl, pH 7.4). 3. Chromatography column (Bio-Rad, catalog # 731-1550). 4. Ni-charged resin (Bio-Rad, catalog # 780-0800). 5. Wash buffer: 25 mM pH 7.4 Imidazole, with PBS. 6. Elution buffer: 250 mM pH 7.4 Imidazole, with PBS. 7. Dialysis kit. 8. Bradford assay kit.

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2.3 Detection of Cellular Receptor Activation by Western Blot

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1. Sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS-PAGE) reagents. 2. Polyvinylidene fluoride (PVDF) membrane. 3. TBS containing 0.1% Tween 20. 4. 5% bovine serum albumen (BSA) and 0.1% Tween 20 diluted in TBS (5%-TBST). 5. Lysis buffer: 50 mM pH 7.5 Tris, 5 mM EDTA, 100 mM NaCl, 1% Triton X-100, 0.1% SDS, and 10% glycerol. 6. Protease and phosphatase inhibitor cocktail (Sigma-Aldrich). 7. Nonreducing 6 SDS-sample buffer: 375 mM pH 6.8 Tris– HCl, 6% SDS, 50% glycerol, and 0.045% bromophenol. 8. 2-mercaptoethanol, proteomics grade. 9. Horseradish antibodies.

peroxidase

(HRP)-conjugated

anti-species

10. Chemiluminescence detection reagents. 2.4 Coimmunoprecipitation of HCMV Glycoproteins and Cellular Receptors

1. NP40 Cell Lysis Buffer (see Note 4). 2. Protease and phosphatase inhibitor cocktail. 3. Dynabeads proteins A/G (ThermoFisher Scientific). 4. IgG elution buffer (ThermoFisher Scientific). 5. 1 M pH 9.0 Tris.

3

Methods

3.1 Purification of HCMV

1. Infect HF at 100% confluency with HCMV (MOI 0.01–0.1) in 4%-DMEM. 2. Incubate infected cells and change media every 2–3 days until a cytopathic effect (CPE) of 90% is achieved (approximately 10–14 days). 3. Change media and continue to incubate infected HF for an additional 3–5 days. 4. Collect the supernatant (see Note 5) and spin at 400  g for 10 min to remove cells and cellular debris. 5. Concentrate virions by centrifugation through a 20% sorbitol cushion at 20,000  g for 1 h at room temperature (RT). 6. Resuspend pellet in TBS and layer onto 20–70% sorbitol step gradients. 7. Band virus by centrifugation at 100,000  g for 1 h at RT. Collect band at the 50–60% density interface, which represents intact enveloped virus.

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3.2 Purification of sgB and sgH 3.2.1 Purify HCMV Bacterial Artificial Chromosome (BAC)

1. Streak bacterial clone containing the HCMV genomic backbone from glycerol stock onto LB agar media with appropriate antibiotic and incubate overnight at 32
C (see Note 6). 2. Next day, inoculate 10 ml LB broth from a single bacterial colony with appropriate antibiotic and incubate for 16–18 h in a shaker incubator at 225 rpm and 32
C. 3. Centrifuge the cells at 3200  g for 10 min. 4. Remove the supernatant and resuspend the pellet in 350 μl S1 buffer with 100 μg/ml RNase A. 5. Transfer the cell paste into a fresh 1.5 ml centrifuge tube and add 350 μl S2 buffer. Mix gently by inverting the tube 6–8 times. 6. Incubate for 4 min to allow cell lysis. 7. Add 350 μl of ice-cold S3 buffer and mix gently by inverting the tube 8–12 times. Incubate on ice for 7 min. 8. Centrifuge at 17,000  g for 10 min at 4
C. 9. Remove the clear lysate using a wide-bore tip as large BAC plasmid DNA are fragile and transfer to a new tube. 10. Centrifuge at 17,000  g for 3 min at 4
C to remove any residual debris. 11. Add 700 μl isopropanol to a 2 ml centrifuge tube and transfer the cleared lysate to the tube. Invert the tube 2–3 times to mix. 12. Centrifuge at 15,000  g for 15 min at 4
C. 13. Discard the supernatant and add 500 μl of 70% ethanol to wash the pellet. 14. Centrifuge at 17,000  g for 4 min at 4
C. 15. Remove the supernatant and dissolve the pellet in 500 μl TEN buffer. Incubate at RT for 1 h. 16. Centrifuge at 15,000  g for 30 s to remove any debris. 17. Transfer the supernatant into a fresh tube and add 1 ml absolute ethanol. Invert 2–3 times to mix them together. 18. Incubate at 20
C for 1 h. 19. Centrifuge at 15,000  g for 10 min at 4
C. 20. Discard the supernatant and add 1 ml 70% ethanol to wash the DNA pellet. 21. Centrifuge at 17,000  g for 1 min at 4
C and remove ethanol. 22. Air-dry any excess ethanol and add 100 μl nuclease-free water to resuspend the BAC DNA.

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1. Amplify the genomic region of gB and gH from isolated BAC using PCR with the following PCR mix: 200 ng BAC DNA, 45 μl Platinum™ PCR SuperMix High Fidelity (ThermoFisher Scientific), 2 μl primers (from 10 μM stock) for sgB or sgH regions, x μl water for a final volume of 50 μl. Put PCR tubes in the thermocycler and use the following program to amplify the region of interest: 94
C: 2 min, 35 cycles (94
C: 30 s, 55
C: 30 s, and 72
C: 3 min), 75
C: 15 min, 4
C: / 2. Run the samples on a 0.8% agarose gel. 3. Cut gB and gH bands and extract DNA with gel extraction kit (Omega). 4. Digest 5 μg of PCR products with AgeI and BamHI restriction enzymes (New England Biolabs). 5. Repeat steps 4 and 5. 6. Digest 5 μg vector pQCXIN with AgeI and BamHI restriction enzymes (New England Biolabs). 7. Repeat steps 4 and 5. 8. Set up the ligation reaction as follows to allow insertion of gB/gH genomic region in the vector: 150 ng vector, 100 ng sgB or sgH, 2 μl 10 buffer, 1 μl T4 DNA ligase, X μl water to make final volume of 20 μl. 9. Incubate the reactions for 1 h at RT. 10. Transform the plasmids into competent high efficiency DH5α E. coli cells (New England Biolabs). 11. Grow the bacteria with appropriate antibiotic selection and isolate plasmids with plasmid extraction kit (Qiagen). 12. Confirm the insertion sequences by DNA sequencing. 13. Prepare glycerol stocks for the clones containing sgB and sgH sequences.

3.2.3 Generating Retroviral Vectors Containing sgB and sgH

1. Isolate plasmid DNA from the bacterial clones containing the sgB and sgH sequences. 2. Transfect in 15 μg of retroviral plasmid with 15 μg of envelop plasmid (VSV-G) into GP2-293 cell line according to the manufacturer’s recommendations (Clontech). 3. Harvest supernatant containing retrovirus after 48 h and filter through a 0.45 μm filter to remove cellular debris. 4. Prepare Expi293F cells for transfection according to the manufacturer’s recommendations (see Note 7). 5. Add the retroviruses to the cells at MOI 10.

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6. After 24 h, measure cell viability, which should be greater than 90%. 7. Add 400 μg/ml of G418 (or appropriate selection pressure if different retroviral vector is used) to the media. 8. Check viability of the cells every 24 h. 9. Change the media every 3 days with specific antibiotic. 10. Once the viability reaches ~20%, decrease the concentration of G418 to a maintenance level of 200 μg/ml. 11. Once the viability recovers to 90–100%, cells are ready to be harvested and stored in the vapor phase of liquid nitrogen. Cells can be stored at a concentration of 1  107/ml with 10% DMSO in Expi293 expression media. 12. Isolate DNA from the cells and confirm sgB and sgH sequences from specific clones. 3.2.4 Affinity Chromatography Purification of sgB and sgH

1. Preincubate 30 ml of Expi293 expression media (ThermoFisher) containing 200 μg/ml G418 in two 125 ml baffled polycarbonate Erlenmeyer flasks with vent cap (Corning) at 8% CO2 on an orbital shaker (125 rpm) for 15 min at 37
C. 2. Add 1  107 Expi293F cells stably expressing sgB or sgH into the flasks and grow for 4 days at 8% CO2 on orbital shaker (125 rpm). 3. After 4 days, take two more 125 ml baffled flasks and repeat step 1. 4. Take 3 ml cells from sgB or sgH expressing Expi293F cells that has been growing for 4 days and add into new flasks. 5. Grow for an additional 4 days at 8% CO2 on orbital shaker (125 rpm). 6. Take the media containing sgB or sgH expressing Expi293F cells and separate into two separate 50 ml conical tubes and spin cells down at 200  g for 10 min. 7. Weigh the cell pellets and resuspend in ice-cold lysis buffer (10 ml/g cell pellet) and mix well with pipette. 8. Keep the cells on ice and sonicate three times with short bursts. 9. Centrifuge at 15,000  g for 30 min and filter the cleared lysates through a 0.45 μm polyethersulfone or cellulose acetate membrane. 10. Prepare sample by mixing the protein extract with an equal volume of lysis buffer. 11. Take two chromatography columns and add appropriate amount of Ni-charged resin (Bio-Rad). 12. Wash the columns with 25 ml cell lysis buffer to equilibrate.

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13. Load the sample onto the column and use gravity flow to allow binding of the soluble glycoproteins with the column. 14. Wash the column with 25 ml washing buffer to remove nonspecific bound protein. 15. Elute the bound His-tagged soluble glycoproteins with 10 ml elution buffer. 16. Perform dialysis in PBS to remove imidazole from the elution buffer. 17. Filter through a 0.2 μm syringe filter. 18. Measure concentration of the glycoproteins using Bradford assay (Bio-Rad). 19. Aliquot and flash freeze the glycoproteins with liquid nitrogen. Store at 80
C. 3.3 Analysis of Cellular Receptor Activation During HCMV Entry by Western Blot

1. Serum starve 2  106 cells for 1–24 h (see Note 8). 2. Cool cells to 4
C for 1.5 h. 3. Add virus at a MOI of 5–10 or sgB/sgH (0.5–5 μg) to treatment groups. 4. Incubate at 4
C for 1 h to allow binding of the virus or soluble glycoproteins to the cell surface (see Note 9). 5. Shift the temperature to 37
C using water bath for 5–30 min to allow for receptor activation (see Note 10). 6. Quickly add double volume of ice-cold PBS. 7. If using nonadherent cells, centrifuge at 1000  g for 2 min at 4
C. 8. Remove the supernatant without disturbing the cell layer or cell pellet and add lysis buffer with protease and phosphatase inhibitors. 9. Add loading buffer to samples and boil for 10 min (see Note 11 for modifications for highly hydrophobic membrane receptors). 10. Load samples into electrophoresis.

polyacrylamide

gel

and

perform

11. Transfer onto PVDF membrane. 12. Block membrane blot in 5%-TBST(see Note 12). 13. Add primary antibody for 1 h at RT. 14. Wash extensively with TBS containing 0.1% Tween-20. 15. Add HRP-conjugated secondary antibody for 1 h at RT. 16. Wash with TBS containing 0.1% Tween-20 and detect band with chemiluminescence detection reagent (see Note 13).

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3.4 Coimmunoprecipitation HCMV Glycoproteins and Cellular Receptors

1. Take 5–10 million cells and follow steps 1–4 as described in Subheading 3.3 with increased MOI of 10–15 (see Note 14). 2. If using nonadherent cells, centrifuge at 1000  g for 5 min at 4
C. 3. Remove the supernatant and gently wash monolayer or pellet twice with PBS. 4. Lyse the cells with gentle pipetting using 500 μl NP40 Cell Lysis Buffer with protease and phosphatase inhibitors (at 1:100 dilution) and incubate for 30 min on ice. 5. Transfer the lysate in Eppendorf tube and spin at maximum speed for 5 min at 4
C. 6. Take 50 μl of the clear lysate as input control. Transfer rest of the clear lysate to fresh Eppendorf tube. 7. Add specific antibody at appropriate dilution according to the manufacturer’s guideline and mix the lysate-antibody mixture overnight at 4
C using a rotatory shaker. 8. Next day, add 40 μl Dynabeads proteins A/G (depending on the species of the primary antibody) into an Eppendorf tube and remove the storage buffer from the beads after placing the tube on magnet. 9. Wash the beads with 400 μl NP40 Cell Lysis Buffer. 10. After removing the lysis buffer, add the protein-antibody mixture to the bead and gently mix them by inverting the tube for 3–4 times. 11. Incubate at 4
C for 2 h on a rotatory shaker. 12. Place the tube on magnet and remove the supernatant. 13. Gently wash the beads twice with NP40 Cell Lysis Buffer. 14. After removing the lysis buffer, add 50 μl IgG elution buffer and incubate at RT for 5 min. 15. Place the tube on magnet and collect the supernatant. 16. Add 5 μl 1 M pH 9 Tris to neutralize the pH. 17. Follow steps 9–16 in Subheading 3.3 for western blot analysis of coimmunoprecipitation.

4

Notes 1. The protocols described in this chapter use HEL fibroblasts (HELs); however, other fibroblast cell types can be used to grow HCMV, such as human foreskin fibroblasts (HFF) or MRC5 cells. 2. Certain strains of HCMV are cell-associated, and thus not present at high levels in the culture media. Under those

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circumstances, HCMV stocks can be generated by mechanical lysis of the infected fibroblasts. 3. To prepare soluble glycoproteins, the transmembrane domain was removed and replaced with a His-tag at the end of both gB and gH sequences during amplification of the genomic regions. 4. A mild lysis buffer with a nonionic detergent must be used as glycoprotein and cellular receptors interactions are often weak and relatively low in abundance. 5. Multiple collections can be done by adding back media to infected flasks and collecting supernatant in 3–4 days. 6. A lower temperature (32
C) is used to grow the Escherichia coli SW102 strain to prevent production of recombination proteins that is inhibited by a temperature-sensitive repressor (cI857). 7. Other mammalian expression systems can be used; however, Expi293 cells have been adapted to be grown in suspension to high density allowing for high pg/cell/day productivity. 8. Serum contains growth factors that can activate cellular receptors, and thus should be kept to a minimum while studying receptor activation. 9. The relative low abundance of glycoprotein and receptor interactions during HCMV entry requires synchronizing virus binding to the cell surface to maximize signal output. Infecting at 4
C will allow HCMV to bind but not initiate signaling until temperature shifted to 37
C. 10. The use of a 37
C water bath is preferred due to the short incubation periods needed to assess signaling events. 11. Highly hydrophobic membrane proteins, such as EGFR, form aggregates at high temperatures which are unable to entry polyacrylamide gels. To avoid aggregate formation, samples must be prepared in loading buffer with low heat (60
C). Certain antibodies cannot detect reduced proteins and, in that case, nonreducing conditions need to be used (i.e., absence of 2-mercaptoethanol) (Fig. 1). 12. Do not use skim milk as a blocking agent. Milk contains phosphatases that may lead to protein dephosphorylation interfering with target identification. Additionally, milk contains casein, a phosphoprotein that may bind anti-phospho antibodies causing nonspecific binding and high background. 13. Because of the relatively low frequency of activated receptors, a chemiluminescent substrate with picogram to femtogram-level protein detection sensitivity may be needed. 14. A higher MOI is needed to achieve enough viral glycoproteins and cellular receptor interactions for coimmunoprecipitation.

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Acknowledgments This work was supported by grants from the Carol M. Baldwin Breast Cancer Research Fund to G.C. Chan, National Institute of Allergy and Infectious Disease (R01AI141460) to G.C. Chan, and National Heart, Lung, and Blood Institute (R01HL139824) to G.C. Chan. References 1. Campadelli-Fiume G, Amasio M, Avitabile E, Cerretani A, Forghieri C, Gianni T, Menotti L (2007) The multipartite system that mediates entry of herpes simplex virus into the cell. Rev Med Virol 17(5):313–326. https://doi.org/ 10.1002/rmv.546 2. Connolly SA, Jackson JO, Jardetzky TS, Longnecker R (2011) Fusing structure and function: a structural view of the herpesvirus entry machinery. Nat Rev Microbiol 9(5):369–381. https://doi.org/10.1038/nrmicro2548 3. Kari B, Gehrz R (1992) A human cytomegalovirus glycoprotein complex designated gC-II is a major heparin-binding component of the envelope. J Virol 66(3):1761–1764 4. Compton T, Nowlin DM, Cooper NR (1993) Initiation of human cytomegalovirus infection requires initial interaction with cell surface heparan sulfate. Virology 193(2):834–841. https://doi.org/10.1006/viro.1993.1192 5. Isaacson MK, Compton T (2009) Human cytomegalovirus glycoprotein B is required for virus entry and cell-to-cell spread but not for virion attachment, assembly, or egress. J Virol 83(8):3891–3903. https://doi.org/10.1128/ JVI.01251-08 6. Nogalski MT, Chan G, Stevenson EV, Gray S, Yurochko AD (2011) Human cytomegalovirus-regulated paxillin in monocytes links cellular pathogenic motility to the process of viral entry. J Virol 85 (3):1360–1369. https://doi.org/10.1128/ JVI.02090-10 7. Smith MS, Bentz GL, Smith PM, Bivins ER, Yurochko AD (2004) HCMV activates PI(3)K in monocytes and promotes monocyte motility and transendothelial migration in a PI(3)Kdependent manner. J Leukoc Biol 76 (1):65–76. https://doi.org/10.1189/jlb. 1203621 8. Wang X, Huong SM, Chiu ML, Raab-Traub N, Huang ES (2003) Epidermal growth factor receptor is a cellular receptor for human cytomegalovirus. Nature 424(6947):456–461. https://doi.org/10.1038/nature01818

9. Wang X, Huang DY, Huong SM, Huang ES (2005) Integrin alphavbeta3 is a coreceptor for human cytomegalovirus. Nat Med 11 (5):515–521. https://doi.org/10.1038/ nm1236 10. Chan G, Nogalski MT, Yurochko AD (2009) Activation of EGFR on monocytes is required for human cytomegalovirus entry and mediates cellular motility. Proc Natl Acad Sci U S A 106 (52):22369–22374. https://doi.org/10. 1073/pnas.0908787106 11. Yurochko AD, Hwang ES, Rasmussen L, Keay S, Pereira L, Huang ES (1997) The human cytomegalovirus UL55 (gB) and UL75 (gH) glycoprotein ligands initiate the rapid activation of Sp1 and NF-kappaB during infection. J Virol 71(7):5051–5059 12. Huber MT, Compton T (1999) Intracellular formation and processing of the heterotrimeric gH-gL-gO (gCIII) glycoprotein envelope complex of human cytomegalovirus. J Virol 73(5):3886–3892 13. Huber MT, Compton T (1998) The human cytomegalovirus UL74 gene encodes the third component of the glycoprotein H-glycoprotein L-containing envelope complex. J Virol 72(10):8191–8197 14. Straschewski S, Patrone M, Walther P, Gallina A, Mertens T, Frascaroli G (2011) Protein pUL128 of human cytomegalovirus is necessary for monocyte infection and blocking of migration. J Virol 85(10):5150–5158. https://doi.org/10.1128/JVI.02100-10 15. Liu J, Jardetzky TS, Chin AL, Johnson DC, Vanarsdall AL (2018) The human cytomegalovirus trimer and Pentamer promote sequential steps in entry into epithelial and endothelial cells at cell surfaces and endosomes. J Virol 92 (21). https://doi.org/10.1128/JVI. 01336-18 16. Adler B, Scrivano L, Ruzcics Z, Rupp B, Sinzger C, Koszinowski U (2006) Role of human cytomegalovirus UL131A in cell typespecific virus entry and release. J Gen Virol 87

HCMV Glycoprotein and Cellular Receptor Interaction (Pt 9):2451–2460. https://doi.org/10.1099/ vir.0.81921-0 17. Wang D, Shenk T (2005) Human cytomegalovirus UL131 open reading frame is required for epithelial cell tropism. J Virol 79 (16):10330–10338. https://doi.org/10. 1128/JVI.79.16.10330-10338.2005 18. Sharma S, Wisner TW, Johnson DC, Heldwein EE (2013) HCMV gB shares structural and functional properties with gB proteins from other herpesviruses. Virology 435 (2):239–249. https://doi.org/10.1016/j. virol.2012.09.024 19. Soroceanu L, Akhavan A, Cobbs CS (2008) Platelet-derived growth factor-alpha receptor activation is required for human cytomegalovirus infection. Nature 455(7211):391–395. https://doi.org/10.1038/nature07209 20. Vanarsdall AL, Wisner TW, Lei H, Kazlauskas A, Johnson DC (2012) PDGF receptor-alpha does not promote HCMV entry into epithelial and endothelial cells but increased quantities stimulate entry by an abnormal pathway. PLoS Pathog 8(9): e1002905. https://doi.org/10.1371/journal. ppat.1002905 21. Isaacson MK, Feire AL, Compton T (2007) Epidermal growth factor receptor is not required for human cytomegalovirus entry or signaling. J Virol 81(12):6241–6247. https:// doi.org/10.1128/JVI.00169-07 22. Altman AM, Mahmud J, Nikolovska-ColeskaZ, Chan G (2019) HCMV modulation of cellular PI3K/AKT/mTOR signaling: new opportunities for therapeutic intervention? Antivir Res 163:82–90. https://doi.org/10. 1016/j.antiviral.2019.01.009 23. Wu Y, Prager A, Boos S, Resch M, Brizic I, Mach M, Wildner S, Scrivano L, Adler B (2017) Human cytomegalovirus glycoprotein complex gH/gL/gO uses PDGFR-alpha as a key for entry. PLoS Pathog 13(4):e1006281. https://doi.org/10.1371/journal.ppat. 1006281 24. Kabanova A, Marcandalli J, Zhou T, Bianchi S, Baxa U, Tsybovsky Y, Lilleri D, Silacci-FregniC, Foglierini M, Fernandez-Rodriguez BM, Druz A, Zhang B, Geiger R, Pagani M, Sallusto F, Kwong PD, Corti D, Lanzavecchia A, Perez L (2016) Plateletderived growth factor-alpha receptor is the cellular receptor for human cytomegalovirus gHgLgO trimer. Nat Microbiol 1(8):16082.

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https://doi.org/10.1038/nmicrobiol. 2016.82 25. Kaur J, Bachhawat AK (2009) A modified Western blot protocol for enhanced sensitivity in the detection of a membrane protein. Anal Biochem 384(2):348–349. https://doi.org/ 10.1016/j.ab.2008.10.005 26. Nemerow GR (2000) Cell receptors involved in adenovirus entry. Virology 274(1):1–4. https://doi.org/10.1006/viro.2000.0468 27. Mercer J, Schelhaas M, Helenius A (2010) Virus entry by endocytosis. Annu Rev Biochem 79:803–833. https://doi.org/10.1146/ annurev-biochem-060208-104626 28. Greber UF (2002) Signalling in viral entry. Cell Mol Life Sci 59(4):608–626 29. Feire AL, Koss H, Compton T (2004) Cellular integrins function as entry receptors for human cytomegalovirus via a highly conserved disintegrin-like domain. Proc Natl Acad Sci U S A 101(43):15470–15475. https://doi.org/ 10.1073/pnas.0406821101 30. Viswanathan K, Smith MS, Malouli D, Mansouri M, Nelson JA, Fruh K (2011) BST2/Tetherin enhances entry of human cytomegalovirus. PLoS Pathog 7(11):e1002332. https://doi.org/10.1371/journal.ppat. 1002332 31. Li Q, Wilkie AR, Weller M, Liu X, Cohen JI (2015) THY-1 cell surface antigen (CD90) has an important role in the initial stage of human cytomegalovirus infection. PLoS Pathog 11 (7):e1004999. https://doi.org/10.1371/jour nal.ppat.1004999 32. Li Q, Fischer E, Cohen JI (2016) Cell surface THY-1 contributes to human cytomegalovirus entry via a macropinocytosis-like process. J Virol 90(21):9766–9781. https://doi.org/ 10.1128/JVI.01092-16 33. Martinez-Martin N, Marcandalli J, Huang CS, Arthur CP, Perotti M, Foglierini M, Ho H, Dosey AM, Shriver S, Payandeh J, Leitner A, Lanzavecchia A, Perez L, Ciferri C (2018) An unbiased screen for human cytomegalovirus identifies Neuropilin-2 as a central viral receptor. Cell 174(5):1158–1171. e19. https://doi. org/10.1016/j.cell.2018.06.028 34. Vanarsdall AL, Pritchard SR, Wisner TW, Liu J, Jardetzky TS, Johnson DC (2018) CD147 promotes entry of Pentamer-expressing human cytomegalovirus into epithelial and endothelial cells. MBio 9(3). https://doi.org/ 10.1128/mBio.00781-18

Chapter 11 Antibody-Independent Quantification of Cytomegalovirus Virion Protein Incorporation Using HiBiT Iris K. A. Jones and Daniel N. Streblow Abstract Human cytomegalovirus (HCMV) is a large double-stranded DNA virus and member of the β-herpesvirus family. HCMV is ubiquitous in the human population and causes lifelong infections. HCMV infection is associated with high morbidity and mortality in immunocompromised individuals and the virus is a major cause of virus-mediated congenital disease. There have been a number of HCMV entry receptors identified that use one of two viral receptor binding complexes, including the gH/gL/gO complex and the pentamer made up of gH/gL/UL128/UL130/UL131a. Cytomegaloviruses (CMVs) are typically host-restricted requiring the use of species-specific modeling and culture conditions. We use rat CMV (RCMV) to study CMV-accelerated vascular disease and chronic allograft rejection. RCMV encodes homologous versions of the entry complex proteins but their incorporation and copy number per virion are still unknown. In this methods article, we describe a novel approach of HiBiT tagging viral proteins in order to detect and quantify protein incorporation into particles. This method is independent of protein-specific antibodies and can be standardized using a commercially available HiBiT protein standard. Using bacterial artificial chromosome (BAC) recombineering, we have constructed two individual viruses containing a HiBiT tag fused to the C0 -terminus of either the UL128 homolog (R129) or the UL130 homolog (R131). Viruses containing these mutations were rescued, purified and analyzed. Our data demonstrate that R129 and R131 are both incorporated into RCMV virions at equimolar ratios relative to genome copy number, supporting this antibody-free approach for quantifying viral protein incorporation and its application toward the identification of domains required for incorporation. Key words Viral entry, Cytomegalovirus, Pentameric entry complex, HiBiT, nanoLuc

1

Introduction Human cytomegalovirus (HCMV) requires a minimum of two membrane glycoprotein complexes for entry into cells including a gH/gL receptor binding complex and the viral fusion protein gB [1]. The gH/gL complex exists in at least three forms: (1) trimeric gH/gL/gO is required for entry into fibroblasts [2]; (2) pentameric gH/gL/UL128/UL130/UL131a is required for entry into epithelial cells, endothelial cells and macrophages [2]; and (3) gH/UL116 [3] has yet unknown functions but the

Andrew D. Yurochko (ed.), Human Cytomegaloviruses: Methods and Protocols, Methods in Molecular Biology, vol. 2244, https://doi.org/10.1007/978-1-0716-1111-1_11, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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homologous complex is required for RCMV entry into fibroblasts. The gH/gL binding region for the ULs is similar to gO but little is known about the structural binding requirements for UL116. Receptor binding induces conformational changes to the gH/gL complex that allow for interactions and triggering of the gB fusion machinery [1, 4]. While many recent advances have increased our knowledge about the gH/gL receptor binding complexes, there is still much to learn about their function, receptor specificity, and stoichiometry. In addition, we have limited information regarding binding of entry proteins from CMV species other than HCMV. Due to strict species specificity of CMVs, many investigators utilize animal models to study mechanisms of CMV dissemination and pathogenesis [5]. We have utilized rat CMV (RCMV) infection in rats to examine the role of CMV infection in cardiac transplant rejection and cardiovascular disease. RCMV encodes proteins involved in the formation of all three of the gH/gL complexes. An interesting feature of the HCMV pentamer complex is the finding that UL128 and UL130 have characteristic chemokine folds at their N0 -terminal region that extend away from the protein body [6, 7]. The functional relationship between this chemokine structure and their role in entry is still unclear. Both proteins also contain unstructured regions at their C0 -terminus with various charged clusters involved in protein–protein interactions important for pentamer complex formation [8, 9]. We were interested in dissecting the pentamer receptor binding versus chemokine activities in viral pathogenesis and cardiac transplant-related disease for the UL128 and UL130 homologs, R129 and R131, respectively. However, an important concern with our mutational strategy was to determine how mutations in the N0 -terminal chemokine or C0 -terminal pentamer binding regions affect pentamer formation. Because of this need, we sought to develop an assay to empirically determine the number of R129 and R131 molecules present in virus particles. However, developing antibodies to detect R129 and R131 is labor intensive and fraught with additional problems. The biggest issue being that the sensitivity of antibody-based assays is set by the binding affinity of the antibody toward the antigen. This feature of antibody binding often limits applications requiring the comparison of different proteins unless they share a common epitope or protein tag. In this chapter, we detail the techniques used for the molecular tagging and detection of the RCMV pentamer complex proteins R129 and R131 using the HiBiT detection system. NanoLuciferase (NanoLuc) is a 19 kDa protein that generates luminescence with a higher dynamic range than conventional firefly luciferase. While the relatively small size of NanoLuc can be useful for quantifying levels of tagged protein expression or as a reporter for expression studies; the size is still considered prohibitive when studying proteins that form tight complexes wherein steric interference may affect protein

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interactions. The adoption of a split form of NanoLuc called NanoBiT was developed that contains two fragments of NanoLuc [5]. The small portion that consists of 11 amino acids of NanoLuc is called HiBiT, and this is the portion that is fused to the protein of interest to be detected. The larger fragment of NanoLuc, called LgBiT, is present in the lytic and extracellular in-solution detection kits, as well as the HiBiT blotting system, from Promega. While these components are encoded and produced separately, they can associate to reconstitute the NanoBiT enzyme to release a bioluminescent signal following addition of substrate. Using this method, we demonstrate that this approach can be applied to full-length and truncated proteins to quantify the number of molecules incorporated into the virion. This approach could be used to quantify levels of virion incorporation of any viral or host protein, and the effect that mutations in those proteins have on their rate of incorporation into virions.

2

Materials

2.1 RCMV BAC Recombineering

1. RCMV Maastricht strain BAC in SW102 E. coli. 2. pc255-GalK/Kan plasmid. 3. Prepare 50 mg/mL Kanamycin (Kan) stock solution in dH2O. Filter sterilize through a 0.22 μm filter. Store at 4  C. 4. Prepare 12.5 mg/mL chloramphenicol (Chlor) stock solution in ethanol, store at 20  C. 5. MacConkey agar plates: add 40 g of Difco MacConkey agar base without lactose plus 4 g D-(+)-Galactose to a 1 L bottle and QS to 1 L with dH2O. Mix and autoclave to sterilize. Allow the solution to cool until warm to the touch, but not cool enough to solidify. Add Chloramphenicol to a final concentration of 12.5 μg/mL and Kanamycin to a final concentration of 50 μg/mL. Pour 20 mL of agar solution per plate and allow to solidify at room temperature overnight. Store at 4  C. 6. Cloning and sequencing primers are listed in Table 1. 7. 2-Deoxy-galactose-1-phosphate (DOG) selection plates: M63 agar plates with 12.5 μg/mL Chloramphenicol. 8. M9 salts: 6 g Na2HPO4, 3 g KH2PO4, 1 g NH4Cl, 0.5 g NaCl, QS to 1 L dH2O. Autoclave to sterilize. 9. PCR master mix for PCR screening of BAC clones: 2 PCR Platinum master mix. 10. PCR master mix for PCR amplification of genes for sequencing and cloning: Platinum Supermix HiFi master mix. 11. DpnI (20,000 U/mL).

HiBiT tag and RCMV sequences for recombination

Reverse screening primer

Forward screening primer

Reverse GalK/Kan homology primer

Forward GalK/ Kan homology primer

Insertion/Detection

Table 1 DNA Primers

50 -ACCTTCTCTGATAAGTTTTCTGAA GGAAAGGAAACATATACACAAACATA TAGAACATAAGCATGTACACGTGTTAGA TATCTAATAAAAACTATACCTACTCAGC AAAAGTTCGATTTA 50 -AACTTTGATGGACAAGATCCAACTATCCTG CAGAGAGTCCCTGCTCTATGTGGA TGTTCAAGGGGAAATTCAGTGTGTG 50 -CCTTCTCTGATAAGTTTTCTGAAGGAA AGGAAACATATACACAAACATATAGAA CATAAGCATGTACACGTGTTAGATA 50 -AACTTTGATGGACAAGATCCAA CTATCCTGCAGAGAGTCCCTG CTCTATGTGGATGTTCAAGGGGAAATTC AGTGTGTGGAAGATAGGTGTTCAGAAG GGCATCACCATCACCATCACGTGAGCGG CTGGCGGCTGTTCAAGAAGATTAGCTAG GTATAGTTTTTATTAGATATCTAACACGT GTACATGCTTATGTTCTATATGTTTGTGT ATATGTTTCCTTTCCTTCAGAAAACTTAT CAGAGAAGGT

50 -CGACTTATTTACCAGATGTACTC ATAACCATCTGTATGCTCAAGATGTGTTG TCTCGGTTGCCTATTCCGAACGGCTAT

50 -CAAAACCTGGCGACGGATGTGAACG AATGGCATAGGAGAAGTAAGAACGGAA ACGTAGTCTTAGGCGTCGGGAACGTCA

50 -CGACTTATTTACCAGATGT ACTCATAACCATCTGTATGCT CAAGATGTGTTGTCTCGGTTGCCTAT TCCGAACGGCTATTCTCTGTTGTCTGGGC CGGCATCACCATCACCATCACGTGAGC GGCTGGCGGCTGTTCAA GAAGATTAGCTGACAACGAT GACGTACGGTGTGACGTTCC CGACGCCTAAGACTACGTTTCC GTTCTTACTTCTCCTATGCCA TTCGTTCACATCCGTCGCCAGG TTTTG

50 -AACTTTGATGGACAAGATCCAACTAT CCTGCAGAGAGTCCCTGCTCTATGTGGA TGTTCAAGGGGAAATTCAGTGTGTGGAA GATAGGTGTTCAGAAGGGCCTGTTGA CAATTAATCATCGGCATAG

RCMV R129(short) HiBiT

50 -TAAGAACGGAAACGTAGTCTTAGGCG TCGGGAACGTCACACCGTACGTCATCG TTGTCACTCAGCAAAAGTTCGATTTA

5 -GTTGTCTCGGTTGCCTATTCCGAAC GGCTATTCTCTGTTGTCTGGGCAGCGGCCTG TTGACAATTAATCATCGGCATAG

0

RCMV R131 HiBiT

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12. Molecular Biology Grade DNase/RNase-free water. 13. PCR machine. 14. Ultrapure agarose. 15. 50 TAE buffer: 242 g Tris base, 57.1 mL glacial acetic acid, 100 mL 0.5 M EDTA (pH 8.0), QS to 1 L dH2O. pH to 8.4. Dilute to 1 with dH2O prior to use. Store at room temperature. 16. SYBR Safe DNA gel stain. 17. DNA gels: 1% Ultrapure Agarose in TAE buffer with 2.5 μL SYBR Safe DNA gel stain per 40 mL gel. 18. Nucleic acids gel electrophoresis unit and power supply. 19. Gel extraction kit. 20. Spectrophotometer machine. 21. 0.1 cm electroporation cuvettes. 22. Electroporation system. 23. Centrifuge and microfuge. 24. 2 YT broth: 31 g broth powder dissolved in 1 L dH2O. Autoclave to sterilize. 25. 15 mL conical tubes. 26. 50 mL conical tubes. 27. 14 mL polypropylene round-bottom culture tubes. 28. Erlenmeyer flasks (125 mL and 500 mL). 29. Plasmid DNA Midiprep kit. 2.2

Virus Rescue

1. BAC-derived DNA for RCMV-R129(short) and RCMV-R131 HiBiT (described in Subheading 3.1). 2. Lipofectamine 2000 (ThermoFisher Scientific). 3. RFL-6 fibroblasts (ATCC) (see Note 1). 4. Opti-MEM (ThermoFisher Scientific). 5. DMEM culture medium supplemented with 5% fetal bovine serum (FBS), 100 U/mL penicillin, 100 mg/mL streptomycin, and 20 mM L-glutamine. 6. 6-well tissue culture dishes. 7. 1.5 mL microcentrifuge tubes. 8. 2 mL polypropylene micro tube with screw cap. 9. DNA/RNA purification kit: GeneJET Viral DNA/RNA purification kit (ThermoFisher Scientific).

2.3 RCMV Growth and Purification

1. Vented T175 tissue culture flasks. 2. Phosphate buffered saline (PBS) pH 7.2 without calcium or magnesium.

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3. 70 μm cell strainers (Fisher Scientific). 4. Ultra-clear ultracentrifuge tubes, 1  3.500 (Beckman Coulter). 5. Ultra-clear ultracentrifuge tubes, 9/16  3.500 (Beckman Coulter). 6. Ultracentrifuge machine and appropriate rotors (e.g., Beckman L7-65). 7. 500 mL Stericup 0.22 μm filter units (Millipore Sigma). 8. 10% D-sorbitol in PBS without Calcium or Magnesium. Filter sterilize through a 0.22 μm filter. 9. Histodenz (Sigma). 10. TNE buffer: 50 mM Tris [pH 7.4], 100 mM NaCl, and 10 mM EDTA. Filter-sterilize through a 0.22 μm filter. 11. 18-gauge 1½00 needles. 12. Microcentrifuge machine. 13. 0.6 mL micro tubes with snap cap. 2.4

Western Blotting

1. Cell culture grade water. 2. 10 Cell lysis buffer (Cell signaling technologies). 3. HALT protease inhibitor cocktail (100) (ThermoFisher Scientific). 4. Sequencing grade modified trypsin. 5. 10-well Novex 10–20% Tricine protein gels with 1.0 mm wells (ThermoFisher Scientific). 6. Novex Tricine SDS running buffer (2) (ThermoFisher Scientific). 7. Mini protein gel electrophoresis tank and power supply. 8. β-mercaptoethanol. 9. Semidry transfer buffer: for 2 L mix 11.6 g tris base, 5.86 g glycine, and 400 mL methanol. QS to 2 L with dH2O. 10. Semidry transfer cell and power supply. 11. 0.45 μm pore size Immobilon PVDF membranes (Millipore). 12. Nano-Glo HiBiT blotting system (Promega). 13. West Pico PLUS chemiluminescent solution (ThermoFisher Scientific). 14. Rat anti-gB monoclonal antibody (OHSU-VGTI Monoclonal Antibody Facility). 15. Rabbit anti-Rat IgG (H+L) HRP (Southern Biotech).

2.5 HiBiT In-Solution Detection

1. Nano-Glo HiBiT lytic detection system (Promega). 2. HiBiT control protein (Promega).

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3. 96-well white walled detection plates. 4. Multimode microplate reader machine. 2.6 RCMV Viral Genome Quantification

1. DNAzol reagent (Invitrogen). 2. 100% and 70% molecular grade ethanol. 3. qPCR master mix: Taqman fast advanced master mix (Applied biosystems). 4. 384-well qPCR plates. 5. Optical adhesive films. 6. RCMV qPCR primers and probe recognizing RCMV-R54. Forward primer: 50 - CCTCACGGGCTACAACATCA (RCMV nucleotides 64,071-64,090); Reverse: 50 -GAGAGTT GACGAAGAACCGACC (Reverse complement of RCMV nucleotides 63,963-63,984); Probe: 50 -VIC- CGGCTTCGA TATCAAGTATCTCCTGCACC -TAMRA (RCMV nucleotides 64,041-64,069). 7. Quantification standard: RCMV viral genomic DNA at a known concentration. 8. Real-time PCR machine.

3

Methods

3.1 Generation of RCMV Containing R131 or R129(short) HiBiT Fusion Tags

The RCMV Maastricht strain genome was captured as a bacterial artificial chromosome (BAC) using homologous recombination by replacing ORFs r144-r146 with a BAC cassette [10]. The BAC is resistant to chloramphenicol and contains an enhanced green fluorescent protein (eGFP) cassette under the control of the HCMV major immediate early promotor. A two-step recombination protocol can be used to molecularly tag RCMV open reading frames without leaving a sequence scar. Individual RCMV R129 and R131 recombinant viruses were constructed by addition of a C0 terminal in-frame fusion tag consisting of 6 Histidine residues and the 11 amino acid HiBiT tag followed by a stop codon (see Note 2). The first recombination step requires the incorporation of a gene cassette expressing galK and kanamycin resistance genes into the RCMV BAC genome at the genomic site for introduction of the tag. Polymerase chain reaction (PCR) is used to generate an amplicon containing 50–100 bp of homology (see Note 3) to the RCMV insertion site flanking the galK/Kan gene cassette. The amplified DNA is treated with Dpn1, purified and then electroporated into competent RCMV-BAC SW102 cells. Clones are positively selected for gain of resistance against Kanamycin. Resistant clones are screened by PCR and sequenced to identify clones with the proper insertion. For the second step in the recombination process, PCR is

TR L

RCMV (224kB) R128

R129

TRR

R131

(b)

R133

R131 HiBiT Putative CC-chemokine domain

HiBiT

55kDa 6xHis

Acidic Cluster

Signal Sequence

Putative entry domain

WT

(a)

R129(short) HiBiT

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R131 HiBiT

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36kDa 28kDa 17kDa

R129(short) HiBiT HiBiTtag

6xHistag

Acidic Cluster

Acidic Cluster

Signal Sequence

10kDa

Putative entry domain

Putative CC-chemokine domain

Fig. 1 HiBiT tag design for R131 and R129(short) viral proteins. (a) Schematic depiction of RCMV genome with R131 HiBiT and R129 HiBiT–tagged proteins and putative protein domains. (b) HiBiT blot detection of R131 HiBiT (28 kDa) and RCMV R129(short) HiBiT (17 kDa) proteins in RCMV-infected cell lysates at the time of maximum cytopathic effect. Nano-Glo HiBiT blotting system was used for luminescence-based detection of proteins. RFL-6 cells infected with a BAC-derived RCMV lacking the fusion HiBiT tag was used as a negative control

used to generate a DNA fragment containing RCMV sequences with the gene-specific in-frame HiBiT tag as shown in Fig. 1a. The amplicon is electroporated into competent bacteria and negatively selected for galK replacement on 2-deoxy-galactose-1-phosphate (DOG) negative selection plates. At this stage BAC clones are grown and checked by PCR and sequencing for correct incorporation of the HiBiT tag. DNA from correct clones is prepared for transfection rescue of infectious virus in mammalian cells. 3.1.1 Generate PCR Fragments for Homologous Recombination

1. Prepare PCR reaction for amplification of galK/Kan cassette to create the construct for the first recombination step using homology primers and High-Fidelity PCR mix in a total volume of 50 μL per reaction. Multiple reactions may need to be performed to ensure that enough amplicon is produced. 2. Perform PCR amplification using the following conditions: 3 min 95  C then 30 s at 95  C–30 s at 58  C–3 min at 72  C for 28 cycles. These PCR conditions should be optimized for each individual amplicon and primer set. 3. Treat the PCR product with 1 μL Dpn1 at 37  C for at least 1 h. 4. Run the PCR product on a 1% agarose TAE gel and cut out band corresponding to PCR product and purify using a commercially available gel extraction kit and elute the final DNA product in 50 μL molecular biology–grade water. 5. Determine concentration spectrophotometer.

of

DNA

product

using

a

HiBiT Virion Protein Quantification 3.1.2 First Recombination Step

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1. Inoculate 5 mL of 2X YT broth plus 12.5 μg/mL Chlor with SW102 cells containing the RCMV BAC and grow overnight at 30  C. 2. Add 500 μL of the overnight culture to 25 mL 2X YT plus 12.5 μg/mL Chlor and incubate at 30  C for approximately 3 h (OD600 is between 0.55 and 0.6). 3. Heat-shock the 25 mL culture at 42  C for exactly 17 min. 4. Cool the culture on ice and transfer into two prechilled 50 mL conical tubes. Culture must be kept ice cold for steps until the electroporation is complete. 5. Pellet the culture at 650  g for 10 min at 0  C. 6. Pour off the supernatant and resuspend the pellet in 10 mL of ice-cold water by shaking gently. 7. Repeat wash steps once. 8. Resuspend the bacterial pellet in 1 mL of ice-cold water by shaking gently. 9. Transfer the resuspended bacterial pellet to a 1.5 mL microcentrifuge tube. 10. Pellet the culture at 5000  g for 1 min. 11. Pipet off the supernatant and wash once more in 1 mL ice-cold water. 12. Pipet off the supernatant and resuspend the bacterial pellet in 240 μL of molecular biology-grade water. 13. Prechill a 0.1 cm cuvette on ice. 14. Into the cuvette add 300 ng of the PCR product and 60 μL of electrocompetent cells, mix by gentle flicking. 15. Pulse the galK/Kan PCR product–competent cell mixture using the following settings: Voltage ¼ 1.8 kV, Capacitance ¼ 25 μF, Resistance ¼ 200 Ohms. 16. Immediately following electroporation, add 1 mL 2X YT broth to the cuvette and then transfer to a fresh 50 mL conical containing an additional 9 mL of 2X YT broth and incubate at 30  C for 2 h. 17. Pellet bacteria and resuspend in 100 μL of 2X YT broth. 18. Plate recovered bacteria on MacConkey/Kan/Chlor plates and incubate at 30  C for 2 days. Positive colonies should turn red and negative colonies will be white. 19. Pick individual colonies and PCR screen to identify positive colonies (positive PCR product should be around 2.5 kb) (see Note 4).

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3.1.3 Second Recombination Step

1. Prepare electrocompetent positive clones by repeating steps outlined in Subheading 3.1.2 (steps 1–8) with the addition of 100 μg/mL kanamycin to culture conditions. 2. Prepare PCR fragments containing the HiBiT tag sequence designed to replace the galK/Kan cassette (see Note 5). 3. Immediately following electroporation, add 1 mL 2X YT broth to the cuvette and then transfer to a fresh 50 mL conical containing an additional 9 mL of 2X YT broth and incubate at 30  C for 5 h. 4. Pellet 1 mL of recovered bacteria by centrifugation at 10,000  g for 1 min. 5. Resuspend bacterial pellet in 1.5 mL of M9 salts. Repeat this wash step once. 6. Plate the washed bacteria in M9 salts on DOG negative selection plates and incubate at 30  C for 2 days. 7. PCR screen to identify positive colonies, sequence the fragments to confirm proper genetic recombination (see Note 6). 8. To prepare BAC DNA for virus rescue transfections, inoculate 100 mL 2X YT broth plus 12.5 μg/mL Chloramphenicol with positive clones. 9. Prepare BAC DNA containing the desired tags by DNA midiprep (see Note 7).

3.2 Transfection of BAC DNA and Rescue of RCMV-R131 and -R129(short) HiBiT

The recovery of CMVs from BAC DNA requires a cell line that is competent for both the successful transfection of highly pure DNA and the ability of the virus to replicate. For the recovery of the Maastricht strain of RCMV, a rat lung fibroblast cell line called RFL-6 fulfills both requirements. Often it is necessary to use culture conditions that promote slower cell growth to prevent the cells from outgrowing the virus as it begins to propagate. We typically grow RFL-6 cells in 5% FBS for this reason. DNA prepared using a midiprep kit ensures sufficient DNA quantity and quality. Successfully transfected cells become GFP+ within 24–48 h and viral spread to neighboring cells should occur within 7 days post transfection. Transfection of a range of DNA concentrations (2–10 μg) will allow for optimization of the conditions necessary for successful rescue. Unless it is possible to identify a virus transfection condition with only one RCMV rescue per well, limiting dilution isolation of culture supernatants harvested at the time of maximum cytopathic effect should be performed to obtain a clonal virus for expansion. Virus mutational analysis and insert validation is accomplished by PCR screening and sequencing of DNA isolated from rescued RCMV.

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1. Seed 6-well plates with RFL-6 cells at a density of 5  105 cells per well in 2 mL of complete DMEM culture medium at 18 h before transfection. 2. Mix Lipofectamine 2000 (10 μL/well) with DNA (2–10 μg) in Opti-MEM (1 mL), in a 1.5 mL microcentrifuge tube. Incubate at room temperature for 15 min. 3. Replace DMEM culture medium with 1 mL of Opti-MEM containing transfection mixture. 4. At 4 h posttransfection, replace transfection medium with 2 mL of fresh DMEM medium. 5. Monitor virus recovery and, at maximum cytopathic effect, collect supernatants and cells into 2.0 mL tubes. 6. Freeze at 80  C for further use. 7. In order to validate RCMV-R129(short) and RCMV-R131 HiBiT insertions, purify viral DNA from 200 μL of supernatant and extract using a DNA/RNA purification kit. 8. PCR amplify the region of insertion as described above in Subheading 3.1.3 (step 7). 9. Sequence PCR products to confirm insertion (at this point the sequence of the entire virus can be performed to confirm virus integrity) (see Note 8). 3.3 RCMV Purification Protocol

There are a number of different protocols for CMV purification. For virion structural studies we routinely infect at least 40 flasks of cells and harvest only supernatant virus at the time of maximum cytopathic effect. Supernatants are clarified of cell debris by centrifugation followed by filtration through a 70 μm filter. The virus is pelleted through a Sorbitol cushion and the pellet is then resuspended and banded using density gradient ultracentrifugation. The banded virus is removed from the gradient and then pelleted to concentrate. This approach yields a highly pure virus preparation with very little cellular contamination. The virus purification workflow is depicted in Fig. 2.

Visualize proteins by western blot

[ Expand RCMV viral clones in 40 T175 flasks of RFL6 cells

Pellet virus from clarified supernatants over 10% Sorbitol

]

Isolate virus by banding over Histodenz gradient

Pellet isolated viral Aliquot and particles over 10% store at -80°C Sorbitol

Fig. 2 Virus Purification and Analysis Workflow Diagram

Quantify HiBiT-tagged proteins by HiBiT lytic detection assay

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1. For each RCMV-HiBiT virus and control, infect 40 confluent T175 RFL-6 with a multiplicity of infection (MOI) equal to 0.25. Incubate at 37  C until full cytopathic effect is achieved. 2. Collect culture medium into 50 mL conical tubes and centrifuge at 1989  g (Beckman GH-3.8 rotor) for 15 min. 3. Filter supernatants through a 70 μm filter unit. 4. Transfer clarified supernatants to SW32 centrifugation tubes (30 mL per tube). 5. Underlay each tube with 5 mL of 10% Sorbitol solution. 6. Add 2 mL of cell-free supernatants to the top of each tube for a total volume of 37 mL per tube. Tubes should be balanced appropriately prior to centrifugation. 7. Centrifuge at 76,755  g (Beckman L7-65 ultracentrifuge; SW32 rotor) for 70 min at 4  C. 8. Pour off the supernatants and resuspend the virus pellets in TNE buffer (4 mL total volume). 9. Overlay the resuspended pellet on top of thawed SW41Ti ultracentrifuge tubes containing 10–50% Histodenz gradient, 2 mL per tube (see Note 9). 10. Centrifuge at 111,132  g (Beckman ultracentrifuge; SW41Ti rotor) for 2 h at 4  C. 11. Virions should be visible as a white band. Collect virion band by fractionating the gradient contents by draining from the bottom of the centrifugation tube using an inserted needle. 12. QS the banded virus fractions in PBS up to 32 mL. 13. Pellet virus as described in Subheading 3.3 (steps 4–7). 14. Pour off the supernatants and resuspend the virus pellet with 300 μL PBS. 15. Store 30 μL aliquots at 80  C until further analysis. 3.4 Virion Protein Detection

Serial dilutions of the RCMV virion preparations were separated by SDS-PAGE and analyzed by immunoblotting for the presence of HiBiT-tagged proteins and viral glycoprotein B (gB). Figure 3 demonstrates that R131 HiBiT and R129 HiBiT are present in their respective virion preparations. Staining for gB was used to normalize virion preparations for further studies. 1. To purified virus preparations, add Tricine loading buffer +2% BME at a ratio of 1:1. 2. Boil samples at 100  C for 5 min and allow to cool to room temperature before loading. 3. Separate samples by electrophoresis on 10–20% Tricine gels at 125 V for 1 h.

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Fig. 3 Detection of Virion-associated Proteins. Histodenz-purified viral preparations of RCMV R131 HiBiT, RCMV R129(short) HiBiT, and RCMV WT were generated as described in Subheading 3.3. Varying amounts of viral preparations were separated by SDS-PAGE and analyzed by western or HiBiT blot as described in Subheading 3.4. The extent of gB staining was used to normalize samples when used in Subheading 3.5

4. Transfer the proteins to a PVDF membrane using a semidry transfer system at 25 V for 25 min. 5. In order to stain for both gB and HiBiT, cut across the membrane at the 36 kDa marker. 6. Detect the presence of gB on the upper blot using a monoclonal α-gB antibody. 7. Block with 5% BSA in TBST buffer for 1 h at room temperature. 8. Incubate the blot with 10 mL of rat α-gB antibody diluted 1:1000 in 5% BSA-TBST for 1 h at room temperature. 9. Wash the blot three times (20 mL) with TBST at room temperature (20, 15, and 5 min). 10. Incubate the blot with an α-rat secondary antibody at 1:5000 in TBST. 11. Wash three times in TBST as in Subheading 3.4 (step 9). Perform one final wash with 10 mL water. 12. Add 2 mL each of pico chemiluminescent substrate solutions and incubate at room temperature for 3 min, shaking gently. 13. Image blot using X-ray film (chemiluminescent gel image systems may also be used). 14. Detect presence of HiBiT tag on the lower blot using HiBiT blotting kit. 15. Wash the blot for 5 min in TBST. 16. Add 1 mL of 10 HiBiT blotting buffer and 50 μL of LgBiT protein to 9 mL of water and mix. 17. Incubate blot in 10 mL of HiBiT blotting buffer containing LgBiT for 1 h at room temperature.

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Fig. 4 R131 and R129 HiBiT proteins are trypsin sensitive. To demonstrate that the virion incorporated R131 and R129 proteins were present on the outside of the virus particle, wild-type RCMV, RCMV R131 HiBiT, and RCMV R129(short) HiBiT viral preparations, normalized to gB content, were treated with PBS or sequence grade Trypsin for 1 h at 37  C. Samples were then separated by SDS-PAGE and blotted for gB and HiBiT. Both gB and HiBiT-tagged proteins were sensitive to trypsin treatment indicating that they were accessible on the virion surface

18. Add 20 μL of HiBiT substrate directly to the 1 blotting buffer with LgBiT and rock at room temperature for 5 min. 19. Image blot for luminescent signal using X-ray film (Chemiluminescent gel image systems may also be used). This detection method was used to confirm the presence of the HiBiT tags in virus particles and to normalize levels of gB staining for wild type RCMV and viruses containing R131 HiBiT or R129(short) HiBiT tags for biochemical analyses. 3.5 Trypsin Sensitivity of Virion-Associated R131 and R129(short) HiBiT

Trypsin treatment of virions is often used to determine whether proteins are present on the outside membrane of the virus particle, accessible to trypsin. In this procedure, we trypsin treated virion preparations that were normalized to levels of gB protein in order to verify that R129 and R131 are on the outside of the virion, similar to the viral glycoprotein gB. Protein analysis procedures were similar to those described in Subheading 3.4. Figure 4 demonstrates trypsin sensitivity of virion-associated gB as well as R129 and R131. 1. For this analysis, create duplicate samples of virion preparations; one of the duplicates will be treated with trypsin and the other will remain untreated. To produce each sample, pipet equivalent gB levels of each virus into an Eppendorf tube and add PBS to a total volume of 15 μL. Add 10 μL of sequencing

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grade modified trypsin (0.5 mg/mL) or the equivalent volume of PBS and incubate the samples at 37  C for 1 h. 2. Add 25 μL of Tricine loading buffer containing 2% BME to each sample and heat at 100  C for 5 min. 3. Separate samples by SDS-PAGE using 10–20% Tricine gels with a run time of 1 h at 125 V. 4. Transfer the proteins to a PVDF membrane using a semidry transfer system at 25 V for 25 min. 5. Detect RCMV gB by immunoblotting and the HiBiT-tagged proteins using the HiBiT blotting system as described above in Subheading 3.4. 3.6 Quantification of Virion-Associated R131 and R129(short) HiBiT–Tagged Molecules Relative to Viral Genome Copy Number

3.6.1 In-Solution Detection of HiBiT

In order to quantify the levels of the pentamer complex proteins per virion, we developed a two-pronged strategy. First, we determined the level of virion incorporated HiBiT-tagged proteins using an in-solution assay and a commercially available HiBiT control protein standard. In the second step, viral genome levels in equivalent amounts of virion preparations was quantified using real-time PCR. Three different starting amounts of each virus were used in order to assess whether input levels affect the detection and calculation of HiBiT molecular copy number per viral genome. We calculated the relative copy number of virion-associated R129 and R131 and found that they were equimolar. This method could be useful to identify functional domains in R129 and R131 and determine how mutations in these proteins affect the levels of virion-associated pentamer and to correlate how the level of pentamer affects entry activity. 1. Add 7.5 μL, 3.75 μL, 1.875 μL of R131 HiBiT, R129(short) HiBiT, and WT RCMV purified viral particles in PBS in duplicate to a white-walled 96-well plate. 2. QS samples to 25 μL with PBS. 3. Prepare a 7-point standard curve using the HiBiT control protein diluted in PBS, ranging from 1000 to 15.625 nM. Add 25 μL of each standard in duplicate to the white-walled 96-well plate. 4. Add 25 μL of PBS to duplicate wells as a background control. 5. Dilute the LgBiT protein 1:100 and the Nano-Glo HiBiT Lytic Substrate 1:50 in Nano-Glo HiBiT Lytic Buffer. Add 25 μL of HiBiT lytic detection reagent to each sample well. 6. Seal the plate and shake at medium-low on a plate shaker in the dark for 10 min. 7. Read sample luminescence on a 96-well plate reader (see Note 10). (Fig. 5a).

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Fig. 5 Determination of R131 and R129(short) HiBiT molecules relative to viral genome copies. (a) In-solution luminescence assay results of HiBiT-tagged viral proteins. Three different volumes of viral particles were diluted to 25 μL and loaded into a 96-well plate in duplicates. (b) Log–log transform of the standard curve from the HiBiT lytic detection assay with mean of duplicate standards shown in red and unknown sample means shown and labeled. R2 of the linear regression of the standard curve was 0.9950. Standard curve was defined as a given number of molecules of HiBiT control protein. (c) Molecules of HiBiT-tagged proteins in unknown samples calculated from standard curve using average luminescence of duplicate sample wells. (d) log(X) transform of qPCR standard curve with mean of technical triplicates shown in red and unknown sample means shown and labeled. R2 of the linear regression of the standard curve was 0.9954. Standard curve was created using RCMV viral DNA at known genome concentrations in 1:10 dilutions ranging from 5  106—5 genome copies per well. (e) RCMV DNA Pol copies per 5 μL of RCMV viral DNA determined by qPCR against a standard curve of known genome copies. (f) Molecules of HiBiT per genome copy as determined in Subheading 3.6.2. R131 HiBiT and R129(short) HiBiT showed similar levels of protein incorporation per viral genome

8. Calculate molecules of HiBiT control protein for each standard. Perform a linear fit on a log-log transform of the standard curve using the mean of the standard curve duplicates (Fig. 5b). Calculate the molecules of HiBiT-tagged protein in the samples based on the mean luminescence reading for each sample. (Fig. 5c).

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1. Add 1 mL DNAzol to 15 μL of purified virus preparations (see Note 11). Add 500 μL 100% ethanol to each sample. Mix by inverting the tube ten times. 2. Centrifuge at 2872  g in a microfuge for 15 min to pellet the DNA. 3. Wash the DNA pellet twice with 1 mL 70% ethanol. After each wash, spin the sample at maximum speed in a table-top centrifuge for 5 min. Carefully pipette off all of the supernatant and air dry the pellet for 15 min. 4. Once the pellet is dry, add 30 μL of molecular biology-grade water to resuspend the DNA. 5. Heat the sample at 55  C in a shaker for 10 min. 6. Dilute the viral DNA 1:3 in molecular biology-grade water. Perform qPCR with primers and probes designed against RCMV DNA polymerase (R54) in technical triplicate. 7. Prepare RCMV qPCR standard consisting of 1:10 dilutions ranging from 1  106 genome copies/μL to 1 genome copy/ μL. Use 5 μL/well for samples and standards. 8. Prepare PCR reaction for detection of RCMV genomic DNA. Load 10 μL PCR mix/well plus 5 μL sample DNA/well. 9. CMV qPCR run cycle parameters: 3 min at 95  C/s, then 40 cycles of 95  C for 1 s, ramped down at a rate of 1.6  C/ s to 60  C, and 60  C for 20 s. These PCR conditions were optimized for this primer and probe set. 10. Utilize TaqMan software to calculate the number of genome copies/sample (Fig. 5d, e). 11. Calculate the number of genome copies/μL in each original prep (RCMV WT, RCMV R131 HiBiT, and RCMV R129(short) HiBiT) as follows: l RCMV DNA pol copies ¼ RCMV genome copies l

Genome copies per sample ¼ Genome copies per 5 μL

l

Genome copies per 5 μL 5

l

l

¼ Genome copies per 1 μL

Genome copies per 1 μL  3 ¼ Original genome concentration from extracted DNA per μL Original genome concentration from extracted DNA per μL  30 μL ¼ Total genome copies in extracted DNA

l

Total genome copies in extracted DNA 15 μL ¼ Total genomes per μL in initial prepared virus 12. Calculate the Molecules of HiBiT-tagged protein per μL over Genome copies per μL for each virus (Fig. 5f).

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Observations

Protein tags are an essential tool of biochemical studies. Here, we have demonstrated that the 11-amino acid HiBiT tag of proteins in CMV BAC-derived viruses enable specific and quantitative detection of proteins. We used this technology to visualize and quantify incorporation of two putative pentamer entry proteins in RCMV, R129, and R131. Furthermore, we were able to quantify the incorporation of these proteins per viral genome by measuring both incorporated proteins and viral genome copies per volume of viral particles. We determined that R129(short) was incorporated into viral particles at an average of 16,622 molecules per viral genome, and that R131 was incorporated at an average of 13,508 molecules per viral genome. These calculations assume that the number of viral particles was equal to the number of viral genomes and future studies should incorporate infectious virus particles into these equations. Future studies utilizing this technique can be performed to identify the functional domains in the C0 -terminal region necessary for formation of the RCMV PEC and determine the effect of mutations on the number of R129 and R131 proteins incorporated into viral particles. Although, we showed here that R129(short) HiBiT is still incorporated into viral particles, we left the predicted necessary charge clusters intact in this mutant (see Fig. 1a). Next, we intend to design C0 -terminal/acidic cluster mutants to determine the impact of these mutations on incorporation of these proteins into the viral particle. This technology could also be employed to determine the molecular copy number of other virion-associated proteins or to quantify viral protein expression levels in cells.

Notes 1. RFL-6 cells were used to transfect, expand, and titer the virus. Cells were maintained in Dulbecco’s modification of Eagle’s Medium (DMEM; Corning) supplemented with 5% fetal bovine serum (FBS), 100 U/mL penicillin, 100 mg/mL streptomycin, and 20 mM L-glutamine (PSG). 2. The amino acid sequence of the HiBiT tag is VSGWRLFKKIS (GTG AGC GGC TGG CGG CTG TTC AAG AAG ATT AGC) [5, 11]. Location of the HiBiT tag within the protein is an important consideration as placement into a region that is buried upon protein folding will decrease detection of naturally folded protein. The 6-His tag was added to allow for eventual pull-down of the tagged proteins and their complexes. The 6-His tag was constructed using alternating CAT-CAC codons. 3. The length of the homology arms should be at least 50 bp each. If the second step of the recombination process (replacing

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GalK/Kan with the gene insert) proves challenging, the homology arms should be extended to 100+ bp each as a first approach. 4. For mixed colonies detected by PCR screening, repeat growth in 2X YT broth plus 12.5 μg/mL Chlor and 100 μg/mL Kan followed by plating onto MacConkey/Kan/Chlor selection plates and rescreen colonies by PCR. If necessary, the concentration of the Kan may be increased up to 200 μg/mL to improve selection. 5. For convenience, we order the second-Step replacement DNA fragment from a commercial vendor. The R129 and R131 HiBiT tag replacement fragments contained a 6-His/HiBiT fusion tag flanked by 50–100 bp homology to the insertion site in the RCMV genome. These gene fragments may be PCR amplified and cloned to provide a stock of the gene fragment for future PCR reactions. 6. For the second recombination step, positive clones arise through recombination and replacement of the GalK gene; however, it is possible that spontaneous mutations that arise in the GalK gene can grow on the DOG plates. PCR and sequencing should reveal whether the colonies contain the proper genetic recombination event. 7. DNA midiprep kit was used to prepare BAC DNA for analysis and transfection into mammalian cells. For RCMV BAC recovery, the phenol–chloroform method of DNA isolation failed to successfully rescue RCMV. BAC DNA preparations should be used within 24 h for best results. 8. PCR amplification and gel purification of the region including the inserted tags prior to sequencing provides cleaner sequencing results. However, it is possible to submit the BAC DNA for sequencing directly. Whole DNA genome sequencing should be performed on viral genomic DNA preparations. 9. A discontinuous 10–50% Histodenz gradient was prepared in TNE buffer (50 mM Tris [pH 7.4], 100 mM NaCl, and 10 mM EDTA). The 50% Histodenz layer was at the bottom of an SW41Ti ultracentrifuge tube and individual 5% steps (1 mL) were layered on top. Each layer was frozen prior to addition of the next to prevent mixing of layers and to retain clean step interface lines. Prior to use the centrifuge gradients were completely thawed. Virus particle preparations were resuspended in 2 mL of TNE buffer (no Histodenz), which was added to the top of the 10% Histodenz layer prior to centrifugation. 10. A Synergy HTX multimode microplate BioTeK plate reader was used to read the HiBiT in-solution assay with the following settings: Luminescence endpoint; Integration time (1.00 s);

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Emission (Hole); Optics: (Bottom); Gain (135); Actual Temperature (22.6  C). 11. DNA extraction is necessary to obtain clean DNA for the qPCR reaction. Performing qPCR on the viral preparation alone results in degradation of the PCR product.

Acknowledgments The methods development presented in this chapter were supported by a grant from the National Institutes of Health NIAID RO1 AI116633. IJ was supported by the OHSU Molecular Microbiology and Immunology Interactions at the Microbe/Host Interface training grant NIH T32 AI007472. We thank Drs. Brock Binkowski, Chris Eggers, and Jessica Rossol-Allison for helpful discussions and advice about the HiBiT system. References 1. Wille PT, Wisner TW, Ryckman B et al (2013) Human cytomegalovirus (HCMV) glycoprotein gB promotes virus entry in trans acting as the viral fusion protein rather than as a receptor-binding protein. MBio 4: e00332–e00313 2. Ryckman BJ, Rainish BL, Chase MC et al (2008) Characterization of the human cytomegalovirus gH/gL/UL128-131 complex that mediates entry into epithelial and endothelial cells. J Virol 82:60–70 3. Calo´ S, Cortese M, Ciferri C et al (2016) The human cytomegalovirus UL116 gene encodes an envelope glycoprotein forming a complex with gH independently from gL. J Virol 90:4926–4938 4. Vanarsdall AL, Howard PW, Wisner TW et al (2016) Human cytomegalovirus gH/gL forms a stable complex with the fusion protein gB in Virions. PLoS Pathog 12:e1005564 5. Schwinn MK, Machleidt T, Zimmerman K et al (2017) CRISPR-mediated tagging of endogenous proteins with a luminescent peptide. ACS Chem Biol 13:467–474 6. Akter P, Cunningham C, McSharry BP et al (2003) Two novel spliced genes in human cytomegalovirus. J Gen Virol 84:1117–1122

7. Malkowska M, Kokoszynska K, Dymecka M et al (2013) Alphaherpesvirinae and Gammaherpesvirinae glycoprotein L and CMV UL130 originate from chemokines. Virol J 10:1 8. Schuessler A, Sampaio KL, Sinzger C (2008) Charge cluster-to-alanine scanning of UL128 for fine tuning of the endothelial cell tropism of human cytomegalovirus. J Virol 82:11239–11246 9. Schuessler A, Sampaio KL, Scrivano L et al (2010) Mutational mapping of UL130 of human cytomegalovirus defines peptide motifs within the C-terminal third as essential for endothelial cell infection. J Virol 84:9019–9026 10. Vomaske J, Denton M, Kreklywich C et al (2012) Cytomegalovirus CC chemokine promotes immune cell migration. J Virol 86:11833–11844 11. Landreman A and Eggers C Quantifying Protein Abundance at Endogenous Levels, https://www.promega.com/resources/ pubhub/2017/quantifying-protein-abun dance-at-endogenous-levels/

Chapter 12 Using a Phosphoproteomic Screen to Profile Early Changes During HCMV Infection of Human Monocytes Liudmila S. Chesnokova and Andrew D. Yurochko Abstract During the binding and infection of monocytes, HCMV binds to at least two major cell surface receptors/ receptor families: the epidermal growth factor receptor (EGFR) to initiate downstream signaling through the EGFR-PI3K pathway, and to β1- and β3-integrins to initiate downstream signaling through the integrin-c-Src pathway (Nogalski et al. PLoS Pathog 9:e1003463, 2013; Chan et al. Proc Natl Acad Sci U S A 106:22369–22374, 2009; Kim et al. Proc Natl Acad Sci U S A 113:8819–8824, 2016; Wang et al. Nature 424:456–461, 2003; Wang et al. Nat Med 11:515–521, 2005; Yurochko et al. Proc Natl Acad Sci U S A 89:9034–9038, 1992). Signaling through these receptors can occur rapidly with phosphorylation observed as early as 15 s after EGF-EGFR interaction, for example (Alvarez-Salamero et al. Front Immunol 8:938, 2017). The ability to detect signaling and the consequences of that signaling are critical for our understanding of how HCMV-receptor engagement promotes infection and modulates the biology of different target cells. In this chapter we describe how we used an ELISA-based antibody platform to perform an assessment of the rapid phosphorylation events that occur in monocytes following infection. This assay can be adapted to other infection systems, time points and cell types as needed. Together, we examined via an ELISA-based antibody array a phosphoproteomic screen to search for potential phosphorylated proteins that might influence HCMV infection. Key words Phosphoproteomic screen, Phosphoproteome, Human peripheral blood monocytes, Monocytes, HCMV

1

Introduction The early steps of herpesvirus infection may be loosely dissected into several steps: virus binding to the cell surface, interaction with an entry receptor, crossing the membrane barrier or virus entry into the interior of the cell, capsid release into the cytoplasm, and delivery of the viral nucleic acids to the nucleus to allow the initiation of the viral gene cascade [1–6, 8–13]. For HCMV infection (especially of monocytes), all of these steps are tightly associated with the cell signaling that is initiated following viral receptor– ligand engagement [6, 14–18]. Most receptor ligand engagements activate protein kinases (both receptor associated and non–

Andrew D. Yurochko (ed.), Human Cytomegaloviruses: Methods and Protocols, Methods in Molecular Biology, vol. 2244, https://doi.org/10.1007/978-1-0716-1111-1_12, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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receptor-associated protein kinases). Phosphorylation and dephosphorylation of the hydroxyl groups of serine, threonine and tyrosine are the essence of the kinase-regulated cell signaling pathways [7]. Several experimental approaches can be used to study changes in patterns of phosphorylation on individual proteins of interest; but to study global changes, one would need to study the phosphoproteome. That is, if a researcher has the knowledge of the specific signaling pathways involved in their infection system, the choice of the targeted western blot or ELISA may be the likely assay of choice. However, to take a generalized view of the whole signaling network or to identify potential new regulated proteins following infection, quantitative mass-spectrometry-based or ELISA-based phosphoproteomic assays would likely be the best choice. In this chapter we focus on the use of Antibody Phospho Arrays in our monocyte system. These arrays have a basic ELISA-like setup and are designed for detecting and quantifying multiple phosphorylated residues on multiple proteins. One does have to note that ELISA-based phosphoproteomic techniques can show crossreactivity due to the very nature of the antibodies used (particularly antibodies to phosphotyrosine). However, in eukaryotic cells, the ratio of serine/threonine/tyrosine phosphorylation is 81%:17%:2% [7], thus the risks of small number of false positives can be managed. The assay, discussed in this chapter, uses over 1300 antibodies specific to phosphorylated residues of several hundred proteins involved in cellular signaling, or to their nonphosphorylated counterparts. Antibodies are printed on a glass slide, and form a microarray grid. Biotinylated soluble cell proteins specifically bind to the array, and the treatment with the fluorescent streptavidin allows one to visualize the binding (see flow chart in Fig. 1). The comparison of fluorescence on two different slides indicates the changes in amount and phosphorylation in different experimental settings. One can therefore use this assay to screen for the changes in phosphorylation patterns on a variety of residues on major signaling proteins. By knowing the nature of these proteins and the activating or deactivating nature of the patterns of phosphorylation, one can potentially decipher patterns of upstream signaling and the functional consequences of those changes [19, 20]. We focus on infection of monocytes, a key cell type in the hematogenous dissemination of HCMV following primary infection, the establishment of long term persistence, and in the process of reactivation from latency [3, 12, 21–24]. With cell type specific differences seen in the nature of viral entry and the signaling ensuing post receptor ligand engagement [25–27], this protocol gives us a chance to investigate the specific proteins that are altered (phosphorylated/ dephosphorylated) following specific receptor–ligand engagement in monocytes. We choose to examine early time periods postinfection because of the more easily defined nature of the

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Fig. 1 Experimental flowchart for use of monocytes in our proteomic screen. Human peripheral monocytes (12
106 per condition) were mock-infected or HCMV-infected. Soluble proteins from both cell extracts were biotinylated and bound to specific antibodies printed on the slides. Bound proteins were visualized with Cy-3 streptavidin. Fluorescence intensity was measured with a 75 mm
25 mm microarray fluorescence scanner. Changes in phosphorylation of individual residues were determined by data analysis with Microsoft Excel program

stimulus (receptor–ligand engagement in the absence of secondary signaling) and thus a more direct link to the mechanisms involved in changes in protein phosphorylation. The experimental setup needs to be carefully considered for each biological system and can readily be adaptable as stated above to multiple systems, time points, and cell types, as long one considers the appropriate biological variables when make statements and conclusions about the generated data.

2

Materials

2.1 HCMV Culture and Infection (See Note 1)

1. Human fibroblasts. 2. CO2 incubator with 5% CO2. 3. HCMV of known MOI (see Note 1). 4. RPMI; no additives to resuspend the virus. 5. Heat-inactivated fetal bovine serum (HI-FBS) to make 4% HI-FBS-DMEM. 6. 20% sorbitol (w/w) containing 1 mM MgCl2, or 20% sucrose (w/w).

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2.2 Isolation of Human Peripheral Blood monocytes (See Note 2)

1. Whole blood from healthy donors. 2. 1% HS-RPMI, 1% male AB human serum (HS) and RPMI. 3. Ficoll Histopaque 1077 (see Note 3). 4. 0.9% irrigation saline. 5. Percoll. 6. 15 and 50 mL polypropylene centrifuge conical tubes (pyrogen free).

2.3 Monocyte Infection

1. CO2 incubator (37  C; 5% CO2). 2. Pipette tips (when intended for adding virus, the tips should be prechilled at 20  C for at least an hour). 3. 1% HS-RPMI.

2.4 Protein Extraction

1. 1
PBS. 2. Standard phosphatase/protease inhibitor cocktail (see Note 4). 3. Full Moon Extraction buffer (Full Moon Biosystems array kit, Cat. no. KAS-2 or KAS-20). 4. Lysis glass or ceramic beads. 5. Vortex.

2.5 Protein Binding to Antibody Arrays and Array Processing

1. Full Moon Antibody array assay kit and antibody arrays. 2. Calibrated pipettes for 2, 20, and 200 μL (see Note 5). 3. Cy3-Streptavidin (see Note 6). 4. You can use the Full Moon Company service for reading the array’s fluorescence intensity (it is free), or you can use a 75 mm
25 mm Microarray fluorescence scanner (the list of array-compatible scanners may be found at the manufacturer’s website: https://www.fullmoonbio.com/datasheets/ AbArrayDetectionGuide.pdf) with suitable software (see Note 7). 5. Milli-Q Grade Water, or distilled deionized water (DDI water) of the same quality (see Note 8). 6. Nano Drop, or a spectrophotometer to measure OD280. 7. Orbital shaker. 8. Tabletop centrifuge. 9. Protein Concentrators, cutoff 10 kDa. 10. 30  C cabinet dryer. 11. 15 mL and 50 mL conical tubes. 12. 100
20 mm cell culture dishes. 13. Vortex.

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Methods

3.1 HCMV Culture and Infection (the Step Takes About 3 Weeks (See Chapter 3 for Additional Detail)

1. Infect confluent HEL fibroblasts at a multiplicity of infection (MOI) appropriate to your system. 2. Keep cells in culture in 4% HI-FBS-DMEM for 2–3 weeks or until the cytopathic effect are seen in a majority of the cells. 3. Collect the supernatant, and clear it by spinning at 800
g for 10 min to get rid of cellular debris. 4. Concentrate HCMV by centrifugation through a 20% sorbitol1 mM MgCl2 (or 20% sucrose) cushion (Beckman Rotor SW32Ti: 22,000 rpm (83,000
g), SW28: 22,000 rpm (87,000
g)) for 1 h. Resuspend the pellet in RPMI, and keep frozen at 80  C, or in a liquid nitrogen tank until use.

3.2 Isolation of Human Peripheral Monocytes (Takes 5– 6 h). See Note 2 and Chapter 6 for Detailed Instructions

Monocytes are isolated from human peripheral blood as previously reported [3, 28–30] and see Chapter 6). All university-required Institutional Review Board and Health Insurance Portability and Accountability Act or other appropriate guidelines should be followed during any study involving human subjects. The main steps of monocyte isolation include separation of red and white blood cells, separation of mononuclear cells from the whole leukocyte pool using a density gradient and then isolation of monocytes from lymphocytes using another density gradient. Once harvested, resuspend purified monocytes in 1% HS-RPMI.

3.3 Monocyte Infection (Takes Less Than 3 h)

Here, we describe the protocol for monocyte infection with HCMV. Use an appropriate protocol suitable for your system. Infect cells at an MOI that is appropriate to your system. We favor MOIs of 3–5 for our phosphoproteomic analyses because most of the monocytes are infected and we see a good signal-tonoise ratio at this MOI that does not really improve with higher MOIs. MOIs of 0.5–1 should also work, but the signal-to-noise ratio may be diminished. The infection process used for monocytes includes steps that are common for any system used. However, highlighted below are the steps used for synchronized infection of monocytes or other cells for distinct short times of infection. 1. Wash cells in all tubes with ice-cold RPMI (centrifugation at 500
g, 5 min); aspirate media, and resuspend cells in 50 μL of ice-cold RPMI. 2. Chill cells on ice for 1 h. 3. Use chilled pipette tips to add prechilled virus to the cells. 4. Bind virus to cells for 1 h on ice (see Note 9). 5. Remove unbound virus by washing the cells twice with ice-cold RPMI (centrifugation at 500
g, 5 min).

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6. Add 100 μL of ice-cold RPMI to each cell pellet, gently resuspend cells, and shift the temperature to 37  C for 15 min. 7. After incubation, wash the cells twice with an ice-cold PBS (centrifugation at 500
g, 5 min), and then resuspend in Full Moon Protein Extraction Buffer (see Note 10) supplemented with a 1
phosphatase and protease inhibitor cocktail (see Note 4). Continue to the next step. 3.4 Protein Extraction (~1.5 h): After This Step, the Sample Can Be Stored at 80  C for up to 2 Weeks

1. At this step, the critical barrier for success of the experiment is to minimize protein degradation and dephosphorylation of the proteins in the sample. Disrupt cells mechanically by six 60 slong vortexing pulses with 10 min intervals with beads. Add fresh phosphatase and protease inhibitor cocktail every 15 min. 2. Keep the samples on ice the entire time you are between vortexing. 3. To clarify supernatant, centrifuge cell extracts for 10 min at 13,000
g at 4  C. 4. Transfer cell extracts to clean prechilled tubes using gel loading tips, and then centrifuge the samples once again at 13,000
g for 10 min. 5. If you decided to freeze the sample at 80  C at this step, the clear supernatant should be again supplemented with a fresh 1
phosphatase/protease inhibitor cocktail. 6. To make sure that the cell extract has a good quality, a fraction of the sample could be taken (before freezing!) to be used for a Western blot analysis.

3.5 Buffer Exchange and Protein Determination (This Step Takes ~30 min): After This Step, the Sample Can Be Stored at 80  C for a Few Days

Replacement of extraction buffer with the labeling buffer is conducted by a batch variant of size-exclusion chromatography according to the array kit protocol. 1. Equilibrate the dry resin inside a spin column with the labeling buffer, after 1 h, remove the excess liquid by centrifugation at 750
g for 2 min at 4  C. 2. Move quickly and add a sample to the center of the column. Do not touch the gel. Insert the column into a new prechilled Eppendorf tube to collect the sample. 3. Collect the proteins by a second centrifugation at 750
g for 2 min at 4  C. 4. Immediately add a fresh protease/phosphatase inhibitor cocktail to the final concentration 1
. 5. Evaluate protein concentration using a spectrophotometer, based on OD280 (Protein OD280 program), or run any protein assay you prefer (Lowry, Bradford, BCA). The goal is to have the protein concentration about 5 mg/mL; if the volume of

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the extraction buffer was too large, and the extract is too dilute, use a protein concentrator with a cutoff 10 kDa, to concentrate the sample (see Note 10). 3.6 Unpacking, and Blocking the Array Slides Prior to the Protein Binding (the Step Takes About 3 h and May Be Done at the Same Time as Subheading 3.5)

1. For consistency and to avoid unnecessary variability, it is best to make sure that all arrays have the same lot number (see Note 11). 2. To preserve the immobilized antibodies, the arrays should be adjusted to room temperature and completely air dried before using. The manufacturer’s protocol is very useful here. Since this step takes several hours, we usually start it before the buffer exchange and protein biotinylation step. 3. Write down the array numbers in your notebook, and assign each slide to a sample. 4. Add 30 mL of Blocking Buffer in a 100
20 mm cell culture dish (see Note 12). 5. Holding the slide on by edges, submerge it in the Blocking Solution. The side with a barcode label must always face up. Incubate on an orbital shaker rotating at slow speed for 1 h at RT. 6. Rinse the slide extensively with DDI water using a 50 mL conical tube. 7. Repeat washing as many times as it takes to stop seeing the antibody grid. 8. Do not allow the slides to dry out. Between steps, keep them in 50 mL conical tubes filled with DDI water.

3.7 Protein Biotinylation. (The Process Takes about 3 h and Could Be Done at the Same Time as the Array Slides Are Being Equilibrated, Blocked and Rinsed)

Biotin, or Vitamin H, is a small molecule that has a high affinity for the protein avidin (KD ~ 1014 M) and, especially, for streptavidin (KD ~ 1015 M) [31]. Wild type streptavidin is a tetramer, which binds four biotin molecules, however, a recombinant streptavidin, broadly used in biotechnology, is usually a monomer, which binds to biotin according 1:1 Langmuir kinetic model [32]. Chemical (primary amine) biotinylation targets a lysine side chain epsilonamino groups and all N-terminal alpha-amino groups. In this array preparation, extensive biotinylation of the proteins in the sample is the key to success since only biotinylated proteins can be detected by fluorescent streptavidin. 1. Dissolve the Full Moon “Biotin Reagent” in N,N-Dimethyl formamide (DMF, included in a Full Moon reagent kit). Mix well by vortexing, then spin the tube in a tabletop centrifuge (see Note 13). Visually make sure that the powder is completely dissolved.

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2. For biotinylation, you need 100–120 μg of the cell proteins in the volume less or equal to 25 μL (see Note 14). Perform the biotinylation according to the array kit protocol. 3.8 Binding of Biotinylated Proteins to the Arrays (~4 h)

1. In 15 mL conical tubes, bring the volume of each of your biotinylated samples up to 6 mL with the coupling buffer, and mix well. 2. Remove a slide from the DDI water; shake off the water drops, then place the slide in a coupling chamber (included in the kit), and slowly pour in the mixture from the previous step. 3. Repeat for each protein sample in your experiment. 4. Set the chambers on an orbital shaker, and incubate at low speed for 2 h at RT. 5. Wash each slide three times with 30 mL of washing buffer. 6. Rinse the slide extensively with DDI water using 50 mL conical tubes, as before. Repeat for a total of ten washes.

3.9 Detection of Bound Biotinylated Proteins with Cy3Streptavidin (~1 h)

Prepare the fluorescent streptavidin solution for all your arrays by diluting it to the detection buffer. For one slide, mix 15 μg of Cy3-streptavidin to 30 mL of the detection buffer (see Note 6). If you have several slides, multiply appropriately. Use a dark glass bottle for making this solution; if using a clear bottle, wrap it in aluminum foil to protect the solution from light. The following steps should be done in a dimmed room to avoid fluorescence bleaching. We covered the cell culture dishes used for the wash buffer and the 50 mL conicals used for the water with aluminum foil. 1. For each slide, pour 30 mL of Cy3-streptavidin-detection buffer mixture into a 100
20 mm cell culture dish, and place a slide with the barcode up. 2. Incubate on an orbital shaker, rotating at low speed for 30 min at RT. 3. Wash each slide three times with 30 mL of washing buffer. 4. To get rid of background fluorescence and increase the signalto-noise ratio, it is very important to remove unbound fluorescent dye completely. Wash slides at least ten times with DDI water (manufacturer’s protocol). We found that 15 washes gave a better signal-to-noise ratio in the final samples.

3.10 Array Quick Drying (~15 min)

To avoid water spots, quickly dry the slides after the last wash. 1. Place the slides into clean 50 mL tubes and spin for 30 s at 50
g in a tabletop centrifuge. 2. Residual moisture should be quickly removed by gently blowing air from a clean compressed air outlet.

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Fig. 2 Sample array images. An example of the arrays received from Full Moon are shown. Each array consists of 16 groups of printed antibodies; the left and the right columns on each slide are identical and thus serve as technical replicates. Each dot represents a distinct antibody and thus a separate potential data point. After reading the slide at the Full Moon Company (using the following parameters: λex ¼ 550 nm; λem ¼ 650 nm), the images of the arrays (as .tif files) are returned and used to assess the numerical data with the appropriate array scanner software. The numbers underneath each individual array represent the unique slide identifiers for the experiments performed

3. Dry slides are ready for reading. Place them back to their boxes and wrap the boxes in foil. Slides should be stored in a dark and dry location at RT. The slides can be analyzed with a 75 mm
25 mm slide-based microarray scanner with appropriate software (the list of array-compatible scanners may be found at the manufacturer’s website: https://www.fullmoonbio. com/datasheets/AbArrayDetectionGuide.pdf). Alternatively, the slides can be shipped to the Full Moon Company for reading; this is a free service. The files with the results can then be downloaded from the Full Moon website and opened with a slide scanner software for quantification (Fig. 2). We have compared the read data performed by Full Moon with compatible scanner and the results were identical. Bioinformatic analysis is available from Full Moon; however, there is a fee for the service (see Note 7). 3.11 Raw Data Analysis (See Note 6)

1. The array scanner software allows one to open the results as a spreadsheet (e.g., in Microsoft Excel). One would then compare the fluorescence reading data (sample data) and the fluorescence minus background (background fluorescence has been subtracted from the sample fluorescence) columns. If the slides were carefully washed of background fluorescent dye, these two numbers/values should be very similar.

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2. Choose the set of data in which the background was subtracted from the Cy3 fluorescence data. Transfer the data into a spreadsheet for analysis. 3. Every slide contains two technical replicates of each array. Calculate the average of the two data sets (the average of the technical replicates (see Note 15)). 4. Calculate the fold changes as a ratio of mean intensity of one array by the second array (that may be infected vs control, plus drug vs no drug, etc.). 5. Compute (if appropriate, depending on the number of repeats) the p-value using a two-tailed, two sample unequal variance T-test, or other appropriate statistical test (see Note 16). 6. Sort the data based on p-value. The data will identify those proteins that show a statistically significant increase in phosphorylation, decrease in phosphorylation and those proteins that remain statistically unchanged. 7. Separate the data for nonphosphorylated proteins. Evaluate the difference between two slides, and consider the normalization step if the difference exceeds 20% (see Note 17). 8. Depending on the questions addressed, one can sort the data based on phosphorylation fold change to determine the residues in which phosphorylation (up or down) was the affected by viral infection. 9. Present the results as a table or as a bar graph (or in any type of format) that shows the comparisons or changes in your system.

4

Notes 1. For this chapter, we have discussed a general protocol that can be used for multiple experimental setups. However, some sections are tailored for the system we employ—infection of primary blood monocytes. With that point in mind, we studied the effects of HCMV on cell signaling in monocytes at a short time frame of 15 min. Any other cell type or time point could be used, as well as any other virus. It should be noted that careful consideration for the culture conditions and the timing may affect the data generated and the interpretation of the results. Our goal was to have no secondary signaling and to examine those rapid events associated with viral entry. To modify the protocol one only has to change the sections associated with cell culture and infection. The protocol for the array itself is independent of the culture conditions. We mainly followed the outline of the manufacturer’s protocol and have added comments from our experience using this phosphoprotein antibody array.

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2. See Chapter 6 in this volume for detailed instructions for harvesting human peripheral blood monocytes from whole blood. 3. In our laboratory, we tested several products suitable for monocyte isolation: Histopaque 1077 and Histopaque-1077 Hybri-Max (both from Sigma-Aldrich), and Ficoll-Paque PLUS Media (GE Healthcare) and found no difference in monocyte quantity and quality following isolation. 4. The manufacturer’s reagent kit does not include any phosphatase or protease inhibitors, however, we decided to use both of them to prevent protein degradation and dephosphorylation. We used the Halt 100
cocktail (Thermo Fisher) at 1
concentration. Some of the protein inhibitors, included in the cocktail, are not stable (e.g., PMSF), so, we added fresh inhibitor every 15 min. 5. Calibrated pipettes are absolutely essential for good results; even a 1–2 μL error during the addition of the reagents can negatively affect the outcome of the experiment. 6. Cy3-streptavidin is not included in the kit and should be bought separately. Depending on the manufacturer, the product may have different fluorescent dye concentration. The calculations in the Full Moon manual refer to concentration of 0.5 mg/mL, but we used Cy3-Streptavidin (GE Healthcare; PA43001) with the concentration of 1 mg/mL, and therefore we had to reevaluate the math. 7. The Full Moon Company offers a service in which they will read the final slides and provide you with the images of the slides (Fig. 2 represents an example of our generated data using these slides). The array scanner software converts these images to numerical values. We have compared reading of the slides from Full Moon to the use of our own slide scanner, and the results were nearly identical in all cases. They also provide a fee for service to bioinformatically analyze the data. Simple data analysis can be performed using a spreadsheet styled program; see Subheading 3.11 of this chapter. 8. Milli-Q grade water is a distillate deionized water (DDI water) purified by ion-exchange chromatography to reach resistivity 18.2 MΩ (million ohm-cm) and conductivity of 0.055 microsiemens. 9. This step allows low affinity virus binding (which increases the virus’ local concentration, and therefore, from a thermodynamic point of view, increases the efficiency of viral entry/ internalization). Furthermore, it allows synchronization of the infection, which is needed for short term incubations (under 1 h).

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10. One should be able to calculate the approximate volume of the extraction buffer to ensure a sufficient protein concentration: for example, 100 μg of soluble proteins can be extracted from approximately one million epithelial cells or three million monocytes. 11. Each array pouch contains two identical slides with printed microarrays, for two experimental settings (e.g., mock vs infected cells or one virus strain vs another virus strain). Data analysis is based on a comparison of phosphorylation on these two arrays. If you want to compare the results with any additional slides, make sure that you use the same lot number to minimize variations. 12. Make sure you are using a 10 cm dish, which is tall enough to hold 30 mL of liquid—this avoids any spills. 13. Almost every step of this protocol includes a mixing step. Use a vortexer or a pipette tip to ensure the complete mixing of the sample. To avoid losing the sample, spin a sample in a tabletop centrifuge after each mixing step for a few seconds. 14. There is a strict requirement for exact protein amounts since the immobilized protein is detected with a fluorescent streptavidin, which binds to biotin residues with a stoichiometry of 1:1 (no signal multiplication). In addition, the fluorescence of a sample needs to be in the range of the calibration curve, since the data for samples that have very low or very high protein amounts might be lost or underestimated. 15. An important question is how many biological replicates a researcher who uses this array needs to run to achieve their research goal (i.e., is it a search for potential hits, validation of a pathway, etc.). We found that if a sample shows strong similarity between technical replicates, then the data is probably going to be reliable. However, until biological replicates are performed, one cannot yet make strong conclusions. Once biological replicates are completed and the data collected is shown to be reproducible between the different biological replicates, solid conclusions can be made. We also always validated multiple proteins of interest via western blot (from additional donors) to provide the high correlation in our system needed to make strong conclusions. Thus from our experience, when we considered the cost of the array vs the number of biological replicates needed, we came to the conclusion that in most cases two biological replicates with two technical replicates (for a total 4 sample replicates) provides a sufficient set of data to determine the changes in the profile of protein phosphorylation in monocytes shortly after the addition of virus.

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16. We discarded all replicates with p-value greater than what is considered statistically significant, since we were worried differences in technical replicates from the same experiment could be caused by issues with the quality of the slide (the printed antibody layer is not always evenly distributed and there could be breakages and holes). This point can be resolved with multiple biological replicates. 17. Theoretically, the nonphosphorylated proteins should always give the same mean fluorescence on both slides from the technical replicates. Under experimental conditions, they are usually very close, but not identical. We never dealt with two very different sets of data, but we always considered the necessity of the normalizing step in a case where the differences would have exceeded 20%.

Acknowledgments This work was supported by grants from the National Institutes of Health: AI056077, AI127335, P30GM110703, P20GM121307, and P20GM121288. References 1. Nogalski MT, Chan GC, Stevenson EV, Collins-McMillen DK, Yurochko AD (2013) The HCMV gH/gL/UL128-131 complex triggers the specific cellular activation required for efficient viral internalization into target monocytes. PLoS Pathog 9:e1003463 2. Chan G, Nogalski MT, Yurochko AD (2009) Activation of EGFR on monocytes is required for human cytomegalovirus entry and mediates cellular motility. Proc Natl Acad Sci U S A 106:22369–22374 3. Kim JH, Collins-McMillen D, Caposio P, Yurochko AD (2016) Viral binding-induced signaling drives a unique and extended intracellular trafficking pattern during infection of primary monocytes. Proc Natl Acad Sci U S A 113:8819–8824 4. Wang X, Huong SM, Chiu ML, Raab-Traub N, Huang ES (2003) Epidermal growth factor receptor is a cellular receptor for human cytomegalovirus. Nature 424:456–461 5. Wang X, Huang DY, Huong SM, Huang ES (2005) Integrin alphavbeta3 is a coreceptor for human cytomegalovirus. Nat Med 11:515–521 6. Yurochko AD, Liu DY, Eierman D, Haskill S (1992) Integrins as primary signal transduction molecule regulating monocyte immediate-

early gene induction. Proc Natl Acad Sci U S A 89:9034–9038 7. Alvarez-Salamero C, Castillo-Gonzalez R, Navarro MN (2017) Lighting up T lymphocyte signaling with quantitative phosphoproteomics. Front Immunol 8:938 8. Knipe DM, Howley PM (2013) Fields virology, 6th edn. Wolters Kluwer/Lippincott Williams & Wilkins Health, Philadelphia, PA 9. Compton T, Feire A (2007) Early events in human cytomegalovirus infection. In: Arvin A, Campadelli-Fiume G, Mocarski E, Moore PS, Roizman B, Whitley R, Yamanishi K (eds) Human herpesviruses: biology, therapy, and Immunoprophylaxis. Cambridge University Press, Cambridge 10. Full F, Ensser A (2019) Early nuclear events after Herpesviral infection. J Clin Med 8 (9):1408 11. Kim JH, Collins-McMillen D, Buehler JC, Goodrum FD, Yurochko AD (2017) Human cytomegalovirus requires epidermal growth factor receptor signaling to enter and initiate the early steps in the establishment of latency in CD34(+) human progenitor cells. J Virol 91: e01206–e01216 12. Stevenson EV, Collins-McMillen D, Kim JH, Cieply SJ, Bentz GL, Yurochko AD (2014)

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HCMV reprogramming of infected monocyte survival and differentiation: a goldilocks phenomenon. Viruses 6:782–807 13. Campadelli-Fiume G, Collins-McMillen D, Gianni T, Yurochko AD (2016) Integrins as herpesvirus receptors and mediators of the host signalosome. Annu Rev Virol 3:215–236 14. Shenk T, Alwine JC (2014) Human cytomegalovirus: coordinating cellular stress, signaling, and metabolic pathways. Annu Rev Virol 1:355–374 15. Smith MS, Bentz GL, Smith PM, Bivins ER, Yurochko AD (2004) HCMV activates PI(3)K in monocytes and promotes monocyte motility and transendothelial migration in a PI(3)Kdependent manner. J Leukoc Biol 76:65–76 16. Collins-McMillen D, Chesnokova L, Lee BJ, Fulkerson HL, Brooks R, Mosher BS, Yurochko AD (2018) HCMV infection and apoptosis: how do monocytes survive HCMV infection? Viruses 10:533 17. Buehler J, Zeltzer S, Reitsma J, Petrucelli A, Umashankar M, Rak M, Zagallo P, Schroeder J, Terhune S, Goodrum F (2016) Opposing regulation of the EGF receptor: a molecular switch controlling cytomegalovirus latency and replication. PLoS Pathog 12: e1005655 18. Peppenelli MA, Miller MJ, Altman AM, Cojohari O, Chan GC (2018) Aberrant regulation of the Akt signaling network by human cytomegalovirus allows for targeting of infected monocytes. Antivir Res 158:13–24 19. Wikibooks TFTP (2018). Signaling inside the cell. https://en.wikibooks.org/w/index.php? title¼Principles_of_Biochemistry& oldid¼3470947 20. Taylor SS, Kornev AP (2011) Protein kinases: evolution of dynamic regulatory proteins. Trends Biochem Sci 36:65–77 21. Chan G, Nogalski MT, Stevenson EV, Yurochko AD (2012) Human cytomegalovirus induction of a unique signalsome during viral entry into monocytes mediates distinct functional changes: a strategy for viral dissemination. J Leukoc Biol 92:743–752 22. Caposio P, Orloff SL, Streblow DN (2011) The role of cytomegalovirus in angiogenesis. Virus Res 157:204–211 23. Crawford LB, Kim JH, Collins-McMillen D, Lee BJ, Landais I, Held C, Nelson JA, Yurochko AD, Caposio P (2018) Human cytomegalovirus encodes a novel FLT3 receptor

ligand necessary for hematopoietic cell differentiation and viral reactivation. MBio 9: e00682–e00618 24. Smith MS, Bentz GL, Alexander JS, Yurochko AD (2004) Human cytomegalovirus induces monocyte differentiation and migration as a strategy for dissemination and persistence. J Virol 78:4444–4453 25. Xiaofei E, Meraner P, Lu P, Perreira JM, Aker AM, WM MD, Zhuge R, Chan GC, Gerstein RM, Caposio P, Yurochko AD, Brass AL, Kowalik TF (2019) OR14I1 is a receptor for the human cytomegalovirus pentameric complex and defines viral epithelial cell tropism. Proc Natl Acad Sci U S A 116:7043–7052 26. Murrell I, Bedford C, Ladell K, Miners KL, Price DA, Tomasec P, Wilkinson GWG, Stanton RJ (2017) The pentameric complex drives immunologically covert cell-cell transmission of wild-type human cytomegalovirus. Proc Natl Acad Sci U S A 114:6104–6109 27. Nguyen CC, Kamil JP (2018) Pathogen at the gates: human cytomegalovirus entry and cell tropism. Viruses 10:704 28. Yurochko AD, Liu DY, Eierman D, Haskill S (1992) Integrins as a primary signal transduction molecule regulating monocyte immediateearly gene induction. Proc Natl Acad Sci U S A 89:9034–9038 29. Chan G, Nogalski MT, Bentz GL, Smith MS, Parmater A, Yurochko AD (2010) PI3Kdependent upregulation of Mcl-1 by human cytomegalovirus is mediated by epidermal growth factor receptor and inhibits apoptosis in short-lived monocytes. J Immunol 184:3213–3222 30. Nogalski MT, Chan GC, Stevenson EV, Collins-McMillen DK, Yurochko AD (2012) A quantitative evaluation of cell migration by the phagokinetic track motility assay. J Vis Exp 70:e4165. https://doi.org/10.3791/4165: e4165 31. Eakin RE, Snell EE, Williams RJ (1941) The concentration and assay of avidin, the injuryproducing protein in raw egg white. J Biol Chem 140:535–543 32. Sedlak SM, Bauer MS, Kluger C, Schendel LC, Milles LF, Pippig DA, Gaub HE (2017) Monodisperse measurement of the biotinstreptavidin interaction strength in a welldefined pulling geometry. PLoS One 12: e0188722

Chapter 13 A Generally Applicable CRISPR/Cas9 Screening Technique to Identify Host Genes Required for Virus Infection as Applied to Human Cytomegalovirus (HCMV) Infection of Epithelial Cells Xiaofei E and Timothy F. Kowalik Abstract Clustered regularly interspaced short palindromic repeats (CRISPR)/Cas9 screens enable virus-host genetic screens to be undertaken in a more robust manner than previously possible and has had a tremendous impact in the field of virus study. Researchers can take advantage of the power of CRISPR genetic screens to discover virus–host interaction genes including host receptors and signaling molecules (Bazzone et al., mBio 10 (1): e02734-18, 2019; E et al., Proc Natl Acad Sci U S A 116(14):7043–7052, 2019; McDougall et al., Curr Opin Virol 29:87–100, 2018; Savidis et al., Cell Rep 16(1):232–246, 2016). In principle, lysis of cells late in the virus infection cycle allows one to screen for essential genes using pooled single-guide RNAs (sgRNAs) that collective target an entire host cell genome simply by identifying mutant cells that are resistant to virus-induced cell death. Here we focus on using this technique on epithelial cells to identify host targets required for human cytomegalovirus (HCMV) infection. Key words CRISPR/Cas9 screening, Human cytomegalovirus, Method, Primary epithelial cells, HCMV

1

Introduction Infection is initiated when a virion attaches to a host cell via interactions with a specific receptor(s) on the cell surface. Following binding of a viral envelope glycoprotein(s) to a cell membrane receptor(s), the virion penetrates the host cell through receptormediated endocytosis or membrane fusion [1–5]. For HCMV, this process is complex as the virus encodes multiple glycoprotein ligand complexes that affect tropism. In addition, its adaptation to cell culture results in genetic changes that affect infectivity. The CRISPR/Cas9 [6, 7] system originates from a prokaryotic immune system that confers resistance to foreign genetic elements such as plasmids and viruses (bacteriophages) and provides an

Andrew D. Yurochko (ed.), Human Cytomegaloviruses: Methods and Protocols, Methods in Molecular Biology, vol. 2244, https://doi.org/10.1007/978-1-0716-1111-1_13, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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acquired immunity for the host [8–10]. The system has been shown to work as a genome engineering or editing tool in human cell culture, animals, and bacteria to mutate or functionally inactivate genes, among other applications [11, 12]. It has also been successfully applied to herpesviruses by engineering, targeting, activating, or repressing specific genes of interest [13, 14]. The delivery tools for transfection or gene transfer are agents that facilitate nucleic acids entry into target cells. The most common strategies for CRISPR/Cas9 delivery are lipoids, viruses, nanoparticles, bacteria, gene guns, electroporation, and nanostrips [15]. Here we use lentivirus as a tool to deliver CRISPR/Cas9 to the host cells. HCMV can infect multiple cell types including epithelial and endothelial cells, fibroblasts, cells within the monocytic lineage, smooth muscle cells, neurons, stromal cells, and hepatocytes, among others. Here we described the method to make epithelial cells-ARPE19 stably expressing CRISPR/Cas9 pooled human gene screen to discover host factors required for HCMV infection [2]. The approach should be applicable to any cell type including primary cells that are difficult to culture or transfect.

2

Materials

2.1 Common Materials

1. Cells: ARPE19 epithelial cells (ATCC) are grown in DMEMF12 supplemented with 10% fetal bovine serum (FBS) and 1% penicillin–streptomycin. Human embryonic lung fibroblasts (HEL; Coriell Repository) and HEK293T cells (ATCC) are grown in DMEM supplemented with 10% FBS and 1% penicillin–streptomycin. H1HeLa cells (ATCC) are grown in minimal essential medium supplemented with 10% fetal calf serum, 1% penicillin–streptomycin, and 2 mM L-glutamine. 2. Virus: HCMV TB40E-GFP virus was generated from a bacterial artificial chromosome (BAC), which was a kind gift of Eain Murphy, Cleveland Clinic, Cleveland, OH (now at SUNYUpstate). This version of TB40E-GFP expresses GFP as a transgene under the control of an SV40 origin/promoter cassette. 3. Plasmids: sgRNA library, part A or B [16] (Addgene), psPAX2 (Addgene); pMD2.G (Addgene).

2.2 Chemical Reagents and Supplies

1. Chemical reagents: TransIT-293 (Mirus), Opti-MeM, hygromycin B, anti-Flag (Sigma, F1804), puromycin, crystal violet, DMSO, 0.25% trypsin–EDTA, PBS, SeaPlaque (Lonza), sodium bicarbonate, methanol, Giemsa (sigma), sorbitol, DNeasy Blood & Tissue Kit (QIAGEN), ethanol, nucleasefree water (Ambion), forward primer and reverse primer are synthesized from IDT, betaine, Herculase II fusion DNA

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polymerase (Agilent), dNTP, agarose, PCR purification kit, sodium acetate. 2. Supplies: 0.45 μm filter, cryovial, 6-well plates, 24-well plates, 10 cm plates, 15 cm plates, micropipettes, pipette tips, serological pipettes (2 ml, 5 ml, 10 ml, and 25 ml), centrifuge tubes polycarbonate. 2.3 Significant Equipment

Incubator, water bath, sonicator, microscope (Zeiss), NanoDrop, Pippin Prep, deep sequencer (Ion Proton), ViiA7 qPCR machine (Applied Biosystems), PCR machine, ultracentrifuge, SW28 rotor.

2.4 Deep Sequencing Reagents (Life Technologies)

5
End repair buffer, End repair Enzyme, 10
ligase buffer, Ion P1 Adapter, Ion Xpress Barcode, dNTP Mix, DNA ligase, Nick repair polymerase, Platinum PCR Super HF, library Amp Primer Mix, Control library, Ion Library TaqMan qPCR Mix, Ion Library TaqMan Quantitation Assay.

3

Methods

3.1 Production for Cas9 Lentivirus

1. Streptococcus pyogenes Cas9 was subcloned from lentiCas9-Blast (Addgene) into the pHAGE-hygromycin lentiviral vector [17]. 2. Day 0: Plate 6–8
106 HEK293T cells in 8 ml D10 (DMEM with 10% FBS and supplements) onto one 10 cm dish. 3. Day 1: Transfect cells: Warm transfection reagent to room temperature (TransIT-293, Mirus). 4. Dispense 1.5 ml Opti-Medium into sterile 15 ml Falcon tube. Add DNA to Opti-Medium: 6 μg human GeCKO v.2 sgRNA library A or library B (Addgene). 4 μg packaging plasmid (e.g., psPAX2, Addgene). 2 μg envelope plasmid (e.g., pMD2.G expressing VSV-G, Addgene) 5. Mix by gently pipetting up and down with 1 ml pipette (see Note 1), then add 36 μl of TransIT-293. 6. Mix gently with 1 ml pipette, incubate at room temp for 15–30 min. 7. Dispense transfection mix onto the medium (8 ml) of one 10 cm dish of cells. 8. Day 2 (24 h after transfection): remove media, cover cells with 6–7 ml fresh growth medium (DMEM with 10% FBS and supplements).

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9. Day 3 (48 h after transfection): collect first batch of viruscontaining supernatant (store at 4  C), cover cells with 6–7 ml fresh growth medium. 10. Day 4 (72 h after transfection): collect second batch of virus containing supernatant (discard cells). 11. Pool the two batches of virus supernatant and filter through a 0.45 μm filter. Virus supernatant is stored at 4  C protected from light (or concentrate viral supernatants by ultracentrifugation in a centrifuge (Beckman) for 1.5 h at 20,000
g and store at 80  C). 3.2 Titering of Cas9 Lentivirus

1. Day 0: Add 0.3
106 HeLa H1 cells in fresh media to the number of wells needed in a 6-well plate. Duplicate or triplicate wells for each lentiviral construct and control should be used. Incubate 6 h at 37  C in a humidified incubator in an atmosphere of 5% CO2. 2. Day 1, Transduction: dilute virus 1:100 to 1:1000,000, and reserve one well for negative control without virus. After 6 h incubation with the diluted virus in a humidified incubator in an atmosphere of 5% CO2, supernatant is replaced with fresh medium. 3. 2 days after transduction, remove media from cells. Replace with 2 ml fresh medium containing 2.5 μg/ml puromycin (see Note 2) to each well. Gently swirl the plate to mix. Puromycin will kill untransduced cells over the course of 1–3 days (or later). Remove dead (non-adherent) cells after adding puromycin and replace with equal volume with fresh media; repeat after another 2–3 days. 4. Cells can be stained with crystal violet (step 6 below) when colonies (including small colonies) are large enough to be seen by eye, usually about 6–10 days after adding puromycin. 5. Remove media and wash once with PBS. 6. Add 1 ml 0.5% crystal violet (Sigma) solution per well, incubate at room temperature for 10–20 min. 7. Wash away crystal violet solution. 8. Count colonies per well.

3.3 Production of ARPE19-Cas9 Stable Cells

1. Day 0: Add 0.2
106 ARPE19 cells in fresh media to the number of wells in a 6-well plate (see Note 3). Incubate for 18–20 h at 37  C in a humidified incubator in an atmosphere of 5% CO2. 2. Day 1, Transduction: Cas9 lentivirus from 3.1 diluted in 1 ml infection medium (2% FBS in DMEM-F12) is used to transduce ARPE19 cells (MOI ¼ 2.0). Two wells are kept as control (no transduction).

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ARPE19-Cas9

ARPE19 mock

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Flag-Cas9

Actin

Fig. 1 Immunoblot of lysates from cells in ARPE19 mock and ARPE19-Cas9 cells. The levels of Cas9 protein expression are assessed by immunoblot analysis using flag antibody (sigma, 1:1000)

3. After a 6 h incubation, supernatant is replaced with fresh medium. 4. Two days postinfection, remove media containing lentiviral particles from wells and replace with fresh media to a volume of 2 ml to each well containing hygromycin B (200 μg/ml) to selected wells. 5. Examine viability every 2 days. Complete cell death after 3–5 days should be observed in the control well. 6. Culture for 10–14 days. Replace the media containing hygromycin B every 3 days until resistant colonies can be identified and expanded. 7. Cas9 expression (160KD) in ARPE19-Cas9 stable cells can then be confirmed by Western blot (Fig. 1). 3.4 Production for Lentiviral sgRNA-Library A and Lentiviral sgRNA-Library B Virus

GeCKOv2 sgRNA library [4, 16] is a complex lentiviral library that targets 19,052 human genes. The GeCKOv2 library contains six unique sgRNAs per gene in two half-libraries (A and B; Addgene). The library backbone is lentiGuide-Puro (Addgene). Libraries A and B each contain three unique sgRNAs per gene. 1. Library A and library B lentivirus stocks are produced in HEK293T cells using the same way as the lentivirus-Cas9 (see Subheading 3.1). 2. Titering of lentivirus-Library A and library B using the same way as the lentivirus-Cas9 (see Subheading 3.2).

3.5 Generation of ARPE19-Cas9-sgRNA Library A and ARPE19Cas9-sgRNA Library B Stable Cells

1. Day 0: Plate 4–6
106 ARPE19-Cas9 cells in 20 ml D10 (DMEM-F12 with 10% FBS and supplements) onto thirtytwo 15 cm dishes. Incubate for 18–20 h at 37  C in a humidified incubator in an atmosphere of 5% CO2.

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2. Day 1, Transduction: lentiviral particles of library A and library B from Subheading 3.4, step 1 are separately diluted in 3 ml infection medium (2% FBS in DMEM-F12, MOI ¼ 0.2) (see Note 4) to transduce to each dish. Fifteen dishes are transduced with library A and 15 dishes are transduced with library B. Two dishes are kept as control without transduction. 3. After a 6 h incubation, media containing lentiviral particles is replaced with fresh medium. 4. Two days postinfection, remove media and replace with fresh media to a volume of 20 ml to each dish containing puromycin (2.5 μg/ml) to selected cells. 5. Examine viability every 2 days to allow sgRNA-guided CRISPR-Cas9-mediated gene deletion within each pool. Watch for death of ARPE19-Cas9 cells (control) following the addition of puromycin. Complete cell death after 3–5 days is typically observed in the control dishes. The surviving cells in the library A and library B transduced cells represent the pool with sgRNA-guided CRISPR-Cas9-mediated gene deletions. 6. Replace the media containing puromycin every 3 days until resistant colonies can be identified and expanded. 7. After 1 or 2 weeks of expansion, trypsinize ARPE19-Cas9 cells transduced with the CRISPR gRNA library. 8. Pool drug selected ARPE19-Cas9 cells transduced with the CRISPR lentiviral library A or B and pellet at 1000
g for 5 min. 9. Resuspend the pellet of selected ARPE19-Cas9 CRISPR lentiviral library in 37  C DMEM-F12 + 10% FBS + 5 μg/ml puromycin and plate 2
106 cells per 15 cm dish. There should be many 15 cm dishes. 10. Incubate for 1 or 2 weeks at 37  C in a humidified incubator in an atmosphere of 5% CO2. 11. Trypsinize and pool all of selected ARPE19-Cas9 cells transduced with the CRISPR gRNA library. 12. Resuspend pellet in 50 ml of 37  C DMEM-F12 + 10% FBS and count cells. 13. At this point, freeze back the library at 1.2
107 cells per cryovial in 50% DMEM +40% FBS + 10% DMSO media or use these cells for screening (see Note 5). 3.6 TB40E-GFP Virus Amplification in Fibroblasts

1. Split HEL fibroblasts into two 15 cm plates: 5.0
106 cells/ plate. 2. Infect HEL fibroblasts with TB40E-GFP virus at an MOI ¼ 0.001.

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3. Observe GFP expression every day until all of the cells express GFP (It usually takes ~2 weeks). 4. After 2 weeks, harvest when monolayer looks considerably infected (most/all cells with extensive CPE). 5. Remove 18 ml of the 20 ml from each 15 cm dish and transfer to conical 50 ml centrifuge tubes, then scrape the remaining cells in each dish and pool the scraped cells and small amount medium into one 50 ml conical tube. 6. Sonicate the tube with scraped cells in minima volume of medium (see Note 6). 7. Spin tubes of scraped cells and medium supernatants for about 10 min at 3000
g to pellet debris. 8. Pool postcentrifugation supernatants from scraped sonicated cells and media supernatants into a large volume clean sterile container. 9. Aliquot and store in 80  C. 3.7 TB40E-GFP Virus Titer Is Determined in HEL Cells by Plaque Assay

1. Set up 24-well plates of HEL cells so they are 90% confluent on day of infection. 2. Make serial dilutions of virus in the 102 to 106 range in DMEM/5%FBS. 3. Aspirate media off the 24-well plate and add 1 ml PBS to each well to wash. Aspirate PBS. 4. Add 0.2 ml virus dilution to each well. 5. Incubate 2 h in incubator. Rock occasionally. 6. While virus is adsorbing, make overlay solutions: Melt 2% SeaPlaque (Lonza) low melting temp agarose and cool in a 42  C water bath. Make 2
DMEM containing FBS and sodium bicarbonate: For 25 ml: 21.5 ml 2
DMEM. 2.5 ml 7.5% sodium bicarbonate. 1 ml FBS. Place at 37  C until ready to use. When infection hour is almost over, mix equal amount of 2% SeaPlaque (at 42 ) with equal amount of 2
DMEM mixture (at 37 ) and place at 42 until ready to use. 7. After a 2 h incubation, remove the virus from each well and add 1.5 ml of overlay (at 42  C) solution to each well. 8. Let plates gel at room temp for about 30 min, then place in incubator. 9. Incubate for 2 weeks.

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10. Add 1 ml/well methanol directly to overlay. Incubate at room temperature 30 min. 11. Remove methanol and shake plates to remove overlays. 12. Rinse monolayers 2
with H2O and add 1 ml/well 20% Giemsa (sigma) for 10–20 min. 13. Remove stain and rinse with H2O. 14. Count plaques: # of plaques
dilution factor (used 0.2 ml virus so multiply by 5 for 1 ml)
well dilution (105, 106 etc). 3.8 TB40E-GFP Adaptation to Epithelial Cells (See Note 7)

1. To get passage 1 TB40E-GFP virus in ARPE19 cells, split ARPE19 cells into two 10 cm plates at 3
106 cells/plate. 2. Use TB40E-GFP virus produced in HEL cells to infect ARPE19 cells at an MOI ¼ 0.01. 3. Observe GFP expression until all the cells express GFP. This usually takes ~1 month (see Note 8). 4. Harvest TB40E-GFP virus produced in ARPE19 cells as epithelial virus Passage 1. 5. Virus titers is determined in HEL cells by the plaque assay (see Subheading 3.7). 6. To get passage 2 TB40E-GFP virus in ARPE19 cells, split ARPE19 cells into two 10 cm plates, each having 3
106 cells. 7. Using the TB40E-GFP epithelial passage 1 virus to infect ARPE19 cells at an MOI ¼ 0.01 to get passage 2 (this step usually takes 3 weeks). 8. Harvest and process TB40E-GFP passage 2 viruses produced in ARPE19 cells (~3 weeks) and titer as described for passage 1. 9. To get more virus, repeat steps 6–8 until get TB40E-GFP passage 6 viruses and titer as described in Subheading 3.7. 10. After PBS wash and 0.25% trypsin–EDTA treatment, split ARPE19 cells into five 10 cm plates, each have 3
106 cells to infect by TB40E-GFP passage 6 (MOI ¼ 0.01) to generate a larger passage 7 virus stock and titer as described in Subheading 3.7. 11. Split ARPE19 cells into ten 15 cm plates to infect by TB40EGFP passage 7 (MOI ¼ 0.01) to get a working stock of passage 8 virus. 12. Observe GFP expression every day until all of the cells express GFP (It usually takes 2–3 weeks). 13. Virus is harvested as described in Subheading 3.6 steps 5–8. 14. In order to concentrate passage 8 virus, put about 20–24 ml of virus into each ultracentrifuge tube. Then slowly underlay with 7 ml of 20% sorbitol cushion.

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15. After balance the tube, ultracentrifuge (SW28 rotor; Beckman) at 20,000
g for 1 h, at 4  C. 16. Aspirate supernatant (media and sorbitol) and add 0.5 ml of 0.1% FBS DMEM-F12 to each pellet. 17. Store virus plus medium at 4 conical tube.



C overnight in a 14 ml

18. Vortex in the next morning and aliquot into 0.4–0.5 ml portions. Aliquots are stocked at 80  C. 19. Virus is titered as described in Subheading 3.7. 3.9 Infection of ARPE19-Cas9-sgRNA Library A and ARPE19Cas9-sgRNA Library B Stable Cells with TB40E-GFP Virus

1. After PBS wash and 0.25% trypsin–EDTA treatment, split ARPE19-Cas9-sgRNA library A cells (0.3
106 cells for each well) into eleven 6-well plates; Split ARPE19-Cas9-sgRNA library B cells (0.3
106 cells for each well) into eleven 6-well plates (see Note 9); Split ARPE19-Cas9 cells (0.3
106 cells for each well) into three 6-well plates (as control). 2. Use TB40E-GFP passage 8 virus to infect library A, library B, and control (MOI ¼ 5.0). 3. Observe GFP every other day. 4. Change refresh medium every week (see Note 10) until see >95% of the cells in infected control plates (GFP positive, Fig. 2). It may take 2–3 weeks for this to happen.

3.10 Monitor for Cell Death (See Note 11)

For the infected control plates (ARPE19-Cas9), cells will be all float off and die eventually. For the library A and library B transduced cells, some of the genes deleted by guide RNA may be important for virus entry or infection, so some of the cells will be resistant to the virus infection. Therefore, a small percentage cells should still alive in library A and B transduced cells. 1. Wait until all the cells infected by TB40E-GFP have died or are floating on the control, ARPE19-Cas9 plates. 2. At that time, most cells (>95%) cells in the library A (Fig. 3) and library B plates will also be dead. Anticipate that 95% of the cells infected in the control well. It takes 2–3 weeks. 8. Wait until all the cells infected by TB40E-GFP have died or are floating on the control ARPE19-Cas9 plate. 9. Allow the surviving cells from library A and B to expand in numbers for 2 more weeks.

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Library A TB40E -GFP Day 60

phase

Fig. 3 ARPE19-Cas9 library A infected by TB40E-GFP at Day 60. TB40E-GFP infected library A at MOI of 5.0. Fluorescent and phase images are taken at Day 60 after infection

10. Wash 3
with PBS and incubate with 0.25% trypsin–EDTA for 5 min. 11. Harvested the surviving cells from all the 6-well plates of library A and B into one tube after centrifuge the cells at 1000
g for 5 min. 12. Remove the supernatant and store the cell pellet at 80  C. 3.12 Isolation of Genomic DNA Using the DNeasy Blood and Tissue Kit (QIAGEN)

1. Resuspended the cell pellet in 200 μl PBS. Add 20 μl proteinase K. 2. Add 200 μl Buffer AL. Mix thoroughly by vortexing, and incubate at 56  C for 10 min. 3. Add 200 μl ethanol (100%) to the sample and mix thoroughly by vortexing. 4. Pipet the mixture from above into the DNeasy Mini spin column placed in a 2 ml collection tube (provided). Centrifuge for 1 min at 6000
g. Discard flow-through and collection tube.

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5. Place the DNeasy Mini spin column in a new 2 ml collection tube (provided), add 500 μl Buffer AW2 and centrifuge for 3 min at 20,000
g to dry the DNeasy membrane. Discard flow-through and collection tube. 6. Place the DNeasy Mini spin column in a clean 1.5 ml microcentrifuge tube and pipet 100 μl nuclease-free water (Ambion) directly onto the DNeasy membrane. 7. Incubate at room temperature for 1 min, and then centrifuge for 1 min at 20,000
g to elute DNA. 3.13 sgRNA Sequences Are Amplified from Integrated Proviruses

1. Quantify DNA. Take 1 μl of the DNA from Subheading 3.12, step 7 to apply on the NanoDrop. 2. PCR. Forward primer. lentiGP-1_F: AATGGACTATCATATGCTTACCGTAACTTGAAAGTATT TCG. Reverse primer: lentiGP-3_R. ATGAATACTGCCATTTGTCTCAAGATCTAGTTACGC. PCR mix: DNA: 15 μl. Betaine: 24.0 μl. lentiGP-1_F: 2.5 μl. lentiGP-3_R: 2.5 μl. 5
Herculase II reaction buffer (Agilent): 20 μl. Herculase II fusion DNA polymerase (Agilent): 1.5 μl. DMSO: 2 μl. H2O: 27.5 μl. dNTP (10 mM): 5 μl. Total 100 μl. Place the tube in a thermal cycler (Applied Biosystems) and run the following program. Step 1: 95  C 2 min. Step 2: 95  C 20 s. Step 3: 58  C 20 s. Step 4: 72  C 3 min. Steps 2–4
36 cycles. Step 5: 72  C 3 min.

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Base Pairs

500 400 300 200 100 -

Fig. 4 2% Agarose gel checking the PCR product for amplifying the guide RNA fragments. The length is 220 bp. 100 bp loading marker was used

3. Gel checking the PCR product (Fig. 4). 2% agarose gel was made to check the PCR product for amplifying the guide RNA fragments. The length is 220 bp. 3.14 Barcode and Adapter Was Added to the PCR Fragment (See Note 13)

1. End repair. amplicon 100 ng: 79 μl. 5
End repair buffer (Life Technologies): 20 μl. End repair Enzyme (Life Technologies): 1 μl. Total 100 μl. Keep sample in room temperature for 20 min, then do PCR purify. The final amount is about 25 μl. 2. Barcoding prepared libraries. To prepare the mix: DNA: 25 μl. 10
ligase buffer (Life Technologies): 10 μl. Ion P1 Adapter (Life Technologies): 2 μl. Ion Xpress Barcode (Life Technologies): 2 μl. dNTP Mix (Life Technologies): 2 μl. Nuclease free water: 49 μl. DNA ligase (Life Technologies): 2 μl. Nick repair polymerase (Life Technologies): 8 μl. Total 100 μl.

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Then place the tube in a thermal cycler (Applied Biosystems) and run the following program: 25  C 15 min. 72  C 5 min 3. PCR purify (QIAGEN Clean Up) the barcode library. Add 5 volumes Buffer PB to 1 volume of the PCR reaction and mix. Add 10 μl 3 M sodium acetate, PH 5.0, and mix. Place QIAquick column in a provided 2 ml collection tube. To bind DNA, apply the sample to the QIAquick column and centrifuge for 30–60 s. Discard flow-through and place the QIAquick column back in the same tube. To wash, add 750 μl Buffer PE to the QIAquick column and centrifuge for 30–60 s. Discard flow-through and place the QIAquick column back in the same tube. Centrifuge the QIAquick column once more in the provided 2 ml collection tube for 1 min to remove residue wash buffer. Place each QIAquick column microcentrifuge tube.

in

a

clean

1.5

ml

To elute DNA, add 30 μl water and centrifuge the column for 1 min. 4. Quantify the DNA. Take 1 μl of the DNA from Subheading 3.14 step 3 to apply onto a NanoDrop to determine the quantity of DNA. 3.15 Size-Selection of the Libraries Using a Pippin Prep Instrument (See Note 13)

1. Thirty microliters of sample is loaded onto the Pippin Prep.

3.16 Deep Sequencing of Samples from Subheading 3.15 step 4

1. PCR to amplify the fragments. Prepare mix.

2. Fragment harvest is set for between 200 and 400 bp. 3. About 40 μl sample solution is used to do purify (QIAGEN clean up). 4. Finally fragment is eluted in 25 μl H2O.

Pippin DNA: 25 μl. Platinum PCR Super HF (Life Technologies): 100 μl. Library Amp Primer Mix (Life Technologies): 5 μl. Mix samples and split into 2 wells. Then place the tube in a thermal cycler (Applied Biosystems) and run the following program: Step 1: 95  C 5 min. Step 2: 95  C 15 s.

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Step 3: 58  C 15 s. Step 4: 70  C 1 min. Steps 2–4
5 cycles 2. Purify (QIAGEN Clean Up). The method is same as Subheading 3.14 step 3. Finally, DNA is eluted in 20 μl H2O. 3. qPCR, Measure the DNA amount by qPCR. First, prepare five sequential tenfold dilutions from the control library (~68 pM stock concentration). Second, prepare an initial 1:20 dilution of the sample library in Nuclease-free water in a nonstick microcentrifuge tube on ice. Third, prepare dilutions of the sample library that target a concentration within the serial dilutions of the control library. It is best to prepare three independent dilutions for qPCR. For a standard 20 μl qPCR reaction, prepare 5 μl of each library dilution per reaction. 4. Set up the PCR reactions. First, prepare a master mix of the following components on ice, as described in the following. Ion Library TaqMan qPCR Mix (Life Technologies), 2
10 μl. Ion Library TaqMan Quantitation Assay (Life Technologies), 20
1 μl. Nuclease-free water to 15 μl. Second, for each reaction, pipet 15 μl of the master mix into a well of the PCR plate. Third, add 5 μl of the diluted control or sample library to each appropriate well. Add 5 μl of nuclease-free water to the no-template control wells. Fourth, seal the plate, centrifuge the plate briefly to spin down the contents and eliminate air bubbles. Fifth, place the plate in a QPCR machine-ViiA™ 7 System (Applied Biosystems) and run the real-time PCR reactions using the following program: Step 1: 50  C 2 min. Step 2: 95  C 20 s. Step 3: 95  C 1 s. Step 4: 60  C 20 s. Steps 2–4
40 cycles

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5. Run samples on the Ion Torrent. Dilute sample to 100 pm. Then using 10 μl of 100 pm sample to run deep sequencing on the Ion Proton sequencer. 3.17

Data Analysis

For analyzing your library results please refer to the original publications [4, 6]. 1. The FASTQ files are trimmed using the Cutadapt (https:// cutadapt.readthedocs.io/en/stable/guide.html) program. 2. The processed FASTQ files are mapped to the sgRNA library key using the Bowtie2 program. 3. Read counts for each sgRNA are calculated using Samtools (http://samtools.sourceforge.net/) and custom Matlab scripts. 4. Selected candidate genes are chosen for further study if they had 20 reads per sgRNA across 3 independent sgRNAs.

4

Notes 1. Do not vortex. 2. The appropriate concentration of puromycin for each cell type is different. The minimum concentration of puromycin that causes complete cell death after 3–5 days should be used for that cell type. If the concentration for the desired cell type is unknown, a titration experiment must be performed. Typically, 2–10 mg/ml are sufficient to kill most untransduced mammalian cell types. 3. The growth rate of cells varies greatly. Adjust the number of cells plated to accommodate a confluency of 70% upon transduction. 4. This lower viral transduction dose is used to decrease the frequency of double or triple transduction events happening in the same cell. 5. Freeze down samples from early passage can prolong their use after thawing. Use low-passage cells where possible. 6. Sonicate the tube with scraped cells in minimal volume of medium can increase yield of free virus. 7. Since high passaged laboratory strains of HCMV grown in fibroblasts often lose or lack the PC on the mature virions, we decided to passage the TB40E-GFP virus in epithelial cell to attempt to avoid losing the PC. 8. Producing the first few passages of TB40E-GFP virus in epithelial cells usually takes a longer time than that observed for growing virus in fibroblasts.

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9. Using 6-well plates can reduce the chance of losing cells due to unexpected contamination which might happen during long term infection. 10. Replace half the volume of media with fresh D10 medium each week. 11. ARPE19 cells can be killed by HCMV, but it takes much longer than that seen in fibroblasts since virus replication in RPE cells is atypically slow, particularly when compared to HCMV replication in human fibroblasts. 12. Some of the cells (cell debris) in HCMV-infected control (ARPE19-Cas9 cells), library A or B transduced cells may remain attached to the plate. That is, although the cells are dead from the long-term HCMV infection they remain stuck as debris to the plates. Thus, one needs to wash and replate the cells to remove this dead cell debris. 13. Protocols for barcoding, adapter linking, and fragment size selection are sequencing platform specific. Conditions described here are for deep sequencing on an Ion Proton Sequencer. The use of different equipment will work fine for these types of experiments; however, one would have to follow those different platform specific guidelines.

Acknowledgments This work was supported by NIH Grants AI109001 (to T.F.K.). References 1. Bazzone LE, King M, MacKay CR, Kyawe PP, Meraner P, Lindstrom D, Rojas-Quintero J, Owen CA, Wang JP, Brass AL, Kurt-Jones EA, Finberg RW (2019) A disintegrin and metalloproteinase 9 domain (ADAM9) is a major susceptibility factor in the early stages of Encephalomyocarditis virus infection. mBio 10(1):e02734-18. https://doi.org/10.1128/ mBio.02734-18 2. E X, Meraner P, Lu P, Perreira JM, Aker AM, McDougall WM, Zhuge R, Chan GC, Gerstein RM, Caposio P, Yurochko AD, Brass AL, Kowalik TF (2019) OR14I1 is a receptor for the human cytomegalovirus pentameric complex and defines viral epithelial cell tropism. Proc Natl Acad Sci U S A 116 (14):7043–7052. https://doi.org/10.1073/ pnas.1814850116 3. McDougall WM, Perreira JM, Reynolds EC, Brass AL (2018) CRISPR genetic screens to discover host-virus interactions. Curr Opin

Virol 29:87–100. https://doi.org/10.1016/j. coviro.2018.03.007 4. Savidis G, McDougall WM, Meraner P, Perreira JM, Portmann JM, Trincucci G, John SP, Aker AM, Renzette N, Robbins DR, Guo Z, Green S, Kowalik TF, Brass AL (2016) Identification of Zika virus and dengue virus dependency factors using functional genomics. Cell Rep 16(1):232–246. https:// doi.org/10.1016/j.celrep.2016.06.028 5. Sobhy H (2017) A comparative review of viral entry and attachment during large and giant dsDNA virus infections. Arch Virol 162 (12):3567–3585. https://doi.org/10.1007/ s00705-017-3497-8 6. Doudna JA, Charpentier E (2014) Genome editing. The new frontier of genome engineering with CRISPR-Cas9. Science 346 (6213):1258096. https://doi.org/10.1126/ science.1258096

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7. Hsu PD, Lander ES, Zhang F (2014) Development and applications of CRISPR-Cas9 for genome engineering. Cell 157(6):1262–1278. https://doi.org/10.1016/j.cell.2014.05.010 8. Barrangou R, Fremaux C, Deveau H, Richards M, Boyaval P, Moineau S, Romero DA, Horvath P (2007) CRISPR provides acquired resistance against viruses in prokaryotes. Science 315(5819):1709–1712. https://doi.org/10.1126/science.1138140 9. Bolotin A, Quinquis B, Sorokin A, Ehrlich SD (2005) Clustered regularly interspaced short palindrome repeats (CRISPRs) have spacers of extrachromosomal origin. Microbiology 151 (Pt 8):2551–2561. https://doi.org/10. 1099/mic.0.28048-0 10. Pourcel C, Salvignol G, Vergnaud G (2005) CRISPR elements in Yersinia pestis acquire new repeats by preferential uptake of bacteriophage DNA, and provide additional tools for evolutionary studies. Microbiology 151 (Pt 3):653–663. https://doi.org/10.1099/ mic.0.27437-0 11. Zarei A, Razban V, Hosseini SE, Tabei SMB (2019) Creating cell and animal models of human disease by genome editing using CRISPR/Cas9. J Gene Med 21(4):e3082. https://doi.org/10.1002/jgm.3082 12. Zhang ZT, Jimenez-Bonilla P, Seo SO, Lu T, Jin YS, Blaschek HP, Wang Y (2018) Bacterial genome editing with CRISPR-Cas9: taking Clostridium beijerinckii as an example.

Methods Mol Biol 1772:297–325. https:// doi.org/10.1007/978-1-4939-7795-6_17 13. Chen YC, Sheng J, Trang P, Liu F (2018) Potential application of the CRISPR/Cas9 system against herpesvirus infections. Viruses 10 (6). https://doi.org/10.3390/v10060291 14. Wang D, Wang XW, Peng XC, Xiang Y, Song SB, Wang YY, Chen L, Xin VW, Lyu YN, Ji J, Ma ZW, Li CB, Xin HW (2018) CRISPR/ Cas9 genome editing technology significantly accelerated herpes simplex virus research. Cancer Gene Ther 25(5-6):93–105. https://doi. org/10.1038/s41417-018-0016-3 15. Liu J, Chang J, Jiang Y, Meng X, Sun T, Mao L, Xu Q, Wang M (2019) Fast and efficient CRISPR/Cas9 genome editing in vivo enabled by bioreducible lipid and messenger RNA nanoparticles. Adv Mater:e1902575. https://doi.org/10.1002/adma.201902575 16. Shalem O, Sanjana NE, Hartenian E, Shi X, Scott DA, Mikkelson T, Heckl D, Ebert BL, Root DE, Doench JG, Zhang F (2014) Genome-scale CRISPR-Cas9 knockout screening in human cells. Science 343(6166):84–87. https://doi.org/10.1126/science.1247005 17. Murphy GJ, Mostoslavsky G, Kotton DN, Mulligan RC (2006) Exogenous control of mammalian gene expression via modulation of translational termination. Nat Med 12 (9):1093–1099. https://doi.org/10.1038/ nm1376

Chapter 14 Quantitative Electron Microscopy to Study HCMV Morphogenesis Clarissa Read, Paul Walther, and Jens von Einem Abstract The generation and release of mature virions from human cytomegalovirus (HCMV) infected cells is a multistep process, involving a profound reorganization of cellular structures and various stages of virus particle morphogenesis in different cellular compartments. Although the general steps of HCMV morphogenesis are known, it has become clear that the detailed molecular mechanisms are complex and dependent on various viral factors and cellular pathways. The lack of a full understanding of HCMV virion morphogenesis emphasizes the need of imaging techniques to visualize the different stages of virion assembly, such as electron microscopy. Here, we describe various electron microscopy techniques and the methodology of high-pressure freezing and freeze substitution for sample preparation to visualize HCMV morphogenesis. These methods are used in our laboratory in combination with a thorough quantification to characterize phenotypic alterations and to identify the function of viral and cellular proteins for the various morphogenesis stages. Key words Tomography, Electron microscopy, HCMV, Cytomegalovirus, Morphogenesis, Secondary envelopment, High-pressure freezing

1

Introduction Mature virus particles of human cytomegalovirus (HCMV), also known as the Human betaherpesvirus 5, are very complex. Nevertheless, their ultrastructure exhibits the typical herpesvirus virion organization [1], consisting of four major components: the core, capsid, tegument and envelope (Fig. 1). The electron dense core is formed by the highly condensed linear dsDNA, which is enclosed by the icosahedral capsid, together termed nucleocapsid. The tegument, a layer of cellular and viral proteins [2], connects the capsid to the viral envelope. The latter is a host cell derived lipid bilayer, which contains embedded viral glycoproteins. Productively infected cells generate, besides infectious virions, noninfectious enveloped particles (NIEP), which are distinguished from virions

Andrew D. Yurochko (ed.), Human Cytomegaloviruses: Methods and Protocols, Methods in Molecular Biology, vol. 2244, https://doi.org/10.1007/978-1-0716-1111-1_14, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Fig. 1 Ultrastructural appearance of (a) nuclear, (b) and (c) cytoplasmic, and (d) extracellular HCMV particles after high-pressure freezing, freeze substitution, and Epoxy resin embedding. (c) Virus particles in the cytoplasm of infected cells are categorized into 1. free, 2. budding, and 3. enveloped capsids. The appearance of budding capsids ranges from capsids in association with curved membranes (a) and capsids that have already underwent various degrees of budding into the vesicle (b–e) to capsids that seem to be completely

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by the lack of the viral genome (core), and enveloped viral protein aggregates, named dense bodies (DB) (Fig. 1b, d). Infection of cells with HCMV induces profound changes of the cell morphology, including an enlargement of the nucleus and the cytoplasm, the formation of a kidney-shaped nucleus and the reorganization of cellular membranes (e.g., Golgi and endosomal membranes) to form the cytoplasmic viral assembly complex (vAC). These large morphological changes reflect viral hijacking of cellular processes in order to generate an optimal environment for viral replication and assembly. Our current knowledge about herpesvirus morphogenesis (reviewed in [3, 4]) and HCMV morphogenesis in particular [5– 9] has been obtained to a large extent through transmission electron microscopy (TEM) studies of infected cells. Unlike light microscopy, TEM enables visualization of virus-induced alterations of the host cell and of viral structures in their cellular context at a nanometer resolution and thus to differentiate various virion maturation stages. Despite the complexity of virus-host cell interactions, ultimately leading to the generation of infectious virions, HCMV morphogenesis can be divided into two major phases: (1) nuclear assembly of nucleocapsids and (2) their cytoplasmic maturation into infectious virions at the vAC. While most enveloped viruses depend on a single envelopment step, HCMV, like all herpesviruses, has evolved two envelopment events: (1) primary envelopment at the inner nuclear membrane in order to mediate transit of capsids from the nucleus into the cytoplasm and (2) secondary envelopment at the vAC, through which capsids acquire their final envelope including viral glycoproteins and glycoprotein complexes and become infectious. Hence, examination of the various budding events at host cell membranes is important for elucidating herpesvirus morphogenesis and the underlying molecular mechanisms. A better understanding of these processes not only expands our knowledge about virus biology but also helps to identify new targets for future development of antiviral agents against HCMV infections. Below, we will briefly summarize the major nuclear and cytoplasmic stages of HCMV morphogenesis with a special emphasis on ultrastructural characteristics. 1.1 Nuclear Stages of HCMV Morphogenesis

Virion morphogenesis begins with the assembly of capsids in the nucleus. Capsids assemble from capsid subunits in a self-assembly mechanism around a protein scaffold [10]. This forms unstable, spherical procapsids, which undergo spontaneous angularization, resulting in an icosahedral capsid shell. In the next step, the viral


Fig. 1 (continued) surrounded with the envelope (f–h) but whose bud neck has not yet underwent scission (f) or whose tegument is not yet homogeneous (g and h, asterisk). NIEP noninfectious enveloped particle, DB dense body; scale bars 100 nm

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genome is packaged into the capsid shell. Prior genome packaging, the internal scaffold needs to be removed from the capsid cavity by proteolytic cleavage [11]. Three different icosahedral capsid forms, designated as A, B, and C capsids, can be distinguished in infected nuclei by TEM (Fig. 1a). B capsids contain the scaffold that appears as a ring structure within the capsid. Successful genome packaging leads to C capsids, which are characterized by their electron dense core. The failure of genome packaging or premature scaffold disassembly results in formation of A capsids, which are devoid of the scaffold and the viral genome (reviewed in [12, 13]). Capsids need to translocate from the nucleus into the cytoplasm in order to continue tegumentation and virion maturation. The most accepted model for nuclear egress is the envelopment– deenvelopment model (reviewed in [14]). Nuclear egress of capsids starts by their primary envelopment at the inner nuclear membrane (INM), resulting in primary enveloped capsids in the perinuclear space (PNS). Fusion of the primary envelope with the outer nuclear membrane than releases nonenveloped capsids into the cytoplasm. Primary envelopment at the INM is mainly orchestrated by the viral nuclear egress complex (NEC), consisting of pUL50 and pUL53. The NEC induces a local disruption of the nuclear lamina covering the INM through the recruitment of cellular and viral kinases [15] and thereby facilitating access of nuclear capsids to the INM to initiate primary envelopment [16, 17]. Cytomegaloviruses appear to have evolved an additional mechanism for nuclear capsid egress by inducing infoldings of the INM into the nucleoplasm. These infoldings greatly enlarge the surface of the INM and seem to be free of nuclear lamina [18]. By 3D electron microscopy it was recently shown that infoldings of the INM form an unexpectedly complex intranuclear membrane network in HCMV infected cells [19]. Budding of capsids at infolded membranes, visualized by electron microscopy, indicates that they indeed serve as sites for primary envelopment. As shown by electron microscopy, primary envelopment of nuclear capsids at the INM involves membrane curvature [14, 19]. The structural mechanism of this membrane curvature process has been understood recently. High resolution cryo-electron microscopy revealed the formation of a lattice of hexagonal building blocks of herpesvirus NEC heterodimers at the nucleoplasmic site of the INM, which initiates negative curvature of the INM into the PNS and the formation of perinuclear vesicles [20–25]. This is further supported by the observation of an electron dense layer in electron microscopy studies of HCMV infected cells, likely representing the NEC lattice, at the nucleoplasmic site of negatively curved portions of the INM and infoldings, eventually forming primary envelopes [19].

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Upon HCMV infection, the vAC forms in the cytoplasm at the indentation of a kidney shaped nucleus. The vAC consists of intertwined membranes of different origin, which are arranged in a cylindrical fashion around the microtubule organizing center (MTOC) [26–31]. The peripheral part of the vAC consists of Golgi-derived membranes, whereas the central part is composed of membranes, which are positive for markers of early and late endosomes [29]. After translocation of capsids from the nucleus to the cytoplasm, they are transported along microtubules toward the vAC [32], where they undergo further tegumentation and secondary envelopment. So far, the vAC is the only cytoplasmic site where secondary envelopment of capsids has been observed [6]. During tegumentation at least 20 viral and an unknown number of cellular proteins accumulate around the capsid, forming the tegument [1, 2]. There is increasing evidence that tegumentation is an organized process, which is regulated by an intricate interplay of protein-protein interactions. This is also reflected in the organization of the tegument, which is arranged in two structurally distinct layers: an obviously ordered and densely packed inner tegument (or capsid associated tegument) and a pleomorphic outer tegument [33]. Because of their bridging function, connecting capsid and envelope, inner and outer tegument proteins play a crucial role as structural components of the virion and as such also for virion morphogenesis [34, 35]. Our current understanding of the secondary envelopment process is deduced from ultrastructural examinations of virus infected cells. Secondary envelopment is initiated when partially tegumented capsids establish contact with the cytosolic face of a cellular vesicle membrane at the vAC. Subsequently, a membrane wrapping process around the capsid is induced, leading eventually to the formation of a bud neck [36] (Fig. 1c). Secondary envelopment is completed by a membrane scission event that closes the viral envelope and separates the virion from the vesicle membrane. The now mature virion is transported within the vesicle to the cell surface where vesicle fusion with the plasma membrane releases the virion from the cell. Although the sequence of morphogenetic steps during secondary envelopment and egress are mainly clarified, researchers have not yet reached a sufficient understanding of the mechanisms underlying tegumentation and secondary envelopment. Discovery of the mechanisms includes an understanding of the function of individual proteins and protein motifs and their ability to form functional complexes during HCMV morphogenesis. This knowledge is required as a basis for the development of new antiviral strategies and is thus of great interest in recent HCMV research [35–43]. A prerequisite for clarification of the cytoplasmic stages of HCMV virion morphogenesis by electron microscopy is a

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clear visualization of cellular and viral structures, especially of the viral envelope and the tegument, which depends on the methodology of sample preparation for electron microscopy. 1.3 Optimized Electron Microscopy Sample Preparation and Imaging Techniques

2 2.1

Most published work about HCMV morphogenesis and mutant virus phenotypes by TEM analyses is based on chemically fixed samples. While conventional chemical fixation with dehydration of the sample by increasing alcohol concentrations preserves structures in many cases sufficiently, improvements, such as the here described sample preparation method of high-pressure freezing (HPF) and freeze substitution (FS), provide superior structural integrity and better contrast of membrane profiles [44–46]. In contrast to chemical fixation, HPF is a very rapid sample immobilization process, which takes only milliseconds, too short for osmotic effects to occur [47]. During freezing, the formation of ice crystals is minimized by application of high pressure. HPF-FS turned out to be particularly suitable for detailed characterization of the stages of HCMV secondary envelopment, by both, 2D and 3D electron microscopy (reviewed in [6, 48, 49]). The clear visualization of tegument densities and membranes in HPF-FS samples is a major improvement for characterizing phenotypes of mutant viruses and for their quantitative analysis [36]. The high quality of the images obtained by this method is also suitable for an automated quantification of structures by deep learning approaches, which is currently a fast developing field [50]. Furthermore, the excellent sample preservation is a prerequisite for more detailed 3D electron microscopy studies. Several 3D electron microscopy methods, including serial sectioning transmission electron microscopy (TEM), scanning transmission electron microscopy (STEM) tomography, and focused ion beam (FIB) -scanning electron microscopy (SEM) tomography have evolved in the last decades to study virus infections (reviewed in [49, 51, 52]). These methods use different principles for 3D data acquisition and differ in their spatial and lateral resolution, their maximum covered sample volume and the required hands-on time. Detailed protocols for serial sectioning TEM [48], FIB-SEM tomography and STEM tomography [49] have been published by our group previously. This chapter details the methodologies of HPF and FS, classical TEM imaging, and STEM tomography in combination with a quantitative analysis focusing on the examination of cytomegalovirus infection and stages of virus morphogenesis.

Materials Cell Culture

1. Cell lines: Human fibroblasts (see Note 1). 2. Cell culture medium: Minimal essential medium (MEM) supplemented with 10% fetal bovine serum (FBS).

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3. Virus strain: TB40E-BAC4 virus and mutant viruses thereof, reconstituted from DNA of respective bacmids (see Note 1). 4. Cell culture plate: A chambered coverslip with 8 wells for cell culture and light microscopy (μ-Slide, ibidi) (see Note 2). 5. Sapphire disks: Diameter of 3 mm and thickness of 50 μm (Engineering Office M. Wohlwend GmbH), coated with a 10–20 nm thick carbon layer, engraved with an asymmetric mark, for example, number “2,” glow discharged and UV-irradiated (see Note 2). 6. Tweezers. 7. Carbon coating device: BAF300 (BAL-TEC AG, Lichtenstein) freeze-etching device for carbon coating of sapphire disks and grids with semithin sections for STEM tomography (see Note 2). 8. Oven (120  C). 9. Electric glow discharger: Pelco EasiGlow (Ted Pella, Inc.) for glow discharging of sapphire disks before seeding of cells, grids for section mounting and STEM tomography grids prior to application of gold fiducials. Furthermore, this device is used for carbon coating of Formvar coated single slot grids (see Note 2). 10. UV light, for example, in sterile bench. 2.2 Electron Microscopy Sample Preparation

1. High-pressure freezer: Wohlwend Compact 01 high-pressure freezing device (Engineering Office M. Wohlwend GmbH) (see Note 2).

2.2.1 High Pressure Freezing

2. Specimen carriers: Aluminum specimen carrier Type A (diameter 3 mm, cavity 0.1/0.2 mm) and aluminum specimen carrier Type B (diameter 3 mm, cavity 0.3 mm) for high-pressure freezer Compact 01 (Engineering Office M. Wohlwend GmbH). 3. Liquid nitrogen. 4. 1-hexadecene (see Note 3). 5. Storage vessels (BEEM® capsules) for sapphire disks (see Note 2). 6. Insulated containers for storage and transfer of high-pressure frozen samples.

2.2.2 Freeze Substitution and Embedding

1. Freeze substitution medium: acetone with 0.2% osmium tetroxide, 0.1% uranyl acetate, and 5% of water for good membrane contrast [45]. Prepare fresh at day of use. For preparation of 10 ml freeze substitution medium, dissolve 10 mg uranyl acetate in 500 μl aqueous 4% osmium tetroxide solution aided by sonication. Finally, add 9.5 ml acetone and mix well.

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2. Freeze-substitution device: EM AFS2 (Leica) freeze substitution device. Other devices, such as self-built devices or easy-toimprovise freeze-substitution devices may work as well [53]. 3. Epoxy embedding medium (Sigma-Aldrich GmbH). 4. Nadic methyl anhydride (NMA, hardener for Epoxy resin) (Sigma-Aldrich GmbH). 5. Dodecenylsuccinic anhydride (DDSA, hardener for Epoxy resin) (Sigma-Aldrich GmbH). 6. Epoxy embedding medium accelerator (SigmaAldrich GmbH). Final Epoxy resin: Mix 60% (w/w) Solution A (155 g Epoxy embedding medium and 250 g DDSA (store at 4  C)) and 40% (w/w) Solution B (200 g Epoxy embedding medium and 178 g NMA (store at 4  C)) for at least 30 min. Add 1.5% (w/w) accelerator and mix again for at least 30 min (see Note 4). 7. Liquid nitrogen. 2.2.3 TEM

1. TEM grids: Single-slot copper grids (slot dimensions 2 mm  1 mm); Formvar-, carbon-coated and glow discharged (see Note 5). 2. Ultramicrotome: Ultramicrotome Ultracut (Leica Microsystems). The ultramicrotome is equipped with a diamond knife with a 45 angle (DiATOME) (see Note 2). 3. Fretsaw equipped with an extra fine blade. 4. Single-edge razor blade. 5. Inverted light microscope. 6. Cuspidal polystyrene foam stick (DiATOME). 7. Eyelash attached to the tip of a wooden pick. 8. Fine tweezers. 9. Filter paper. 10. Grid storage box.

2.2.4 STEM Tomography

1. TEM grids: Parallel bars (200 mesh or 300 mesh for smaller sections) to avoid shadowing of structures at high tilt angles. 2. Inverted light microscope. 3. Fretsaw equipped with an extra fine blade. 4. Single-edge razor blade. 5. Ultramicrotome: Ultramicrotome Ultracut (Leica Microsystems) equipped with a diamond knife with a 35 angle (DiATOME) (see Note 2). 6. Cuspidal polystyrene foam stick (DiATOME).

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7. Eyelash attached to the tip of a wooden pick. 8. Fine tweezers. 9. Filter paper. 10. Embryo dish. 11. Wooden pick to transfer sections into embryo dish. 12. Hot plate. 13. Poly-L-lysine: 10% poly-L-lysine in H2Odd. 14. Colloidal gold particles with a diameter of 25 or 15 nm (Aurion) as fiducial markers (see Note 6). 15. Electron beam evaporation device BAF300 (BAL-TEC AG, Liechtenstein) (see Note 2). 16. Plasma cleaner. 2.3 Image Acquisition and Analysis

1. Transmission electron microscope: JEM-1400 (Jeol) equipped with a TEM CCD camera (Veleta, Olympus).

2.3.1 TEM

3. Image processing software: We used Adobe® Photoshop® for image processing, however, free software such as Gimp and Image J can also be used.

2. Imaging software: iTEM 5.2.

4. Software for statistical analyses: GraphPad Prism or others (see Note 2). 2.3.2 STEM Tomography

1. Scanning transmission electron microscope: JEM-2100F (Jeol) with a field emission gun at 200 kV equipped with a Jeol high tilt holder and a Jeol bright field detector. 2. Tomogram acquisition software: EM-tools (TVIPS). 3. Data processing: IMOD software package for tomogram reconstruction and Avizo® Fire 6.3 (Visualization Science Group Inc.) for segmentation and data display (see Note 7). 4. Quantifications and measurements: IMOD and Etomo, which are part of the IMOD software package (see Note 7). 5. Software for statistical analyses: GraphPad Prism (see Note 2).

3 3.1

Methods Cell Culture

3.1.1 Preparation of Sapphire Disks

1. Sonicate the sapphire disks for 15 min each in 60% sulfuric acid, soap water, and twice in absolute ethanol to clean them. 2. Allow the cleaned disks to dry. 3. Carbon coat the cleaned disks with 10–20 nm carbon on one side by using a carbon coating device.

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4. Dry and bake coated disks in an oven for at least 8 h or overnight at 120  C to increase the carbon coat stability. 5. Engrave an asymmetric mark into the carbon coat, for example, the number “2,” by using a sharp tweezer or needle. This is necessary to identify the carbon-coated side of the disk. 6. Glow discharge sapphire disks for better hydrophilicity (see Note 8). 7. Expose the disks 30 min to UV light for sterilization. 3.1.2 Cell Culture and Infection

1. Add 100 μl cell culture medium into each well (see Note 9). 2. Carefully immerse 3–4 carbon coated, glow discharged and sterilized sapphire disks per well with the carbon coat facing up. The number “2” must be readable (see Note 10). 3. Add human fibroblasts (25,000 cells/well) in 100 μl cell culture medium into each well. Distribute the cells evenly within the well and on the sapphire disks by gently rocking the plate (see Note 11). 4. Let the cells adhere for at least 24 h until a confluency of at least 90% is achieved (see Note 12). 5. Infect cells at an infection rate of approx. 60% (MOI 1) by adding virus. 6. Incubate cells overnight at 37  C. 7. Replace virus-containing medium with fresh medium. 8. Incubate infected cells at 37  C until 120 h post infection (hpi) (see Note 13). 9. Examine the cell culture by light microscopy prior to further sample preparation (see Note 14).

3.2 Electron Microscopy Sample Preparation

1. Prepare the high-pressure freezer for sample preparation according to the manual (e.g., liquid nitrogen available, cool down) (see Note 15).

3.2.1 High Pressure Freezing

2. Label and cool down freezing vessels for sapphire disk storage. 3. Soak specimen carriers type A and B in 1-hexadecene. 4. Place one specimen carrier type B with the flat surface facing up into the high-pressure freezer sample holder. All of the following steps need to be performed quickly to ensure that the cells are frozen at their current stage and to avoid that the cells dry before freezing. Steps 5–7 take an experienced user 10–20 s. 5. Grip a sapphire disk with cells with a tweezer, shortly blot it on filter paper to remove most of the medium, dip the disk into 1-hexadecene and place it with cells facing up onto the specimen carrier type B.

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6. Cover the cells with a 1-hexadecene soaked specimen carrier type A with the 100 μm cavity facing down to enclose the cell monolayer, avoid trapping air in the sandwich. 7. Tightly close the HPF holder and conduct the freezing process. 8. After extraction of the sample holder from the high-pressure freezer, instantly immerse the tip of the HPF sample holder in liquid nitrogen and extract the frozen sapphire disk to store it in liquid nitrogen until further processing by freeze substitution. From now on always precool the tweezer in liquid nitrogen before touching the frozen sapphire disk (see Note 16). 3.2.2 Freeze Substitution and Embedding

1. Fill the tank of the substitution device with liquid nitrogen so that it is sufficient for a substitution process over 17 h (see Note 17). 2. Set the temperature of the chamber to constant

90  C.

3. Prepare the substitution medium as specified in Subheading 2.2.2, item 1. 4. Fill one 0.5 ml reaction tube per sapphire disk with 500 μl freeze substitution medium and precool to 90  C for at least 30 min in the device’s chamber. 5. Carefully but quickly transfer the sapphire disks from liquid nitrogen through the liquid nitrogen atmosphere into the precooled freeze substitution medium without letting them thaw (see Note 18). 6. Start the freeze substitution process (see Note 19). 7. After reaching room temperature, replace the freeze substitution medium with fresh acetone and incubate for 30 min at room temperature. 8. Wash sapphire disks two more times with acetone for 30 min to remove precipitates. Remove remaining specimen carriers from the tubes after the last washing step. 9. Replace acetone with a 1:3 Epoxy resin–acetone dilution and incubate the sapphire disks for at least 1 h at room temperature (see Note 20). 10. Replace the Epoxy resin–acetone dilution with a 3:1 Epoxy resin–acetone dilution and incubate for at least 3 h at room temperature. 11. Incubate the sapphire disks in pure Epoxy resin overnight at room temperature. 12. Add approximately 0.3 ml of freshly prepared Epoxy resin into a 0.5 ml reaction tube. Pulse-centrifuge for 30 s to remove air bubbles.

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13. Align sapphire disk in the reaction tubes with the cells facing the tube’s lid. The number “2” on the disk is readable when looking through the opening of the tube (see Note 21). 14. Polymerize the Epoxy resin in the reaction tubes at 60  C for at least 48 h with open lids. 15. Release the Epoxy resin block from the tube by cooling the reaction tube with liquid nitrogen for approx. 30 s. Usually during this process the larger part of the resin cone with embedded cells and carbon layer separates automatically from the sapphire disk that usually stays attached to the smaller tip of the resin cone (see Note 22). 3.2.3 Ultrathin Sectioning for TEM

1. Determine the area of interest on the carbon coated surface of the large part of the resin block by using an inverse light microscope (see Note 23). 2. Trim the block so that you generate a trapezoid area of interest by using a razor blade (see Note 24). 3. Insert the trimmed block into the microtome holder. 4. Cut ultrathin sections (see Note 25). 5. Mount the ultrathin sections on carbon- and formvar-coated and glow discharged TEM grids (see Note 26). 6. Dry the grids at room temperature and store them in grid storage boxes until imaging.

3.2.4 Preparation of STEM Tomography Samples (See Note 27)

1. Check the quality of sample preparation by TEM of ultrathin sections cut from the same resin block before semithin sections for STEM tomography are prepared. 2. Coat TEM grids with parallel grid bars with poly-L-lysine for better attachment of thick sections by dipping the grids into 10% poly-L-lysine. 3. Dry grids afterward on filter paper. Grids can then be stored until use. 4. Freshly glow discharge grids for better hydrophilicity at the day of sample sectioning according to the device you are using. 5. Prepare your sample block as described in steps 1–3 of Subheading 3.2.3. 6. Cut semithin sections (see Note 28). 7. Transfer sections with a wooden pick into 80  C warm water (container on hot plate) to stretch sections. 8. Mount one section onto the center of one poly-L-lysine–coated and glow discharged TEM grid with an eyelash and let it dry. 9. Put the grid with the section on a glass slide onto the hot plate for 5 min to obtain a flat section.

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10. Glow discharge the grid with the section. 11. Immerse grid with section in a solution of 10% poly-L-lysine in water and allow to dry on filter paper. 12. Centrifuge the gold fiducial suspension at 2500  g for 1 min to pellet gold aggregates. 13. Horizontally immerse grid with section in the gold fiducial suspension for 15 s to let gold fiducials adhere to both sides of the section. 14. Allow to dry horizontally for 15 s and then dab with filter paper. 15. Coat grids with 5 nm carbon from both sides using an electron beam evaporation device to provide electric conductivity. 16. Clean samples by using a plasma cleaner for approx. 5 s before proceeding with imaging (see Note 29). 3.3 Selection of Infected Cells and Data Acquisition 3.3.1 TEM

1. Insert TEM grid into the TEM. 2. Look for infected cells. HCMV infected cells can be easily detected because they are usually the largest cells on the section, often have a kidney-shaped nucleus and contain viral capsids in the nucleus. 3. Choose infected cells at a late stage of HCMV infection (see Fig. 2). 4. Acquire an image of the entire cell (see Note 30). 5. Study of secondary envelopment: Acquire an overview image of the vAC. The vAC is often located at the recess of a reniform nucleus (Fig. 2). Proceed to step 7 (see Note 31). 6. Study of nuclear stages: acquire an overview image of the nucleus. 7. Study of secondary envelopment: Choose a magnification that allows you to clearly identify HCMV virions or capsids in their different stages of secondary envelopment (pixel size  ~7 nm). Record the entire area of the vAC with this magnification by acquiring several tiles (see Fig. 2). 8. Study of nuclear stages: Choose a magnification that allows you to clearly identify the different capsid types in the nucleus (pixel size  ~10 nm). 9. Acquire images of interesting details with the highest magnification possible, for example, of particles that are representative for the phenotype of the investigated virus.

3.3.2 STEM Tomography

1. Look for infected cells in the section (see Note 32). 2. Choose the region of interest and move it to the center of the image. Choose a magnification according to the size of the region of interest and field of view (see Note 33).

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Fig. 2 How to identify HCMV infected cells in a late stage of infection by TEM? (a and b) Late infected cells are usually the largest cells on the section and often show an enlarged reniform nucleus (n). They are characterized by numerous capsids in the nucleus at large DNA replication centers (rc). In many cases virus induced ER-derived structures (white arrowheads in b) are observed in the cytoplasm (c) around the vAC and the nucleus. The vAC is an area of the cytoplasm that is mostly devoid of mitochondria (m) and lacks the

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3. Bring the sample to eucentricity (see Note 34). 4. Set up the tilt series image collection process in the software. Select the bright-field detector for data acquisition. Take care that the sensitivity of the detector is set to a value that provides good brightness and contrast also at high tilt angles (see Note 35). 5. Acquire tilt-series images (see Note 36). 1. Select cells for quantification:

3.4 Image Analysis/ Quantification

(a) Choose cells that are in a similar stage of infection (see Note 13 and Fig. 2).

3.4.1 TEM

(b) Cells should contain a high number of virus particles for subsequent statistical analyses (see example below). 2. Perform the quantification after setting up categories. (a) Determine defined structural criteria that characterize the different stages of HCMV morphogenesis or morphological alterations of cellular and viral structures. (b) Ideally, analyze several cells from at least two independent experiments. Example for quantification of stages of secondary envelopment. l

Virus particles at the area of the vAC are categorize into three categories: (1) naked, (2) budding, and (3) enveloped capsids [36]. Naked particles are partially tegumented capsids that are either not associated with any membranes or in contact with membranes, which show no curvature into the direction of the capsid. Budding particles are those that are in close contact with membranes, which show curvature into the direction of the capsid or additionally wrap around the capsid. There are usually several differently advanced stages of budding (see examples in Fig. 1c). Enveloped particles meet the following three criteria: the particle has a regular circular shape; the particle has an envelope that surrounds the tegument; the tegument appears as a homogenous layer between the capsid and the envelope (see Note 37).


Fig. 2 (continued) virus-induced electron dense ER-structures. It is furthermore characterized by the presence of many virus particles, such as virions, dense bodies and NIEPs, and numerous vesicles and membrane structures of different sizes. Ribbons of cis-Golgi cisternae (g) mark the transition of the vAC toward the rest of the cytoplasm. (b) For quantitative analysis of secondary envelopment, as in our example, an overview image of the entire cell and the vAC area is acquired first. Then, four or more images that cover the entire area of the vAC (boxes 1–4) are acquired with higher resolution to allow for categorization of virus capsids (black arrowheads in higher magnifications). Scale bars in overview images 10 nm. Scale bar in higher magnification images 2 μm

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3.4.2 STEM Tomography

l

Numbers of naked, budding, and enveloped capsids are counted in cross sections of the vACs from at least ten different cells and are given as their ratios in percent (see Note 38).

l

The documentation of the quantification for each individual cell allows for statistical analyses that is performed by applying the Kruskal-Wallis test followed by a Dunn’s multiplecomparison test.

Tomogram reconstruction: l

Reconstruct the tomogram from tilt series images by weighted back projection with standard settings of the IMOD software package (see Note 39). Quantification of stages of HCMV morphogenesis:

3.5

Conclusions

l

This is done as described under Subheading 3.4.1 (see Note 40).

l

The size of structures or distances between structures can be determined from the tomogram by using the IMOD program 3dmod (example in [36]).

Visualization of viral and cellular structures during HCMV morphogenesis requires very high resolutions, which can only be achieved by electron microscopy. We have just begun to understand the underlying mechanisms that drive the generation of infectious virus progeny. Structural analysis of virus infected cells and mutant viruses have been fundamental in our understanding of which viral proteins or cellular pathways are involved in HCMV morphogenesis and at which particular morphogenesis stage those viral or cellular proteins function. In the last decades, we and others, have improved sample preparation techniques to achieve a conservation of structures very close to their natural state by minimizing preparation artefacts, such as shrinkage, deformation, or collapse of structures. One important technique, setting the standard for electron microscopy sample preparation, is the here described method of HPF in combination with FS and standard Epoxy resin embedding. The high sample quality in combination with standard TEM analysis leads to images with detailed structural content, which is optimal to study HCMV morphogenesis or the morphogenesis of other viruses. While HPF and FS are still rather specialized techniques requiring respective specialized devices, the standard equipment of electron microscopy facilities consists of ultramicrotomes and TEM. Access to a TEM even allows for acquisition of 3D data, for example, by serial sectioning TEM (for detailed protocol see reference [48]) or TEM tomography that uses the TEM similar to the here described technique for STEM tomography. Other

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methods for 3D data acquisition are STEM (this chapter and others [9, 54–58]) and FIB-SEM tomography [19, 59, 60]. Both techniques require, however, specialized electron microscopes. It is important to note that there is an even larger toolbox of various 3D electron microscopy techniques, which were not focus of this chapter [61–65]. All these techniques come with specific advantages and disadvantages which are reviewed in [49, 66]. The choice of which of the 3D techniques to use for your studies depends not only on the size of the structures of interest, the cellular context, and the required resolution but also on the unique technical characteristics of each method (i.e., requirement of special devices and expertise). The last and, in our opinion, the most important aspect for electron microscopic studies on HCMV morphogenesis is to combine electron microscopy with quantitative analyses of the phenotypic characteristics. Electron microscopy is a single cell analysis technique that usually visualizes only a small volume of an entire cell and therefore is subject to a high variability on the single cell level. Overall, this chapter describes a selection of protocols that are effective for preparation of samples for electron microscopy, acquisition of ultrastructural data in 2D and 3D and their quantitative analysis for the examination of HCMV morphogenesis.

4

Notes 1. Other cell types and virus backgrounds can be used, but may require adjustments to the protocol provided. 2. Material, software or devices from other suppliers likely also work, but may require some adjustments to the protocol. 3. 1-hexadecene does not serve as a cryoprotectant. Instead, it functions in proper transfer of pressure and heat during the freezing process by providing good contact between the specimen carriers and the sample. 4. Due to the high viscosity of reagents, it is more practical to weight them. 5. Conventional TEM grids may also work; however, grid bars may cover cells of interest, which thus cannot be analyzed by TEM. That is especially important when sections contain only a few infected cells. 6. Use gold fiducials with 25 nm diameter for tomography of entire cells and large areas of cells (width of tomogram: several μm). Use gold fiducials with 15 nm diameter for imaging of cellular details (width of tomogram: few μm). If the distribution of gold fiducials on the section surface is too dense, dilute the gold fiducial suspension.

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7. The IMOD software package for tomogram reconstruction [67, 68] can be downloaded as freeware from the IMOD website (http://bio3d.colorado.edu/imod). IMOD is also suitable for segmentation and data display instead of Avizo. 8. Glow discharging of sapphire disks needs to be done the same day as the seeding of the cells. 9. This protocol describes the procedure for 8-well chamber slides (μ-slide, ibidi). 10. Push the sapphire disks gently down to the bottom of the well so that they do not float. Take care that the sapphire disks are distributed in the center as well as in the edges of the well. Cover all sapphire disks with culture medium by gently rocking the plate. 11. It is helpful to prepare for each virus infection two wells of cells for follow-up fluorescence microscopy examinations that allow for determination of the infection rate or phenotypes. That is, we usually prepare two wells—one well with and the other without sapphire disks. 12. If desired, cells can be synchronized by, for example, serum starvation, prior to infection with virus. 13. Dependent on the question asked, shorter or longer infection times are possible. Keep in mind that investigating HCMV morphogenesis by electron microscopy requires sufficient numbers of virus particles. Accumulation of virus particles in the vAC and nucleus can be detected already at 96 hpi. 14. Check the quality and quantity of your virus infection and only continue with sample preparation when the experiment looks as expected. 15. This protocol is optimized for the Wohlwend Compact 01 high-pressure freezing device and may require modifications if other freezing devices are used. 16. The samples can be stored in liquid nitrogen for at least a year. 17. This protocol has been adapted from previous protocols [6, 36, 49] and is now used with the freeze substitution device Leica EM AFS2. The liquid nitrogen tank of this device is filled to 50% for a 17 h substitution. 18. Make sure the sapphire disks are submerged in the freezing medium. Sometimes, specimen carriers do not separate from the sapphire disks. Substitution also works when you put the disks into the freeze substitution medium together with the specimen carriers. They will separate early enough during substitution. 19. In this protocol, freeze substitution is performed from 90  C to 0  C within 17 h. Following temperature profile is used:

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90 to 80  C within 8 h, 80 to 60  C within 4 h, 60 to 35  C within 3 h, 35 to 0  C within 2 h, constant 0  C for 1 h, 0  C to room temperature within 1 h. The temperature progression profile has been adopted from [45]. Substitution is improved when the temperature is slowly increased especially at very low temperatures since the substitution process is decelerated at low temperatures. Therefore, the samples are kept the longest time between 90  C to 80  C. During the raise from 0  C to room temperature, the reaction of osmium tetroxide with the sample can take place and good contrast is achieved. Alternatively, there are shorter freeze substitution protocols, which generate similar result [53]. 20. Use freshly prepared Epoxy resin with a low viscosity especially for the final embedding, but also for the preembedding steps, to reach optimal infiltration of the samples. 21. Perpendicular orientation of the sapphire disks to the long axis of the tube/parallel alignment relative to the tube lid is crucial for ultrathin sectioning. Place a piece of polymerized Epoxy resin (tip of the Epoxy resin cone that is produced by embedding of intact sapphire disks) into the tip of the reaction tube to support pieces of broken sapphire disk. 22. After cooling the embedded sapphire disk to liquid nitrogen temperature, you can usually hear a cracking sound that indicates that the tip of the polymerized Epoxy resin with the sapphire disk cracked off, leaving the embedded cells and the carbon layer in the large part of the polymerized Epoxy resin. If the sapphire disk is still attached to the large part of the resin block, carefully remove the disk with a tweezer. 23. The number “2” is inversed in this view. It helps to define areas of high cell density as area of interest. Remember that infected cells are enlarged compared to noninfected cells. This helps defining the area of interest. We usually saw the block into two halves along the long axis before trimming. The entire procedure of trimming and ultrathin sectioning is described in detail in [48, 49]. 24. The size of the trapezoid area of interest should have maximum dimensions of 1 mm  0.75 mm. Larger trapezoids mean that more cells are contained in one ultrathin section; however, the sectioning process is more difficult. A more detailed description of the trimming and sectioning procedure is provided in reference [48, 49]. 25. For routine analysis of HCMV secondary envelopment, we usually section the samples parallel to the carbon coat. For normal TEM, it is recommended to cut sections with a thickness of 70–90 nm. You can also generate 3D data with the same sample preparation procedure as it has been described so far in

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this chapter. To generate 3D data you need to cut and collect serial sections instead of single ultrathin sections. A detailed serial sectioning protocol is published in [48]. 26. For analysis of secondary envelopment, it is recommended to collect the sections at a height between 1.5 and 2.5 μm measured from the carbon coat. This guarantees that the vAC is sectioned through its center (often indicated by visibility of one or even both centrioles) where it exhibits its largest diameter and usually shows numerous virus particles. When the study also focuses on the structure of extracellular virions, sections at a height of 0.1–0.5 μm from the carbon coat should be collected. These sections usually contain large numbers of extracellular virus progeny that is trapped below the cell. 27. This protocol is an updated version of a previously published protocol describing STEM tomography [9, 49, 56]. 28. STEM tomography allows for imaging of sections with a thickness of up to 1 μm [55, 56]. We usually generate sections with a nominal thickness of 800 nm for practical reasons. However, tomography can also be performed in TEM mode (electron tomography). This allows for imaging of sections with a thickness of up to 500 nm. The advantage of TEM tomography over STEM tomography is that it can basically be performed in every electron microscopy lab. The only prerequisite is that the sample holder can be tilted. 29. It may not be necessary to plasma clean the sample as we have observed with a Jeol JEM2100F electron microscope. 30. It is recommended to keep track of which images were taken from which cells. This allows for characterization of the morphological phenotype of individual cells and assessment of the biological variability of the phenotype. 31. Secondary envelopment has been observed only in the area of the cytoplasmic vAC [6]. Keep in mind that the vAC can exhibit a different shape or can be less clearly defined depending on the virus strain, cell type or mutant virus used. 32. The STEM mode usually allows for imaging with high magnifications. Therefore, it is rather difficult to find infected cells on the section and to get a good overview of the cells only. Thus, use the TEM mode to acquire an overview of the sections and to select infected cells before switching to STEM mode. Furthermore, sections for tomography are usually considerably thicker than normal TEM sections. Thus, structures might not be easily recognizable. This complicates the identification of infected cells. However, nuclear capsids are usually well visible even in semithin sections and a good marker for infected cells.

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33. Keep in mind that tilting your section may cause shifting of the region of interest out of the field of view. You can account for this by choosing a slightly larger field of view, which equals a lower magnification. 34. The imaging software usually finds the eucentric height automatically. To set it manually, tilt the sample first to an angle of 5 . Move the region of interest back into the field of view by adjusting the z-value of the sample holder (height of the holder in the column). Repeat the procedure with tilt angles 10 , 30 , and 72 each. 35. It has been shown that the use of the bright-field image for tomogram acquisition is preferable because it shows better resolution in the depth of the section compared to the darkfield image [57, 69]. 36. We usually tilt from 72 to 72 in 1.5 increments on the Jeol JEM2100F. Attention: Such high tilt angles are not possible for every holder due to geometrical reasons. 37. In some cases, not the entire membrane profile is well visible (dependent on the cutting angle relative to the membrane). In these cases, the shape of the particle together with the tegument appearance is sufficient for making a decision. 38. Cells containing less than 10 particles in the vAC cross section should be included in the quantification with care. Due to their low capsid number, they are often not representative regarding the ratios of naked, budding and enveloped capsids. It must be mentioned if cells are excluded from quantification. 39. The simultaneous iterative reconstruction technique (SIRT) can be used if better contrast is needed. 40. Quantification of secondary envelopment, as described in Subheading 3.4.1. can also be performed with STEM tomography data. However, selection of infected cells is more difficult in the STEM mode and the volume that is recorded usually covers only a small part of the area of the vAC. Nevertheless, the volume imaged by STEM tomography may contain higher numbers of virus particles due to the semithick section than would be visible in the same area of an ultrathin section. References 1. Mocarski E, Shenk T, Pass R (2006) Cytomegaloviruses. In: Knipe DM, Howley PM, Griffin DE et al (eds) Fields virology, 5th edn. Lippincott Williams & Wilkins, Philadelphia, PA, pp 2701–2771 2. Varnum SM, Streblow DN, Monroe ME et al (2004) Identification of proteins in human cytomegalovirus (HCMV) particles: the HCMV proteome. J Virol 78:10960–10966.

https://doi.org/10.1128/JVI.78.20.1096010966.2004 3. Mettenleiter TC (2004) Budding events in herpesvirus morphogenesis. Virus Res 106:167–180. https://doi.org/10.1016/j. virusres.2004.08.013 4. Mettenleiter TC (2002) Herpesvirus assembly and egress. J Virol 76:1537–1547. https://doi. org/10.1128/JVI.76.4.1537-1547.2002

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5. Severi B, Landini MP, Govoni E (1988) Human cytomegalovirus morphogenesis: an ultrastructural study of the late cytoplasmic phases. Arch Virol 98:51–64. https://doi. org/10.1007/bf01321005 6. Schauflinger M, Villinger C, Mertens T et al (2013) Analysis of human cytomegalovirus secondary envelopment by advanced electron microscopy: HCMV morphogenesis. Cell Microbiol 15:305–314. https://doi.org/10. 1111/cmi.12077 7. Seo J-Y, Britt WJ (2007) Cytoplasmic envelopment of human cytomegalovirus requires the postlocalization function of tegument protein pp28 within the assembly compartment. J Virol 81:6536–6547. https://doi.org/10. 1128/JVI.02852-06 8. Gibson W, Bogner E (2013) Morphogenesis of the cytomegalovirus Virion and subviral particles. In: Reddehase MJ (ed) Cytomegaloviruses: from molecular pathogenesis to intervention, 2nd edn. Caister Academic Press, Norfolk, pp 230–246 9. Walther P, Wang L, Ließem S, Frascaroli G (2010) Viral infection of cells in culture: approaches for electron microscopy. In: Mu¨ller-Reichert T (ed) Methods in cell biology. Academic, Cambridge, MA, pp 603–618. https://doi.org/10.1016/S0091-679X(10) 96025-1 10. Gibson W (1996) Structure and assembly of the virion. Intervirology 39:389–400. https://doi.org/10.1159/000150509 11. Yu X, Trang P, Shah S et al (2005) Dissecting human cytomegalovirus gene function and capsid maturation by ribozyme targeting and electron cryomicroscopy. Proc Natl Acad Sci U S A 102:7103–7108. https://doi.org/10. 1073/pnas.0408826102 12. Cardone G, Heymann JB, Cheng N et al (2012) Procapsid assembly, maturation, nuclear exit: dynamic steps in the production of infectious Herpesvirions. Adv Exp Med Biol 726:423–439. https://doi.org/10.1007/ 978-1-4614-0980-9_19 13. Tandon R, Mocarski ES, Conway JF (2015) The A, B, Cs of herpesvirus capsids. Viruses 7:899–914. https://doi.org/10.3390/ v7030899 14. Mettenleiter TC, Mu¨ller F, Granzow H, Klupp BG (2013) The way out: what we know and do not know about herpesvirus nuclear egress. Cell Microbiol 15:170–178. https://doi.org/ 10.1111/cmi.12044 15. Milbradt J, Auerochs S, Sticht H, Marschall M (2009) Cytomegaloviral proteins that associate with the nuclear lamina: components of a

postulated nuclear egress complex. J Gen Virol 90:579–590. https://doi.org/10.1099/ vir.0.005231-0 16. Muranyi W, Haas J, Wagner M et al (2002) Cytomegalovirus recruitment of cellular kinases to dissolve the nuclear lamina. Science 297:854–857. https://doi.org/10.1126/sci ence.1071506 17. Sharma M, Kamil JP, Coughlin M et al (2014) Human cytomegalovirus UL50 and UL53 recruit viral protein kinase UL97, not protein kinase C, for disruption of nuclear lamina and nuclear egress in infected cells. J Virol 88:249–262. https://doi.org/10.1128/JVI. 02358-13 18. Buser C, Walther P, Mertens T, Michel D (2007) Cytomegalovirus primary envelopment occurs at large Infoldings of the inner nuclear membrane. J Virol 81:3042–3048. https:// doi.org/10.1128/JVI.01564-06 19. Villinger C, Neusser G, Kranz C et al (2015) 3D analysis of HCMV induced-nuclear membrane structures by FIB/SEM tomography: insight into an unprecedented membrane morphology. Viruses 7:5686–5704. https://doi. org/10.3390/v7112900 20. Bigalke JM, Heuser T, Nicastro D, Heldwein EE (2014) Membrane deformation and scission by the HSV-1 nuclear egress complex. Nat Commun 5:4131. https://doi.org/10. 1038/ncomms5131 21. Bigalke JM, Heldwein EE (2015) Structural basis of membrane budding by the nuclear egress complex of herpesviruses. EMBO J 34 (23):2921–2936. https://doi.org/10.15252/ embj.201592359 22. Bigalke JM, Heldwein EE (2015) The great (nuclear) escape: new insights into the role of the nuclear egress complex of herpesviruses. J Virol 89:9150–9153. https://doi.org/10. 1128/JVI.02530-14 23. Hagen C, Dent KC, Zeev-Ben-Mordehai T et al (2015) Structural basis of vesicle formation at the inner nuclear membrane. Cell 163:1692–1701. https://doi.org/10.1016/j. cell.2015.11.029 24. Klupp BG, Granzow H, Fuchs W et al (2007) Vesicle formation from the nuclear membrane is induced by coexpression of two conserved herpesvirus proteins. Proc Natl Acad Sci U S A 104:7241–7246. https://doi.org/10. 1073/pnas.0701757104 25. Lorenz M, Vollmer B, Unsay JD et al (2015) A single herpesvirus protein can mediate vesicle formation in the nuclear envelope. J Biol Chem 290:6962–6974. https://doi.org/10.1074/ jbc.M114.627521

EM of HCMV Infection 26. Sanchez V, Greis KD, Sztul E, Britt WJ (2000) Accumulation of virion tegument and envelope proteins in a stable cytoplasmic compartment during human cytomegalovirus replication: characterization of a potential site of virus assembly. J Virol 74:975–986. https://doi. org/10.1128/JVI.74.2.975-986.2000 27. Sanchez V, Sztul E, Britt WJ (2000) Human cytomegalovirus pp28 (UL99) localizes to a cytoplasmic compartment which overlaps the endoplasmic reticulum-golgi-intermediate compartment. J Virol 74:3842–3851. https://doi.org/10.1128/jvi.74.8.38423851.2000 28. Homman-Loudiyi M, Hultenby K, Britt W, Soderberg-Naucler C (2003) Envelopment of human cytomegalovirus occurs by budding into Golgi-derived vacuole compartments positive for gB, Rab 3, trans-Golgi network 46, and mannosidase II. J Virol 77:3191–3203. https://doi.org/10.1128/ JVI.77.5.3191-3203.2003 29. Das S, Vasanji A, Pellett PE (2007) Threedimensional structure of the human cytomegalovirus cytoplasmic virion assembly complex includes a reoriented secretory apparatus. J Virol 81:11861–11869. https://doi.org/10. 1128/JVI.01077-07 30. Das S, Pellett PE (2011) Spatial relationships between markers for secretory and endosomal machinery in human cytomegalovirus-infected cells versus those in uninfected cells. J Virol 85:5864–5879. https://doi.org/10.1128/ JVI.00155-11 31. Alwine JC (2012) The human cytomegalovirus assembly compartment: a masterpiece of viral manipulation of cellular processes that facilitates assembly and egress. PLoS Pathog 8: e1002878. https://doi.org/10.1371/journal. ppat.1002878 32. Ogawa-Goto K, Tanaka K, Gibson W et al (2003) Microtubule network facilitates nuclear targeting of human cytomegalovirus capsid. J Virol 77:8541–8547. https://doi.org/10. 1128/JVI.77.15.8541-8547.2003 33. Yu X, Shah S, Lee M et al (2011) Biochemical and structural characterization of the capsidbound tegument proteins of human cytomegalovirus. J Struct Biol 174:451–460. https:// doi.org/10.1016/j.jsb.2011.03.006 34. Guo H, Shen S, Wang L, Deng H (2010) Role of tegument proteins in herpesvirus assembly and egress. Protein Cell 1:987–998. https:// doi.org/10.1007/s13238-010-0120-0 35. Smith RM, Kosuri S, Kerry JA (2014) Role of human cytomegalovirus tegument proteins in virion assembly. Viruses 6:582–605. https:// doi.org/10.3390/v6020582

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36. Read C, Schauflinger M, Nikolaenko D et al (2019) Regulation of human cytomegalovirus secondary envelopment by a C-terminal tetralysine motif in pUL71. J Virol 93:e02244-18. https://doi.org/10.1128/JVI.02244-18 37. Chevillotte M, Landwehr S, Linta L et al (2008) Major tegument protein pp65 of human cytomegalovirus is required for the incorporation of pUL69 and pUL97 into the virus particle and for viral growth in macrophages. J Virol 83:2480–2490. https://doi. org/10.1128/JVI.01818-08 38. Meissner CS, Suffner S, Schauflinger M et al (2011) A leucine zipper motif of a tegument protein triggers final envelopment of human cytomegalovirus. J Virol 86:3370–3382. https://doi.org/10.1128/JVI.06556-11 39. Schauflinger M, Fischer D, Schreiber A et al (2011) The tegument protein UL71 of human cytomegalovirus is involved in late envelopment and affects multivesicular bodies. J Virol 85:3821–3832. https://doi.org/10. 1128/JVI.01540-10 40. Ahlqvist J, Mocarski E (2011) Cytomegalovirus UL103 controls virion and dense body egress. J Virol 85:5125–5135. https://doi. org/10.1128/JVI.01682-10 41. Phillips SL, Bresnahan WA (2012) The human cytomegalovirus (HCMV) tegument protein UL94 is essential for secondary envelopment of HCMV virions. J Virol 86:2523–2532. https://doi.org/10.1128/JVI.06548-11 42. Kim Y-E, Oh SE, Kwon KM et al (2016) Involvement of the N-terminal DUB domain of human cytomegalovirus UL48 tegument protein in auto-ubiquitination, virion stability, virus entry. J Virol 90(6):3229–3242. https:// doi.org/10.1128/JVI.02766-15 43. Brock I, Kruger M, Mertens T, von Einem J (2013) Nuclear targeting of human cytomegalovirus large tegument protein pUL48 is essential for viral growth. J Virol 87:6005–6019. https://doi.org/10.1128/JVI.03558-12 44. Moor H (1987) Theory and practice of high pressure freezing. In: Steinbrecht RA, Zierold K (eds) Cryotechniques in biological electron microscopy. Springer, Berlin, pp 175–191. https://doi.org/10.1007/978-3-642-728150_8 45. Walther P, Ziegler A (2002) Freeze substitution of high-pressure frozen samples: the visibility of biological membranes is improved when the substitution medium contains water. J Microsc 208:3–10. https://doi.org/10. 1046/j.1365-2818.2002.01064.x 46. Buser C, Walther P (2008) Freezesubstitution: the addition of water to polar

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solvents enhances the retention of structure and acts at temperatures around 60 C. J Microsc 230:268–277. https://doi.org/10. 1111/j.1365-2818.2008.01984.x 47. Szczesny PJ, Walther P, Mu¨ller M (1996) Light damage in rod outer segments: the effects of fixation on ultrastructural alterations. Curr Eye Res 15:807–814. https://doi.org/10.3109/ 02713689609017621 48. Schauflinger M, Villinger C, Walther P (2013) Three-dimensional visualization of virusinfected cells by serial sectioning: an electron microscopic study using resin embedded cells. In: Bailer S, Lieber D (eds) Virus-host interactions. Methods in molecular biology (methods and protocols). Humana Press, Totowa, NJ, pp 227–237. https://doi.org/10.1007/978-162703-601-6_16 49. Villinger C, Schauflinger M, Gregorius H et al (2013) Three-dimensional imaging of adherent cells using FIB/SEM and STEM. In: Kuo J (ed) Methods in molecular biology (methods and protocols). Humana Press, Totowa, NJ, pp 617–638. https://doi.org/10.1007/978-162703-776-1_27 50. Shaga-Devan K, Walther P, von Einem J et al (2019) Detection of herpesvirus capsids in transmission electron microscopy images using transfer learning. Histochem Cell Biol 151:101–114. https://doi.org/10.1007/ s00418-018-1759-5 51. Romero-Brey I, Bartenschlager R (2015) Viral infection at high magnification: 3D electron microscopy methods to analyze the architecture of infected cells. Viruses 7:6316–6345. https://doi.org/10.3390/v7122940 52. Risco C, de Castro IF, Sanz-Sa´nchez L et al (2014) Three-dimensional imaging of viral infections. Annu Rev Virol 1:453–473. https://doi.org/10.1146/annurev-virology031413-085351 53. McDonald KL, Webb RI (2011) Freeze substitution in 3 hours or less. J Microsc 243:227–233. https://doi.org/10.1111/j. 1365-2818.2011.03526.x 54. Abdellatif MEA, Sinzger C, Walther P (2018) Investigating HCMV entry into host cells by STEM tomography. J Struct Biol 204:406–419. https://doi.org/10.1016/j. jsb.2018.10.007 55. Aoyama K, Takagi T, Hirase A, Miyazawa A (2008) STEM tomography for thick biological specimens. Ultramicroscopy 109:70–80. https://doi.org/10.1016/j.ultramic.2008.08. 005 56. Ho¨hn K, Sailer M, Wang L et al (2011) Preparation of cryofixed cells for improved 3D

ultrastructure with scanning transmission electron tomography. Histochem Cell Biol 135:1–9. https://doi.org/10.1007/s00418010-0765-z 57. Walther P, Bauer A, Wenske N et al (2018) STEM tomography of high-pressure frozen and freeze-substituted cells: a comparison of image stacks obtained at 200 kV or 300 kV. Histochem Cell Biol 150:545–556. https:// doi.org/10.1007/s00418-018-1727-0 58. Yakushevska AE, Lebbink MN, Geerts WJC et al (2007) STEM tomography in cell biology. J Struct Biol 159:381–391. https://doi.org/ 10.1016/j.jsb.2007.04.006 59. Villinger C, Gregorius H, Kranz C et al (2012) FIB/SEM tomography with TEM-like resolution for 3D imaging of high-pressure frozen cells. Histochem Cell Biol 138:549–556. https://doi.org/10.1007/s00418-012-10206 60. Kizilyaprak C, Daraspe J, Humbel B m. (2014) Focused ion beam scanning electron microscopy in biology. J Microsc 254:109–114. https://doi.org/10.1111/jmi.12127 61. Burel A, Lavault M-T, Chevalier C et al (2018) A targeted 3D EM and correlative microscopy method using SEM array tomography. Development 145:dev160879. https://doi.org/10. 1242/dev.160879 62. Denk W, Horstmann H (2004) Serial blockface scanning electron microscopy to reconstruct three-dimensional tissue nanostructure. PLoS Biol 2:e329. https://doi.org/10.1371/ journal.pbio.0020329 63. Rosier DJD, Klug A (1968) Reconstruction of three dimensional structures from electron micrographs. Nature 217:130–134. https:// doi.org/10.1038/217130a0 64. Wacker I, Schroeder RR (2013) Array tomography. J Microsc 252:93–99. https://doi.org/ 10.1111/jmi.12087 65. White JG, Southgate E, Thomson JN, Brenner S (1986) The structure of the nervous system of the nematode Caenorhabditis elegans. Philos Trans R Soc Lond Ser B Biol Sci 314:1–340. https://doi.org/10.1098/rstb. 1986.0056 66. Peddie CJ, Collinson LM (2014) Exploring the third dimension: volume electron microscopy comes of age. Micron Oxf Engl 61:9–19. https://doi.org/10.1016/j.micron.2014.01. 009 67. Kremer JR, Mastronarde DN, McIntosh JR (1996) Computer visualization of threedimensional image data using IMOD. J Struct Biol 116:71–76. https://doi.org/10.1006/ jsbi.1996.0013

EM of HCMV Infection 68. Mastronarde DN (2008) Correction for non-perpendicularity of beam and tilt axis in tomographic reconstructions with the IMOD package. J Microsc 230:212–217. https://doi. org/10.1111/j.1365-2818.2008.01977.x

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Chapter 15 Detection of Cytomegalovirus Interleukin 10 (cmvIL-10) by Enzyme-Linked Immunosorbent Assay (ELISA) Vivian P. Young, Margarette C. Mariano, Lionel Faure, and Juliet V. Spencer Abstract Since its introduction in 1971, the enzyme-linked immunosorbent assay (ELISA) has revolutionized medicine by enabling detection of both antigens and antibodies in a variety of samples. We describe here a customized sandwich ELISA developed for the detection of Human Cytomegalovirus interleukin-10 (cmvIL-10). CmvIL-10 is a virally encoded cytokine and ortholog of human interleukin 10 (hIL-10). While cmvIL-10 and hIL-10 are similar in structure and function, overall amino acid sequence identity is only 27%, resulting in antigenically distinct proteins. The cmvIL-10 ELISA is specific and does not detect hIL-10. The assay is sensitive enough to detect cmvIL-10 in both culture supernatants and patient serum. The ability to quantify cmvIL-10 levels during HCMV infection could provide valuable information about immune evasion strategies and viral control of host signaling pathways. Key words CMV, cmvIL-10, Cytokine, Human cytomegalovirus, ELISA, Immunoassay

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Introduction The use of labeled antibodies to quantify hormone levels was so innovative and impactful that it was recognized with a Nobel Prize. In 1977, Rosalyn Yalow became the first American-born woman to win the Nobel Prize in Physiology or Medicine for the development of the radioimmunoassay (RIA). Yalow and her longtime collaborator, Solomon Berson, who regrettably passed away before the prize was awarded, developed a very sensitive in vitro method to measure concentrations of hormones in blood by using antibodies labeled with radioisotopes [1]. While the RIA quickly became an essential tool in medicine, the harmful effects of radiation exposure could not be ignored, and storage and disposal of radioactive waste became a serious challenge. In 1971, two groups independently reported an advancement on the method—the use of enzymelabeled antibodies to detect target proteins. Eva Engvall and Peter Perlmann used alkaline phosphatase-conjugated antibodies to

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detect rabbit immunoglobulin [2], while Anton Schuurs and Bauke van Weemen employed horseradish peroxidase–conjugated antibodies to detect human chorionic gonadotropin [3]. The use of enzyme–substrate reactions enabled detection of target proteins that was just as sensitive as the RIA, but without the dangers associated with radioisotope use. Today, ELISA, or enzyme-linked immunosorbent assay, is one of the most widely used performed diagnostic tests in the world, and commercial ELISA kits can be purchased for the detection of virtually any target protein. For novel target proteins, it is possible to develop an ELISA as long as specific antibodies are available. The assay described here uses goat polyclonal antibodies to detect cmvIL-10, an ortholog of human interleukin 10 (hIL-10), which is encoded by the UL111A gene of human cytomegalovirus (HCMV) [4, 5]. CmvIL-10 is secreted from infected cells late in infection and has potent immune suppressive properties such as inhibition of inflammatory cytokine synthesis and suppression of major histocompatibility complex (MHC) class I and II levels [6], which impair overall immune responses [7]. CmvIL-10 acts by binding to the cellular IL-10 receptor and the three-dimensional structure is highly conserved with that of hIL-10 [8]. However, actual amino acid sequence identity is quite low (27%) [4], resulting in proteins that are antigenically distinct [9, 10]. To detect cmvIL-10 in supernatants from virus infected cells, we developed a sandwich ELISA in which goat polyclonal antibodies serve as both the capture and detection reagents [11]. Together with recombinant purified cmvIL-10 as a protein standard, this assay enables quantification of the viral cytokine during infection.

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Materials Prepare all solutions using distilled water and store at room temperature, unless otherwise noted. Adhere to waste disposal regulations.

2.1

Buffers

1. 10 phosphate buffered saline (PBS): Place 800 mL of distilled water in a large beaker. Add 80 g NaCl, 2.0 g KCl, 14.4 g KH2PO4, and 2.4 g KH2PO4, stirring to dissolve. Adjust pH to 7.4. Add distilled water to a total volume of 1 L (see Note 1). 2. 1 PBS: Combine 50 mL 10 PBS with 450 mL distilled water. Sterile filter and store at 4  C (see Note 2). 3. Wash Buffer: PBS + 0.05% Tween 20. Combine 50 mL 10 PBS with 450 mL distilled water. Add 250 μL Tween 20. Mix well and store at 4  C.

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4. Reagent Diluent/Blocking Buffer: PBS + 1% Bovine Serum Albumin (BSA). Combine 50 mL 10 PBS with 450 mL distilled water. Add 5.0 g BSA and stir to dissolve. Sterile filter and store at 4  C (see Note 3). 5. Protein Standard Dilution Buffer: PBS + 0.1% BSA. Combine 1 mL Reagent Diluent with 9 mL sterile PBS. Store at 4  C. 2.2 Antibodies and Protein Standards

1. Capture antibody: cmvIL-10 Affinity Purified Polyclonal Antibody, Goat IgG (R&D Systems, Minneapolis, MN). Reconstitute in 500 μL of sterile PBS for a final concentration of 200 μg/mL (see Note 4). 2. Detection antibody: cmvIL-10 Biotinylated Affinity Purified Polyclonal Antibody, Goat IgG (R&D Systems). Reconstitute in 250 μL of sterile PBS for a final concentration of 200 μg/mL (see Note 5). 3. Protein Standard: Recombinant cmvIL-10 (R&D Systems). Reconstitute in 250 μL of Protein Standard Dilution Buffer (sterile PBS containing 0.1% BSA) for a final concentration at 100 μg/mL (see Note 6).

2.3 Other Reagents and Consumables

1. Streptavidin-HRP: Dilute as recommended by manufacturer, typically 1:40 in Reagent Diluent. 2. Substrate Solution (R&D Systems). Contains two substrate solutions (A & B) that are combined just prior to use (see Note 7). 3. Stop Solution: 1 M H2SO4. 4. 96-well ELISA plates: Nunc MaxiSorp™ high protein-binding capacity, flat-bottom plates. 5. 96-well 2 mL Deep Well plate for dilutions (see Note 8). 6. Adhesive plate seals. 7. Reagent reservoirs, 30–100 mL capacity. 8. 15 mL conical tubes for dilution. 9. 1.7 mL Eppendorf tube for dilution. 10. P1000, P200, and P10 pipette tips. 11. Paper towels.

2.4

Equipment

1. Multichannel pipettor, eight channel, 50–300 μL capacity. 2. Standard pipettors (P1000, P200, and P10). 3. Multimode or absorbance plate reader with ability to read optical density at 450 nm.

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Methods Carry out all procedures at room temperature, unless otherwise specified. Reagents that are stored at 4  C should be warmed to room temperature for 15 min prior to use.

3.1 Day 1: Coat the Plate

1. Coat the ELISA plate with capture antibody. For a full 96-well plate, prepare 10 mL of Capture Antibody at a final working concentration of 2 μg/mL. Place 10 mL PBS into a 15 mL conical tube, then add 100 μL of the stock Capture Antibody solution (200 μg/mL). Invert gently to mix and then pour antibody solution into a reagent reservoir. 2. Use a multichannel pipettor to dispense 100 μL to each well of the 96-well Nunc MaxiSorp™ plate. 3. Cover with an adhesive plate seal, ensuring edges are firmly sealed. 4. Incubate the plate overnight at 4  C.

3.2

Day 2: The Assay

1. Allow the ELISA plate, Wash Buffer, and Blocking Buffer to equilibrate to room temperature for 15 min. 2. Remove the Capture Antibody solution from the ELISA plate by inverting the plate over a sink and using a quick wrist flick to shake out fluid. Prepare a stack of paper towels and gently but firmly tap the plate down smartly onto the paper towels to ensure fluid has been removed from the wells and top of plate is dry (see Note 9). 3. Place about 100 mL of Wash Buffer into a reagent reservoir. Dispense 200 μL to each well of the plate. Remove fluid from the plate by flicking as described above. Repeat twice for a total of three washes. 4. Place about 25 mL of Reagent Diluent/Blocking Buffer into a reagent reservoir. Dispense 200 μL to each well of the plate. Replace plate seal and incubate at room temperature for 1 h. 5. Remove Reagent Diluent/Blocking Buffer from the plate by flicking. Wash three times (see Note 10). 6. Add samples and standards to the plate (see Fig. 1). Use 1000 pg/mL as the top standard and decrease protein concentration twofold down to 15.625 pg/mL (see Note 11). Prepare standards in the 96-well 2 mL Deep Well plate by placing 995 μL of Reagent Diluent into the first well (A1). 7. Add 500 μL to wells B1–H1. 8. Set Deep Well plate aside briefly and make a 200 ng/mL solution of the Recombinant cmvIL-10 Protein Standard (100 μg/mL stock) by placing 998 μL Reagent Diluent and

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Fig. 1 Standard assay plate layout. This image represents a 96-well ELISA plate. Standards are placed in columns 1–3, such that each standard is plated three times (triplicates). Standards consist of seven concentrations in a twofold dilution series. The last standard is a blank and contains Reagent Diluent only. Samples are placed in columns 4–6 in triplicate. Additional samples may be placed in columns 7–9 and 10–12 for a total of 24 samples in triplicate per plate

2 μL Protein Standard into an Eppendorf tube. Invert to mix, then add 5 μL of this 200 ng/mL solution to well A1 of the Deep Well plate. 9. Use a P1000 set for 500 μL to pipet up and down to mix several times (gently to avoid making bubbles), then transfer 500 μL into well B1. Pipet up and down to mix several times, then transfer 500 μL into well C1. Continue in this fashion until well G1. 10. Use the multichannel pipettor to transfer 100 μL from each well in column 1 of the Deep Well plate to each well in column 1 of the ELISA plate. Repeat, transferring 100 μL into columns 2 and 3 of the ELISA plate so that standards are now plated in triplicate. 11. Add samples directly to the plate for total volume of 100 μL in each well (see Note 12). Cover plate with adhesive seal and incubate at room temperature for 2 h. 12. Discard samples and standards by flicking into sink. Wash 3 times with a volume of 200 μL per well as described in step 3. 13. Add Detection Antibody to each well. Place 10 mL Reagent Diluent into a conical tube and add 10 μL Detection Antibody (200 μg/mL stock) to make a working solution at a final concentration of 0.2 μg/mL. Invert tube gently to mix, place into reagent reservoir, and dispense 100 μL to each well of the ELISA plate. Cover plate with adhesive seal and incubate at room temperature for 2 h.

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14. Discard Detection Antibody from the plate by flicking into sink. Wash 3 times with a volume of 200 μL per well as described in step 3. 15. Add Streptavidin-HRP that has been diluted 1:40 to working concentration (or as indicated by manufacturer’s instructions). Prepare a conical tube with 10 mL Reagent Diluent and add 250 μL Streptavidin-HRP. Invert tube gently to mix, place into reagent reservoir, and dispense 100 μL to each well of the ELISA plate. 16. Cover plate with adhesive seal and then with foil to avoid exposure to light. Incubate at room temperature for 20 min. 17. Discard the Streptavidin-HRP from the plate by flicking into sink. Wash 3 times with Wash Buffer with a volume of 200 μL per well as described in step 3. 18. Add freshly prepared Substrate Solution. Combine 5.5 mL Color Reagent A with 5.5 mL Color Reagent B in a conical tube, invert to mix, then place into reagent reservoir. Dispense 100 μL to each well of the ELISA plate. 19. Cover plate with foil to avoid exposure to light. Incubate at room temperature for 20 min. A blue color will develop in wells containing cmvIL-10, and color intensity is proportional to protein concentration. 20. Add 50 μL Stop Solution to each well of the ELISA plate. Note that the Substrate Solution is not removed, so each well now contains a total volume of 150 μL. 21. Determine the optical density (OD) by reading the plate at 450 nm. 22. Average the triplicate values for each standard and create a standard curve by plotting OD against concentration using a four-parameter logistic curve fit. Concentrations for samples (average of triplicate) can be interpolated from the standard curve (see Fig. 2).

4

Notes 1. Either 10 PBS or PBS purchased from a reputable vendor is an acceptable substitute. For labs doing a lot of ELISAs, making a 10 PBS stock and diluting to 1 as needed is more economical. 2. The Corning® 500 mL Vacuum Filter/Storage Bottle System, 0.2 μm Pore 33.2 cm2 Nylon Membrane, or a similar system is used for filter sterilization. 3. There are many possible blocking solutions, and many types of bovine serum albumin on the market. Bovine serum albumin,

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Fig. 2 Detection of cmvIL-10 in culture supernatants via ELISA. HEK293 cells were transfected with pcDNA-cmvIL-10 plasmid DNA or mock transfected as described [6] and supernatants harvested after 48 h. For virus infection, human neonatal foreskin fibroblast cells were infected with HCMV AD169 at an MOI of 1 for 96 h and supernatants collected as described [11]. Error bars represent standard error of three replicate samples

heat shock fraction (Sigma-Aldrich, St. Louis, MO) has been a reliable blocking reagent for this assay in our laboratory. 4. Allow lyophilized antibody to rehydrate with added PBS for at least 15 min at room temperature before use. One vial of 100 μg antibody, reconstituted in 500 μL PBS is sufficient for coating five ELISA plates. If five plates will not be needed within 1 week, the antibody solution should be aliquoted and stored at 20  C. 5. As noted above, allow lyophilized antibody to rehydrate with added PBS for at least 15 min at room temperature before use. One vial of 25 μg biotinylated antibody is sufficient for 25 ELISA plates. The antibody solution should be aliquoted and stored at 20  C. 6. Allow lyophilized protein to rehydrate for 15 min at room temperature prior to use. One vial of 25 μg Recombinant cmvIL-10 is sufficient for many assays. The protein should be aliquoted and stored at 20  C. 7. The substrate reagent kit contains two solutions, Color Reagent A (H2O2) and Color Reagent B (tetramethylbenzidine). Color Reagent B may cause skin, eye, or respiratory irritation. Follow the manufacturer’s instructions and avoid breathing fumes. These solutions should be combined in equal volumes just prior to use. For example, for one 96-well plate, prepare 11 mL of Substrate Solution by combining 5.5 mL of Color Reagent A and 5.5 mL of Color Reagent B. Mix gently and dispense 100 μL per well.

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8. While not essential, these Deep Well reagent blocks work well for preparing the standards and also for arranging samples. They enable rapid transfer of samples in triplicate to the ELISA plate, which is important to prevent the plate from drying out. 9. All the liquid should be removed at each stage, either by the flicking method or via aspiration. It is essential, however, that the plate does not dry out. Therefore, the next reagent to be added should be ready before the liquid is removed from the plate at any stage. 10. Because the plate should not dry out, it is important to have the protein standards and samples prepared and ready before completing the wash steps at this stage. If necessary, keep wash buffer on the plate until the materials for the next step are ready. 11. The range of 1000–15.625 pg/mL works well for most applications, however, the standard curve can be modified as needed to extend the assay range. We have used a high standard of 10,000 pg/mL with fivefold dilution for detection of cmvIL10 in both culture supernatants and human plasma [11]. For culture supernatants, use these samples undiluted and follow this protocol using PBS + 1% BSA as the Reagent Diluent to prepare protein standard dilutions as noted. For detection of cmvIL-10 in plasma, we found that tenfold dilution of plasma samples in PBS yielded the most accurate and reproducible results, particularly when coupled with protein standard that had been prepared using PBS containing 10% plasma from an HCMV seronegative donor as the Reagent Diluent. 12. Using the Deep Well plate to prepare the protein standard dilutions allows all the standards to be added at once with a multichannel pipettor. It is not recommended to attempt this assay without a multichannel pipettor, as it will take too long to add each standard and sample, and the plate will dry out. For samples, especially culture supernatants, these can be added directly the Deep Well plate and quickly transferred to the corresponding wells of the ELISA plate.

Acknowledgements This work was supported by grants from the National Institute for Allergy and Infectious Disease of the National Institutes of Health (AI111232) and the Avon Foundation for Women (02-2014-052) to J.V.S.

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References 1. Yalow RS, Berson SA (1959) Assay of plasma insulin in human subjects by immunological methods. Nature 184(Suppl 21):1648–1649 2. Engvall E, Perlmann P (1971) Enzyme-linked immunosorbent assay (ELISA). Quantitative assay of immunoglobulin G. Immunochemistry 8:871–874 3. Van Weemen BK, Schuurs AH (1971) Immunoassay using antigen-enzyme conjugates. FEBS Lett 15:232–236 4. Kotenko SV, Saccani S, Izotova LS, Mirochnitchenko OV, Pestka S (2000) Human cytomegalovirus harbors its own unique IL-10 homolog (cmvIL-10). Proc Natl Acad Sci U S A 97:1695–1700 5. Lockridge KM, Zhou SS, Kravitz RH, Johnson JL, Sawai ET, Blewett EL et al (2000) Primate cytomegaloviruses encode and express an IL-10-like protein. Virology 268:272–280 6. Spencer JV, Lockridge KM, Barry PA, Lin G, Tsang M, Penfold ME et al (2002) Potent immunosuppressive activities of cytomegalovirus-encoded interleukin-10. J Virol 76:1285–1292

7. Chang WL, Barry PA (2010) Attenuation of innate immunity by cytomegalovirus IL-10 establishes a long-term deficit of adaptive antiviral immunity. Proc Natl Acad Sci U S A 107:22647–22652 8. Jones BC, Logsdon NJ, Josephson K, Cook J, Barry PA, Walter MR (2002) Crystal structure of human cytomegalovirus IL-10 bound to soluble human IL-10R1. Proc Natl Acad Sci U S A 99:9404–9409 9. Brodeur ND, Spencer JV (2010) Antibodies to human IL-10 neutralize ebvIL-10-mediated cytokine suppression but have no effect on cmvIL-10 activity. Virus Res 153:265–268 10. Eberhardt MK, Chang WL, Logsdon NJ, Yue Y, Walter MR, Barry PA (2012) Host immune responses to a viral immune modulating protein: immunogenicity of viral interleukin-10 in rhesus cytomegalovirusinfected rhesus macaques. PLoS One 7:e37931 11. Young VP, Mariano MC, Tu CC, Allaire KM, Avdic S, Slobedman B et al (2017) Modulation of the host environment by human cytomegalovirus with viral interleukin 10 in peripheral blood. J Infect Dis 215:874–882

Chapter 16 Techniques for Characterizing Cytomegalovirus-Encoded miRNAs Nicole L. Diggins, Lindsey B. Crawford, Hillary M. Struthers, Lauren M. Hook, Igor Landais, Rebecca L. Skalsky, and Meaghan H. Hancock Abstract microRNAs (miRNAs) are small noncoding RNAs that regulate gene expression at the posttranscriptional level by binding to sites within the 30 untranslated regions of messenger RNA (mRNA) transcripts. The discovery of this completely new mechanism of gene regulation necessitated the development of a variety of techniques to further characterize miRNAs, their expression, and function. In this chapter, we will discuss techniques currently used in the miRNA field to detect, express and inhibit miRNAs, as well as methods used to identify and validate their targets, specifically with respect to the miRNAs encoded by human cytomegalovirus. Key words microRNA (miRNA), Northern Blot, Stem-loop real-time PCR, RISC-immunoprecipitation (RISC-IP), Photoactivatable ribonucleoside-enhanced cross-linking and Immunoprecipitation (PAR CLIP), Luciferase assay, Locked nucleic acids (LNA), Human cytomegalovirus (HCMV)

1

Introduction The discovery that animal cells and later DNA viruses [1] encode small, noncoding regulatory RNA molecules known as microRNAs (miRNAs) gave rise to the study of an entirely new method of gene regulation. Since their initial discovery [2], the miRNA family has expanded exponentially, aided by the development of a variety of identification techniques, from in silico prediction and cross-species identification [3] to small RNA cloning [4] and highly sensitive deep-sequencing approaches [5–9]. Many of these miRNAs are conserved across species and regulate important biological processes including cellular differentiation, DNA repair, metabolism, apoptosis, immunity, aging, and cancer [10]. The vast majority of miRNAs encoded by viruses have been identified in the herpesvirus family, including HSV-1

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(18 pre-miRNAs), EBV (25 pre-miRNAs), KSHV (12 pre-miRNAs), HCMV (15 pre-miRNAs), MCMV (18 pre-miRNAs), and RCMV (24 pre-miRNAs) [5, 6, 8, 11– 17]. Virally encoded miRNAs are hypothesized to play a key role in mediating viral pathogenicity through their interactions with both viral and host targets [18]. The effect of an individual miRNA on a given target may be quite small; however, recent studies show that multiple viral miRNAs can cooperate to target several distinct components belonging to a regulatory pathway or cellular process to likely play a larger role in modulating the cell or promoting infection [19–22]. The combined functions for the majority of viral miRNAs are unknown. Following a brief introduction to miRNA biogenesis and mode of action, this chapter will focus on the methods routinely used to express, detect, and inhibit CMV miRNAs as wells as methods used to identify and validate miRNA targets. The canonical pathway for cellular biogenesis of miRNAs [10, 23] begins with RNA polymerase II–mediated transcription of a single-stranded primary miRNA transcript (pri-miRNA) which subsequently folds into one or more hairpin structures. In the nucleus, pri-miRNAs are processed by the endonuclease, Drosha, generating a ~60 nucleotide (nt) precursor miRNA hairpin species (pre-miRNA) that is exported to the cytoplasm by Exportin 5. There, the pre-miRNA is cleaved by a second endonuclease, Dicer, generating an 18–22 nt double-stranded miRNA duplex. One strand is then incorporated into the RNA-induced Silencing Complex (RISC) that minimally contains an Argonaute (Ago) protein and a mature miRNA. The second strand, or passenger strand, is usually degraded, but in some instances may also be incorporated into RISC generating two RISC populations from one double-stranded duplex [10]. Understanding the biogenesis of miRNAs has allowed for the development of several strategies to ectopically express miRNAs in cells, which are detailed below (see Subheading 3.1). After their incorporation into RISC, miRNAs posttranscriptionally regulate cellular gene expression through the miRNAdirected and sequence-specific binding of RISC to RNA targets. This feature is the basis of several related methods aimed at identifying miRNA targets. These methods are based on the affinity purification of ribonucleoprotein (RNP) complexes followed by RNA purification and identification of miRNA targets by microarray or deep sequencing. In contrast with the RISC-Immunoprecipitation (RISC-IP) [24–27] and Streptavidin pull-down of biotinylated miRNA mimic strategies [28–30] (detailed below, see Subheading 3.3.2), other methods such as HITS-CLIP (highthroughput sequencing of RNAs isolated by cross-linking and immunoprecipitation) [31], PAR-CLIP (photoactivatable ribonucleoside enhanced cross-linking and immunoprecipitation, [32–35]

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(detailed below, see Subheading 3.3.1) and iCLIP (individualnucleotide resolution UV cross-linking and immunoprecipitation) [36] use UV cross-linking to stabilize the RNP complex. These cross-linking methods have the potential to significantly expand the number of potential CMV miRNA targets. miRNA target sites are most commonly located in the 0 3 untranslated region (UTR) of a mRNA, although sites in the 50 UTR and the coding sequence have also been observed [10, 30]. This feature has been exploited to develop sensitive luciferase assays (see Subheading 3.2.2) aimed at measuring the posttranscriptional effect of miRNAs on their potential targets. The fate of the target mRNA depends on whether the miRNA incorporated into RISC is fully or partially complementary with the target mRNA. Extensive complementarity generally leads to Ago-mediated mRNA cleavage, while partial complementarity via the seed sequence, a region encompassing nucleotides (nt) 2 through 8 of the mature miRNA [1, 10], leads to translational repression and/or mRNA decay through mechanisms which are not completely understood. Since mRNA levels are not always affected by miRNAs, protein-based quantitative methods such as SILAC (stable isotopes labeling by amino acids in cell culture) coupled with mass spectroscopy have also been developed to identify miRNA targets [7, 37]. These methods provide a valuable counterpoint to the RNP affinity purification methods mentioned above for target identification.

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Materials

2.1 Expression of miRNAs in Cells

1. pSIREN-retroQ-ZsGreen (Clontech, see Note 1).

2.1.1 Expressing miRNAs Using an Expression Vector (pSIREN)

3. Purified viral DNA (10–50 ng/μL).

2. BamHI and EcoRI restriction enzymes and digestion buffers. 4. Forward and reverse PCR primers designed 50, 200, or 500 base pairs upstream and downstream of the miRNA of interest, respectively (see Note 2). 5. PCR reagents. 6. Thermocycler. 7. 1% agarose gel. 8. Ethidium bromide. 9. Agarose gel electrophoresis apparatus and reagents. 10. PCR fragment purification kit. 11. DNA ligation kit. 12. Electrocompetent E. coli bacteria strain (e.g., DH5α). 13. LB-ampicillin dishes.

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14. LB broth + ampicillin. 15. Miniprep kit. 16. Maxiprep kit. 17. Opti-MEM I Reduced Serum media (Invitrogen). 18. Lipofectamine 2000 (Invitrogen, see Note 3). 19. HEK293 cells and growth media (see Note 3). 20. Fluorescent microscope. 21. Amaxa® Cell Line Nucleofector® Kit R (Lonza). 22. Nucleofector® II Device (Lonza). 2.1.2 Expressing miRNAs Using GFP-Expressing Adenovirus

1. pAdTrack-CMV (Addgene). 2. pAdEasier electrocompetent cells (Addgene). 3. PCR reagents. 4. 1% agarose gel. 5. Ethidium bromide. 6. Agarose gel electrophoresis apparatus and reagents. 7. NotI, XhoI, PmeI, and PacI restriction enzymes and buffers. 8. PCR fragment purification kit. 9. DNA ligation kit. 10. Electrocompetent E. coli bacteria strain (e.g., DH5α). 11. LB-Kanamycin plates. 12. LB Broth + Kanamycin. 13. 293m cells and growth media. 14. Tissue culture treated T25 flasks. 15. Nucleobond® BAC 100 Maxiprep kit (Macherey-Nagel). 16. QIAEX®II gel extraction kit (Qiagen). 17. Lipofectamine 2000 (Invitrogen). 18. Opti-MEM I Reduced Serum media (Invitrogen). 19. Phosphate buffered saline.

2.2 Detecting miRNAs 2.2.1 miRNA Northern Blot Urea–Acrylamide Gel Components

1. Urea–acrylamide gel: In a 500 mL glass bottle, mix 150 mL 19:1 acrylamide (30%), 30 mL 10 TBE (890 mM Tris base, 890 mM boric acid, and 20 mM EDTA pH 8.0), and 126 g of urea. Place in 68  C water bath, stirring if necessary, until components are dissolved. Make up to 300 mL using nuclease-free water. Pass through a 0.45 μM Steriflip filter (Millipore) and store at 4  C, shielded from light. 2. Ammonium persulfate: 10% solution in water, store at 4  C. 3. N,N,N,N0 -tetramethylethylenediamine (TEMED), store at 4  C.

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4. Running buffer: 1 TBE (diluted from 10 TBE using distilled water). 5. Formamide, store at 4  C. 6. 6 RNA loading dye: 10 mM Tris–HCl pH 7.6, 0.03% bromophenol blue, 0.03% xylene cyanol, 60% glycerol, 60 mM EDTA. Store at 20  C. 7. Mini PROTEAN 3 System (Bio-Rad) or other mini-gel electrophoresis system. 8. 10% SDS (see Note 4). 9. 95% ethanol. 10. Ethidium bromide (10 mg/mL solution) (see Note 5). 11. UV transilluminator with camera. 12. Fluorescent ruler. 13. Small RNA marker. 14. Total RNA samples from cells expressing the miRNA of interest (see Note 6). Transfer Components

1. Gene Screen PLUS membrane (PerkinElmer) or other positively charged nylon membrane. 2. Whatman paper (0.34 mm thickness). 3. Transfer buffer: 1 TBE. 4. Mini Trans-blot Electrophoretic transfer cell. 5. UV Stratalinker 1800.

Probe Labeling Components

1. 10 kinase buffer A. 2. Adenosine 50 triphosphate (γ-32P), 10uCi/uL (see Note 7). 3. Polynucleotide kinase, 10 U/μL. 4. 37  C heat block. 5. G-25 Quick Spin Columns (Millipore Sigma). 6. 1 TE buffer: 10 mM Tris–HCl, pH 7.6, 1 mM EDTA, pH 8.0. 7. Tabletop centrifuge. 8. Oligonucleotide probe designed antisense to the miRNA of interest. The sequences of known HCMV miRNAs are readily available online within the miRBase database (http://www. mirbase.org/) [14–17].

Hybridization Components

1. PerfectHyb solution (Sigma). 2. Hybridization tubes. 3. Hybridization oven.

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4. 20 SSC: 3 M NaCl, 300 mM sodium citrate. 5. Wash solution 1: 2 SSC, 0.05% SDS, dissolved in water. 6. Wash solution 2: 0.1 SSC, 0.1% SDS, dissolved in water. Detection Components

1. Autoradiography film. 2. Film cassette. 3. Intensifier screen. 4. Film developer.

2.2.2 Stem-Loop RT-PCR for Detection of miRNAs Reverse Transcription Components

1. 200 μM stock of RT primer or 5 stock (Applied Biosystems) or similar. 2. 10 RT buffer (100 mM Tris, pH 8.3, 500 mM KCl, and 5.5 mM MgCl2). 3. 10 μM dNTP stock. 4. Multiscribe Enzyme 50 U/μL (Applied Biosystems). 5. RNasin PLUS RNase inhibitor, 40 U/μL (Promega). 6. Thermocycler. 7. RNA template (50 ng/μL) (see Note 8).

Taqman Assay Components

1. 2 Universal PCR Master Mix (Applied Biosystems) or similar. 2. 50 μM Forward and Reverse primers or 20 mastermix (Applied Biosystems) or similar. 3. 10 μM probe. 4. Real-time PCR machine.

2.3 Identifying mRNA Targets of Viral miRNAs

1. DPBS 1, Dulbecco’s phosphate buffered saline. Store at 4  C. 2. Nuclease-free water. 3. 4-thiouridine (4SU), 1 M in DMSO (Sigma). Store at 20  C.

2.3.1 Photoactivatable Ribonucleoside-Enhanced Cross-Linking and Immunoprecipitation (PAR-CLIP)

4. Protein G Dynabeads (ThermoFisher Scientific). Store at 4  C.

Immunoprecipitation Components

6. Complete, mini, EDTA-free Protease Inhibitor Cocktail tablets (Sigma) or similar. Store at 4  C.

5. Lysis buffer: 50 mM HEPES, pH 7.5, 150 mM KCl, 2 mM EDTA, 1 mM NaF, 0.5% NP40, 0.5 mM DTT (added just before use), and 1 Protease inhibitor cocktail (added just before use). Keep on ice.

7. Antibody directed against one of the RISC complex components (see Note 9). 8. RNase T1. Store at 20  C. 9. IP Wash Buffer: 50 mM HEPES, pH 7.5, 300 mM KCl, 0.05% NP40, 0.5 mM DTT (added just before use), 1 protease inhibitor cocktail (added just before use). Keep on ice.

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10. High Salt Wash Buffer: 50 mM HEPES, pH 7.5, 500 mM KCl, 0.05% NP40, 0.5 mM DTT (added just before use), 1 protease inhibitor cocktail (added just before use). Keep on ice. 11. Alkaline phosphatase, calf intestinal (New England Biolabs). Store at 20  C. 12. NEBuffer 3 (New England Biolabs). Store at 20  C. 13. Cross-link Wash Buffer: 50 mM Tris–HCl, pH 7.5, 20 mM EGTA, 0.5% NP40. 14. PNK Buffer: 50 mM Tris–HCl, pH 7.5 50 mM NaCl, 10 mM MgCl2, 5 mM DTT (added just before use, prepare buffer with and without). Keep on ice. 15. ATP, 100 mM (Fisher). Store at 20  C. 16. 1 SDS-PAGE loading buffer: 50 mM Tris, pH 6.8, 2 mM EDTA, 10% glycerol, 2% SDS, 0.1% bromophenol blue, 100 mM DTT (added just before use). 17. 4–12% Criterion XT Bis-Tris gel, 12 wells (Bio-Rad). Store at 4  C. 18. Precision Plus Protein Marker. Store at 20  C. 19. 20 MOPS Buffer (diluted to 1 using Molecular grade water). 20. D-tube dialyzer midi tube and accessory kit (Sigma). 21. 5 μm Acrodisc Syringe Filter with Supor® membrane (Pall). 22. Magnetic particle concentrator. 23. UV Stratalinker 2400. 24. Tabletop centrifuge capable of refrigeration. 25. 95  C heat block. 26. Criterion midi electrophoresis gel apparatus (Bio-Rad, Hercules, California, USA). 27. Agarose gel electrophoresis apparatus. 28. 150 mm dishes of cells expressing miRNA(s) of interest (see Note 10) (see Subheading 3.1). RNA Extraction and cDNA Library Components

1. 2 Proteinase K buffer: 100 mM Tris–HCl, pH 7.5, 150 mM NaCl, 12.5 mM EDTA, 2% SDS. 2. Proteinase K, recombinant, PCR grade (Sigma). Store at 4  C. 3. Acidic phenol–chloroform–isoamyl alcohol (25:24:1). Store at 4  C. 4. Chloroform. 5. 3 M NaCl. 6. GlycoBlue Coprecipitant, 15 mg/mL (ThermoFisher). Store at 20  C (see Note 11).

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7. 100% ETOH. Keep an aliquot on ice. 8. 95% ETOH. Keep an aliquot on ice. 9. TruSeq Small Library Prep Kit (Illumina). Store at 20  C (see Note 12). 10. T4 RNA Ligase 2, truncated (supplied with 5 First-strand buffer) (New England Biolabs). Store at 20  C (see Note 13). 11. Superscript III Reverse Transcriptase. Store at 20  C. 12. 5% Criterion TBE gel, 18 wells (Bio-Rad). Store at 4  C. 13. 10 TBE buffer: 890 mM Tris base, 890 mM boric acid, 20 mM EDTA, pH 8 (dilute to 1 in nuclease-free water). 14. 6 DNA loading dye (Fisher). 15. SYBR Safe (Fisher). 16. 0.5 M NaCl (dilute from 3 M in nuclease-free water). 17. UV transilluminator. 2.3.2 RNA-Induced Silencing Complex (RISC) Immunoprecipitation (IP) Immunoprecipitation and Pull-Down Components

1. DPBS 1, Dulbecco’s phosphate buffered saline. Store at 4  C. 2. Lysis buffer: 20 mM Tris–HCl, pH 7.5, 2.5 mM MgCl2, 200 mM NaCl, 0.05% NP-40, 1 mM DTT (added just before use), 1 Protease Inhibitor Cocktail (added just before use). Keep on ice. 3. Complete, Mini, EDTA-free Protease Inhibitor Cocktail Tablets (Roche Applied Science) or similar. Store at 4  C. 4. Streptavidin-Agarose (Invitrogen), ANTI-c-myc Agarose Conjugate (Sigma), Protein A beads, or similar. Store at 4  C. 5. Antibody directed against one of the RISC complex components (see Note 9). 6. 10 mg/mL yeast tRNA. 7. 10 mg/mL BSA. 8. RNasin Plus RNase Inhibitor 40 U/μL. Store at 20  C. 9. Cell scrapers. 10. Tabletop centrifuge capable of refrigeration. 11. 100 mm dishes of cells expressing miRNA(s) of interest (see Note 14) (see Subheading 3.1).

RNA Isolation Components

1. TRIzol Reagent or similar. Store at 4  C. 2. Chloroform. 3. Isopropanol. 4. GlycoBlue Coprecipitant (Applied Biosystems) (see Note 11). Store at 20  C. 5. 75% ETOH: Leave an aliquot on ice. Store at 20  C. 6. Nuclease-free water.

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1. HEK293 cells (see Note 3). 2. Exogenous miRNA source (see Note 15). 3. Dual luciferase reporter vector with test, negative, and positive control 30 UTRs cloned downstream of the reporter: pSICheck-2 vector (Promega) (see Note 16). In the case of the pSICheck-2 vector, which contains both Renilla and Firefly luciferase genes, the 30 UTR of the potential target is cloned downstream of the Renilla luciferase gene. 4. Black-walled clear-bottom 96-well tissue culture treated plate (see Note 17). 5. Lipofectamine 2000 (Invitrogen, see Note 3). 6. Opti-MEM I Reduced Serum Media.

Luciferase Assay Components

1. Dual-Luciferase Reporter Assay System.

2.4 Inhibition of HCMV miRNAs Using Locked Nucleic Acids (LNA)

1. LNA: miRCURY LNA™ microRNA Power Inhibitors (see Note 19) from Exiqon (www.exiqon.com/mirna-inhibitors). Resuspend 5 nmol in 50 μL or 250 μL RNase-free water to obtain a 100 μM or 20 μM stock, respectively. Aliquot and store at 80  C. Do not freeze–thaw more than five times.

2. Luminometer (see Note 18).

2. miRNA mimic (see Subheading 3.1). 3. Luciferase reporter plasmid (see Subheading 3.3.3 for design and cloning). 4. Lipofectamine 2000 (Invitrogen, see Note 3). 5. HEK293 cells and growth media. 6. Opti-MEM I Reduced Serum Media.

3

Methods

3.1 Expression of miRNAs in Cells

The simplest method for expressing CMV miRNAs in cells is through native infection with CMV. This strategy can be very valuable to monitor the effects of physiologically relevant concentrations of miRNAs during the course of CMV infection (up- or downregulation of target genes, virus growth, etc.). However, isolating the effect of a particular miRNA in this setting is complicated by the simultaneous expression of multiple viral genes and miRNAs. To explore the effects of CMV-encoded miRNAs on cellular gene expression in a more defined environment, exogenous miRNA expression is necessary. The choice between the two transfection-based expression methods or viral vector infection methods described below will depend on the objective of the study: if the goal is to express the miRNA at high levels while keeping the cost low, using expression vectors or viral vector

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strategies are appropriate. The downside of this method is that precisely adjusting the quantity of miRNA expressed in the cell is not possible. This situation may be problematic when comparing the effect of individual or combinations of miRNAs, or when performing titration curves. Alternatively, transfection of miRNA mimics allows for the delivery of precise amounts of miRNAs into the cell [38], but the cost can be limiting if large experiments are envisioned. Finally, lentiviral- or adenoviral-mediated miRNA expression might be the only possibility in difficult to transfect cells or when long-term expression is required, but can require significant optimization (see Note 20). 3.1.1 Expressing miRNAs Using a pSIREN Expression Vector

The following protocol explains how to clone and express miRNAs using the pSIREN vector as well as methods to introduce the expression vector into adherent and nonadherent cell types.

Cloning into pSIREN Vector

1. Amplify the region surrounding the miRNA of interest using forward and reverse primers containing BamHI and EcoRI restriction sites, respectively, and 10–50 ng purified viral DNA (see Note 21) using appropriate PCR procedures. 2. Verify the amplification of the predicted product by loading 2 μL of the PCR reaction on a 1% agarose gel and visualize the DNA with ethidium bromide. 3. Purify the remaining PCR product using a PCR fragment purification kit (see Note 22). 4. Digest 1 μg pSIREN vector and the purified PCR fragment using the BamHI and EcoRI restriction enzymes following manufacturer’s instructions. 5. Purify the digested pSIREN and PCR fragment using the appropriate purification kit (see Note 22). 6. Ligate the digested PCR fragment to pSIREN according to the instructions of the ligation kit. 7. Transform the ligation reaction in competent bacteria following standard procedures. 8. Pick colonies and grow small cultures (3–5 mL) in LB-ampicillin media; extract plasmid DNA using a miniprep kit. 9. Verify the presence of an insert of the correct size by colonyPCR or BamHI–EcoRI digestion. 10. Confirm positive clones by sequencing using the Forward and/or Reverse PCR primer. 11. Perform a maxiprep on a validated clone to obtain working quantities of plasmid DNA.

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1. 24 h before transfection, plate 1  105 HEK293 cells in 1 mL media per well in a 12-well plate or 2  106 cells in 10 mL media per 100 mm dish. 2. For each well of a 12-well plate, add 0.5 μg pSIREN-miRNA DNA to 75 μL Opti-MEM I Reduced Serum media and 2 μL lipofectamine 2000 to 75 μL Opti-MEM I Reduced Serum media. For a 100 mm dish, add 5 μg pSIREN-miRNA DNA to 500 μL Opti-MEM I Reduced Serum Media and 20 μL Lipofectamine 2000 to 500 μL Opti-MEM I Reduced Serum Media. 3. Incubate for 5 min at room temperature, combine DNA- and lipofectamine-containing media, mix gently, and incubate 20 min. 4. Add transfection mix to the cells dropwise. 5. After 24 h, assess plasmid transfection efficiency by evaluating GFP expression using a fluorescent microscope or flow cytometry (see Note 23). 6. Remove medium and harvest cells using the appropriate protocol (see Subheadings 3.2 and 3.3).

Nucleofection

1. Twenty-four hours before nucleofection, prepare, for each nucleofection, low passage suspension cells by spinning down 1–3  106 cells and resuspend in fresh media at 1  106 cells/ mL. 2. Alternatively, thaw and wash primary suspension cells and resuspend in fresh media at appropriate density (see Note 24). 3. After 24 h, mix the appropriate Amaxa® Cell Line Nucleofector® Kit R buffer and supplement per manufacturer’s instructions (see Note 24) to make 100 μL working buffer per nucleofection. Aliquot 100 μL of working buffer per nucleofection, and add 1 μg/1  106 cells endotoxin-free maxiprepped pSIREN plasmid to each aliquot. Pipet gently to mix. 4. Prepare one recovery well per nucleofection. Place 1 mL media/1  106 nucleofected cells in each recovery well, that is, if performing three nucleofections, each with 3  106 cells, prepare three wells of a 6-well plate, each containing 3 mL recovery media. 5. Spin down cells, aspirate supernatant. 6. Resuspend cells in 15 mL PBS per nucleofection, and aliquot the cells in PBS for each nucleofection to a new 15-mL conical or Eppendorf tube. 7. Spin down cells and aspirate supernatant without disturbing the cell pellet.

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8. Gently and slowly (but thoroughly) resuspend cells for one nucleofection in 100 μL working buffer containing plasmid, using p200 pipette and tips. Transfer cells in working buffer with plasmid to Nucleocuvette. 9. Electroporate using the Nucleofector® II device using program V-001 (see Note 24). 10. Immediately add a small aliquot of media from the recovery well to the cuvette using a provided micro-transfer pipette and transfer the cells from the cuvette to their recovery well. 11. Repeat steps 7–9 for each nucleofection. 12. Allow cells to recover for an appropriate length of time according to your experimental protocol. 13. Harvest cells by centrifuging and removing supernatant (see Subheadings 3.2 and 3.3 miRNAs for a sample of downstream applications). 3.1.2 Expressing miRNAs Using GFP-Expressing Adenovirus

The following protocol explains how to clone and express miRNAs from GFP-expressing adenoviruses using the pAdEasy recombination-based system [39]. Adenovirus infection is an efficient means to deliver a miRNA expression cassette in difficult-totransfect cells such as the nonadherent monocytic cell lines THP-1 and Kasumi-3 as well as primary human CD34+ hematopoietic progenitor cells (HPCs).

Cloning into the pAdTrack-CMV Vector

1. Amplify the region surrounding the miRNA of interest using forward and reverse primers containing NotI and XhoI restriction sites (see Note 25), respectively, at the 50 -end and 10–50 ng purified viral DNA (see Note 21) using appropriate PCR procedures. 2. Verify the amplification of the predicted product by loading 2 μL of the PCR reaction on a 1% agarose gel and visualize the DNA with ethidium bromide (see Note 5). 3. Purify the remaining PCR product using a PCR fragment purification kit (see Note 22). 4. Digest 4 μg of pAdTrack-CMV vector and the PCR amplification product using NotI and XhoI restriction enzymes according to the manufacturer’s protocols. 5. Purify the digested pAdTrack-CMV and PCR fragment using appropriate purification kit(s). 6. Ligate the digested PCR fragment into pAdTrack-CMV according to the instructions of the ligation kit. 7. Prepare pAdEasier electrocompetent cells (see Note 26).

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8. Linearize the pAdTrack-CMV plasmid containing your sequence of interest with PmeI according to the manufacturer’s protocols. 9. Purify plasmid using PCR fragment purification kit. 10. Transform at least 500 ng of purified pAdTrack-CMV containing your sequence of interest into the pAdEasier electrocompetent cells. Recover in LB for 30 min and plate bacteria over 3–5 LB plates containing kanamycin overnight at 37  C. 11. Pick small colonies and grow in LB + Kanamycin broth at 37  C overnight. 12. Perform minipreps according to the manufacturer’s protocols and digest plasmids with PmeI. Run at least 20 μL of the digested minipreps on a 1% agarose gel and visualize DNA with ethidium bromide (see Note 5). Correctly recombined clones should yield a band of ~3–4.5 kb as well as a larger band running near 30 kb. 13. Retransform 2 μL of miniprepped DNA for each correct clone into electocompetent bacteria (DH5α). Adenovirus Production

1. Prepare one T25 flask of 293m cells per adenovirus transfection by plating 7.5  105 cells/flask 24 h prior to transfection. Cells should be approximately 50–70% confluent at time of transfection. 2. Prepare at least 4 μg of recombinant adenoviral DNA using the Nucleobond® 100 BAC maxiprep kit. Digest 4 μg of DNA using PacI according to manufacturer’s protocol. Run digested adenoviral DNA on a 1% agarose gel and visualize DNA with ethidium bromide (see Note 5). Excise the larger ~30 kb band and purify using the QIAEX®II gel purification kit. 3. Perform a standard Lipofectamine 2000 transfection according to the manufacturer’s protocol. Use up to 4 μg of PacIdigested, purified adenoviral DNA with 20 μL Lipofectamine 2000 in 500 μL Opti-MEM media. 4. Remove growth media from the 293m cells, wash once with PBS and add 2.5 mL Opti-MEM media per T25 flask prior to addition of transfection mix. 5. The next morning, replace transfection media with growth media. 6. GFP-positive cells should be observed within 7–10 days posttransfection. 7. When close to 100% of the cells are GFP-positive, scrape cells and transfer to a 50 mL conical tube. Spin to pellet cells and resuspend in 2 mL PBS. Freeze cells at 80  C and thaw in water bath at 37  C. Vortex vigorously and repeat freeze–thaw

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3 more times (four times total). Do not let the supernatants warm above melting temperature. Spin samples briefly and store at 80  C. 8. Infect 2 T25 flasks of 293m cells using 30–50% of the supernatant from step 7. Within 3–5 days one-third to one-half of the cells should become detached. Scrape cells and prepare supernatant as in step 7 (see Note 27). Each round of amplification should increase the number of adenoviral particles by ten-fold. 9. To generate a high-titer, large adenoviral stock, prepare 10 T150 flasks of 293m cells and infect and harvest as in step 8. 10. Concentrate virus by ultracentrifugation of clarified supernatant over a 20% sorbitol cushion at 72,000  g for 2 h. 11. Aliquot and freeze stocks at 80  C. 12. Titer virus by serial dilution and infection of 293m cells using a TCID50 assay. Read titers at 1 week postinfection and calculate PFU/mL. 13. For infection of suspension cells (i.e., Kasumi-3, THP-1, or primary HPCs), experimentally optimize the infection conditions for the downstream assay, balancing infection, viability, and miRNA expression. A good starting point is a serial dilution time course using virus at an MOI of 10 to 1000 and assessing the percent of cells that are both viable and GFP+ at 24, 48, and 72 hpi. 3.1.3 Designing and Expressing miRNA Mimics

Some HCMV miRNA double-strand RNA mimics are commercially available as duplexes or pre-miRNA hairpins. For miRNAs currently unavailable or to design miRNA duplexes for newly discovered miRNAs the following protocol can be used [28, 29].

Design miRNA Mimics

1. Identify the sequence of the mature or guide strand of the miRNA of interest. The sequences of known HCMV miRNAs are readily available online within the miRBase database (http://www.mirbase.org/) [14–17]. 2. Include the two nucleotides directly downstream of the mature strand at the 30 end (see Fig. 1). These nucleotides are easily identifiable by examining the miRNA stem-loop structure. 3. Reverse-complement the mature strand sequence to serve as the antisense strand and add the 2 nucleotides directly downstream as in step 2. At this point the sequences of the two strands should be completely complementary except for the two-nucleotide overhangs on the 30 end of each strand. 4. Make a mutation in the fourth nucleotide from the 30 end of the antisense strand (see Note 28). 5. Send RNA sequences to chosen manufacturer (IDT).

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Fig. 1 Design of custom miRNA mimics. 2 nucleotides directly downstream of the mature miRNA (shown here in bold) are added by examining the stem-loop structure in mirBase. The sequence of the passenger strand is determined by reverse complementing the mature strand, followed by the addition of 2 nucleotides on the 30 -end. A mismatch is then made in the passenger strand at the fourth nucleotide from the 30 -end to facility incorporation of the mature strand into RISC. Phosphates are added to the 50 -ends of each strand. A biotin moiety can be added to the 30 -end of the mature strand to facilitate isolation of miRNA containing complexes using affinity purification techniques

6. Request the addition of phosphates to the 50 ends of each strand. If biotinylating the mature strand, request the addition of a biotin moiety at the 30 end of the mature strand (see Note 29). 7. Have the two RNA species annealed by the manufacturer. Alternatively, the RNAs may be annealed by heating the two RNA species to 95  C and slowing cooling to room temperature. Transfection of miRNA Mimics

A variety of reagents for transfection of miRNAs and siRNAs are commercially available. Conditions for transfection of both primary cells and established cell lines are typically provided by the manufacturer. 1. Lipofectamine 2000, described in detail in Subheadings 3.1.1 and 3.4, provides a high level of transfection for a variety of cell types and gives the most consistent results when transfecting both plasmids and RNAs together. We typically use 1–2 pmol of miRNA mimic and 1 μL of Lipofectamine 2000 to transfect HEK293 cells in a 24-well format. 2. We have also successfully used Lipofectamine RNAiMax when transfecting miRNA mimics or siRNAs alone. We typically use 0.8 μL of RNAiMax to transfect HEK293 cells in a 24-well format. For subsequent use in RISC-IP (see Subheading 3.3.2), begin optimization using 100–400 pmol of the biotinylated miRNA mimic and 20 μL of Lipofectamine 2000 to transfect HEK293 cells in one 100 mm dish.

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3.2 Detecting miRNAs 3.2.1 miRNA Northern Blot

Urea–Acrylamide Gel Electrophoresis

In this protocol, small RNAs are separated through an urea–acrylamide gel and transferred electrophoretically to a nylon-based membrane. An oligonucleotide probe designed antisense to the miRNA of interest is labeled with γ-32P ATP and hybridized to the membrane [40]. Denaturing urea–acrylamide gels allow for the efficient separation of small, single-stranded RNA species. Therefore, both the pre- and mature forms of a miRNA can be simultaneously visualized (see Fig. 2). When compared to the stem-loop RT-PCR assay described elsewhere in this chapter, Northern blotting requires more total RNA and is less sensitive, which may be problematic for detecting miRNAs expressed at low levels. However, this technique is quantitative and technically straightforward and is an important validation step for miRNAs identified by more high-throughput methods. Carry out all procedures at room temperature, unless otherwise noted. 1. Make sure that all glass plates have been cleaned with 10% SDS, rinsed with water and with 95% ethanol. Allow the glass plates to dry before casting gel. 2. Prepare a Mini PROTEAN 3 casting tray using 1.5 mm spacers. Mix 10 mL of urea–acrylamide, 12.5 μL of TEMED, and 50 μL of 10% APS. Pour into casting tray along with appropriate comb and allow gel to set.

Fig. 2 miRNA Northern blot analysis of RNA isolated from fibroblasts mock infected (M) or infected with CMV (I) for 72 h or 293 cells transfected (T) with a pSIREN vector for 72 h. Both the pre- (~60 nucleotide) and mature (~22 nucleotide) forms of a miRNA can be detected. The signal at the very top of the membrane likely represents the larger sized pri-miRNA transcripts from which the miRNA precursors are derived

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3. Prepare gel tank apparatus and fill with 1 TBE. Rinse wells with 1 TBE using a syringe. Prerun apparatus at 180 V for 1 h. 4. Add 1 volume of formamide and 6 RNA loading dye to 20 μg of concentrated RNA (see Note 6). Heat to 100  C for 5 min and snap cool on ice. Load samples, along with 5 μL of small RNA marker and run gel at 180 V until the bromophenol blue dye front is approximately two thirds of the way down the gel. 5. Carefully remove gel from the Mini PROTEAN 3 glass plates. Stain the gel with 3 μL of ethidium bromide (see Note 5) diluted in 50 mL of water for 15 min. An image of the gel can be taken at this stage using an UV transilluminator and a fluorescent gel ruler lined up along the RNA marker lane (see Note 30). Electrophoretic Transfer

1. Cut GeneScreen membrane and four pieces of Whatman paper to the correct size for the gel. Prewet the membrane by immersing in water and then equilibrate in 1 TBE for 15 min. 2. Prewet the Mini Trans-blot sponges and Whatman paper in 1 TBE. 3. Prepare the transfer as follows: Place a prewetted sponge on the left side of the cassette. Add two pieces of prewetted Whatman paper. Carefully place the gel on this Whatman paper. Cover the gel with a piece of GeneScreen membrane and two more pieces of Whatman paper. Remove all bubbles from the system by carefully rolling a pipette across the stack. Add the second sponge to the top of the stack and close the cassette. Place the cassette into the electrophoresis unit ensuring the membrane is closest to the positive (red) electrode. Fill the apparatus with 1 TBE and run at the highest voltage possible (15–20 V) for 1 h. 4. Disassemble the transfer system and carefully remove the membrane. An image of the membrane can be taken using an UV transilluminator taking care to note the position of the RNA marker bands on the membrane using a pencil (see Note 30). Cross-link the RNA and membrane using a UV Stratalinker using the auto cross-link function (1.2  105 μJ).

Prehybridization

1. Warm the PerfectHyb buffer to 42  C and swirl to completely dissolve any precipitate. Warm the hybridization oven to 38  C. 2. Prewet the membrane with water and lightly blot to remove excess liquid. 3. Incubate the membrane in 5 mL of PerfectHyb solution in a hybridization tube with continuous rotation at 38  C for at least 30 min.

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Label Probe

1. Add 1 μL of 1 pmol/μL oligonucleotide probe, 2 μL 10 forward buffer, 2 μL γ-32P ATP, 1 μL polynucleotide kinase and 14 μL water to an eppendorf tube. 2. Mix components well and incubate at 37  C for 1 h (see Note 31). 3. About 5–10 min before the end of the probe incubation, prepare the Quick Spin columns as follows: Gently invert the column several times to resuspend the matrix. Remove the cap and break off the bottom tip. Allow the packaged buffer to drain by gravity flow and discard. Place the column in a collection tube and centrifuge at 1100  g using a tabletop centrifuge for 2 min. Repeat, discarding the eluted buffer. 4. Add 20 μL of TE buffer to the labeling reaction, mixing well. 5. Keeping the column in an upright position in a new collection tube, carefully apply the labeling reaction to the center of the column bed. 6. Centrifuge at 1100  g for 4 min and collect the eluate. 7. Add an additional 30 μL of TE to the column bed. 8. Centrifuge at 1100  g, mixing the two eluates. 9. Count 1 μL of the eluate using a scintillation counter. A typical labeling reaction should produce between 70 and 90,000 CPM/μL.

Hybridization and Washes

1. Add all of the labeling reaction to the 5 mL of PerfectHyb in the hybridization tube, making sure not to touch the membrane. Rotate at 38  C overnight. 2. The next day, rinse the blot in wash solution 1 two times at room temperature. Properly dispose of the radioactive waste (see Note 7). 3. Wash an additional two times in wash solution 1 at 38  C for 10 min each. 4. Wash the blot two times in 5 mL wash solution 2 at 38  C for 10 min each.

Film Development

1. Carefully remove the blot from the hybridization tube and remove excess solution by blotting. 2. Wrap the blot in plastic wrap and expose to autoradiography film in a dark room. An intensifier screen is useful if the miRNA of interest is expressed at low abundance. 3. Store film cassette at 80  C. The length of time for film exposure will depend on the amount of labeled probe hybridized to the blot. Normally the film can be developed after 24 h of exposure (see Note 32).

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3.2.2 Stem-Loop RT-PCR for Detection of miRNAs

Detection of miRNAs in total RNA preparations can be performed using stem-loop RT-PCR as described in [41]. miRNA species are too short for the standard primer pair probe design, thus this technique relies on the hybridization of the last 6 nucleotides of the 30 -end of the miRNA of interest to an RT primer, which lengthens the cDNA product to approximately 65 nucleotides. The stem-loop structure prevents binding of the RT primer to the pre- or pri-miRNA due to steric hindrance; therefore, no preor pri-miRNA should be amplified. miRNA-specific forward primers and probes allow for amplification of the cDNA product with both specificity and sensitivity (see Note 33). This assay requires only small amounts of input RNA and is easily amenable to highthroughput analyses. Primer and probe sets for most cellular and HCMV miRNAs are now available for purchase from commercial sources. Custom primer and probe sets can also be designed for novel viral miRNAs (see Note 34).

Anneal the RT Primer (See Note 35)

1. Mix 25 μL of 200 μM RT primer with 10 μL of 10 RT buffer and 65 μL water in a PCR tube. 2. Cycle at 95  C for 30 min, 72  C for 2 min, 37  C for 2 min— ramping down 1 degree per second from 72  C and 25  C for 2 min—ramping down 1 degree per second from 37  C. 3. The primer concentration after annealing is 50 μM. This can be frozen at 20  C (see Note 36). The working concentration is 5 μM.

Reverse Transcription

1. For each reaction, mix together the following: 0.15 μL 10 μM dNTP mix, 1 μL Multiscribe enzyme, 1.5 μL 10 RT buffer, 0.19 μL RNasin, 0.3 μL 5 μM RT primer, and 9.86 μL water. 2. Aliquot 13 μL per tube. Add up to 100 ng of RNA in 2 μL (see Note 37). 3. Cycle at 16  C for 30 min, 42  C for 30 min, and 85  C for 5 min. RT mix can be kept at 4  C for short periods of time (days). Store at 20  C for longer periods of time.

Taqman qPCR Assay

1. Mix together the following for each sample: 10 μL 2 Universal Master Mix, 7.46 μL water, 0.36 μL each of 50 μM Forward and Reverse primers, 0.32 μL 10 μM probe, and 1.5 μL RT sample (see Note 38). 2. Cycle at 50  C for 2 min, 95  C for 10 min, 95  C for 15 s, and 60  C for 1 min. Repeat the last two steps 40 times.

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3.3 Identifying mRNA Targets of Viral miRNAs 3.3.1 Photoactivatable Ribonucleoside-Enhanced Cross-Linking and Immunoprecipitation (PAR-CLIP)

One of the more challenging aspects of miRNA biology is determining which mRNA transcripts are regulated by an individual miRNA. Numerous bioinformatic prediction algorithms have been developed based on complementarity between the miRNA of interest and sites within the 30 UTRs of transcripts; however, these often lead to high numbers of false positives. In addition, many of these algorithms rely upon sequence conservation across species, and thus cannot accurately predict targets for nonconserved viral miRNAs. Techniques including microarray and SILAC have been used to identify targeted transcripts and proteins, respectively, that are differentially expressed upon the exogenous addition of the miRNA of interest or antisense inhibitors to the miRNA. However, changes in levels are often small and difficult to detect. Moreover, distinguishing direct from indirect targets of the miRNA can be difficult. Recently, biochemical approaches, including RISC-IP, PAR-CLIP, and HITS-CLIP, have been developed that directly identify targeted transcripts that have been incorporated into RISC [32, 33, 41–43]. These approaches usually identify transcripts containing canonical seed matches and further experimental validation is exceptionally successful compared with in silico prediction-based approaches [7]. This section of the chapter will outline in detail the PAR-CLIP procedure. PAR-CLIP utilizes a photoreactive nucleoside analog, 4-thiouridine (4SU), which is incorporated into newly synthesized RNAs when exogenously supplied to metabolically active cells. Upon exposure to UV light, 4SU-labeled RNAs are cross-linked to interacting RNA-binding proteins. RNAs cross-linked to RISC are then isolated by affinity purification of Argonaute (i.e., Ago2). Extracted RNA fragments are converted into a cDNA library and deep sequenced [32, 44, 45]. Bioinformatic analysis of PAR-CLIP datasets can be performed using a number of open source software tools [46–55]. There are two major advantages to using PAR-CLIP over other similar methods: (1) improved capture of RNAs and (2) the ability to reliably define signal from background RNA. Cross-linking RNAs to RNA-binding proteins increases the RNA fragments captured compared to performing a standard RISC-IP [42, 44, 45]. Because cross-linking is irreversible, more stringent conditions can be used to isolate RISC-bound RNAs than conventional IP methods. Moreover, incorporation of 4SU into nascent RNAs enhances cross-linking efficiency and leads to a characteristic T-toC mutation during generation of cDNA libraries, which can distinguish cross-linked RNAs from non-cross-linked RNAs and reduce background signal [32, 33]. When designing PAR-CLIP experiments, the cell type used and the source of the miRNA must be taken into consideration. Infection with HCMV can make detection of cellular RNA targets difficult due to the vast quantity of HCMV-encoded transcripts

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expressed during lytic infection. Lentiviral- or adenoviral-mediated miRNA expression is recommended over transient miRNA expression for PAR-CLIP experiments (described in Subheading 3.1), considering the scale required to obtain enough RNA for analysis (see Note 10). Prepare all solutions using nuclease-free ultrapure water. It is of utmost importance to perform all manipulations under RNase-free conditions to preserve the interaction between the RNA and protein and to prevent degradation of the RNA. Carry out all procedures on ice unless otherwise specified. Preparing Cell Pellets

1. Grow desired number of adherent cells expressing miRNA to ~80% confluency (see Note 10). 2. Add 4SU to a final concentration of 100 μM and treat the cells for 18–22 h. 3. Gently remove media from each dish (see Note 39) and wash cells by slowly adding 10 mL ice cold PBS to the side of the dish. Repeat wash once, leaving 10 mL ice cold PBS on each dish. 4. UV irradiate uncovered with 150,000 mJ/cm2 total energy of 365 nm UV light in a UV Stratalinker (see Note 40). Remove cells from dish by pipetting and place into a prechilled 50 mL conical on ice. 5. Repeat steps 4 and 5 for the remaining dishes. 6. Centrifuge at 1000  g for 5 min at 4  C (see Note 41). 7. Remove the supernatant, resuspend pellets in ice cold PBS, and combine pellets from multiple tubes into a single 15 mL conical tube. Centrifuge at 1000  g for 5 min at 4  C. 8. Remove excess PBS and flash-freeze cell pellets, storing at 80  C until ready to process.

Cell Lysis and Preparation of Magnetic Beads

1. Thaw pellet(s) on ice until completely thawed (see Note 42). 2. Prepare the magnetic beads for immunoprecipitation. Thoroughly mix Dynabeads and transfer 10 μL beads per 1 mL of cleared lysate to a 1.5 mL Eppendorf tube (see Note 43). 3. Place the tube on a magnetic particle concentrator to collect beads. Remove the supernatant and wash beads with 1 mL lysis buffer. Resuspend the beads in lysis buffer two times the original bead volume. 4. Add Ago2 antibody (see Note 9) to the beads at a volume of 10 μL per 90 μL original bead volume (see Note 43). Rotate for at least 1 h at 4  C, or until ready to perform immunoprecipitation.

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5. Collect beads on the magnetic rack. Remove the supernatant and wash the beads twice with 1 mL lysis buffer. Resuspend beads in two bead volumes of lysis buffer. 6. Once cell pellet is completely thawed on ice, resuspend the cell pellet in 3 cell pellet volumes of lysis buffer (see Note 43). Incubate cell lysate on ice for 10 min. 7. Centrifuge at 12,000  g for 10 min at 4  C (if using a tabletop centrifuge, transfer lysates to 1.5 mL Eppendorf tubes after the lysis step). Save 25 μL of the supernatant at 20  C for Western Blot analysis (see Note 44). 8. Filter lysate through a 5-μm membrane syringe filter and collect in a 15 mL conical tube on ice. RNase T1 Treatment and Immunoprecipitation

1. Adjust a water bath to 22  C (see Note 45). 2. Add RNase T1 to the cleared lysates to a final concentration of 1 U/μL. 3. Incubate reaction in the 22  C water bath for 15 min, manually mixing every 5 min (see Note 45). 4. Stop the reaction by placing the tube on ice for 5 min. 5. Thoroughly resuspend the prepared magnetic beads in lysis buffer and add to the RNase T1–treated lysates. Incubate for >1 h at 4  C on a rotator. 6. Place the magnetic rack on ice. Place one empty 1.5 mL Eppendorf tube per sample onto the rack. Transfer 1 mL of lysatebead suspension at a time into the tube and collect the beads. Discard the supernatant. Repeat until all of the beads from the 15 mL suspension are collected into the 1.5 mL Eppendorf tube. 7. Wash the beads three times with 1 mL IP wash buffer. Gently pipet to mix. Place the tube back on the magnetic rack to collect the beads and discard the supernatant between washes. 8. Remove excess wash buffer with a pipette and resuspend beads in the original bead volume of IP wash buffer. 9. Add RNase T1 to a final concentration of 100 U/μL. Incubate reaction for 15 min in a 22  C water bath. Manually mix the tube every 5 min (see Note 45). 10. Stop the reaction by placing the tube on ice for 5 min. 11. Place the tube on the magnetic rack, collect beads, and discard the supernatant. Wash beads three times with 1 mL of high salt wash buffer as in step 7.

Dephosphorylation, Labeling, and Phosphorylation

1. Resuspend beads in the original bead volume of 1 NEB buffer 3 containing CIP at a final concentration of 0.5 U/μL. Incubate at 37  C for 10 min to dephosphorylate RNAs. Manually mix every 3–5 min.

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2. Place the tube on the magnetic rack, collect beads, and discard the supernatant. Wash the beads twice with 1 mL of cross-link wash buffer. 3. Place the tube on the magnetic rack, collect beads, and discard the supernatant. Wash the beads twice with 1 mL of PNK buffer containing DTT. 4. Place the tube on the magnetic rack, collect beads, and discard the supernatant. Resuspend the beads in the original bead volume of PNK buffer containing DTT. 5. Add ATP to a final concentration of 100 μM and incubate at 37  C for 30 min. Manually mix every 10 min. 6. Place the tube on the magnetic rack, collect beads, and discard the supernatant. Wash beads five times with 1 mL of PNK buffer without DTT. 7. Place the tube on the magnetic rack, collect beads, and discard the supernatant. Resuspend beads in 65 μL of 1 SDS loading buffer. Add SDS loading buffer to the pre-IP sample (saved at 20  C) (see Note 44). 8. Boil the samples for 5 min at 95  C, then vortex beads for 10 s to release immunopurified Ago-RNA complexes into the loading buffer. At this point, either proceed to the SDS-PAGE step or save the samples at 80  C. SDS-PAGE and Electroelution

1. Collect the beads on the magnetic rack and transfer the supernatant (containing immunopurified Ago-RNA complexes) into a new tube. 2. Prepare a Criterion midi electrophoresis gel apparatus with a 4–12% Bis-Tris Protein gel and fill with 1 MOPS running buffer. 3. For each sample, load 2 lanes of the gel with the IP sample (30 μL in each well) and load adjacent lanes with a protein marker. Leave empty lanes between different samples or run separate gels. On the opposite side of the gel, load pre-IP and IP samples plus a protein marker for Western blot analysis (see Note 44). Run the gel at 100 V for 45–60 min until the bromophenol blue loading dye reaches the bottom of the gel. 4. Using a clean razor blade, excise the region of the gel corresponding to the expected size of the RNA-binding protein (i.e., for Ago2, ~100 kD) (see Note 44). 5. Prepare a D-tube dialyzer tube by adding 800 μL 1 MOPS buffer to the inner chamber and incubating for 5 min at room temperature. Carefully remove the MOPS buffer without piercing the membrane. Transfer the excised gel band plus 800 μL fresh 1 MOPS buffer to the dialyzer tube.

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6. Prepare an agarose gel chamber with an electroelution rack and fill the chamber with enough buffer to fully cover the dialyzer tube. Place the tube in the electroelution rack, oriented so that the membrane of the dialyzer tube is perpendicular and exposed to the flow of current. Run at 100 V for 1.5 h. 7. Reverse current for 2 min to release protein from the membrane. Transfer the 1 MOPS solution from the inner chamber of the dialyzer tube to two Eppendorf tubes with 400 μl in each tube. RNA Extraction

1. Following dialysis, add 1 volume of 2 proteinase K buffer plus proteinase K to a final concentration of 1.2 mg/mL to each Eppendorf tube containing RNA-binding protein complexes. Incubate at 55  C for 30 min and invert to mix every 5–10 min. 2. Perform phenol–chloroform extraction to purify RNAs. To each tube, add 1 volume of acidic phenol–chloroform–isoamyl alcohol (25:24:1), vortex, and centrifuge at 12,000  g for 15 min at 4  C. Transfer the upper aqueous phase to a new tube. 3. Add one equal amount of chloroform, vortex, and centrifuge at 12,000  g for 10 min at 4  C. Transfer the upper aqueous phase to a new tube. 4. Precipitate the RNAs by adding 1/10 volume 3 M NaCl, 1 μL of GlycoBlue (see Note 11), and 3 volumes 100% ethanol. Incubate for 30 min to 1 h at 80  C or overnight at 20  C. Pellet RNAs by centrifugation at 16,000  g for >30 min at 4  C (see Note 46). 5. Carefully discard the supernatant. Wash RNA pellets once with ice cold 95% EtOH and centrifuge at 16,000  g for 10 min at 4  C. 6. Carefully discard the supernatant, briefly air-dry RNA pellets on ice (1–2 min), and resuspend in a total volume of 10 μL of nuclease-free H2O. At this point, RNA can be stored at 80  C.

Generation of cDNA Libraries

cDNA Library purification (See Note 47)

PAR-CLIP sequencing libraries are generated using 5 μL of immunopurified RNA and commercially available kits, such as the Illumina TruSeq Small RNA Library Prep kit. Adaptor ligation, reverse transcription, and PCR amplification are performed according to the manufacturer’s instructions (see Notes 12 and 47). 1. Prepare a Criterion midi gel apparatus with a 5% TBE polyacrylamide gel and fill with 1 TBE buffer. Add DNA loading dye to each sample load cDNA libraries along with a 25-base pair DNA ladder. Run at 100 V for 1 h.

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2. Carefully remove the gel and stain with 20 μL SYBR Safe diluted in 30–40 mL TBE buffer and gentle shaking for 10 min. Visualize on a UV transilluminator. 3. Using a clean razor blade, excise the region of the gel containing the sample between 120–200 base pairs and place into a new tube. 4. Add 500 μL 0.5 M NaCl and rotate at 4  C overnight. 5. Centrifuge at 12,000  g at 4  C for 10 min to pellet gel pieces. Transfer the supernatant to new tube. 6. Add 2–3 volumes of 100% EtOH. Add 1 μL GlycoBlue (see Note 11). Precipitate the cDNA at 80  C overnight. 7. Centrifuge at 16,000  g for 30 min at 4  C. Carefully discard the supernatant, making sure not to disturb cDNA pellet. 8. Wash the pellet with 1 mL ice cold 95% EtOH. Centrifuge at 16,000  g for 15 min at 4  C. 9. Discard the supernatant and air dry the pellet. 10. Resuspend in nuclease-free water. Determine the concentration and purity using an Agilent Bioanalyzer or similar (see Note 48). 11. Submit purified cDNA for Illumina sequencing. 3.3.2 RNA-Induced Silencing Complex (RISC) Immunoprecipitation (IP)

RISC-IP takes advantage of the complex that is formed when a miRNA within the RISC complex interacts with its target. Affinity purification of the RISC complex followed by RNA isolation and microarray or deep sequencing analysis of bound transcripts leads to the direct identification of targeted transcripts. While PAR-CLIP is a more powerful tool for the identification of miRNA targets, RISP-IP is less costly and time-consuming than PAR-CLIP analysis. As such, RISC-IPs can be used to validate the enrichment of known target(s) during infection or in the presence of exogenous miRNA or alternatively can be used to show de-enrichment of a target upon infection with a miRNA mutant virus or in the presence of miRNA inhibitors. Several considerations must be taken into account when designing a RISC-IP screen including the source of miRNA, whether through infection or exogenous expression (described in Subheading 3.1), and the methods of isolating the RISC complex, for example by using antibodies directed against one of the RISC components (eg-Ago1, Ago2, Ago3, or Ago4), cell lines expressing tagged components of the RISC complex (eg-c-myc-taggedAgo2), or miRNAs that have been biotinylated (described in Subheading 3.1.3).

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Prepare all solutions using nuclease-free ultrapure water. It is of utmost importance to perform all manipulations carefully to preserve the interaction between the RNA and protein and to prevent degradation of the RNA. Carry out all procedures on ice unless otherwise specified. Preparing the Cell Lysate

1. Determine the number of tubes and tissue culture scrapers needed to collect cell lysates and place on ice. Prepare DPBS 1 and lysis buffer and place on ice until cold. Immediately before lysing cells, remove a small aliquot of lysis buffer (enough for 1 mL/100 mm tissue culture dish) and add RNasin Plus RNase Inhibitor to a final concentration of 1 U/μL. Place on ice. 2. Gently remove media from one 100 cm tissue culture dish using an aspirating pipette. Keeping the dish in an upright position, remove the remaining media with a P1000 pipette. 3. Wash cells by slowly adding 10 mL ice cold DPBS to the side of the dish and gently tip to cover all cells. Remove DPBS wash as in step 2. 4. Add 1 mL ice-cold lysis buffer plus RNasin Plus RNase Inhibitor to the dish covering the cells and scrape to collect cells and buffer. Work quickly to minimize time spent at room temperature. Collect lysate and transfer to an ice-cold tube and place on ice. 5. Repeat steps 2–4 to harvest the remaining dishes. 6. Vortex each tube at maximum velocity for 10 s and return to ice for 10 min. Vortex again for 10 s and then centrifuge at 12,000  g for 15 min at 4  C. While spinning, place additional tubes on ice to chill. 7. Carefully transfer the supernatant to the new chilled tubes and place on ice. Remove 1/20 of the volume to be used as the total RNA sample and transfer to a tube containing 500 μL TRIzol. Vortex and freeze at 80  C. Removing a small aliquot of cell lysate at this step to serve as the total protein sample is also recommended (see Note 49).

Preparing the Streptavidin-Agarose, ANTI-c-myc Agarose Conjugate, or Protein A Beads

1. Thoroughly mix the Streptavidin-Agarose, ANTI-c-myc Agarose, or protein A to resuspend the beads and transfer 25 μL/ IP sample to a single tube. Centrifuge at 12,000  g for 30 s and discard supernatant. 2. Wash the beads by adding 500 μL lysis buffer without RNasin, gently mix, and centrifuge at 12,000  g for 30 s and discard supernatant. Repeat wash and resuspend beads in lysis buffer without RNasin (100 μL/IP sample). For each 100 μL, add 10 μL tRNA (10 mg/mL) and 10 μL BSA (10 mg/mL). Rotate 2 h at 4  C to block. Once the beads are rotating in

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block, begin the immunoprecipitating section below if isolating the RISC complex using an antibody against one of the components. 3. Wash the beads by adding 300 μL lysis buffer, gently mix, and centrifuge at 12,000  g for 30 s and discard supernatant. Repeat for a total of two washes, and resuspend in lysis buffer (100 μL/IP sample). Place on ice. Immunoprecipitating

1. If isolating the RISC-complex using antibodies directed against one of the components, add the antibody to the cell lysates from step 7 in Subheading “Preparing the Cell Lysate” (see Note 9). If using Streptavidin-Agarose to purify biotinylated miRNAs or ANTI-myc Agarose to purify myc-tagged Ago2, proceed to the pull-down section below. 2. Rotate 2 h at 4  C. Proceed to the pull-down section.

Pull-Down

1. Gently mix the Streptavidin-Agarose, ANTI-c-myc Agarose Conjugate or protein A from step 3 in Subheading “Preparing the Streptavidin-Agarose, ANTI-c-myc Agarose Conjugate, or Protein A Beads” to resuspend beads and add 100 uL to the cell lysates. 2. Rotate for 2 h at 4  C. 3. Centrifuge for 1 min at 5000  g 4  C. 4. Carefully remove supernatant and resuspend in 500 μL lysis buffer. Centrifuge for 1 min at 5000  g 4  C. Repeat wash three additional times for a total of four washes. After the third wash, transfer suspended beads to a new chilled tube for the final wash. 5. After the final wash, leave the beads in approximately 50 μL lysis buffer. Removing a small aliquot of the immunoprecipitate at this step can serve as a control to ensure efficient immunoprecipitation of RISC complex proteins like Ago (see Note 50). Add 500 μL of TRIzol to the beads and remaining lysis buffer, mix well, and freeze at 80  C until ready to process.

RNA Isolation from Pull-Downs

Adapted from TRIzol method for RNA isolation. 1. Add 0.1 mL chloroform to each total and IP sample, shake vigorously for 15 s, and incubate for 2–3 min at room temperature. 2. Centrifuge the samples at no more than 12,000  g for 15 min at 4  C. 3. Transfer the aqueous phase to a fresh tube. 4. Precipitate RNA by adding 0.25 mL isopropanol and 1 μL GlycoBlue Coprecipitant (see Note 11), mix well, and incubate for 10 min at room temperature.

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5. Centrifuge samples at no more than 12,000  g for 15 min at 4  C. 6. Remove supernatant and wash the pellet once with 0.5 mL of 75% ETOH. Mix by vortexing and centrifuge at no more than 7500  g for 5 min at 4  C. 7. Repeat step 6 once for a total of two washes (see Note 51). 8. Completely remove the wash, dry briefly, and resuspend IP and total samples in 15 μL and 50 μL nuclease-free water, respectively. 9. Determine RNA concentration and purity by using an Agilent Bioanalyzer or similar (see Note 48). Analysis

1. Determine the success of the affinity purification by using a Western blot analysis to compare the amount of RISC proteins isolated in the purified fraction versus those in the total sample (see Note 52). Alternatively, the enrichment of the miRNA and a validated mRNA (if known) in the purified fraction compared with the total fraction can be determined by Taqman RT-PCR (see Subheading 3.2.2) (see Note 53). 2. Determine transcript levels in total and IP samples by microarray using an appropriate microarray platform and analysis software (see Note 54). 3. Determine fold-enrichment to identify potential miRNA targets. Cellular mRNA transcripts that are associated with the RISC complex are identified by dividing the amount of mRNA transcripts in the IP fraction by the amount of mRNA in the total RNA fraction in order to account for direct effects of the miRNA expression on transcript levels within the cell. To identify cellular mRNA transcripts specifically enriched within the RISC-complex containing exogenously added miRNA and to exclude those transcripts isolated by association with cellular miRNAs (this is particularly important when transfecting in miRNAs and controls using an expression vector and immunoprecipitating or pulling down via a component of the RISCcomplex), the enrichment profile in miRNA transfected cells is compared to cells transfected with the negative control such that the enrichment value of any given mRNA transcript ¼ (IPmiRNA/TotalmiRNA)/(IPNeg/TotNeg) [30]. A similar determination is used when pulling down RISC using biotinylated miRNAs in conjunction with streptavidin-agarose. mRNA transcripts are then ranked according to level of enrichment with the highest enriched transcripts considered potential targets of miRNA.

Characterizing Cytomegalovirus-ncoded miRNAs 3.3.3 Reporter Assays for Identification of Viral miRNA Targets

Transfection

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Reporter assays have proven invaluable in evaluating gene expression and have been adapted and utilized extensively to identify and characterize potential miRNA targets. Typically, the 30 UTR of a predicted miRNA target is cloned downstream of a reporter gene and is cotransfected into cells along with the miRNA of interest or negative control miRNA. Decreases in reporter activity in cells transfected with the miRNA relative to cells transfected with the negative control indicate miRNA-mediated regulation. A wide variety of reporter constructs and detection systems are commercially available. Carry out all procedures at room temperature, unless otherwise specified. 1. Plate HEK293 cells at 1–2  104 cells/well in 100 μL media in a black-walled clear-bottom 96-well tissue culture treated plate (see Note 55). 2. 24 h later, prepare (per well): 100 ng luciferase reporter vector plus 0.3–0.4 pmol miRNA in 25 μL Opti-MEM I Reduced Serum Media (see Note 56) 3. For each well, add 0.4 μL Lipofectamine 2000–25 μL OptiMEM I Reduced Serum Media. A mastermix is recommended for multiple samples to ensure that a similar amount of transfection reagent is added to each well. Gently mix by inverting and incubate for 5 min at room temperature (see Note 57). 4. Add 25 μL of the Lipofectamine 2000 mastermix in step 3 to the luciferase reporter and miRNA in step 2, gently mix, and incubate for 20 min at room temperature. 5. Transfer the 50 μL reaction in step 4 to each well dropwise and gently swirl plate. Incubate plate 12–24 h at 37  C, 5% CO2.

Luciferase Assay

1. Carefully remove media and lyse cells by adding 20 μL 1 Passive Lysis Buffer to each well (see Note 58). Incubate the plate for 15 min on rocking platform at room temperature. 2. Prepare the necessary quantities of Luciferase Assay Reagent II (LAR II) and Stop & Glo reagents according to the manufacturer’s recommendations (50 μL/well + 0.5 mL extra) (see Note 59). 3. Prepare a plate-reading luminometer with two injectors according to the manufacturer’s suggestions. Measurements are performed for each well sequentially as follows: inject 50 μL LAR II, delay for a 2 s preread, measure firefly luciferase activity for 2 s, inject 50 μL Stop & Glo, delay for a 2 s preread, and measure Renilla luciferase activity for 2 s (see Note 60). 4. Calculations: For each sample, calculate the ratio of Renilla signal/firefly luciferase signal where Renilla is the test reporter and firefly luciferase is the control. Then calculate the average

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and standard deviation among replicates (see Note 61). Decreases in reporter activity in cells transfected with the miRNA relative to cells transfected with the negative control indicate miRNA-mediated regulation. 3.4 Inhibition of HCMV miRNAs Using Locked Nucleic Acids (LNA)

Several miRNA inhibition strategies based on antisense nucleic acids have been developed including miRNA sponges, decoys, and antagomirs [56]. Each contains sequence complementary to the mature miRNA and prevents the miRNA from binding its intended target. Chemical modifications of these inhibitors, such as the substitution of phosphate bonds with phosphorothioate bonds, methylation of the oxygen at position 2 in the ribose (20 -O-methyl), and the addition of an extra bridge between the carbons of the ribose ring (locked nucleic acid, LNA) have greatly increased their stability, affinity, and specificity. Among the chemically modified species, LNAs have recently emerged as the most efficient miRNA inhibitors due to their potency, versatility, and efficacy both in vitro and in vivo [57, 58]. For instance, several clinical trials using LNAs as small as 8 nucleotides have demonstrated strong efficacy against hepatitis C virus in primates in vivo [59]. These findings have been confirmed in our lab, with LNAs designed against HCMV miRNAs displaying potent, versatile and reliable inhibitory activity (data not shown). The following protocol describes how LNAs can be used to inhibit a miRNA mimic in the context of a luciferase assay. LNAs can also be used to antagonize miRNAs derived from the cell or virally expressed miRNAs following infection, though significant optimization is required. Inhibiting the expression of HCMV miRNAs individually or in combination through the use of LNAs is an important tool to evaluate the roles of miRNAs during infection, complementing the more time-consuming generation of miRNA mutant viruses. Carry out all procedures at room temperature. The following protocol is designed for HEK293 cells in a 96-well plate format and can be adapted for other cell types and tissue culture formats (see Notes 62–66). 1. Seed 1  104 HEK293 cells/well in a black-walled 96-well plate (see Note 62) in 0.1 mL medium. 2. 24 h later, prepare (per well): 0.3 pmol miRNA, 0.6 pmol LNA, and 100 ng luciferase reporter vector in 25 μL OptiMEM I Reduced Serum Media (see Note 63). 3. For each well, add 0.25 μL Lipofectamine 2000 to 25 μL OptiMEM I Reduced Serum Media (see Notes 64 and 65). A mastermix is recommended for multiple samples to ensure that a similar amount of transfection reagent is added to each well. Gently mix by inverting and incubate for 5 min at room temperature.

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4. Combine the reaction mixes from steps 2 and 3, mix gently, and incubate for 20 min. 5. Add mix dropwise into wells for a final volume of 150 μL and gently swirl plate to distribute. Incubate plate at 37  C, 5% CO2. 6. Perform luciferase assay 12–24 h posttransfection (see Subheading 3.3.3) (see Note 66).

4

Notes 1. We use the pSIREN-retroQ-ZsGreen for CMV miRNA expression because (1) the promoter is a RNA polymerase III–dependent promoter not derived from CMV, and (2) it constitutively expresses GFP allowing one to easily monitor transfection efficiency. 2. Cloning the viral region surrounding the miRNA locus allows for the expression of the full pri-miRNA and the formation of a native hairpin. Mature miRNAs are considered to be more efficiently processed from a full hairpin than from a truncated one. Since the exact extent of the pri-miRNA is not known in most cases, comparing clones encompassing regions of different sizes around the miRNA locus allows for selection of the region that best expresses the miRNA in one round of cloning. 3. This transfection protocol has been optimized for HEK293 cells. miRNAs can be transfected using other methods (including electroporation) and/or in other cell lines but each approach must be individually optimized. 4. SDS powder should be handled in a manner to minimize formation of aerosols. Wear proper protective equipment including a facemask when measuring SDS powder. 5. Ethidium bromide is an agent that intercalates into nucleic acids and is therefore believed to be a mutagen. Although the concentrations used in the laboratory are not associated with health risks, ethidium bromide should be handled with care. Wear proper protective equipment when handling ethidium bromide. 6. Due to the small well size of the Mini PROTEAN gel system (wells can hold approximately 40 μL), RNA should be concentrated to at least 1 μg/μL. 7. When using and disposing of radioactive materials, be sure to follow all federal, state, and local regulations. Make sure to wear proper protective equipment. 8. Proper RNA handling techniques should be observed at all times to prevent degradation.

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9. PAR-CLIP and RISC-IP has been performed successfully using commercially available antibodies against Ago2 and RISC-IP has also been performed successfully using serum raised against Ago2 in rabbits. Optimization is required to determine the concentration of antibody required for optimal immunoprecipitation. 10. Typically, 2–5 mL of wet cell pellet is required for PAR-CLIP experiments. This protocol has been optimized for 20–50,150 mm cell culture dishes of HEK293 cells transduced with a lentiviral expression vector to stably express the miRNA of interest (see Note 20). Optimization is required for other cell types and/or methods of miRNA expression. 11. The addition of GlycoBlue during precipitation aids in visualization of the pellet during washing. However, addition will interfere with the quantification of RNA unless the same concentration of GlycoBlue is included in the buffer used to blank the spectrophotometer. 12. For the TruSeq Small RNA Library Prep kits, there are four kits available (sets A–D) that work with this protocol. These kits are identical except for the 12 indices supplied, which allow for pooling samples and multiplexing during RNA-seq analysis. If multiplexing is desired, use a different index for each sample. 13. T4 RNA Ligase 2, deletion mutant is not supplied with the TruSeq small RNA Library Prep kit. 14. A pilot experiment should be performed to determine the number of 100 mm plates required to obtain a sufficient amount of RNA for subsequent analysis by microarray. 15. Some HCMV miRNA mimics are commercially available for purchase as mature miRNAs or as pre-miRNA hairpins. If unavailable, miRNAs can be cloned into expression vectors as described in Subheading 3.1.1 or can be designed and custommade by a variety of sources as described in Subheading 3.1.3. 16. A variety of reporter vectors are commercially available. DualLuciferase reporters are highly recommended as the effects of the test reporter can be compared to the control reporter. The control reporter provides an internal control that takes into account well-to-well variation due to differences in cell viability, transfection efficiency, pipetting error, and assay efficiency. 17. Performing experiments in a 96-well format is highly recommended. While this protocol is designed for 96-well plates, it can be scaled up for other cell culture formats. 18. For increased accuracy and high-throughput analyses, the use of a plate-reading luminometer equipped with two reagent injectors is highly recommended. If injectors are not available, perform injections manually and read one well at a time (see Note 60).

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19. In addition to negative controls, Exiqon has designed LNAs for all of the HCMV miRNAs deposited in mirBase. LNAs can also be custom made to target novel miRNAs or ones not present in mirBase. 20. The strategy for cloning a miRNA into a lentiviral vector is very similar to the pSIREN and adenoviral vector cloning strategy described here. Refer to the manufacturer’s instructions for cloning details. Make sure to choose a lentiviral vector where miRNA expression is driven by a promoter that is not derived from CMV, since this could affect the level of expression in infected cells. 21. Purify viral DNA by isolating viral particles (e.g., using sorbitol cushion method) followed by DNA extraction (e.g., DNAzol or phenol–chloroform). Alternatively, total DNA from virusinfected cells can be isolated and used as template. 22. If the PCR reaction displays several amplification products, gel-purify the correct band using a DNA gel purification kit. 23. In our hands, transfection efficiency in HEK293 cells is close to 100% as estimated by GFP expression. 24. Transfection of suspension cells is more challenging than adherent lines such as HEK293 cells. Nucleofection is a significant improvement over lipid-based methods for suspension cells (i.e., Kasumi-3 or THP-1) or primary cells (i.e., CD34+ HPCs, monocytes), but requires optimization and experimental validation to balance nucleofection efficiency with cell survival. Lonza offers optimization kits which allow comparison of multiple buffers and a selection of nucleofection conditions which provides a good starting point for experimental optimization. 25. The pAdTrack-CMV vector contains a limited multiple cloning site with alternative restriction sites in addition to NotI and XhoI. 26. It is best to prepare the pAdEasier electrocompetent cells the same day you will use them. 27. Prior to generation of a high titer viral stock, it is worthwhile to ensure expression of the miRNA of interest. Viral DNA can be isolated from the supernatant of infected cells and used to amplify the region of interest for sequencing. Additionally, RNA can be isolated from a small-scale infection and miRNA levels measured using stem-loop qRT-PCR (see Subheading 3.2.2). 28. The introduction of a mutation in the fourth nucleotide from the 30 -end of the antisense strand results in the preferential incorporation of the mature or guide strand into the RISC complex.

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29. miRNAs that are biotinylated can be used in subsequent protocols including RISC-IP to identify miRNA targets. Only the mature strand should be biotinylated. 30. Carefully line up your gel and membrane with the fluorescent ruler when taking images using the UV transilluminator. The small RNA marker generally does appear on the membrane, and the bands should be clearly noted based on the position of the ruler for future reference. 31. This step can be performed during the prehybridization of the membrane. 32. Often, both pre- (~60 nucleotide) and mature (~22 nucleotide) forms of a miRNA can be detected by Northern blotting, so noting the position of the RNA markers is important (see Fig. 2). Commonly, radioactive signal can also be detected at the very top of the membrane. This signal likely represents the larger sized pri-miRNA transcripts from which the miRNA precursors are derived. 33. In these assays, expression of each miRNA is assessed using a unique RT primer. The efficiency of the RT reaction for each miRNA is unknown and therefore the expression of multiple miRNAs relative to one another cannot be accurately assessed. 34. To design custom primers and probes, it is very important to accurately determine the sequence of the mature miRNA, especially with regards to the 30 end. The RT primer is designed using the sequence GTCGTATCCAGTGCAGGGTCCGAGG TATTCGCACTGGATACGAC . For each miRNA, the final additional 30 nucleotides of the RT primer are unique and antisense to the last 6 nucleotides of the mature miRNA. The forward primers are designed as the first 15 nucleotides of the mature miRNA with 3–5 additional nucleotides at the 50 -end to result in a final annealing temperature of approximately 60  C. The reverse primer is the same for each reaction (GTG CAGGGTCCGAGGT ) and binds within the RT primer sequence. Finally, the probe sequence used with the RT primer outlined above has a sequence of TGGATACGAC followed by six nucleotides designed antisense to the last nucleotides of the mature miRNA (identical to the RT primer). The probe should be synthesized with a 50 fluorescein (FAM) moiety, and minor groove binder (MGB) nonfluorescent quencher on the 30 -end [41]. Examples of custom-designed primer and probe sets can be found in [6]. 35. If Taqman RT-PCR assays are purchased from Applied Biosystems, RT primers do not need to be annealed. 36. We have found that 50 μM annealed primers are very stable through multiple freeze–thaw cycles.

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37. Mock-treated samples are an important control for the RT-PCR reaction. Cross-reactivity of a viral primer and probe set with a cellular miRNA is unlikely but has been observed [6]. In addition, copy number for each miRNA can be estimated by preparing an RT reaction using a known concentration (106 to 108) of HPLC-purified oligonucleotide with the identical sequence to your miRNA of interest. If generating a standard curve using a miRNA duplex, denature the RNA prior to reverse transcription by incubating at 85  C for 5 min followed by 60  C for 6 min. 38. To determine the copy number of each miRNA in your samples, perform a five-step ten-fold serial dilution of the RT reaction containing a known number of copies of the oligonucleotide standard (usually 106 to 109 copies to be determined for each miRNA depending on their abundance). Prepare a Taqman reaction for each serial dilution and generate a standard curve by plotting Ct value versus copy number. Determine the copy number of the miRNA in each sample using the standard curve. 39. Confluent HEK293 cells are adherent but are easily detached from cell culture dishes. It is recommended to pour off cell culture media and slowly pipet PBS to the side of the dish to reduce loss of cells prior to cross-linking. 40. Work with six 150 mm cell culture dishes at a time in order to fit all of the dishes in the UV Stratalinker for the crosslinking step. 41. Prechill the centrifuge to ensure that cells stay cold throughout the centrifugation steps. 42. This protocol can easily be performed with four cell pellets at one time. If there are more than four samples, it is recommended to stagger cell lysis steps and immunoprecipitation steps or perform multiple experiments to minimize the number of samples handled at one time. 43. For 3 mL of wet cell pellet, 9 mL of lysis buffer and 90 μL beads + 10 μL antibody will be used. Adjust accordingly for a different volume of cell pellet. 44. It is recommended to perform Western blot analysis in parallel to SDS-PAGE of the IP samples in order to ensure successful immunoprecipitation and that the correct portion of the gel is excised for electroelution. This should be done in the same gel so that the presence of Ago2 in the Western blot reflects the region of the gel containing the immunoprecipitated samples excised for electroelution. If performing a Western blot, load the following on the opposite side of the gel from the IP samples: a protein marker, the remaining 5 μL of the IP sample, and 15–20 μL of the pre-IP lysate (saved from step 7 of

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Subheading “Cell Lysis and Preparation of Magnetic Beads” in Subheading 3.3.1). Perform a Western blot for Ago2 on this portion of the gel using standard practices. Ago2 should be detected in the ~100 kD region. As the Western blot will likely not be done until after the electroelution step, the rest of the gel may be saved at 20  C until confirmation that the correct gel region was excised. 45. The timing and temperature of RNase treatment are critical. Over digestion can lead to RNA fragments that are too small for RNA-seq analysis. 46. If a pellet is not observed, repeat the centrifugation. An overnight precipitation at 80  C is recommended. 47. Amplification of cDNA libraries should be in the linear range to ensure library complexity and to avoid overamplifying highly expressed RNAs. Therefore, a pilot PCR is performed with 50% of the ligated sample (equally distributed between four or five PCR tubes) to determine the necessary cycle number (e.g., 15, 18, 21, 24 cycles). The final PCR is performed using the same conditions, but adjusting the cycle number as predetermined. 48. Determine whether the quality, concentration, and total amount of RNA are sufficient for subsequent analyses. Highquality RNA is required for the labeling reactions. The quantity and purity of each RNA sample is analyzed by applying a small amount for analysis on a bioanalyzer, which determines purity based on the ratio of the 28S to 18S ribosomal subunits. However, since RNA fractions obtained from RISC immunoprecipitation and pull-down based methods are not expected to contain ribosomal RNAs, basing the quality and purity of RNA on this ratio can be misleading. 49. The total protein sample can be compared with a protein sample taken after affinity purification to confirm the success or troubleshoot the RISC-IP. Western blot analyses for the immunoprecipitated protein or another component of the RISC complex can be performed comparing the total and IP protein samples with the expectation that components of the RISC complex will be pulled down. 50. The IP protein sample can be compared with the total protein sample by Western blot as described above in Note 49. 51. Two washes are required to remove any residual phenol–chloroform; residual phenol–chloroform will interfere with the subsequent steps required for microarray analysis. 52. Beads can be suspended directly in protein sample buffer for analysis by Western blot.

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53. These steps are helpful in determining the success of the procedure. If the RISC-IP procedure was performed successfully, components of the RISC-complex will be detected in the IP sample by Western blot and the miRNA and any known targets will be enriched in the IP sample as detected by stem-loop RT-PCR and real-time PCR, respectively. The entire RISC-IP procedure can also be performed on a small scale and used to confirm potential miRNA targets. For example, to confirm potential targets identified by a large RISC-IP screen, the RISC-IP can be repeated on a smaller scale followed by stemloop RT-PCR, to confirm the incorporation of the miRNA into the RISC complex and subsequent recruitment of the target mRNA. 54. Consulting with your institutional microarray core regarding amount and purity of RNA required, array platform, and postarray normalization, as well as overall experimental design is recommended. 55. Cells should be no greater than 30–50% confluent at the time of transfection. HEK293 will adhere to plastic and remain adherent, however steps should be taken to minimize washing or replacing media. If using a transfection reagent other than Lipofectamine 2000, consult the manufacturer’s guide in respect to cell plating density, amount and type of miRNA to use, and general protocol. 56. The amount of miRNA and vector used may need to be optimized on an individual basis. When transfecting two or more miRNAs simultaneously, the final concentration of miRNA in all samples should be equivalent. A negative control or GFP siRNA can be used to ensure that the same amount of miRNA or siRNA is present within each well. Transfecting more than one miRNA expression vector per well is not recommended. 57. Continue to the following step within 20 min. 58. Prepare 1 Passive Lysis Buffer from the 5 stock provided in the Dual-Luciferase Reporter Assay System by diluting 1 part 5 stock in 4 parts distilled water and mixing well. At the time of lysis, cells should be no more than 95% confluent to achieve optimum lysis. 59. Prepare LAR II by resuspending the LAR II substrate in 10 mL Luciferase Assay Buffer II, aliquoting in small usable quantities, and freezing up to 1 year at 70  C. We regularly prepare Stop & Glo by combining the vial of Stop & Glo substrate with the bottle of Stop & Glo buffer, mixing well, aliquoting in small usable volumes, and freezing at 70  C. For small-scale assays, resuspend the Stop & Glo substrate in the Stop & Glo buffer at a ratio of 1:50.

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60. The intensity of the firefly and renilla luciferase signal varies depending on the cell line, the amount of reporter plasmid, the transfection efficiency and other factors. The measurement times may need to be optimized and can range from 1 to 10 s or more. Keep a preread delay of 2 s after injection and the same measurement time for firefly and renilla luciferase. If performing injections manually, proceed one well at a time: inject LAR II reagent and read the firefly luciferase signal for 12 s after a 3 s delay, then inject Stop & Glo reagent and read the renilla luciferase signal for 12 s after a 3 s delay. 61. As the effects of miRNAs are often modest, performing assays in triplicate and running multiple experiments is highly recommended. 62. Fibroblasts and THP-1 cells are seeded at 1  104 cells/well in a 96-well format. For THP-1 cells, differentiation and plate attachment can be induced by adding 50 ng/mL phorbol-12myristate-12-acetate (PMA) to the culture media. 63. When cotransfecting an LNA with a miRNA mimic, use a 1:1 to 10:1 LNA–miRNA concentration ratio. Controls consist of combinations of nontargeting miRNAs, negative control (NEG), and LNAs (see Fig. 3 for an example of controlled experiment). 64. miRCURY LNA™ microRNA Power Inhibitors enter HEK293 cells even in the absence of transfection reagent and can be added directly to the cell culture before or after the transfection of the other components, and at the same

Fig. 3 LNA-US5-2 inhibits miR-US5-2 targeting of US7 30 UTR. The pSICHECK2-US7 30 UTR dual luciferase reporter construct (which contains a target site for miR-US5-2 that efficiently decreases luciferase expression) was cotransfected with the indicated combinations of miR-US5-2 and LNA-US5-2 or their corresponding negative control, miR-NEG and LNA-NEG. Left panel: 0.1 pmol/well miR-US5-2 inhibits luciferase expression threefold while LNA-NEG has no effect. Right panel: LNA-US5-2 completely blocks the effect of miR-US5-2 in the range 0.2–2 pmol/well and partially inhibits miR-US5-2 activity at 0.02 pmol/well

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concentration as in a regular transfection. LNAs are stable and remain active in the HEK293 cell culture for at least 2 days. It is not clear if these LNAs would exhibit similar properties in other cell types. 65. Luciferase assays can be very sensitive to transfection reagentmediated cell toxicity. When using other transfection reagents and cell lines, make sure to optimize the quantity of transfection reagent. 66. At the proper LNA–miRNA ratio, the LNA antisense to the miRNA (but not the nontargeting LNA) will block the effect of the miRNA as measured by luciferase assay (see Fig. 3). References 1. Bartel DP (2018) Metazoan MicroRNAs. Cell 173(1):20–51. https://doi.org/10.1016/j. cell.2018.03.006 2. Lee RC, Feinbaum RL, Ambros V (1993) The C. elegans heterochronic gene lin-4 encodes small RNAs with antisense complementarity to lin-14. Cell 75(5):843–854 3. Grey F, Antoniewicz A, Allen E, Saugstad J, McShea A, Carrington JC, Nelson J (2005) Identification and characterization of human cytomegalovirus-encoded microRNAs. J Virol 79(18):12095–12099. https://doi.org/10. 1128/JVI.79.18.12095-12099.2005 4. Pfeffer S, Sewer A, Lagos-Quintana M, Sheridan R, Sander C, Grasser FA, van Dyk LF, Ho CK, Shuman S, Chien M, Russo JJ, Ju J, Randall G, Lindenbach BD, Rice CM, Simon V, Ho DD, Zavolan M, Tuschl T (2005) Identification of microRNAs of the herpesvirus family. Nat Methods 2(4):269–276. https://doi.org/10.1038/nmeth746 5. Meyer C, Grey F, Kreklywich CN, Andoh TF, Tirabassi RS, Orloff SL, Streblow DN (2011) Cytomegalovirus microRNA expression is tissue specific and is associated with persistence. J Virol 85(1):378–389. https://doi.org/10. 1128/JVI.01900-10 6. Hancock MH, Tirabassi RS, Nelson JA (2012) Rhesus cytomegalovirus encodes seventeen microRNAs that are differentially expressed in vitro and in vivo. Virology 425 (2):133–142. https://doi.org/10.1016/j. virol.2012.01.009 7. Thomas M, Lieberman J, Lal A (2010) Desperately seeking microRNA targets. Nat Struct Mol Biol 17(10):1169–1174 8. Stark TJ, Arnold JD, Spector DH, Yeo GW (2012) High-resolution profiling and analysis of viral and host small RNAs during human

cytomegalovirus infection. J Virol 86 (1):226–235. https://doi.org/10.1128/JVI. 05903-11 9. Fu M, Gao Y, Zhou Q, Zhang Q, Peng Y, Tian K, Wang J, Zheng X (2014) Human cytomegalovirus latent infection alters the expression of cellular and viral microRNA. Gene 536 (2):272–278. https://doi.org/10.1016/j. gene.2013.12.012 10. Bartel DP (2009) MicroRNAs: target recognition and regulatory functions. Cell 136 (2):215–233. https://doi.org/10.1016/j.cell. 2009.01.002 11. Dolken L, Perot J, Cognat V, Alioua A, John M, Soutschek J, Ruzsics Z, Koszinowski U, Voinnet O, Pfeffer S (2007) Mouse cytomegalovirus microRNAs dominate the cellular small RNA profile during lytic infection and show features of posttranscriptional regulation. J Virol 81 (24):13771–13782. https://doi.org/10. 1128/JVI.01313-07 12. Buck AH, Santoyo-Lopez J, Robertson KA, Kumar DS, Reczko M, Ghazal P (2007) Discrete clusters of virus-encoded micrornas are associated with complementary strands of the genome and the 7.2-kilobase stable intron in murine cytomegalovirus. J Virol 81 (24):13761–13770. https://doi.org/10. 1128/JVI.01290-07 13. Cullen BR (2011) Viruses and microRNAs: RISCy interactions with serious consequences. Genes Dev 25(18):1881–1894. https://doi. org/10.1101/gad.17352611 14. Griffiths-Jones S (2004) The microRNA registry. Nucleic Acids Res 32(Database issue): D109–D111. https://doi.org/10.1093/nar/ gkh023

340

Nicole L. Diggins et al.

15. Griffiths-Jones S, Grocock RJ, van Dongen S, Bateman A, Enright AJ (2006) miRBase: microRNA sequences, targets and gene nomenclature. Nucleic Acids Res 34(Database issue):D140–D144. https://doi.org/10. 1093/nar/gkj112 16. Griffiths-Jones S, Saini HK, van Dongen S, Enright AJ (2008) miRBase: tools for microRNA genomics. Nucleic Acids Res 36(Database issue):D154–D158. https://doi.org/10. 1093/nar/gkm952 17. Kozomara A, Griffiths-Jones S (2011) miRBase: integrating microRNA annotation and deep-sequencing data. Nucleic Acids Res 39 (Database issue):D152–D157. https://doi. org/10.1093/nar/gkq1027 18. Umbach JL, Cullen BR (2009) The role of RNAi and microRNAs in animal virus replication and antiviral immunity. Genes Dev 15 (23):1151–1164 19. Chen Y, Fachko D, Ivanov NS, Skinner CM, Skalsky RL (2019) Epstein-Barr virus microRNAs regulate B cell receptor signal transduction and lytic reactivation. PLoS Pathog 15(1): e1007535. https://doi.org/10.1371/journal. ppat.1007535 20. Kim S, Seo D, Kim D, Hong Y, Chang H, Baek D, Kim VN, Lee S, Ahn K (2015) Temporal landscape of MicroRNA-mediated hostvirus crosstalk during productive human cytomegalovirus infection. Cell Host Microbe 17 (6):838–851. https://doi.org/10.1016/j. chom.2015.05.014 21. Hook LM, Grey F, Grabski R, Tirabassi R, Doyle T, Hancock M, Landais I, Jeng S, McWeeney S, Britt W, Nelson JA (2014) Cytomegalovirus miRNAs target secretory pathway genes to facilitate formation of the virion assembly compartment and reduce cytokine secretion. Cell Host Microbe 15(3):363–373. https://doi.org/10.1016/j.chom.2014.02. 004 22. Albanese M, Tagawa T, Buschle A, Hammerschmidt W (2017) MicroRNAs of Epstein-Barr virus control innate and adaptive antiviral immunity. J Virol 91(16). https://doi. org/10.1128/JVI.01667-16 23. Yang JS, Lai EC (2011) Alternative miRNA biogenesis pathways and the interpretation of core miRNA pathway mutants. Mol Cell 43 (6):892–903. https://doi.org/10.1016/j. molcel.2011.07.024 24. Karginov FV, Conaco C, Xuan Z, Schmidt BH, Parker JS, Mandel G, Hannon GJ (2007) A biochemical approach to identifying microRNA targets. Proc Natl Acad Sci U S A 104 (49):19291–19296. https://doi.org/10. 1073/pnas.0709971104

25. Keene JD, Komisarow JM, Friedersdorf MB (2006) RIP-Chip: the isolation and identification of mRNAs, microRNAs and protein components of ribonucleoprotein complexes from cell extracts. Nat Protoc 1(1):302–307. https://doi.org/10.1038/nprot.2006.47 26. Tan LP, Seinen E, Duns G, de Jong D, Sibon OC, Poppema S, Kroesen BJ, Kok K, van den Berg A (2009) A high throughput experimental approach to identify miRNA targets in human cells. Nucleic Acids Res 37(20):e137. https://doi.org/10.1093/nar/gkp715 27. Easow G, Teleman AA, Cohen SM (2007) Isolation of microRNA targets by miRNP immunopurification. RNA 13(8):1198–1204. https://doi.org/10.1261/rna.563707 28. Orom UA, Lund AH (2007) Isolation of microRNA targets using biotinylated synthetic microRNAs. Methods 43(2):162–165. https://doi.org/10.1016/j.ymeth.2007.04. 007 29. Orom UA, Nielsen FC, Lund AH (2008) MicroRNA-10a binds the 5’UTR of ribosomal protein mRNAs and enhances their translation. Mol Cell 30(4):460–471. https://doi.org/10. 1016/j.molcel.2008.05.001 30. Grey F, Tirabassi R, Meyers H, Wu G, McWeeney S, Hook L, Nelson JA (2010) A viral microRNA down-regulates multiple cell cycle genes through mRNA 5’UTRs. PLoS Pathog 6(6):e1000967. https://doi.org/10. 1371/journal.ppat.1000967 31. Licatalosi DD, Mele A, Fak JJ, Ule J, Kayikci M, Chi SW, Clark TA, Schweitzer AC, Blume JE, Wang X, Darnell JC, Darnell RB (2008) HITS-CLIP yields genome-wide insights into brain alternative RNA processing. Nature 456(7221):464–469. https://doi.org/ 10.1038/nature07488 32. Hafner M, Landthaler M, Burger L, Khorshid M, Hausser J, Berninger P, Rothballer A, Ascano M Jr, Jungkamp AC, Munschauer M, Ulrich A, Wardle GS, Dewell S, Zavolan M, Tuschl T (2010) Transcriptome-wide identification of RNA-binding protein and microRNA target sites by PAR-CLIP. Cell 141(1):129–141. https://doi.org/10.1016/j.cell.2010.03.009 33. Hafner M, Landthaler M, Burger L, Khorshid M, Hausser J, Berninger P, Rothballer A, Ascano M, Jungkamp AC, Munschauer M, Ulrich A, Wardle GS, Dewell S, Zavolan M, Tuschl T (2010) PAR-CliP--a method to identify transcriptome-wide the binding sites of RNA binding proteins. J Vis Exp 41:2034. https://doi.org/10.3791/ 2034

Characterizing Cytomegalovirus-ncoded miRNAs 34. Skalsky RL, Corcoran DL, Gottwein E, Frank CL, Kang D, Hafner M, Nusbaum JD, Feederle R, Delecluse HJ, Luftig MA, Tuschl T, Ohler U, Cullen BR (2012) The viral and cellular microRNA targetome in lymphoblastoid cell lines. PLoS Pathog 8(1): e1002484. https://doi.org/10.1371/journal. ppat.1002484 35. Skinner CM, Ivanov NS, Barr SA, Chen Y, Skalsky RL (2017) An Epstein-Barr virus MicroRNA blocks Interleukin-1 (IL-1) signaling by targeting IL-1 receptor 1. J Virol 91 (21):e00530–e00517. https://doi.org/10. 1128/JVI.00530-17 36. Konig J, Zarnack K, Rot G, Curk T, Kayikci M, Zupan B, Turner DJ, Luscombe NM, Ule J (2011) iCLIP--transcriptome-wide mapping of protein-RNA interactions with individual nucleotide resolution. J Vis Exp 50:2638. https://doi.org/10.3791/2638 37. Vinther J, Hedegaard MM, Gardner PP, Andersen JS, Arctander P (2006) Identification of miRNA targets with stable isotope labeling by amino acids in cell culture. Nucleic Acids Res 34(16):e107. https://doi.org/10.1093/ nar/gkl590 38. Tirabassi R, Hook L, Landais I, Grey F, Meyers H, Hewitt H, Nelson J (2011) Human cytomegalovirus US7 is regulated synergistically by two virally encoded microRNAs and by two distinct mechanisms. J Virol 85 (22):11938–11944. https://doi.org/10. 1128/JVI.05443-11 39. He TC, Zhou S, da Costa LT, Yu J, Kinzler KW, Vogelstein B (1998) A simplified system for generating recombinant adenoviruses. Proc Natl Acad Sci U S A 95(5):2509–2514. https://doi.org/10.1073/pnas.95.5.2509 40. Reinhart BJ, Slack FJ, Basson M, Pasquinelli AE, Bettinger JC, Rougvie AE, Horvitz HR, Ruvkun G (2000) The 21-nucleotide let-7 RNA regulates developmental timing in Caenorhabditis elegans. Nature 403 (6772):901–906. https://doi.org/10.1038/ 35002607 41. Chen C, Ridzon DA, Broomer AJ, Zhou Z, Lee DH, Nguyen JT, Barbisin M, Xu NL, Mahuvakar VR, Andersen MR, Lao KQ, Livak KJ, Guegler KJ (2005) Real-time quantification of microRNAs by stem-loop RT-PCR. Nucleic Acids Res 33(20):e179. https://doi. org/10.1093/nar/gni178 42. Riley KJ, Steitz JA (2013) The "observer effect" in genome-wide surveys of proteinRNA interactions. Mol Cell 49(4):601–604. https://doi.org/10.1016/j.molcel.2013.01. 030

341

43. Ramanathan M, Porter DF, Khavari PA (2019) Methods to study RNA-protein interactions. Nat Methods 16(3):225–234. https://doi. org/10.1038/s41592-019-0330-1 44. Garzia A, Meyer C, Morozov P, Sajek M, Tuschl T (2017) Optimization of PAR-CLIP for transcriptome-wide identification of binding sites of RNA-binding proteins. Methods 118-119:24–40. https://doi.org/10.1016/j. ymeth.2016.10.007 45. Spitzer J, Hafner M, Landthaler M, Ascano M, Farazi T, Wardle G, Nusbaum J, Khorshid M, Burger L, Zavolan M, Tuschl T (2014) PAR-CLIP (photoactivatable Ribonucleosideenhanced crosslinking and immunoprecipitation): a step-by-step protocol to the transcriptome-wide identification of binding sites of RNA-binding proteins. Methods Enzymol 539:113–161. https://doi.org/10.1016/ B978-0-12-420120-0.00008-6 46. Corcoran DL, Georgiev S, Mukherjee N, Gottwein E, Skalsky RL, Keene JD, Ohler U (2011) PARalyzer: definition of RNA binding sites from PAR-CLIP short-read sequence data. Genome Biol 12(8):R79. https://doi. org/10.1186/gb-2011-12-8-r79 47. Khorshid M, Rodak C, Zavolan M (2011) CLIPZ: a database and analysis environment for experimentally determined binding sites of RNA-binding proteins. Nucleic Acids Res 39 (Database issue):D245–D252. https://doi. org/10.1093/nar/gkq940 48. Yang JH, Li JH, Shao P, Zhou H, Chen YQ, Qu LH (2011) starBase: a database for exploring microRNA-mRNA interaction maps from Argonaute CLIP-Seq and Degradome-Seq data. Nucleic Acids Res 39(Database issue): D202–D209. https://doi.org/10.1093/nar/ gkq1056 49. Sievers C, Schlumpf T, Sawarkar R, Comoglio F, Paro R (2012) Mixture models and wavelet transforms reveal high confidence RNA-protein interaction sites in MOV10 PAR-CLIP data. Nucleic Acids Res 40(20): e160. https://doi.org/10.1093/nar/gks697 50. Anders G, Mackowiak SD, Jens M, Maaskola J, Kuntzagk A, Rajewsky N, Landthaler M, Dieterich C (2012) doRiNA: a database of RNA interactions in post-transcriptional regulation. Nucleic Acids Res 40(Database issue): D180–D186. https://doi.org/10.1093/nar/ gkr1007 51. Uren PJ, Bahrami-Samani E, Burns SC, Qiao M, Karginov FV, Hodges E, Hannon GJ, Sanford JR, Penalva LO, Smith AD (2012) Site identification in high-throughput RNA-protein interaction data. Bioinformatics

342

Nicole L. Diggins et al.

28(23):3013–3020. https://doi.org/10. 1093/bioinformatics/bts569 52. Chen B, Yun J, Kim MS, Mendell JT, Xie Y (2014) PIPE-CLIP: a comprehensive online tool for CLIP-seq data analysis. Genome Biol 15(1):R18. https://doi.org/10.1186/gb2014-15-1-r18 53. Chou CH, Lin FM, Chou MT, Hsu SD, Chang TH, Weng SL, Shrestha S, Hsiao CC, Hung JH, Huang HD (2013) A computational approach for identifying microRNA-target interactions using high-throughput CLIP and PAR-CLIP sequencing. BMC Genomics 14 (Suppl 1):S2. https://doi.org/10.1186/ 1471-2164-14-S1-S2 54. Wang T, Xie Y, Xiao G (2014) dCLIP: a computational approach for comparative CLIP-seq analyses. Genome Biol 15(1):R11. https://doi.org/10.1186/gb-2014-15-1-r11 55. De S, Gorospe M (2017) Bioinformatic tools for analysis of CLIP ribonucleoprotein data. Wiley Interdiscip Rev RNA 8(4). https://doi. org/10.1002/wrna.1404

56. Stenvang J, Petri A, Lindow M, Obad S, Kauppinen S (2012) Inhibition of microRNA function by antimiR oligonucleotides. Silence 3 (1):1. https://doi.org/10.1186/1758-907X3-1 57. Grunweller A, Hartmann RK (2007) Locked nucleic acid oligonucleotides: the next generation of antisense agents? BioDrugs 21 (4):235–243. https://doi.org/10.2165/ 00063030-200721040-00004 58. Veedu RN, Wengel J (2010) Locked nucleic acids: promising nucleic acid analogs for therapeutic applications. Chem Biodivers 7 (3):536–542. https://doi.org/10.1002/cbdv. 200900343 59. Lanford RE, Hildebrandt-Eriksen ES, Petri A, Persson R, Lindow M, Munk ME, Kauppinen S, Orum H (2010) Therapeutic silencing of microRNA-122 in primates with chronic hepatitis C virus infection. Science 327(5962):198–201. https://doi.org/10. 1126/science.1178178

Chapter 17 Development of a huBLT Mouse Model to Study HCMV Latency, Reactivation, and Immune Response Lindsey B. Crawford and Patrizia Caposio Abstract Immunodeficient mice engrafted with human tissues provide a robust model for the in vivo investigation of human-restricted viruses such as human cytomegalovirus (HCMV). Several humanized mouse models have been developed and improved over the last 30 years. Here, we describe a protocol for the transplant of human hematopoietic stem cells with autologous fetal liver and thymic tissues into NOD.Cg-PrkdcscidIL2rγ tm1Wjl mice to create a humanized bone marrow–liver–thymus model (huBLT) that can be infected with HCMV. The presence of human thymus allows the development of a functional human immune system, including HLA-restricted human T-cells and B-cells. Indeed, following infection, huBLT mice generate virus-specific CD4+ and CD8+ T-cell responses. Additionally, both HCMV-specific IgM and IgG B-cell responses can be detected. This huBLT model provides the first animal model to explore the adaptive human immune response to HCMV infection. Key words Human Cytomegalovirus, Humanized mice, huBLT, Hematopoietic progenitor cells, Thymus, T-cell response, B-cell response

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Introduction Human cytomegalovirus (HCMV) is a human-specific beta-herpesvirus that infects the majority of the world’s population establishing a lifelong latency in the infected host [1]. In healthy subjects, HCMV infection is usually asymptomatic, however, transplant recipients undergoing immunosuppressive therapy, HIV-infected individuals, and the developing fetus are susceptible to severe viral infection and complications [2]. HCMV infects a variety of cell types, including hematopoietic and stromal cells of the bone marrow, allowing the virus to spread to and persist within multiple organs in the host. Of the hematopoietic lineage cells, the hematopoietic to myeloid cell lineage is the most important with respect to HCMV latency, reactivation, and persistence [3]. The strict species specificity of HCMV and the lack of a suitable animal model system have driven the field to use humanized mouse

Andrew D. Yurochko (ed.), Human Cytomegaloviruses: Methods and Protocols, Methods in Molecular Biology, vol. 2244, https://doi.org/10.1007/978-1-0716-1111-1_17, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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models in which immunodeficient mice are engrafted with human cells and/or tissues to study the immunobiology of the virus. The first humanized mouse model used in this context assessed the ability of the HCMV strain Toledo to replicate within human tissues using SCID-huThy/Liv mice (severe combined immunodeficient mice engrafted with human fetal thymus and liver tissues) [4]. The same model was used by Brown et al. to evaluate and compare the replicative capacity of a low passage versus a high passage Toledo, as well as the laboratory adapted HCMV strains AD169 and Towne [5]. More recently, Dulal et al. reported a functional analysis of the HCMV UL/b0 region using SCID-hu mice transplanted with human fetal thymus and liver tissues directly inoculated with purified virus [6]. The SCID-huThy/Liv mouse model has been useful to address some questions regarding the in vivo replication of the virus, however this model has several limitations including lack of long-term human cell engraftment, low diversity in the types of cells engrafted, lack of distribution of human cells in the mouse and inability to generate human immune response [7]. Over the past decade, second generation humanized mouse models have been developed in which immunodeficient mice are engrafted with human hematopoietic progenitor cells (HPCs) with the goal of recapitulating a functional immune system. The optimal mouse strains for engraftment of human HPCs are enhanced immunodeficient mice such us NOD/scid, Rag1null, or Rag2null that express a mutant IL-2 receptor common gamma chain (IL2rγ) gene [8]. The absence of the IL2rγ-chain leads to severe impairment in adaptive immune system development and function as well as preventing NK cell development [7]. Immunodeficient mice bearing a mutated IL2rγ gene support greatly enhanced human HPC engraftment. Analysis of human hematopoietic cells demonstrated that these mice reconstituted monocytes, macrophages, B-cells, and limited T-cells [8]. The limit in T-cell maturation is believed to be due to the education of these cells in the mouse thymus in the context of mouse MHC I and II. Improvement in T-cell and NK cell development can be achieved by combining the humanized mouse model in enhanced immunodeficient strains with cotransplantation of human fetal thymus and liver tissue. This provides maturation of human T-cells in the context of human thymic tissue alongside robust reconstitution of monocytic and B-lineage cells and facilitates the study of humantropic pathogens [9, 10]. We previously reported the first humanized mouse model in which NOD.Cg-PrkdcscidIL2rγ tm1Wjl (NSG) mice engrafted with human CD34+ HPCs were infected with HCMV supporting a latent viral infection [11]. Subsequent mobilization of HPCs using granulocyte-colony stimulating factor (G-CSF) induces viral reactivation in human macrophages in this model. These huNSG

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mice have been extremely useful to analyze HCMV infection and compare viral mutants [12–16]. However, due to the lack of full and functional immune cell maturation, huNSG mice are unable to develop complete T-cell responses and do not support antibody maturation. These limitations were overcome with the development of humanized mice that have been reconstituted with human fetal liver and thymus tissue in addition to matched CD34+ HPC transplant (huBLT). The huBLT mouse model represents a significant improvement over the huNSG model, since huBLT mice exhibit improved systemic reconstitution of human hematopoietic cells including myeloid lineage cells, NK cells and CD4+ and CD8+ T-cells due, in part, to the presence of human thymic epithelium. We have adapted this system to develop a unique model of HCMV-specific immune responses (see Fig. 1) [17]. This chapter describes the current techniques used in our laboratory to generate huBLT mice, infect the animals with HCMV and induce viral reactivation, and analyze virus-specific cellular and humoral immune responses.

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Materials

2.1 Tissue Preparation

1. Deidentified human fetal liver and thymus, matched donor, 12–24 weeks gestation (Advanced Bioscience Resources), process immediately following receipt. 2. HPC PBS: PBS supplemented with 1% each Penicillin/Streptomycin, Amphotericin B, and FBS. Store at 4  C. 3. HPC media: IMDM supplemented with 1% each Penicillin/ Streptomycin, Amphotericin B, and FBS. 4. Stem Cell Media: HPC media supplemented with 20 ng/mL each human recombinant IL-3, IL-6, SCF, and FLT3L. 5. DNase I from bovine pancreas (CAS 9003-98-9): make a 100 stock at 300 K/mL in HPC PBS, store at 20  C. 6. Hyaluronidase from sheep testes (CAS 37326-33-3): make a 100 stock at 2400 U/mL in HPC PBS, store at 20  C. 7. Collagenase from Clostridium histolyticum (CAS 9001-12-1): make a 100 stock at 507 U/mL in HPC PBS, store at 20  C. 8. Disposable, sterile petri dishes (100 mm). 9. Disposable, sterile 70 μM cell strainers. 10. Disposable, sterile #10 scalpel. 11. Rocking platform housed in a 37  C incubator. 12. Ficoll-Paque.

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Human Fetal Thymus and Liver

sub-lethal irradiation

HCMV

12 weeks

huBLT Mouse with mature T-cells

Adult NSG Mouse

8 weeks

CD34-

HCMV

EBV

Unstim. LCLs

+ control

ELISA HCMV Antibody Titer

ELISPOT

HCMV Lysate

qPCR Spleen or Liver

IgG IgM Viral DNA

CD34+

Time Post-Infection

- G-CSF

+ G-CSF

Fig. 1 Generation of humanized BLT-NSG (huBLT) mice for HCMV infection, reactivation, and immune response analysis. Human fetal liver is digested, leukocytes isolated by Ficoll and the CD34+ hematopoietic stem and progenitor cell (HPC) fraction is separated from CD34 cells by magnetic bead isolation. CD34 cells can be used to generate autologous, EBV-transformed LCLs. Donor-matched thymus is collected in parallel with a sample of the liver and surgically transplanted under the kidney capsule of an adult NSG mouse. Postsurgical transplant, mice are sublethally irradiated and injected (IV) with previously isolated CD34+ HPCs. Following engraftment and T-cell maturation, at 12+ weeks following transplant, humanized BLT (huBLT) mice are infected with HCMV, and maintained for an additional 8 weeks to establish viral latency. Infected huBLT mice can then be analyzed for T-cell responses by ELISPOT, B-cell responses by measurement of antibodies by ELISA, and for viral latency and reactivation (following G-CSF and AMD3100 treatment) by qPCR. (This figure was adapted in part from our previous work [17], and the graphs shown for ELISA and qPCR represent potential data from these assays)

13. RBC lysis buffer: 8.3 g NH4Cl, 1.7 g NaHCO3, and 0.2 mL 0.5 M EDTA pH 8.0 and 1 L H2O. Sterile filter with a 0.22 μM filter and store at room temperature for up to 1 year. 14. CD34 microbeads (see Note 1). 15. LS columns (see Note 1). 16. Magnetic-bead Isolation Magnet appropriate for LS columns (see Note 1). 17. 70 μM preseparation filters appropriate for LS columns. 18. Freezing Media: 80% FBS plus 20% DMSO, store at 4  C, dilute 1:1 with cells in culture media.

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2.2 Immunodeficient Mouse Supplies

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1. Vivarium clean room equipped with biosafety cabinet. 2. Sterilized HEPA-filtered microisolator mouse housing with sterilized food, water, bedding, and equipment. 3. Disposable, sterile gauze (200  200 ). 4. Disposable, sterile drapes (2ply, standard medical drape), cut to size. 5. Disposable, sterile alcohol wipes. 6. 1 mL syringes. 7. 5 mL syringes. 8. 28 g sterile needles. 9. 27 g sterile needles. 10. 25 g sterile needles. 11. Separate 200 μL filter pipette tips and micropipette for vivarium use.

2.3 Generation of huBLT Mice

1. Adult NOD.Cg-Prkdcscid IL2rgtm1Wjil/SxJ (NSG) mice (Jackson Labs 005557). 2. Betadine. 3. 0.5% irrigation saline. 4. Small sterile surgical drape (transparent, nonadhesive). 5. Surgical suture (6–0, sterile, absorbable). 6. Sterile surgical gloves. 7. Surgical tape. 8. Surgical heat source (see Note 2). 9. Harman Mosquito Forceps (2). 10. Thumb Dressing Forceps (4.500 , serrated). 11. Reflex 7 mm wound clip applier. 12. 7 mm wound clips. 13. Reflex wound clip remover. 14. Ring Handle Ear punch (2 mm, 400 handle). 15. Rosenthal Needle (16  1 5/16), also known as a 16 g trocar. 16. Fine point forceps. 17. Straight scissors (5–1/200 ). 18. 7 mm wound clips. 19. Cotton-tipped applicators. 20. Autoclave sterilization pouches for items 9–19 above. 21. Sterile, disposable petri dishes (100 mm) for holding surgical instruments.

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22. Sterile, disposable petri dishes (35 mm) for supplying soaked chow in cages. 23. #15 blade sterile scalpel. 24. Small electric shaver (see Note 3). 25. Chemical depilatory (see Note 3). 26. Portable anesthesia unit equipped for controllable oxygen and isoflurane delivery. 27. Anesthesia chamber. 28. Mouse nose cone. 29. Postoperative surgery analgesics for subrenal implant as approved by IACUC, such as Buprenex SR (Buprenorphine, slow release, SR Veterinary Technologies), use at 0.1 mg/ kg/mouse, store at RT. 30. Postoperative surgery analgesics for osmotic pump implant as approved by IACUC, such as and Carprofen (Zoetis), 50 mg/ mL diluted to 1 mg/mL, store at 4  C and use at 100 μL/ mouse. 31. RadDisk/Rad Box rodent microisolation irradiation disk. 32. Cs-source irradiator approved for use with animals. 33. Thioglycollate (Brewer’s Media) at 4% in dI water, autoclaved and stored at RT in the dark. 34. Osmotic pump, such as Alzet 1000. 35. G-CSF (Neupogen at 300 μg/mL, Amgen). 36. AMD3100 (1,10 -[1,4-phenylenebis(methylene)]bis-1,4,8,11tetraazacyclotetradecane octahydrochloride (CAS 15514831-5). 2.4

Cell Culture

1. TB40E-GFP strain HCMV viral stocks. 2. NHDF: ATCC PCS-201-010. 3. MRC5: ATCC CCL-171. 4. Culture media for cell expansion: DMEM containing 10% FBS and 1% PSG. 5. Culture media for virus expansion: DMEM containing 2% FBS and 1% PSG. 6. Trypsin–EDTA, 0.05%.

2.5 Collection of Blood Samples to Monitor Human Cell Engraftment

1. Tailveiner Restrainer (standard mouse size). 2. Heat lamp. 3. Round-bottom 96-well plates containing 100 μL/well EDTA (100 mM). 4. Adhesive plate covers.

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2.6 Necropsy and Sample Collection

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1. Forceps. 2. #10 blade scalpel. 3. Surgical scissors. 4. Autoclave sterilization pouches for items 1–3 above. 5. CO2 or other IACUC-approved euthanasia method. 6. 1.5 mL epi tubes containing 100 μL EDTA (100 mM). 7. 12-well plates containing DMEM supplemented 10% FBS and 1% PSG. 8. 1.5 mL epi tubes containing 500 μL RNAlater reagent. 9. Tubes containing 10% neutral buffered formalin (scale tubes and formalin volumes appropriately to fix all tissues in 24–72 h). 10. 70 μM cell strainers. 11. 5 mL syringes. 12. Freezing Media: 80% FBS plus 20% DMSO, dilute 1:1 with cells in culture media.

2.7 Extraction of DNA/RNA from Tissue

1. DNAzol reagent. 2. 2 mL tubes (appropriate for bead homogenization). 3. 2 mm glass beads (presterilized by autoclave). 4. 100% EtOH. 5. Nuclease-free water. 6. 3 M NaOAc, pH 5.5. 7. Precellys 24 homogenizer.

2.8 qPCR for Viral Genomes

1. TaqMan FastAdvance Master Mix. 2. HCMV UL141 Forward primer: 50 - GATGTGGGCCGA GAATTATGA-30 . 3. HCMV UL141 Reverse primer: 50 -ATGGGCCAGGAGTGTG TCA-30 . 4. HCMV UL141 FAM-tagged probe: CGAGGGAGAGCAAG TT. 5. FastOptical 96-well reaction plate. 6. MicroAmp optical adhesive film. 7. qPCR appropriate thermocycler.

2.9

Flow Cytometry

1. Positive control sample: human lymphocytes (Ficoll-isolated and viably frozen). 2. Negative control sample: nonengrafted NSG mouse bled in parallel with experimental mice. 3. RBC lysis buffer (see Subheading 2.1).

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4. PBS. 5. Viability Dye: Zombie Aqua Viability Dye or equivalent for staining live/dead cells compatible with formalin fixing. 6. FACS buffer: 10 stock: 0.2 M KH2PO4, 1.5 M NaCl, 2.5% BSA, 0.5% Na Azide, diluted to 1 in sterile dI water, store at 4  C. 7. FACS block: 5% each human and mouse serum in FACS buffer. 8. Cell surface antibodies to phenotype human engraftment in the periphery of huBLT mice: anti-human antibodies: CD3 (UCHT1, PerCP-Cy5.5), CD4 (OKT4, PE-Cy7), CD8a (RPA-T8, PE), CD14 (M5E2, Brilliant Violet 785), CD19 (HIB19, Brilliant Violet 421), CD33 (WM53, APC), CD45 (HI30, APC-Cy7); anti-mouse CD45 (30-F11, Alexa Fluor 700). 9. Fixing Solution: 2% neutral buffered formalin in water, store at RT. 10. Flow cytometer equipped with 405, 488, and 633 nm lasers. 2.10 Immunological Analysis of T-Cells

1. Sterile, round bottom 96-well plates.

2.10.1 Production of Cytokines by T-Cells Analyzed by ELISPOT

3. PBS.

2. Human IFNγ ELISPOT kit. 4. ELISpot plate reader. 5. Immunological positive control stimuli: Staphylococcus aureus enterotoxin B (SEB). 6. Alternative immunological positive control stimuli: anti-CD3. 7. HCMV Lysate (generated from purified HCMV viral stocks, lysed in-house, and diluted to 5 μg/mL). 8. HCMV peptides (pp65 or IE).

2.10.2 Production of Cytokines by T-Cells Analyzed by Intracellular Staining

1. Supplies for flow cytometry (see Subheading 2.9). 2. Perm/Fix kit. 3. Antibodies for cytokine staining, such as anti-human IFNγ (B27) and TNFα (Mab11) with fluorophores appropriate for panel design. 4. Brefeldin A. 5. Immunological stimuli (see Subheading 2.10.1).

2.10.3 Additional Isolation of T-Cell Subsets

1. Supplies for flow cytometry (see Subheading 2.8). 2. BD FACS Aria with appropriate antibodies and controls, and/or magnetic bead separation.

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1. EBV strain B958, concentrate using standard protocols. 2. LCL generation media: RPMI supplemented with 20% FBS, 1% PSG, and cyclosporine A. 3. LCL long term culture media: RPMI supplemented with 10% FBS, 1% PSG, and 1% HEPES.

2.11 HCMV Antibody Analysis by ELISA

1. Clear polystyrene high-protein-binding EIA plates. 2. PBS. 3. HCMV Lysate or recombinant CMV gB antibody. 4. Coating buffer: 0.015 M Na2CO3 plus 0.035 M NaHCO3 in water. 5. Wash buffer: PBS supplemented with 0.25% Tween 20. 6. Blocking buffer: wash buffer supplemented with 0.89% BSA. 7. Secondary antibodies: anti-IgA/M/G, anti-IgG, and/or antiIgM. 8. Chromogen OPB substrate. 9. ELISA plate reader.

3

Methods This protocol involves extensive manipulation of mice, unscreened primary human tissues and human pathogens. All work should be done in a standard biosafety cabinet in a contained BSL2 or BSL2+ facility under a preapproved Institutional Biosafety Committee (IBC) protocol. Unscreened primary human tissues should be treated under BSL2+ conditions and assumed to contain human pathogens. All animal work should be performed in an area approved for both BSL2 agents and for immunodeficient mice following preapproved Institutional Animal Care and Use Committee (IACUC) protocols.

3.1 Preparation of Human Fetal Tissue for Transplant and Isolation of Primary CD34+ HPCs

1. Spin down tissues (300  g, 5 min, RT) and aspirate off media. Wash both liver and thymus with 30 mL HPC PBS, spin down and aspirate off PBS. 2. Add 5 mL of HPC media and carefully decant the media and tissue into sterile petri dishes. Using a sterile scalpel, cut one piece of liver to sufficient size to prepare the implants for all recipient mice (typically 2 the size of the thymic tissue excluding thymic connective tissue, or two 1 mm3 pieces per mouse). Transfer the thymus and liver pieces to new conical tubes, parafilm tops and store at 4  C until surgery (see Note 4).

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3. Mince the remaining liver into small pieces using a sterile scalpel and place back into the original 50 mL conical. Wash the petri dish with 5 mL of media and add to the tube. Repeat wash and transfer until all tissue is returned to the tube and volume is 20–35 mL depending on the size of the tissue. Add digestion enzymes (DNase I, Hyaluronidase and Collagenase to 1 final concentration), mix well by rocking, and incubate at 37  C on a rocking platform for 1–2.5 h until the tissue is digested (see Note 5). 4. Forcefully pipette digested liver up and down within the conical tube ~5 with a 10 mL pipette to break up tissue clumps and filter through 70 μM cell strainers into new conical tubes. Wash cell strainers thoroughly with HPC PBS, spin down, and aspirate off supernatant from cell pellet. Wash the cell pellet with HPC PBS once more. 5. Lyse red blood cells by resuspending cell pellet in 5–15 mL RBC Lysis Buffer, pipette gently to mix and incubate at RT for 5 min. Bring volume up to 50 mL with HPC PBS, spin down and aspirate off supernatant from the cell pellet. Wash cell pellet with 50 mL HPC PBS once more. 6. Resuspend each cell pellet in 10 mL HPC media and bring up to appropriate volume with HPC PBS (see Note 6). Overlay cells on top of Ficoll-Paque gradients (30 mL cell suspension to 15 mL Ficoll per tube) and spin at 500  g, 20 min, RT, without brake. 7. Transfer metaphase layers to a new 50 mL conical tube and wash with HPC PBS. Combine into one conical tube, bring volume to 50 mL with HPC PBS, count total cells and spin down. 8. Aspirate all supernatant and resuspend the cells in 300 μL HPC PBS per 108 cells. Positively select CD34 fraction using CD34microbeads following the manufacturer’s instructions. For every 108 cells, add 100uL FcR Blocking Reagent and 100 μL CD34 microbeads. Incubate for 30 min on ice. 9. Wash cells 1 in HPC PBS and positively select CD34+ HPCs by passing cells over an LS column following the manufacturer’s instructions (see Note 1). Wash cells three times, each with 3 mL HPC PBS and elute positive fraction in 5 mL HPC PBS (see Note 7). 10. Spin down all collected cells, count, and viably freeze for later injection. 11. Save an aliquot of 105 cells for phenotypic analysis. 12. Save the CD34 fraction for generation of autologous endothelial cells, fibroblasts, and EBV-transformed LCLs.

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3.2 Generation of huBLT Mice 3.2.1 Preoperative Preparation

3.2.2 Surgical Implantation of Human Tissue into Mice

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1. Prepare mice for surgery by shaving and use of a chemical depilatory (see Note 3) to prepare a clean surgical site Prepare an area approximately the size of a quarter located above the kidney of choice (see Note 8). 2. Determine mouse weight for postsurgical monitoring and ear-punch for unique identification. 1. Before starting, determine the surgical workflow. It is helpful to have two people to perform surgery, one for anesthesia, tissue and trocar prep, and mouse handling; and a second, who remains sterile, for surgical manipulation and tissue implantation. 2. Place reserved fetal thymic and liver pieces from Subheading 3.1, step 2 above into a sterile petri dish and dissect into 1 mm3 pieces using a scalpel (see Note 9). 3. Anesthetize the mouse using the isoflurane PAM chamber at 2.5% O2 and 2–3% isoflurane. Inject Buprenex via subcutaneous injection and transfer the mouse to the surgical area (see Note 10). 4. Place the mouse on the covered heat source (see Note 2) with the right (lateral) side facing up (see Note 8) and nose away from the surgeon. Betadine the surgical area and cover with a sterile surgical drape. 5. Use scissors and forceps to make a vertical incision (ventral to the spine) approximately 1 cm long between the bottom of the rib cage and top of the hip (see Note 8). Locate the kidney and make a separate, slightly smaller incision through the peritoneal cavity and muscle wall just above the kidney (visual references for surgical techniques and access to the kidney in this manner are previously published [18–20]). 6. Using forceps and saline-dampened cotton swaps, slide the kidney out and rest it on top of the peritoneal cavity. Do not apply direct pressure to the kidney at any time to prevent damage. Secure the kidney by grasping just below the kidney through the skin and peritoneal cavity with mosquito forceps. Keep the kidney moist by frequently dripping sterile saline over the surface using a moistened cotton swab. 7. Make a small (~1.5 mm long) incision through the kidney capsule using a #15 blade scalpel and carefully create a pocket under the capsule but above the kidney tissue using a 27 g needle. 8. Assemble the trocar and load with pieces of fetal liver and thymus (each ~1 mm3 in size) (see Note 11). Gently insert the trocar into the pocket on the surface of the kidney and slide as far as possible up the length of the kidney (to anterior). Be

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careful to stay as superficial as possible to avoid excessive damage and bleeding resulting from puncture of the kidney tissue, but avoid tearing through the fragile kidney capsule as this results in loss of tissue engraftment. 9. Carefully depress the plunger on the trocar, while slowly withdrawing the trocar about halfway. Tissue pieces should be visible under the kidney capsule. To prevent backsliding of the tissue pieces, the flat side of a needle can be placed between the tissue pieces and the trocar, through the surface of the capsule, while the trocar is removed completely. Blot minimal bleeding with a dry cotton swab. 10. Release the mosquito forceps and gently pull apart the peritoneal cavity incision to release the kidney back into the body cavity. If necessary, gently push the kidney through using a damp cotton swab. Close the cavity using suture via a simple running stitch and/or interrupted suture (depending on the size of incision and IACUC guidelines). Close the skin incision using 2–3 wound clips. 11. Place the mouse on a plastic wrapped heat pack in a clean housing cage prestocked with soaked chow and monitor until it is recumbent and awake following anesthesia. Remove the heat pack and return the mouse to the housing area. 12. Repeat steps 3–11 for additional mice. Disinfect surgical tools with a hot bead sterilizer and get new cotton swabs for betadine and saline between each mouse. Surgical drape and suture that remain in good condition can be reused for additional mice. 3.2.3 Postoperative Monitoring

1. Perform health checks and replace soaked chow (or other additional care measures as per specific IACUC) twice daily. 2. Weigh mice at 3 and 7–10 days following surgery and check that weight is within specific postsurgery guidelines or follow up as appropriate. 3. Remove wound clips at 7–10 days following surgery.

3.2.4 Irradiation and Injection of Autologous CD34+ HPCs

1. At 10–14 days following surgery, irradiate mice by whole-body gamma irradiation using a 137Cs source for a total dose of 200cGy, following IACUC guidelines (see Note 12). 2. Thaw autologous CD34+ HPCs and culture overnight in stem cell media. 3. Twenty to twenty four hours following irradiation, wash HPCs and resuspend at the desired concentration in 100 μL PBS per mouse. Inject ~105 cells per mouse via intravenous injection.

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3.3 Monitoring Human Cell Engraftment by Survival Bleed and Flow Cytometry

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Human cells are visible in some cohorts as early as 3 weeks posttransplant, however functional T-cells are not mature until approximately 12 weeks. We typically screen mice for engraftment beginning at 8 weeks following HPC injection and every 4 weeks thereafter. 1. Collect a sample of peripheral blood by tail vein bleed (see Note 13) from each mouse and for an engraftment negative control, one additional nonhumanized control NSG mouse (see Note 14). 2. Spin down cells in 96-well plates (500  g, 3 min, RT), decant supernatant and resuspend in 200 μL RBC lysis buffer for 15 min. Add positive control lymphocytes to one well at this point. 3. Wash plate twice with 200 μL PBS to remove RBCs and stain with viability dye following the manufacturer’s instructions. 4. Wash twice with 200 μL FACS buffer and block for 30–60 min in 100 μL FACS block. 5. Stain with appropriate antibodies in FACS block for 30–60 min, then fix with 2% neutral buffered formalin for 15 min at RT. 6. Wash twice with FACS buffer and analyze on a BD LSRII flow cytometer (or equivalent) equipped with appropriate lasers and filters. 7. Analyze data using BD FACS Diva (or equivalent) and set gates using the human lymphocyte positive control (muCD45 negative and human cell marker(s) positive) and the nonhumanized NSG control mouse (human cell marker(s) negative, muCD45 positive). 8. Distribute mice into cohorts, normalizing for level of human cell engraftment and T-cell reconstitution.

3.4 Infection of huBLT Mice with HCMV and Reactivation In Vivo 3.4.1 Preparation of HCMV-Infected Human fibroblasts and Infection of huBLT Mice

1. Expand human fibroblasts (2 T150 flasks per mouse) to 90% confluent. 2. Infect at an MOI of 0.01 with HCMV and culture until full CPE is reached, but cells remain attached. 3. One day before harvest, collect one flask and pretiter by TCID50 for 24 h to estimate injection titer (see Note 15). 4. Prestimulate mice by intraperitoneal (IP) injection of 1 mL 4% thioglycollate media. 5. The following, day harvest HCMV-infected fibroblasts by normal trypsinization of cells. 6. Wash collected cell pellets twice with PBS and resuspend in 2 mL of PBS per mouse. 7. Inject each mouse with 2 mL (~2 T150 flasks of cells) via IP injection using a 25 g needle.

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3.4.2 Reactivation of HCMV In Vivo

1. After 8 weeks of infection, following the establishment of latency, reactivate the virus by treatment with G-CSF and AMD3100. 2. Prepare mice for surgery as described in Subheading 3.2.1. Prepare an area approximately the size of a nickel located just ventral to the spine along the shoulder. 3. Prepare osmotic pumps by filling with 100 μL G-CSF per pump. Check that pumps are accurately loaded by weighing pumps before and after loading to confirm the correct dose. Proceed to implant within an hour postloading. 4. Anesthetize the mouse using the isoflurane PAM chamber at 2.5% O2 and 2–3% isoflurane. Inject Carprofen via subcutaneous injection, inject AMD3100 via IP injection, and transfer the mouse to the surgical area (see Note 10). 5. Place the mouse on the covered heat pack with the right side facing up and nose toward the dominant hand of the surgeon. Betadine the surgical area and cover with a sterile surgical drape. 6. Use scissors and forceps to make a horizontal (lateral to the spine) incision approximately 1 cm long just ventral to the spine. Prewet straight forceps in sterile saline and loosen a pocket under the skin and above the connective tissue on an angle toward the posterior. 7. Dip the pump in sterile saline and slide into the pocket pump end first. Close the skin incision using wound clips. 8. Place the mouse on a plastic wrapped heat pack in a clean housing cage prestocked with soaked chow and monitor until it is recumbent and awake following anesthesia. Remove the heat pack and return the mouse to the housing area. 9. Repeat steps 4–8 for additional mice. Maintain mice for one additional week to allow for viral reactivation.

3.5 Analysis of HCMV Infection and Immune Responses 3.5.1 Necropsy of HCMV-Infected huBLT Mice 3.5.2 Analysis of Viral DNA in Tissues (DNA Extraction and qPCR)

1. Euthanize mice via CO2 using IACUC approved methods. 2. Collect blood, lymphoid organs (spleen, liver and bone marrow), and any additional relevant tissues (lymph nodes, submandibular gland, lungs, heart, etc.). Collect samples as appropriate for downstream analysis (see step 1 for each section below). 1. Collect sections of tissues (see Note 16) in 500 μL RNAlater in epi tubes and store at 80  C until processing. 2. Thaw samples stored in RNAlater at room temperature and transfer each piece to a 2 mL screw-top tube with O-ring

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containing 500 μL DNAzol and ~30 glass beads. Parafilm the tube top. 3. Disassociate tissue using a homogenizer (Precellys24) at 4445  g for 23 s followed by a 15 s pause. Repeat three times. 4. Spin down the sample (10 min, 12,000  g at 4  C) and transfer DNA plus DNAzol supernatant to a new tube containing 500 μL ethanol. 5. Invert 3 to mix and incubate at RT for ~10 min, then spin down (6000  g, 10 min, 4  C) and remove the supernatant by pipetting. 6. Wash 1 in 0.8–1 mL 75% ethanol, vortex and spin down (4000 g, 2 min, 4  C). Remove supernatant by pipetting and repeat the wash step. 7. Spin down sediment (4000  g, 2 min, 4  C), remove residual ethanol, and dry at RT for 5 min. 8. Resuspend DNA in 300 μL nuclease-free water and incubate in a 55  C shaker for 30 min. 9. Spin down the sample (12,000  g, 10 min, RT) and transfer supernatant to a new tube containing 30 μL 3 M NaOAc. 10. Add 600 μL 100% ethanol and precipitate the DNA at 80  C for 30 min. 11. Spin down the sample at 12,000  g, 30 min, 4  C. 12. Wash by resuspending pellet in 500 μL 70% ethanol, vortex to mix, incubate at RT for 5 min. Spin down the sample (12,000  g, 10 min, 4  C) and discard the supernatant. 13. Spin down again (12,000  g, 2 min, 4  C or RT), remove residual ethanol and dry at RT for 5 min. 14. Resuspend in 50 μL water and determine concentration. Dilute as necessary for a final concentration of 100 ng/μL. 15. Set up a qPCR mastermix. For each well, use: 15 μL TaqMan FastAdvance Master Mix, 0.9 μL each Forward and Reverse primer, 0.5 μL probe, and 10 μL DNA. 16. Set up copy number controls using predetermined standards made from diluted HCMV BAC DNA. Standards should be run in duplicate and samples in triplicate. 17. Run the Taqman assay using the following program: activation for 1 cycle (50  C for 2 min, 95  C 20 s) followed by detection for 40 cycles (95  C 1 s, 60  C 20 s). 3.5.3 Flow Cytometry of Human Cell Populations in Lymphoid Organs

1. Collect sections of tissues and bone marrow into media in a 12-well plate and store at RT for same day processing. 2. Collect blood into an epi tube containing 100 mM EDTA.

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3. For analysis of tissue engraftment, smash tissues through a 70 μM cell strainer using the syringe plunger from a 5 mL syringe. 4. Wash the cell strainer thoroughly with PBS and spin down cells (300  g, 5 min, RT). 5. Optional: isolate lymphocytic fraction by Ficoll-Paque gradient before analysis. 6. Plate 106 cells per well on a 96-well plate and process samples for flow cytometry as described in Subheading 3.3. 3.5.4 Analysis of Cytokines Produced by T-Cells

1. Collect and process sections of spleen and liver, and blood as described above in Subheading 3.5.3. 2. Separate isolated cells over Ficoll-Paque by overlaying 5 mL of Ficoll-Paque with 10 mL of cell suspension in a 15 mL conical tube. 3. Spin at 500  g, 15 min, RT without brake. 4. Collect lymphocyte layer, wash, and count cells. 5. Analyze 2  105 total cells by ELISPOT or intracellular flow cytometry staining using commercial reagents. 6. Cells are stimulated with negative control, positive control (SEB toxin or anti-CD3), HCMV lysate, and HCMV-specific peptides. 7. For standard analysis, we use MabTech human IFNγ ELISPOT according to the manufacturer’s instructions and analyze cytokine secretion at 48–56 h postplating. Dried plates are imaged using an AID ELISpot Reader. 8. For ICS, cells are incubated in RPMI + 10% FBS + 1% PSG overnight with stimuli, then an additional 4 h in the presence of Brefeldin A. 9. Samples are then washed, plated, and stained using surface markers for flow cytometry according to the protocol described in Subheading 3.3. 10. Cells are fixed using a commercial Perm/Fix kit and stained for intracellular cytokines, washed and analyzed by flow cytometry. 11. To generate autologous LCLs for peptide presentation as previously described [21], begin by thawing viable CD34 fraction cells isolated from autologous fetal liver tissue. 12. Wash and resuspend the cells at 2  107 per mL in LCL media containing cyclosporine A, add concentrated EBV diluted in an equivalent volume of media, and plate 1 mL/well in a 24-well plate. 13. After 10 days, maintain the cells with twice-weekly media changes for an additional 6 weeks before use.

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14. Once transformed (following 6–8 weeks of culture), LCLs can be further expanded for bulk use. 15. LCLs are preloaded with peptides for presentation to purified T-cell subsets. 16. Isolate T-cell subsets using magnetic beads or FACS for desired populations. 17. Mix isolated T-cells and peptide loaded LCLs and analyze by ELISPOT (steps 5–7 above) or ICS (steps 8–10 above). 3.5.5 Analysis of Antibody Production

1. Collect blood from live time-point bleeds (see Subheading 3.3) or terminal analysis (see Subheading 3.5.1) into an epi tube containing 100 μL of 100 mM EDTA. 2. Spin down blood at 10,000  g, 10 min, RT and remove plasma. 3. Store plasma at 80  C until further analysis. 4. Resuspend blood in an equal volume of PBS (equivalent to the volume of plasma removed) and process for DNA/RNA (see Subheading 3.5.2), flow cytometry (see Subheading 3.5.3), and/or T-cell analysis (see Subheading 3.5.4). 5. Precoat EIA plates with purified HCMV virions diluted in PBS or recombinant CMV gB diluted in coating buffer and adhere overnight. 6. Wash five times with wash buffer and block for 90 min at RT in blocking buffer. 7. Serially dilute plasma in twofold increments in blocking buffer and plate at 100 μL per well, before incubating at 37  C for 45 min. 8. Wash 5 in wash buffer and detect with 50 μL secondary antibody at 37  C for 30 min. 9. Wash 5 in wash buffer and quantify using chromogen OPB substrate before reading at 450 nm using an ELISA plate reader. 10. Calculate endpoint antibody titers using log–log transformation on the linear portion of the curve.

3.6

Conclusions

The immunobiology of HCMV combined with viral latency and reactivation is a complex series of processes dependent on both viral and cellular factors. Use of huBLT mice facilitates the study of these interconnected mechanisms by providing a small animal model that allows for viral replication in the context of relevant human cells in the presence of a functional human immune system. This model thus provides an opportunity to experimentally manipulate the viral and cellular systems to determine the mechanisms involved. The protocols outlined in this chapter provide a foundation to prepare

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and utilize the huBLT model for HCMV infection, the establishment of latency and subsequent viral reactivation, and to assess HCMV-specific immune responses in an in vivo setting.

4

Notes 1. This protocol is optimized using the Miltenyi Biotech human CD34 magnetic bead isolation protocol and following the manufacturer’s instructions. Alternative positive selection methods using other magnetic beads or FACS-based isolation are available. All methods should be confirmed by postisolation analysis of the both the positive and negative fractions for, at minimum: viability, % CD34+, contaminating T-cells (i.e., % CD3+), and any other markers of interest. 2. During surgery and postrecovery, mice need to be kept warm. Consult your institutional veterinarian for IACUC approved methods. Either a water-jacket blanket or hand-warmer heat packs (~2  300 up to 130  C max., the kind that fit in gloves) can be used. Place one each under the mouse during surgery and in the postsurgery recovery cage. 3. Surgery requires a clean site of entry into the body cavity. Get preapproval on your IACUC protocol for the desired method. Preshaving and using a chemical depilatory (such as VEET or NAIR, sensitive skin-formula without added moisturizers or scents) gives a good surgical area free of hair and stubble. Preshave mice up to 3 days in advance of surgery. If surgery is delayed up to 10 days after pre-op, the surgical site can be updated by reapplication of the chemical depilatory. Be sure to follow strict timing when using a chemical depilatory and wash and dry the area posttreatment to prevent overtreatment and scabbing of the skin area. 4. When working with same day arrival tissues (tissues that arrive in the afternoon) or limited personnel, the tissue is fine if stored at 4  C overnight for surgery on the following day. 5. Critical step: ensure that the liver tissue is cut through the capsule and pieces are no more than 5mm3 prior to digestion. Excessive connective tissue does not contain HPCs and can be scraped clean and discarded or digested separately. Tissue digestion should proceed until digestion is progressed, but tissue is not degraded. Depending on the size of the tissue pieces (the amount of mechanical disruption), this is usually 1–3 h. Digestion should not exceed 3.5 h under any circumstances or substantial decreases in viability will be seen. 6. The number of Ficoll tubes used for lymphocyte isolation depends on the size of the original tissue and completeness of

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RBC lysis. An average (3–5 g tissue) uses 3–4 tubes of Ficoll for complete separation. If Ficoll is overloaded, excess RBCs remain in the metaphase and can interfere with downstream processes. 7. To improve CD34+ HPC yield by an additional ~10%, pass the flow-through (CD34) fraction from the first column over a second LS column at each stage. Wash the secondary column with additional clean PBS and then elute the positive cells from the second column and combine with the initial CD34+ fraction. 8. Other protocols [18–20] use the left kidney for surgery. The orientation of both kidneys in the mouse is the same, however, the left kidney is often placed slightly lower and is more accessible from underneath the rib cage. However, we prefer to use the right kidney to avoid any interference with the left-located spleen and related surgical complications. Either kidney provides sufficient engraftment. 9. For a large cohort of mice, it is helpful to dissect the thymus and liver into 2–4 large pieces and work with one set at a time keeping the other sets on ice until needed to prevent tissue degradation and stickiness of the tissue. For each mouse, dissect off one 1 mm3 section at a time for ease of manipulation. 10. The surgical area should be already prepared with a covered heat source (see Note 2) and aligned nose cone to delivery continuous anesthesia throughout the surgical procedure. Surgical tools and supplies (cotton applicators, suture, wound clips) should be located in an accessible but sterile area. 11. A liver–thymus–liver sandwich is effective and traditional; however, if liver is in short supply, a thymus–liver–thymus sandwich, liver–thymus duo, or thymus only is capable of engraftment. 12. Irradiation of adult NSG mice using 200cGy total body irradiation at least 1 week following surgery does not significantly cause morbidity or associated weight loss and no mortality has been seen in over 600 mice in our hands. 13. The method, volumes and frequency for survival bleeds when screening of huBLT mice depends on IACUC approval. A general rule is no more than 10% body weight every month (we factor about 200 μL per mouse per month). If sampling less (e.g., flow cytometry screening requires as little as 50 μL blood for an engrafted animal), the frequency can be increased. 14. Flow cytometry analysis requires both a positive and negative control for accurate analysis of engraftment in huBLT mice. For a negative control (nonengrafted control), bleed a comparable nonmanipulated NSG mouse at the same time as the

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experimental cohort and process in parallel. Previously, viably frozen human lymphocytes or freshly collected human blood/ PMBCs can be used as a positive/staining control. 15. When comparing different viral mutants or viral strains in one experiment, it is essential to match the input amount of virus for each experimental group as closely as possible. Since viral growth varies, pretitering the input virus before injection allows for normalization of changes in growth kinetics. Therefore, on the day before mouse infection, one representative flask of each virus is harvested and titered by TCID50 for 24 h. Input virus amounts can then be normalized between experimental groups by varying the amount of each cell and virus pool to be injected. 16. Viral seeding of huBLT tissues occurs in foci within the tissue, likely associated with cluster of human monocytic cells. To accurately identify virus and changes in viral load, section the tissues and process as independent pieces rather than homogenizing the whole tissue. Splenic tissue is typically sectioned into two individual ~1 mm3 sections per mouse. Liver tissue from one small lobe is divided equally into four sections (~2  3  1 mm) per mouse and processed individually.

Acknowledgments We would like to thank L. Drew Martin, DVM, DACLAM for veterinary oversight in setting up these protocols; and Christopher Parkins, Andrew Pham, and Rebecca Tempel, PhD for technical assistance on various cohorts of huBLT mice. This work was supported by NIH funding P01 AI127335. References 1. Cannon MJ, Schmid DS, Hyde TB (2010) Review of cytomegalovirus seroprevalence and demographic characteristics associated with infection. Rev Med Virol 20(4):202–213. https://doi.org/10.1002/rmv.655 2. Griffiths P, Baraniak I, Reeves M (2015) The pathogenesis of human cytomegalovirus. J Pathol 235(2):288–297. https://doi.org/10. 1002/path.4437 3. Goodrum F (2016) Human cytomegalovirus latency: approaching the Gordian knot. Annu Rev Virol 3(1):333–357. https://doi.org/10. 1146/annurev-virology-110615-042422

4. Mocarski ES, Bonyhadi M, Salimi S, McCune JM, Kaneshima H (1993) Human cytomegalovirus in a SCID-hu mouse: thymic epithelial cells are prominent targets of viral replication. Proc Natl Acad Sci U S A 90(1):104–108 5. Brown J, Kaneshima H, Mocarski E (1995) Dramatic interstrain differences in the replication of human cytomegalovirus in SCID-hu mice. J Infect Dis 171(6):1599–1603 6. Dulal K, Cheng T, Yang L, Wang W, Huang Y, Silver B, Selariu A, Xie C, Wang D, Espeseth A, Lin Y, Wen L, Xia N, Fu T-M, Zhu H (2016) Functional analysis of human cytomegalovirus UL/b0 region using SCID-hu mouse model. J

HCMV-Infected huBLT Mice Med Virol 88(8):1417–1426. https://doi. org/10.1002/jmv.24484 7. Shultz LD, Brehm MA, Garcia-Martinez JV, Greiner DL (2012) Humanized mice for immune system investigation: progress, promise and challenges. Nat Rev Immunol 12 (11):786–798 8. Shultz LD, Lyons BL, Burzenski LM, Gott B, Chen X, Chaleff S, Kotb M, Gillies SD, King M, Mangada J, Greiner DL, Handgretinger R (2005) Human lymphoid and myeloid cell development in NOD/LtSz-scid IL2R gamma null mice engrafted with mobilized human Hemopoietic stem cells. J Immunol 174(10):6477–6489. https://doi.org/10. 4049/jimmunol.174.10.6477 9. Shultz LD, Keck J, Burzenski L, Jangalwe S, Vaidya S, Greiner DL, Brehm MA (2019) Humanized mouse models of immunological diseases and precision medicine. Mamm Genome 30(5-6):123–142. https://doi.org/ 10.1007/s00335-019-09796-2 10. Crawford LB, Streblow DN, Hakki M, Nelson JA, Caposio P (2015) Humanized mouse models of human cytomegalovirus infection. Curr Opin Virol 13:86–92. https://doi.org/10. 1016/j.coviro.2015.06.006 11. Smith MS, Goldman DC, Bailey AS, Pfaffle DL, Kreklywich CN, Spencer DB, Othieno FA, Streblow DN, Garcia JV, Fleming WH, Nelson JA (2010) Granulocyte-Colony stimulating factor reactivates human cytomegalovirus in a latently infected humanized mouse model. Cell Host Microbe 8(3):284–291 12. Umashankar M, Petrucelli A, Cicchini L, Caposio P, Kreklywich CN, Rak M, Bughio F, Goldman DC, Hamlin KL, Nelson JA, Fleming WH, Streblow DN, Goodrum F (2011) A novel human cytomegalovirus locus modulates cell type-specific outcomes of infection. PLoS Pathog 7(12):e1002444. https://doi.org/10. 1371/journal.ppat.1002444 13. Caviness K, Bughio F, Crawford LB, Streblow DN, Nelson JA, Caposio P, Goodrum F (2016) Complex interplay of the UL136 isoforms balances cytomegalovirus replication and latency. MBio 7(2):e01986. https://doi.org/10. 1128/mBio.01986-15 14. Theobald SJ, Khailaie S, Meyer-Hermann M, Volk V, Olbrich H, Danisch S, Gerasch L, Schneider A, Sinzger C, Schaudien D, Lienenklaus S, Riese P, Guzman CA, Figueiredo C, von Kaisenberg C, Spineli LM, Glaesener S, Meyer-Bahlburg A, Ganser A, Schmitt M, Mach M, Messerle M, Stripecke R (2018) Signatures of T and B cell development, functional responses and PD-1 upregulation

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after HCMV latent infections and reactivations in nod.Rag.Gamma mice humanized with cord blood CD34+ cells. Front Immunol 9:2734. https://doi.org/10.3389/fimmu.2018. 02734 15. Crawford LB, Kim JH, Collins-McMillen D, Lee B-J, Landais I, Held C, Nelson JA, Yurochko AD, Caposio P (2018) Human cytomegalovirus encodes a novel FLT3 receptor ligand necessary for hematopoietic cell differentiation and viral reactivation. MBio 9:2. https://doi.org/10.1128/mBio.00682-18 16. Vashee S, Stockwell TB, Alperovich N, Denisova EA, Gibson DG, Cady KC, Miller K, Kannan K, Malouli D, Crawford LB, Voorhies AA, Bruening E, Caposio P, Fru¨h K (2017) Cloning, assembly, and modification of the primary human cytomegalovirus isolate Toledo by yeast-based transformation-associated recombination. mSphere 2(5):e00331–e00317. https://doi.org/10.1128/mSphereDirect. 00331-17 17. Crawford LB, Tempel R, Streblow DN, Kreklywich C, Smith P, Picker LJ, Nelson JA, Caposio P (2017) Human cytomegalovirus induces cellular and humoral virus-specific immune responses in humanized BLT mice. Sci Rep 7(1):937. https://doi.org/10.1038/ s41598-017-01051-5 18. Aryee K-E, Shultz LD, Brehm MA (2014) Immunodeficient mouse model for human hematopoietic stem cell engraftment and immune system development. Methods Mol Biol 1185:267–278. https://doi.org/10. 1007/978-1-4939-1133-2_18 19. Vatakis DN, Bristol GC, Kim SG, Levin B, Liu W, Radu CG, Kitchen SG, Zack JA (2012) Using the BLT humanized mouse as a stem cell based gene therapy tumor model. J Vis Exp 70:e4181. https://doi.org/10.3791/ 4181 20. Zhen A, Rezek V, Youn C, Rick J, Lam B, Chang N, Zack J, Kamata M, Kitchen S (2016) Stem-cell based engineered immunity against HIV infection in the humanized mouse model. J Vis Exp 113:e54048. https://doi. org/10.3791/54048 21. Lavender KJ, Pang WW, Messer RJ, Duley AK, Race B, Phillips K, Scott D, Peterson KE, Chan CK, Dittmer U, Dudek T, Allen TM, Weissman IL, Hasenkrug KJ (2013) BLT-humanized C57BL/6 Rag2/γc/-CD47/ mice are resistant to GVHD and develop B and T cell immunity to HIV infection. Blood 122 (25):4013–4020. https://doi.org/10.1182/ blood-2013-06-506949

Chapter 18 Rodent Models of Congenital Cytomegalovirus Infection Berislav Lisnic´, Jelena Tomac, Djurdjica Cekinovic´, Stipan Jonjic´, and Vanda Juranic´ Lisnic´ Abstract Human cytomegalovirus (HCMV) is a leading viral cause of congenital infections in the central nervous system (CNS) and may result in severe long-term sequelae. High rates of sequelae following congenital HCMV infection and insufficient antiviral therapy in the perinatal period makes the development of an HCMV-specific vaccine a high priority of modern medicine. Due to the species specificity of HCMV, animal models are frequently used to study CMV pathogenesis. Studies of murine cytomegalovirus (MCMV) infections of adult mice have played a significant role as a model of CMV biology and pathogenesis, while MCMV infection of newborn mice has been successfully used as a model of perinatal CMV infection. Newborn mice infected with MCMV have high levels of viremia during which the virus establishes a productive infection in most organs, coupled with a robust inflammatory response. Productive infection in the brain parenchyma during early postnatal period leads to an extensive nonnecrotizing multifocal widespread encephalitis characterized by infiltration of components of both innate and adaptive immunity. As a result, impairment in postnatal development of mouse cerebellum leads to long-term motor and sensor disabilities. This chapter summarizes current findings of rodent models of perinatal CMV infection and describes methods for analysis of perinatal MCMV infection in newborn mice. Key words Cytomegalovirus, Brain, Congenital infections

1

Introduction

1.1 HCMV and Congenital Infection

Human cytomegalovirus (HCMV), also known as human herpesvirus 5 (HHV-5), is a widespread virus for which the most recent estimates of mean global seroprevalence rates range from 78% to 88% in the general population and 83% to 89% in women of reproductive age [1]. However, despite such an extensive global presence of the virus, approximately 80% of women worldwide appear to be completely unaware of the risks associated with the congenital HCMV infection [2, 3]. The apparent disregard of HCMV by the general public is probably caused by the fact that the virus usually causes mild or no symptoms in healthy immunocompetent individuals, even though it can infect and/or replicate in a broad range of cell types, such as fibroblasts, endothelial cells,

Andrew D. Yurochko (ed.), Human Cytomegaloviruses: Methods and Protocols, Methods in Molecular Biology, vol. 2244, https://doi.org/10.1007/978-1-0716-1111-1_18, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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epithelial cells, smooth muscle cells, monocytes, macrophages, dendritic cells, polymorphonuclear leukocytes, CD34+ hematopoietic progenitor cells, stromal cells, neurons, hepatocytes, as well as placental trophoblast and cytotrophoblast cells [4–6]. In contrast with normal healthy adults, however, the young are often more vulnerable to HCMV infection during prenatal, perinatal, and postnatal periods, when their immune system has not yet reached sufficient maturity to effectively combat HCMV infection. In fact, HCMV is the most frequent viral cause of congenital infections with an annual prevalence between 0.1% and 2% of newborns [7], and a principal nongenetic cause of congenital and developmental disabilities [8]. HCMV can be transmitted from seropositive mothers to unborn children during pregnancy or delivery, as well as to a newborn child following birth. Infection with HCMV in these cases occurs either because of the transplacental spread of the virions to the developing fetus, of exposure of the newborn to the virus in the maternal genital tract during birth or exposure of the infant to the virus during breastfeeding [9]. The long-term sequelae following HCMV infection, however, appears to be primarily a consequence of in utero infection, while the dissemination of the virions via breast milk seems to present a hazard predominantly to prematurely born infants [10]. Once the infection has taken place, symptomatic disease occurs in only approximately 10% of the infected infants [11, 12], who develop signs of disease affecting auditory, vision, hematopoietic, and hepatobiliary systems and skin, as well as the central nervous system [12, 13]. Of the affected systems, HCMV infection of the developing CNS is particularly threatening, since it may result in the often irreversible, long-term neurological sequelae, including, but not limited to, progressive hearing loss, cerebral palsy, and mental retardation [12]. The remaining ~90% of the infected neonates, which are asymptomatic at birth and during the newborn period, are also at risk for developing long-term neurological deficits later in life [12, 14]. Regrettably, the strict species specificity of cytomegaloviruses precludes scientific examination of HCMV pathogenesis in experimental animals, and none of the disease models developed thus far completely recapitulate the properties of congenital CMV infection in humans [15]. While the rhesus macaque disease model of perinatal RhCMV infection features many similarities with congenital HCMV infection in humans, the main drawback of this model is the paucity of RhCMV-seronegative macaques and the often prohibitively high cost of laboratory animals and experimental setups. These issues in CMV research are, to a certain extent, circumvented by employing nonprimate disease models of congenital HCMV infection, each of which holds its strengths and weaknesses. For example, the guinea pig model of congenital GPCMV infection

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remains unique among rodent models due to the ability of the GPCMV to traverse the guinea pig placenta [16]. Additionally, congenitally infected guinea pigs present systemic viremia with an effect on the CNS [16, 17], whereas the infection of the cochlea can result in the development of sensorineural hearing loss (SNHL) [15]. Practical limitations of this model are the need for high doses of the virus for infection of dams, with consequent significant fetal loss and small litters (average of three pups per pregnant animal with pregnancy lasting between 65 and 70 days). Experiments in the guinea pig model of congenital GPCMV infection can therefore require a substantial number of experimental animals. The rat CMV (RCMV) model, on the other hand, is frequently used for studying CMV-associated vascular diseases [18], even though in the early studies RCMV could not be recovered from rat embryos originating from RCMV-infected mothers [19]. Despite this early setback, Loh and colleagues have reported a novel RCMV strain, isolated from placental tissue, that can infect rat fetuses [20, 21]. In addition, an in utero model of RCMV infection of the developing rat brain, based on intraventricular administration of viral suspension to E15 embryos, was recently shown to share similarities with human congenital CMV infection of the CNS [22]. However, it remains to be determined how useful the new RCMV strain(s), or the unconventional and simulated routes of virus spread, are in the modeling of all relevant aspects of congenital CMV infection. Pathogenesis of the CMV infection is most frequently studied using mouse cytomegalovirus (MCMV) in mouse disease models of CMV infection, mainly due to a well-characterized MCMV genome and proteome, availability of numerous viral mutants and the relative ease of the use of mice as experimental animals. One significant disadvantage of the mouse model, however, is the inability of MCMV to cross the placental barrier and infect mouse embryos. We and others have attempted to overcome this obstacle by employing alternative routes of infection of either mouse embryos or newborn mice. Keeping in mind that in congenital HCMV disease, the extent of damage to the CNS is the primary determinant of the outcome of the infection, most investigators have attempted to design their experimental systems in a way to model the CNS disease in mice in the most representative manner. Similarly to the rat model of CNS infection described above [22], MCMV infection of the developing CNS can therefore be established by direct virus inoculation into cerebral hemispheres or lateral ventricles of either mouse embryos or newborn mice [23, 24]. In addition, other routes of virus administration can be employed in the modelling of CMV disease of the CNS in mice, such as intraplacental inoculation of the virus [25], intraperitoneal infection of newborn mice [25, 26], or infection of suckling mice by milk extracted from MCMV-infected dams [10].

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In contrast to other murine models that require either cryoanesthesia of pregnant mouse dams or newborn mice and ultrasound-guided inoculation of virus into cerebral ventricles, or treatment of dams with TNFα to induce fetal infection following intraplacental inoculation of the virus [24, 25], the model of perinatal infection of newborn mice developed in our laboratory is based on intraperitoneal inoculation of the virus without the requirement for any additional pretreatment [26]. In addition, intraperitoneal inoculation of the virus results in systemic infection in which virus replication can be detected in parenchymal tissues, including the brain, as well as in blood cells and plasma (Fig. 1). Furthermore, and in contrast to intracerebral inoculation of MCMV, which results in localized infection corresponding to virus injection site in the brain parenchyma [27, 28], our model

Fig. 1 MCMV infection of newborn mice. Mouse offspring are infected with low doses of MCMV by injecting the virus intraperitoneally on day of the delivery (within 24 h after birth) (cartoon/graphics). Intraperitoneal inoculation of the virus results in systemic infection during which the virus enters the CNS. MCMV-infected cells are observed in the brain parenchyma (MCMV pp89+ cells). Productive infection in the CNS is detectable within days 7–17 p.i. (panel a, middle). Virus presence in the brain induces development of encephalitis characterized with numerous pathohistological lesions that reside in the brain parenchyma for months after the productive infection is abolished (panel a, bottom). Immune response in the MCMV-infected newborn brain is predominately derived from T lymphocytes among which CD8+ T cells highly outnumber CD4+ T cells (panel b). MCMV-infected newborn mice present with developmental abnormalities of the cerebellum; the size of the cerebellum is significantly lower than that seen in uninfected controls (panel c, top). Reduced cerebellar area in MCMV-infected mice is coupled with increased thickness of the external granular layer (EGL) of the cerebellum (panel c, middle) and impaired morphology of Purkinje cells (panel c, down) in cerebella as compared to control animals. (Some parts of the figure are taken from and modified with permission from the Journal of Immunology (44))

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most closely recapitulates the presumed route of CMV entry into the developing CNS via a systemic viremia [29]. Finally, newborn mice are neurodevelopmentally equivalent to human fetuses in the late third trimester of pregnancy [30]. These and other properties [31] make the mouse model of CMV disease an attractive and robust alternative to other animal models, including the primate model of CMV disease. 1.2 Developmental Abnormalities in the Brains of MCMV-Infected Newborn Mice

Newborn mice infected with MCMV present brain pathology and developmental abnormalities of the cerebellum, a finding similar to those described in studies utilizing either cranial ultrasound or magnetic resonance imaging in fetuses and infants congenitally infected with HCMV [26, 32]. In addition, MCMV infected newborn mice exhibit hearing loss associated with inner ear inflammation, loss of spiral ganglia neurons, and loss of cochlear vasculature as well [33–35]. Interestingly, a recent study identified impairment of olfaction as a consequence of a severe infection in the olfactory bulb as another sequela of congenital CMV infection in mouse model of intraplacental and intracranial infection of few day old pups [34]. The characteristic phenotype of the developing cerebellum in neonatally infected newborn mice includes reduced cerebellar foliation, decreased cerebellar area, and increased thickness of the external granular layer (EGL) [26] (Fig. 1). These morphological impairments result from reduced proliferation of granule neurons in the EGL, delayed migration of postmitotic neurons from EGL into deeper parts of the cerebellar cortex and impaired morphology of Purkinje cells in the cerebella of MCMV-infected newborn mice. In the same model, virus-infected cells and infiltrates of CD3+ mononuclear immune cells could also be observed in the spiral ganglion and stria vascularis of the inner ear [33, 34]. The exact mechanism of observed MCMV induced malformations is not yet fully understood. Impaired responsiveness to brainderived neurotrophins (BDNF) due to decreased expression of the BDNF specific receptor TrkB or a robust inflammatory response in the CNS mediated by expression of several proinflammatory cytokines and chemokines may play essential roles. Although developmental abnormalities correlated temporally with virus replication in the CNS, they were symmetric and affected the whole brain with no evident colocalization of the virus with the described abnormalities [26]. Cochlear tissue isolated from the inner ear of perinatally infected mice had increased expression levels of proinflammatory cytokines that persisted for months after infection [33]. Furthermore, MCMV-infected mice treated with common antiinflammatory agents, glucocorticoids, had decreased expression of proinflammatory cytokines and diminished developmental abnormalities of cerebellum while the level of virus replication was not impacted [36]. Finally, treatment of MCMV-infected

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newborns with anti-TNFα decreased the frequency of infiltrating activated mononuclear cells in the brain and the levels of proinflammatory cytokines, and also resulted in the correction of neurodevelopmental parameters and normalization of the expression of neurodevelopmentally essential genes, while not impacting replication of the virus within the brain [37]. Thus, it appears that the immune response, rather than the virus replication, is the main culprit responsible for neurodevelopmental malformations associated with congenital CMV infection. 1.3 Immune Response in MCMV-Infected Newborn Mice

Systemic MCMV infection in perinatally infected newborn mice affects nearly all tissues and organ systems and induces a robust inflammatory response in which inflammatory lesions are comprised of both polymorphonuclear and mononuclear cell infiltrates. Numerous cytomegalic cells and cells with eosinophilic inclusions coupled with focal necrosis can be observed in specific organs, most frequently in the liver. Sera of these mice contain elevated levels of TNFα that do not correlate temporally with the peak of virus replication in various organs, suggesting that tissue damage in peripheral organs of MCMV-infected newborn mice is cytokinemediated [38]. Although the expression of TNFα was also elevated in the brains of infected newborn mice, no sign of necrosis or tissue damage was detected [26]. This observation is likely attributable to efficient control of the immune response within the CNS; components of both the innate and adaptive immune response can be detected in the brain (Fig. 1). NK cells can be isolated from the CNS of infected newborn mice by day 7 postinfection (unpublished data) and, although NK cells represent the minor immune cell population in infected brain, a robust inflammatory response characterized by upregulation of proinflammatory genes associated with interferon production (IRF-1, IRF-7, USP18, and LRG-47) and chemokine secretion (TNFα, CxCL3, IFNγ, IL1β, MCP1, RANTES, CCL5, CCL21, and CXCL10) [26, 37] coincides with the appearance of NK cells and inflammatory monocytes in the brain parenchyma. The results mentioned above suggest that infiltrating NK cells and monocytes could be prominent producers of cytokines in the MCMV-infected newborn brain. MCMV-infected newborn mice, depleted of NK cells in the early postnatal period, showed significantly higher viral burden in the CNS following intracranial (i.e.) inoculation of the virus [27]. Studies performed on newborn mice infected with an attenuated MCMVs that express ligands for the activating NK cell receptor NKG2D Rae-1γ and MULT-1, show reduced virus replication in various organs, including brain, compared to wild type virus [39, 40], arguing for an active role of NK cells in the control of MCMV infection in newborn mice. A recent work, utilizing intracerebral inoculation, demonstrated that C57Bl/6 mice, which are less susceptible to MCMV infection, are also more

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resistant to sensorineural hearing loss caused by MCMV in comparison with MCMV-sensitive Balb/c strain [41]. C57Bl/6 mice owe their resistance to Ly49H, an activating NK cell receptor absent in BALB/c mice, that recognizes virally encoded protein m157 [42]. Intracranially infected BALB/c newborns, as well as C57Bl/6 newborns depleted of Ly49H+ NK cells, displayed a loss of outer hair cells of the organ of Corti that correlated with sensorineural hearing loss. Even though this finding may appear to argue in favor of the protective capacity of NK cells in perinatal MCMV infection of the CNS it remains to be seen whether this is the case when virus reaches the brain via systemic route. In the second week postinfection, monocytes are also recruited to the CNS. Activated microglia cells and infiltrating monocytes can be found close to virus-infected cells within encephalitic lesions, finding that argues for an active role of these cells in control of MCMV infection in the newborn mouse brain [37, 43]. Furthermore, neutralization of TNFα resulted in the diminished frequency of Ly6Chi CCR2+ inflammatory monocytes in the blood and their consequent diminished recruitment to the brain from the periphery as well as diminished activation of brain microglia [37]. Notwithstanding the robust recruitment of NK cells and monocytes, CD8+ T lymphocytes play a prominent role in the elimination of virus from the infected brain (Fig. 1). Starting from the second week postinfection, activation of T cells and the influx into infected CNS are readily observed [43, 44]. Infiltration of these cells into the infected brain correlates with a significant reduction of virus replication in the CNS. Moreover, depletion of CD8+ T lymphocytes in neonatal mice resulted in a significant increase in viral genome copy numbers in the CNS, and these mice succumbed to infection by postnatal day 15 [43]. Even more impressive is the fact that virus-specific CD8 T cells remain in the brain for over a year after perinatal infection and the resolution of productive CMV replication [44]. The phenotype of these cells was consistent with tissue-resident T cells and required active viral replication for their generation. In vivo significance of T lymphocytes isolated from the brain of MCMV-infected newborn mice at later stages, postinfection was confirmed by the adoptive transfer of brain-isolated mononuclear cells into γ-irradiated, immunocompromised animals. Mice that received CNS-isolated CD8+ T cells had a lower viral burden in the liver, lungs, and spleen, as compared to control mice [43]. In another work, purified populations of brain and splenic virusspecific CD8+ T cells were transferred either into newborns perinatally infected with MCMV, or into B-cell deficient adult mice intracranially challenged with MCMV. Both splenic and brain resident CD8+ T cells provided equal levels of protection [44]. Interestingly, not all brain resident CD8+ T cells were positive for CD103, a marker usually associated with tissue residence; however,

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CD103+ tissue resident cells were more responsive and proliferated more after stimulation and also offered better protection in adoptive transfer experiments [44]. Finally, long-term treatment of perinatally infected mice with depleting antibodies that lead to partial depletion of brain resident CD8 T cells resulted in increased reactivation of virus from latency and affected the activation status of microglia (more MHC II following depletion) which could be a consequence of uncontrolled and more frequent virus reactivation [44]. CD4 T cells also infiltrate the brain (Fig. 1) following perinatal infection of the mice and persist long after infection. Our recent study demonstrated that they contribute to the control of the virus both directly and indirectly through aiding in the generation and maintenance of tissue-resident CD8 T cells [45]. As with CD8 T cell depletion, depletion of CD4 T cells resulted in an increased frequency of latent virus reactivation. 1.4 Role of Antibodies in the Control of MCMV Infection in Newborn Mice

In adult mice, virus-specific antibodies appear to be dispensable for the resolution of acute CMV infections but are essential for the prevention of virus spread following reactivation [46, 47]. A pivotal role for virus-specific antibodies has been confirmed in congenital HCMV infection where preconceptual immunity reduces the rate of virus transmission from mother to child but fails to provide complete protection from infection of the fetus [48, 49]. This incomplete protection from virus transmission is considered to be the consequence of reinfection with a new serotype(s) of HCMV or reactivation of an endogenous virus [50, 51]. In a model of perinatal MCMV infection, antibodies have a proven role in the control and prevention of infection in the developing CNS [52]. When administered to infected newborn mice early after infection, antibodies provided protection and limited virus replication in the brain parenchyma, and when administered at later time points postinfection, even at the time of peak virus infection in the brain, the antibodies significantly reduced virus titers in the CNS [52] (Fig. 2). As a consequence, mice treated with antibodies had less pronounced inflammatory lesions in the brain parenchyma and improved postnatal cerebellar development as compared to MCMV-infected, untreated animals [52]. In addition, we have shown that female mice vaccinated before pregnancy with the highly attenuated virus carrying the ligand for NK cell activating receptor NKG2D induces an antibody response in dams that is sufficient to limit MCMV infection in MCMV-infected offspring [39, 40]. Experimental data therefore strongly argue in favor of a protective role for maternal antibodies in perinatal MCMV infection.

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Fig. 2 Antibody-mediated protection of MCMV infection in newborn mice. Transplacentally transferred antibodies from immunized dams provide efficient protection from MCMV infection to newborns since no

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Materials Standard equipment and material for cell culture, virus preparation, infection, and sacrifice of animals and tissue harvesting are necessary (hoods, incubators, centrifuges, microtomes, cryostat, pipettes, tubes, dishes, small surgical instruments, etc.). Additional primary materials are listed below: 1. Viruses: MCMV (most commonly used strains are Smith, K181, Bac-derived Smith strain). 2. Mice: most commonly used are BALB/c or C57BL/6. 3. Cell lines: primary or immortalized mouse embryonic fibroblasts (MEF). 4. Minimal essential medium (MEM). 5. Dulbecco’s Modified Eagle Medium (DMEM). 6. RPMI-1640. 7. Fetal calf (FCS) and newborn bovine serum (NBS). 8. Stainless steel wire mesh or bead homogenizer. 9. Cell strainer. 10. Methylcellulose medium (viscous medium): To 320 mL distilled water add 8.8 g of 0.16 M methylcellulose (4000 centipoises, Sigma). Stir with a large magnetic stirrer. Leave medium for 4 days at 4
C until methylcellulose has completely dissolved. Autoclave for 30 min at 121
C and then supplement the solution with 40 mL of 10 MEM, 40 mL FCS, 4  104 U penicillin, 40 mg streptomycin, 4 mL 1 M HEPES, and 0.88 g of Na2CO3. 11. Phosphate-buffered saline (PBS) commercial or made in house. 10 stock: 2.0 KCl, 2.0 g of KH2PO4, and 80 g of NaCl dissolved in 700 mL of ddH2O. 11.4 g of Na2HPO4 anhydrous dissolve in 200 mL of ddH2O with stirring and heating. Combine both solutions. Adjust pH to 7.4 with 5 M NaOH. Adjust volume with ddH2O to 1 L. 12. TBS: 1 M Tris–HCl, pH 7.4, 1 M NaCl. 13. Trypsin for cell culture: 0.25% trypsin plus 1 mM EDTA.

ä Fig. 2 (continued) productive infection was observed in organs of MCMV-infected newborn mice delivered from mothers infected with MCMV prior to pregnancy (panel a). Passively transferred virus-specific antibodies administered to MCMV-infected newborn mice prior to virus replication in the CNS reduce productive infection in the brain to undetectable levels (panel b). (Some parts of the figure are copyright © American Society for Microbiology, Journal of Virology, December 2008 vol. 82 no. 2412172-12,180 (53))

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14. 15% sucrose/VSB buffer: 50 mM Tris–HCl, 12 mM KCl, 5 mM Na2EDTA, adjust pH to pH ¼ 7.8 with HCl. 15. FACS (fluorescence-activated cell sorting) medium: PBS, 10 mM EDTA, 20 mM HEPES, pH 7.2, 2% FCS, 0.1% NaN3. 16. Erythrocyte lysing buffer (Tris/NH4Cl): 90 mL of 0.16 M NH4Cl, 10 mL of 0.17 M Tris–HCl, pH 7.65. Adjust final pH to 7.2 with HCl. 17. Ca-stock solution (500 mM calcium solution): 1.095 g CaCl2·6H2O in 10 mL pure RPMI. 18. Ca-RPMI: 100 mL 10% FCS supplemented RPMI, 1 mL 100 Ca-stock solution. 19. Collagenase D: Dilute the stock solution to 1 mg/mL in Ca-RPMI. 20. 3% acetone. 21. Paraformaldehyde. 22. Xylol. 23. NP 40 detergent: dissolve 500 μL of NP40 in distilled water. 24. Percoll (Amersham): 100% Percoll: dissolve 9 mL of Percoll with 1 mL 10 PBS (see Note 14). 25. Cresyl violet (0.1 g of cresyl violet in 100 mL water and 300 mL glacial acetic acid). 26. Relevant and properly validated antibodies [53] for the detection of various immune cell subsets. 27. IE1.01 antibody (Center for Proteomics, Croatia, cat. no. HR-MCMV-12) for MCMV or rIE1 for RCMV or any other antibody that stains MCMV or RCMV-infected cells in paraffin-embedded or frozen tissue sections, preferably recognizing an immediate-early antigen. 28. m04.10 (Center for no. HR-MCMV-01).

Proteomics,

Croatia,

cat.

29. 2.4 G2 antibody against murine Fc receptor. 30. FACS medium, commercially available or made in house (PBS with the addition of 1% BSA and 0.1% sodium azide). 31. Tetramers (Produced by NIH tetramer core facility, Bethesda, SAD (http://www.niaid.nih.gov/reposit/tetramer/overview. html). (a) H-2Ld-IE1168–176 (168-YPHFMPRNL-176): MHC class I molecule coupled with MCMV peptide IE1/m123pp89. (b) H-2Dd-m164257–265 (257- AGPPRYSRI -265): MHC class I molecule coupled with MCMV peptide m164.

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(c) H-2Dd-m04243–251 (243- YGPSLYRRG -COOH-251): MHC class I molecule coupled with MCMV peptide m04. 32. Ethylcarbazole (Sigma-Aldrich): 4 mL deionized water, 2 drops of acetate buffer, 1 drop of AEC chromogen (ethyl carbazole), 1 drop 3% hydrogen peroxide. 33. DAB (diaminobenzidine) (DAKO): 1 mL substrate buffer, 1 drop of DAB chromogen. 34. Propidium iodide (Sigma-Aldrich): dilute 10 μL in 990 μL of FACS media. 35. Entellan® mounting media.

3

Methods

3.1 Analysis of the MCMV Pathogenesis in Newborn Mice

The methods described below outline the procedures for investigating the MCMV pathogenesis primarily in the brain of newborn mice. Most of described methods, however, can also be applied to other organs and adult mice. When this is not the case, we have provided separate protocols. Following intraperitoneal (i.p.) and intravenous (i.v.—for adult mice only) inoculation, the virus establishes systemic infection, during which practically all organs can be affected, and the virus can be isolated from both blood cells and plasma [26, 52]. Productive MCMV infection in the CNS can be detected between days already at day 6–7 with the peak of virus replication between days 9 and 14 postinfection [26]. Immunohistochemical analysis of the brain tissue from infected newborn mice reveals no preference for the site of infection and development of widespread encephalitis. Both neurons and glial cells are permissive for MCMV infection [15, 25] and while lytic infection in the brain terminates within 3 weeks postinfection, pathohistological and functional lesions of the CNS remain detectable for several months. The preservation of neuronal lesions argues for either virus persistence in the brain that continually primes the immune response or consequent development of immunopathology in this immunologically privileged organ. The pathohistological lesions in virus-infected newborn mice closely resemble the ones described in autopsied cases of encephalitis in congenitally infected infants, stillbirths and fetuses [54] and are comprised of infiltrating monocytes and activated microglia cells spread throughout the brain parenchyma and also within specific sites of infection [26, 43, 52].

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MCMV is readily attenuated by sequential passaging in the culture of mouse embryonic fibroblasts [55], and the original virulence can be restored by propagating the virus in murine salivary glands [56]. Consequently, stocks of the more virulent salivary gland derived virus (SGV) might be the material of choice when performing lethal-dose experiments in adult animals [57]. On the other hand, less virulent, cell culture–derived virus should be used for infection of newborn mice due to their high sensitivity to MCMV infection. Cell culture–derived MCMV can be produced in either permissive primary murine cell lines such as primary embryonic fibroblasts (MEFs) and adherent mouse newborn cells (MNCs) [58]; or in immortalized cell lines, such as NIH 3T3, BALB/c 3T3, M2-10B4, and B12. Many immortalized cell lines divide much faster than primary ones and continue to divide even after infection with MCMV. This fact should be considered when seeding the cells for virus production. Additionally, many immortalized cell lines require media supplemented with 10% FCS, which contrasts with primary cells that require only 3% FCS. The protocol detailed below is suitable for production of wild-type or MCMV mutant strains in primary MEFs grown in DMEM supplemented with 3% FCS (see Notes 1 and 2). All steps should be carried out in a laminar flow hood. 1. Using six 15-cm culture dishes and freshly prepared or cryopreserved primary MEF cells, seed each 15-cm culture dish with approximately 5  106 primary MEF cells in a complete medium. Grow the cells until the cell monolayer reaches ~80–90% confluence (~11  106 cells/dish). Primary MEF cells that have been passaged for more than a total of two or three times before infection should not be used for the production of virus stocks. 2. Prepare the inoculum suspension by bringing together and thoroughly mixing ~3.3  106 PFU of MCMV from a purified virus stock and a complete medium in a total volume of 36 mL. If purified virus stock is unavailable, use 50–100 μL of unpurified or crude virus preparation with unknown titers for preparation of inoculum suspension. 3. Remove the medium from primary MEF cells prepared in step 1 and overlay the cell monolayer in each plate with 6 mL of inoculum suspension prepared in step 2, thereby performing the infection at an MOI ¼ ~0.01 PFU/cell. Incubate the infected cells for 4–6 h at 37
C on a horizontal surface in a CO2 incubator. Periodically gently tilt the dishes to ensure uniform contact of cells with the virus inoculum and prevent drying out the cells.

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4. Following incubation, add 14–24 mL of complete medium, prewarmed to 37
C, to each of the culture dishes and incubate the infected cells for 3–5 days at 37
C in a CO2 incubator (see Note 3). 5. Using a cell scraper, detach cells from the culture dish surfaces and transfer supernatant and primary MEF cells from each plate into a separate, prechilled, 50 mL centrifuge tube on ice. Separate the supernatant (containing virions) from the cellular material by centrifugation at 2000  g for 30 min at 4
C. Refrain from using trypsin for cell detachment, since the residual trypsin activity might digest the proteins present on the surface of the virions. 6. Transfer the supernatants into sterile, cold ultracentrifuge tubes on ice, and pellet the virus from the supernatants by ultracentrifugation at 50,000  g at 4
C for 90 min. 7. Decant almost the entire volume of the supernatant, but make sure to leave ~300 μL of the residual medium overlaying the pelleted virus in each ultracentrifuge tube. Then place the ultracentrifuge tubes on ice or in a cold room at 4
C overnight. 8. On the next day, thoroughly resuspend the virus pellets in the residual medium and pool the resuspended pellets together in a single ultracentrifuge tube and place it on ice. Depending on the volume of the leftover medium and the number of ultracentrifugation tubes used, the total volume of pooled virus suspension may exhibit noticeable lab-to-lab variability, but in general, varies between 2.5 and 5 mL. 9. Add 15–18 mL of the cold, 15% sucrose cushion solution into each of the two new, sterile, prechilled ultracentrifuge tubes on ice. Gently overlay equal volumes of the virus suspension from step 8 onto the surface of the sucrose cushion solution in each of the tubes. Pellet the virions by centrifugation at 70,000  g at 4
C for 90 min. 10. Completely remove the supernatant and overlay the pellets with 300 μL of ice-cold Mg2+-free and Ca2+-free PBS. Leave the tubes on the ice at +4
C overnight to facilitate subsequent resuspension of the pelleted virions. Make sure that PBS is completely covering the pellets in each tube. 11. The next day, thoroughly resuspend all virus pellets using a micropipette and pool all resuspended pellets into a cold microcentrifuge tube on ice. Optionally, further dissociate microscopic virus clumps by sonicating the pooled virus suspension on ice with a total of three 10-s, 150 W, 20 kHz ultrasonic pulses.

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12. Gently mix the virus suspension using a micropipette, distribute 25–100 μL aliquots to separate microcentrifuge tubes on ice, and store the freshly prepared virus stock at 80
C. If maintained at a constant 80
C and not subjected to freeze– thaw cycles, the stock virus suspension prepared in this manner is stable for years. 13. On the next day, thaw one aliquot and determine the virus titer as described in Subheading 3.1.3. For infection of newborn mice, virus stocks prepared according to the above protocol should be further diluted by a factor of 200 in a sterile DMEM, RPMI or PBS and obtained diluted virus stock titrated again. Titers of the diluted stocks should range from 105 PFU/mL to 106 PFU/mL. 3.1.2 Infection of Newborn Mice

In general, the pathogenesis of the CMV infection can be studied following infection of the experimental animals via the intravenous (i.v.), intraperitoneal (i.p.), footpad (f.p.), intranasal (i.n.), oral (o), intracranial (i.c.), or other routes. Even though the footpad, i.n. or the o route of infection may more closely mimic the natural transmission of the virus, i.n. and o routes appear to be less efficient or more challenging to control [10, 59]. For the reasons mentioned above and elsewhere in this protocol, we therefore recommend the infection of newborn mice via the i.p. route, which is known to result in systemic viremia in both adult and newborn mice [60, 61]. For adult mice, the most frequently used dose of cell-culture produced virus is 2–5  105 PFU/mouse in 100–500 μL of PBS or pure media. Furthermore, viral dose for the i.p. infection of newborns should be determined according to the experimental design, mouse strain used and the desired readout. Newborn mice can develop symptoms of the CMV infection following injection of as little as 100 PFU into their bodies. Administering significantly higher doses of the virus (1000 PFU/newborn mouse) can cause severe disease manifesting as stunted growth, hair loss, and a lethal outcome in a high proportion of infected newborns. Moreover, newborn mice of various mouse strains, such as the commonly used BALB/c and C57BL/6 strains, differ in their sensitivity to MCMV infection. Unlike in adults, newborn C57Bl/6 mice are more sensitive to MCMV than BALB/c newborns. Therefore, we suggest an experimenter to test the optimal dose in the desired strain. A good starting dose is 100–400 PFU/mouse in a maximal volume of 50 μL PBS, DMEM or RPMI. In addition to these considerations, know that adults of certain mouse strains tend to eat their newborns, especially if they have been touched or otherwise disturbed by the experimenter. To prevent cannibalism, plant the infected newborns to foster mothers of other mouse

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strains after infection. In our experience, BALB/c females make excellent foster mothers. Make sure to label the planted newborns if they have the same coat color as foster mothers. 1. Prepare the working virus suspension and insulin syringes in advance and keep the virus suspension on ice. 2. Fill the insulin syringe with the appropriate volume of working material. 3. Gently lift and constrain the newborn mouse with the nondominant hand in such a way to hold the animal belly up, by the neck with thumb and ring finger, and pulling gently by the tail with index and middle finger. The animal’s head should therefore be proximal, and the tail distal from the experimenter’s body. 4. Do not perform the infection by penetrating the needle directly through the abdominal wall and into the peritoneal cavity. Instead, with the dominant hand, pick up the insulin syringe containing the required amount of virus suspension, and insert the needle carefully under the skin in the upper thoracic region. Take special care not to puncture the liver and avoid disturbing the syringe plunger, in order to avoid unintentional and undesirable release of the virions to nontarget sites. 5. Slide the needle toward the peritoneal cavity. Again, be careful not to injure other organs, such as the liver or the urinary bladder. With the top of the needle in the correct position, gently press the syringe plunger to deliver the desired volume of the virus suspension to the peritoneal cavity. After inoculation, wait 4–5 s before removing the needle to avoid virus solution leaking out of body through puncture site during needle withdrawal. The same insulin syringe/needle combination can be used to administer the virus to all animals belonging to the same level (i.e., same virus concentration) of a given factor (i.e., virus strain) in the experimental design. 6. Administer the equal volume of virus-free RPMI, DMEM, or PBS to the control group of newborns (mock-infected). 7. Detach the used needle from the used insulin syringe and discard both according to the local guidelines and laws for the disposal of infectious and sharp materials. 8. Return the newborns to their mothers or foster mothers. 3.1.3 Determination of Virus Titers in Organs and Virus Stock Solutions Using Standard Plaque-Forming Assay

Even though various cell lines and types are permissive to MCMV infection, growth properties and plaque appearance make primary MEFs the first choice for the determination of virus titers. This protocol describes the use of primary MEFs grown in DMEM with 3% FCS as targets for determining the number of infectious particles in samples of virus suspensions. When comparing several

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different viruses or mouse strains, titrate one type of organ for all mice in all experimental groups at the same time (e.g., all spleens from the same experiment). Following euthanasia of mice and organ collection, freeze and store whole organs in microcentrifuge tubes at 20
C in 1 mL of complete medium. Avoid repeated freeze–thaw cycles of frozen organs or virus suspensions. 1. The day before titration, seed MEFs in 48-well plates in a complete medium at a cell density of 200,000 cells/well. We suggest seeding at least 30 wells with cells when titrating an individual virus stock solution, and at least 15 wells when titrating a single organ homogenate. Incubate the plates at 37
C in a CO2 incubator. 2. The next day, before use, thaw and keep mouse organs or virus stocks on ice. 3. Prepare organ homogenates by passing the organs through the 450-μm stainless-steel wire-mesh or 70 μm to 100 μm cell strainer into the 60-mm cell-culture dish on ice. Wash off the residual material from the mesh with 1–2 mL of complete medium and collect the entire organ homogenate into a 2 mL microcentrifuge tube. Alternatively, place the organs into 2 mL centrifuge tubes, add 1 mL of complete medium, appropriate amount of metal beads and prepare organ homogenates using bead homogenizer. Either way, place the fresh organ homogenates on ice as soon as they are prepared and immediately proceed with the next step of the protocol (see Note 4). 4. Preparing serial dilutions of virus stock. Label a total of nine microcentrifuge tubes with numbers from 2 to 9. Fill the microcentrifuge tube labeled with number 2 with 990 μL, and the remaining eight microcentrifuge tubes, labeled with numbers 3 through 10, with 900 μL of the complete ice-cold medium. Dilute the original virus suspension 100-fold by transferring 10 μL of the virus stock to 990 μL of the medium in tube number 2 and mix well. Prepare consecutive dilutions (103-fold to 1010-fold) of the virus stock by transferring 100 μL of the previously prepared dilution to subsequent microcentrifuge tube containing 900 μL of complete medium, and mixing well in between steps. Keep the microcentrifuge tubes on ice during the whole procedure. 5. Preparing serial dilutions of an organ homogenate. Label a total of four microcentrifuge tubes with numbers from 1 to 4, and add to each 900 μL of complete ice-cold medium. Prepare the ten-fold dilution of the organ homogenate by transferring 100 μL of the organ homogenate to 900 μL of the medium in tube number 2 and mix well. Prepare consecutive dilutions (102-fold to 104-fold) by transferring

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100 μL of the previously prepared dilution to subsequent microcentrifuge tube containing 900 μL of complete medium and mixing well in between steps. Keep the microcentrifuge tubes on ice during the whole procedure. 6. Remove media from MEFs prepared in step 1, overlay them in triplicate with 100 μL of each serial dilution of organ homogenate and incubate the plates for 60 min at 37
C in a CO2 incubator. During this step, the virus attaches to the cells. When titrating organ homogenates, to increase sensitivity, you can perform centrifugal enhancement by centrifuging the plates at 800  g for 30 min on room temperature (see Note 5). 7. Overlay the cell monolayers in each well with 500–1000 μL of methylcellulose nutrient overlay and incubate the plates at 37
C in a CO2 incubator for 3–4 days. Methylcellulose nutrient overlay is a very viscous medium that prevents horizontal spread of the virus. Due to high viscosity, it cannot be dispensed using a pipette and should be poured over cells from sterile beakers or graduated cylinders. 8. Count virus plaques using an inverted microscope. Count only those wells where individual plaques are well-separated and visible (no more than 100 plaques per well is a good rule of thumb). Calculate the number of PFUs per organ/aliquot/ gram of tissue, taking the dilution factor into account. 3.1.4 Extensive Titration

When investigating virus reactivation from latency, infectious virus particles in the target organs might be present in low numbers. In such cases, and in order to increase sensitivity and enable the detection of infectious particles, we recommend the use of an entire organ homogenate for titration. 1. The day before titration, seed MEFs into all wells on a 24-well plate, in a complete medium, and at a cell density of 500,000 cells/well. Prepare one fully seeded 24-well plate for every organ analyzed. 2. On the day of titration, thaw and keep mouse organs on ice. 3. Prepare organ homogenates by pressing the organs against the 450-μm stainless-steel wire-mesh; 70 μm to 100 μm cell strainer into the 100-mm cell culture dish on ice or using bead-mill homogenizer. If using mesh wire or cell strainer, wash off the residual material from the mesh with 5 mL of complete medium and collect the entire organ homogenate into a sterile 15 mL Falcon tube on ice. 4. Remove media from MEFs prepared in step 1, overlay cells in all 24 wells with 200 μL of the organ homogenate, and incubate the plates for 60 min at 37
C in a CO2 incubator. During this step, the virus attaches to the cells.

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5. Centrifuge the plates at 800  g for 30 min at RT (see Note 5). 6. Remove the plates from the centrifuge and incubate at 37
C in a CO2 incubator for an additional 30 min. 7. Overlay with 500–1000 μL of methylcellulose medium. Methylcellulose nutrient overlay is a very viscous medium that prevents the horizontal spread of the virus. Due to high viscosity, it cannot be dispensed using a pipette and should be poured over cells from sterile beakers or graduated cylinders. 8. Incubate the plates in a CO2 incubator at 37
C for 3–4 days until viral plaques become visible. 9. Count virus plaques using an inverted microscope. Calculate the number of PFUs per organ/aliquot/gram of tissue. 3.1.5 Determination of Virus DNA by Quantitative PCR

Quantitative PCR (qPCR) is sometimes used instead of standard virus plaque assay or extensive titration to detect viral DNA in organs or body fluids. However, while qPCR is a highly sensitive method, one important thing should be considered: the amount of viral DNA (number of genomes) is several orders of magnitude higher than the number of infectious virions. This discrepancy is most likely due to the existence of multicapsid virions and inefficient virus entry: without centrifugal enhancement there is usually 500 times more viral DNA than infectious virus particles, as determined by virus plaque assay, while this number falls to around 25 after centrifugal enhancement [62, 63]. Therefore, qPCR is reserved for cases when either the amount of virus is very low (e.g., in virus reactivation from latency), or when the sample in which virus should be determined is very small, both of which preclude the applicability of standard plaque assay. Detailed protocol, as well as validated primers and probes for the detection of MCMV genome or transcripts by qPCR, are already described elsewhere, so only a few crucial considerations will be included in this chapter [64]. We usually perform qPCR detection of viral DNA in experiments where the virus is reactivating from latency. Depending on the organ and the initial load of the virus, preamplification, as well as running qPCR for 50 cycles instead of the usual 40, might be needed to enhance the sensitivity of the assay [44]. We routinely use whole organ homogenates prepared as described in Subheading 3.1.3. The remaining homogenate can be used for extensive titration if desired. 1. Thaw organs on ice and, upon thawing, homogenize them in 1 mL of pure DMEM media in a bead-mill homogenizer. 2. Take 50–200 μL of homogenate and isolate total DNA. In general, for most organs, 100–200 μL of homogenate will give sufficient DNA. An exception is spleen, which, because of its high cellularity, contains more DNA/g of tissue. Take only 50 μL of spleen homogenate.

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3. Isolate DNA (we recommend Blood & Tissue kit from Qiagen). 4. Perform qPCR using primers and probes for viral DNA and mouse genes to allow cross-comparison between the samples (we use premade GAPDH assay from ThermoFisher). Make sure to include no-template control and mock-infected organs as negative controls as well as acutely infected organ (e.g., spleen or liver at 3 days postinfection) as a positive control. 5. The mouse genome should be detectable in the 8–25 cycles. If no virus DNA is detectable, include preamplification step for both mouse and viral DNA. Preamplification of both targets can be performed using the same primers or ready-made assays and in the same tube. Make sure to include no template control for the preamplification step, as well. 3.2 Immunohistochemical Detection of MCMV-Infected Cells and Immune Cells in Tissues

Histopathological evaluation of mouse organs is routinely performed to investigate the presence, extent and duration of lesions caused by the MCMV infection and is also used for detection of MCMV-infected cells. Following organ collection, tissue samples should undergo proper fixation, processing, embedding and cutting procedures. The details, advantages and disadvantages of specific approaches, materials and reagents used during tissue preparation can be found in modern textbooks [65] and are beyond the scope of this protocol. In general, however, IHC examinations of tissues affected by MCMV infection are performed using either frozen or paraffin-embedded tissue sections (see Note 6). For the purposes of detecting immune or MCMV-infected cells, we suggest adding blocking serum, primary and secondary antibodies, enzyme–avidin complex, and chromogenic substrate with a pipette directly onto the slides in total volume of 200 μL per slide. All washing steps should be performed in ample amounts of deionized water to prevent background staining. In case of high background, we suggest an increase in the number of washing steps and their duration. For the detection of MCMV-infected cells in paraffinembedded tissue, we routinely use anti-IE1 antibody for intranuclear staining or anti-m04 for cytoplasmic staining, both produced in our laboratory.

3.2.1 Preparation of Paraffin-Embedded Tissue Sections for IHC

1. Using Histokinette, deparaffinize slides by rinsing them in 3 xylol, followed by washing steps in a series of solutions containing decreasing concentrations of ethanol: Xylol 3 8 min, 100% ethanol 2 4 min, then 90%, 80%, 70%, and 50% ethanol, each for 4 min. 2. Wash slides in deionized water for 1 min. 3. Perform antigen retrieval, if needed (see Note 7).

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4. Wash with deionized water several times for few mins (e.g., 3  2 min). 5. Block tissue peroxidase in commercially available blocking solution or 30 mL PBS + 30 mL methanol + 1 mL H2O2 for 30 min. Blocking solution should be made right before use. 6. Wash with deionized water several times for few mins (e.g., 3  2 min). 7. Prevent unspecific antibody binding with blocking solution and incubate for 25 min at room temperature (see Notes 8 and 9). 8. Add primary antibody (e.g., mouse anti-MCMV pp89) and incubate for 60 min at 37
C or overnight at +4
C. 9. Wash with PBS several times for few minutes (e.g., 3  2 min). 10. Add biotinylated secondary antibody (e.g., goat anti mouse IgG) and incubate for 30 min/RT (see Notes 9 and 10). 11. Wash with PBS several times for few minutes (e.g., 3  2 min). 12. Add enzyme–avidin complex (e.g., Streptavidin-POD, streptavidin-AP) and incubate for 30 min at RT. Enzyme–avidin complex should be prepared 15 min before use. 13. Wash with PBS several times for few minutes (e.g., 3  2 min). 14. Add chromogenic substrate (DAB, AEC, or any other) according to manufacturer’s instructions. 15. Wash under running water few minutes. 16. Wash with deionized water several times for few minutes (e.g., 3  2 min). 17. Counterstain with hematoxylin and mount with permanent medium according to manufacturer’s instructions and taking into account chromogenic substrate used. 3.2.2 Preparation of Frozen Tissue Sections for IHC

1. Dry sections for 15–20 min at RT. 2. Rinse slides for 3–4 min in cold PBS. 3. Fix slides in cold acetone (4
C) for 5 min or 2.5% PFA for 20 min (see Note 11). 4. Wash with PBS several times for few mins (e.g., 3  2 min). 5. Block tissue peroxidase in commercially available blocking solution or 30 mL PBS + 30 mL methanol + 1 mL H2O2 for 30 min. Blocking solution should be made right before use. 6. Wash with PBS several times for a few mins (e.g., 3  2 min). 7. Add blocking solution and incubate for 25 min at RT (see Note 8). 8. Add primary antibodies incubate for 60 min at RT or overnight at +4
C (see Note 9).

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9. Wash with PBS several times for few mins (e.g., 3  2 min). 10. Add biotinylated secondary antibody and incubate for 30 min at RT. 11. Wash with PBS several times for few mins (e.g., 3  2 min). 12. Add SA-POD and incubate for 30 min at RT. 13. Wash with PBS several times for few mins (e.g., 3  2 min). 14. Add chromogen (DAB, AEC, or any other) and incubate according to manufacturer’s instructions. 15. Wash thoroughly in deionized water. 16. Counterstain with hematoxylin and mount with permanent medium according to manufacturer’s instructions and considering chromogenic substrate used. 3.3 Assessment of Developmental Abnormalities in Brains of MCMV-Infected Newborn Mice and Consequent Neurological Impairments

Pathohistological lesions and morphometric analyses are performed on cresyl violet–stained paraffin-embedded brain sections. For morphometrical analyses (circumference of the cerebellum, thickness of external granular layer, etc.) any free or commercial morphometry software is suitable. We use CellSens (Olympus).

3.3.1 Histomorphometrical Analyses of the Brain—Cresyl Cresyl Violet Staining

Cresyl violet staining can be used for both paraffin-embedded and frozen sections. Because of its high affinity for Nissl’s bodies it is highly suitable for staining of neuronal structures in central nervous tissue. Cresyl violet solution can be made in advance and stored at 4
C. Prior to use it needs to be filtered through filter-paper. 1. Deparaffinize the sections in xylol 2  10 min, then hydrate the slides in 100% ethanol 2  5 min, 95% ethanol 3 min, and 75% ethanol 3 min. 2. Wash under tap water several times for few minutes (e.g., 3  2 min). 3. Repeat washing with ample amounts of deionized water. 4. Stain the slides in 0.1% cresyl violet for 5 min on room temperature (see Note 12). 5. Wash with ample amounts of deionized water several times for few minutes (e.g., 3  2 min). 6. Rinse slides in 90% ethanol for 15 min to wash off excess dye.

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7. Dehydrate the slides in 100% ethanol 2  5 min. 8. Dehydrate the slides in xylol 2 5 min. 9. Mount the slides with permanent mounting medium (e.g., Entellan). Measurement of Cerebellar Area and External Granular Layer Thickness

1. Measure cerebellar area on serial cresyl violet–stained sections under the microscope on small magnification so whole cerebellum is visible. 2. Determine the thickness of the external granular layer by marking eight points along the primary cerebellar fissurae on serial cresyl violet–stained sections on the microscope, under higher (e.g., 20) magnification.

3.3.2 Balance Beam Test

1. Use a narrow beam, 90 cm long and 2 cm in diameter. 2. Position a beam 10 cm above the ground and place a mouse in the middle of a beam. 3. Measure the distance the mouse has crossed in 2 min (see Note 13). 4. Repeat the process.

3.4 Assessment of Inflammatory Response in MCMV-Infected Newborn Brain

Following virus entry into newborn mouse CNS, the initial innate immune response is mediated by resident glia cells which secrete proinflammatory cytokine and recruit other immune cells from the periphery [26, 36, 37, 45, 61]. While the primary role of brain infiltrating immune cells is the control of the virus replication, a growing body of evidence indicates that their activity is a major cause of pathological lesions and long-term neurological sequalae observed in CMV-infected individuals. Multiparametric flow cytometry is the most used method for analysis of phenotype and functionality of immune cells since it allows for fast, precise, and simultaneous detection of multiple parameters. The main prerequisite for performing flow cytometric analysis is dissociation of the tissue to obtain single cell suspension. Parenchymatous soft tissues (like spleen and brain) can be dissociated mechanically by passing through mesh wire or cell strainer. Tissues containing collagen capsule and stroma benefit from pretreatment with collagenase D to enzymatically disrupt extracellular matrix before passing it through mesh wire or cell strainer. Bear in mind that all these treatments stress the cells and can alter their phenotype. Additionally, enzymatic treatment may cleave off some surface markers, so it is generally a good idea to test the sensitivity of your desired markers to your selected tissue disruption method. Tissues will also contain numerous nonimmune cells that can express markers used to identify functionality of immune cells. For this reason, mononuclear immune cells are often enriched by Percoll and/or staining the cell suspension with an antibody against specific marker of leukocytes CD45.

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Dead cells can be a major source of unspecific antibody binding so the use of viability dies is highly recommended. Moreover, nowadays when there is such a wide range of dyes to satisfy every requirement for excitation and emission maximum, there is no excuse not to include them in your analysis. Viability dyes come in two flavors: DNA binding (or intercalating) dyes and aminereactive dyes that bind to amines. DNA-binding dyes, like propidium-iodide (PI), SYTOX, 7-AAD, or DAPI, fluoresce when bound to double-stranded nucleic acids. Live cells, with intact plasma and nuclear membranes, are thus left unstained [66]. These dyes are relatively cheap and simple to use: just add them a few minutes before analysis. However, their use often results in strong background fluorescence, especially if the analyzed cells contain a lot of RNA or in samples with lots of dead and degraded cells. Propidium iodide is also notoriously sticky and will adhere to sample tubing in the flow cytometer and can thus get transferred from one sample into the next. These dyes are also unsuitable for use in fixed samples. Amine-reactive dyes are also cell membrane impermeable, like DNA-binding dyes. In contrast with DNA-binding dyes, aminereactive dyes bind irreversibly to amines making these dyes suitable for dead cell exclusion in fixed samples. Important note to remember is to add them before sample fixation, along with antibodies against surface antigens. Amine-reactive dyes will also stain live cells (there are still some amines on the cell surface) but to a significantly lesser extent than dead cells [67, 68]. 3.4.1 Isolation of Mononuclear Cells from Brain and Similar Soft Tissues (e.g., Liver)

1. Infect animals as described above. 2. Place the brain in a cell-culture dish in ~10 mL of cold media (RPMI or DMEM) on ice. 3. Cut the organ into small pieces using sterile scissors or a scalpel and then pass through metal wire-mesh or cell strainer to obtain single cell suspension. 4. Centrifuge at 500  g for 5 min. 5. Resuspend the cell pellet in cold media and add Percoll to desired density. The total volume of media and Percoll should be 10 mL (see Note 14). 6. Slowly overlay the suspension onto 10 mL of diluted Percoll (see Note 14). 7. Centrifuge at 700  g for 30 min at 4
C without brake. 8. Remove buffy middle layer and transfer into a new tube. 9. Add media to bring volume to 10 mL and pellet the cells at 200  g for 10 min at 4
C to remove Percoll (see Note 15). 10. Resuspend pellet in desired volume of media or PBS.

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1. Prepare single cell suspension of splenocytes by passing the spleen through wire mesh or cell strainer. To facilitate passing through cell strainer you may cut the spleen into smaller pieces with scissors. We often use syringe plunger to push the spleen through the cell strainer. 2. Pellet the cells by centrifugation at 500  g for 5 min at room temperature. 3. Lyse erythrocytes by resuspending cell pellet in 5 mL of erythrocyte lysis buffer and incubating on ice for 5 min. Stop the lysis by addition of 10 mL media. Pellet the cells at 200  g for 5 min at room temperature. 4. Resuspend in 10 mL media.

3.4.3 Isolation of Mononuclear Cells from Blood

1. Prepare tubes for blood collection by adding 60 μL of 0.5 M EDTA to prevent clotting. 150–300 μL of blood can be obtained from one mouse by cardiac puncture procedure or from orbital vein from adult mice, while less blood can be obtained from the tail vein. 2. Dilute the blood with double volume of PBS and overlay over 3–5 mL of lymphoprep. 3. Spin at 800  g for 10 min with no brake. 4. Collect middle layer (buffy coat) with pipette and transfer to new tube. 5. Add media/PBS to total volume of 10 mL to wash lymphoprep. 6. Spin at 500  g for 5 min at room temperature. 7. Lyse erythrocytes with erythrocyte lysis buffer, if needed as described above.

3.4.4 Isolation of Mononuclear Cells from Lungs

1. Remove lungs, place in one well of a 6-well plate and cut them to small pieces with scissors. 2. Add 3 mL collagenase D (1 mg/mL) and incubate at 37
C for 60 min. 3. Pass the tissue through wire mesh or cell strainer and wash with 5 mL RPMI. 4. Pellet the cells by centrifugation at 500  g for 5 min. 5. Resuspend in 4 mL of RPMI. Overlay over 30% Percoll in a new falcon tube. 6. Spin at 800 g for 30 min with no brake. 7. Remove supernatant carefully and resuspend in 5 mL of RPMI. 8. Spin at 500  g for 5 min. 9. Lyse erythrocytes by addition of 2 mL of erythrocyte lysis buffer if needed as described above.

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10. Stop the lysis by addition of 5 mL of RPMI. 11. Pellet the cells at 500  g for 5 min. 12. Resuspend the pellet in 1 mL of RPMI. 3.4.5 Labeling of Surface Targets for Flow Cytometry Analysis

Surface staining can be performed on fixed or unfixed samples. If surface staining is performed on unfixed, live samples, all steps should be performed on ice to slow down cellular metabolism and turnover of surface markers. Make sure to protect your fluorescently labeled antibodies from direct sunlight to prevent fluorochrome photobleaching and degradation. In order to save expensive antibodies, we usually stain up to five million lymphocytes in 50 μL. 1. Prepare single cell suspensions as described above and determine the concentration of the cells in the suspension (by hemocytometer or automatic cell counter). 2. Place the cells in 1.2 mL microtubes or in multiwell plates with rounded or V-bottom. Plan for 1–5 million cells per staining, depending on the rarity of immune cell population you wish to analyze. If needed, the cells can be pelleted at 500  g for 5 min and then resuspended in FACS medium at a more suitable density. 3. Pellet the cells at 500  g for 5 min, remove the supernatant by aspiration or by flicking off the excess fluid and resuspend them in small volume (e.g., 50 μL) of Fc-blocking solution (see Note 16) and incubate on ice for 15 min. 4. Prepare Fc blocking solution in FACS medium to prevent unspecific binding of antibodies due to Fc receptors present on numerous immune cells. We use 2.4 G2. 5. Resuspend the cells in Fc block solution and incubate at +4
C for 15 min. 6. Prepare solution of your primary surface antibodies in FACS medium. If you wish to use amine-reactive viability dye, you may include it in this step as well. 7. Wash cells thoroughly with ample amounts of FACS medium (1 mL for microtubes or 200 μL for multiwell plates). 8. Pellet the cells at 500  g for 5 min, remove the supernatant by aspiration or by flicking off the excess fluid and resuspend in antibody solution. Incubate for 20–30 min on ice, protected from direct light (e.g., covered with aluminum foil). 9. Prepare solution of your secondary antibodies (if applicable) and repeat steps 5 and 6. 10. Resuspend in 200 μL of FACS media for analysis on flow cytometer and proceed to the analysis as soon as possible. If DNA-binding viability dye is used, prepare a dilution of the dye

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in FACS media or PBS according to manufacturer’s recommendation and add it right before analysis on flow cytometer. If the flow cytometer is not available within the next few hours, we recommend you fix the cells and store them protected from light (see the protocol and further recommendations below). 3.4.6 Fixation of Cells for Flow Cytometry

Cell fixation in flow cytometry is used to “freeze” the cells in certain state, to allow for permeabilization for intracellular antigen detection and for safety reasons when working with infectious or human samples. There are many commercially available fixatives one can use but most commonly used are formaldehyde. Once fixed, the cells can be kept for several weeks and analyzed at your convenience. We usually fix the cells following the labeling of surface antigens with antibodies; however, fixation can be performed before. Bear in mind that fixation can alter the availability and structure of the epitope. Fixed cells have a smaller size and higher autofluorescence than unfixed samples, so if you are using cells to prepare single-stained controls for compensation, use fixed cells as well.

Fixation of Cells for Flow cytometry Using Formaldehyde

1. Pellet the cells at 500  g for 5 min, remove the supernatant by aspiration or by flicking off the excess fluid and resuspend them in 100 μL of 0.5–4% formaldehyde in PBS. For general purposes and for most antigens we have tested, 2% formaldehyde usually works well. 2. Incubate at room temperature, protected from light (if there are fluorescently labeled antibodies or proteins present in the sample) for 20 min. 3. Wash with 900 μL (for microtubes) or 100 μL (for 96-well plates) of FACS media and spin down at 800  g for 5 min. 4. Proceed to permeabilization or resuspend the cells in FACS media for storage or subsequent analysis on flow cytometer.

3.4.7 Permeabilization of Cells and Intracellular Staining for Flow Cytometry

Permeabilization of Cells Using Methanol

If analysis of intracellular markers is required, the cells need to be fixed first and then permeabilized in order to allow antibodies to reach them. Although numerous commercially available permeabilization reagents are available, permeabilization with methanol or saponin is very efficient and quite cheap. The decision on which method for permeabilization to use depends on antibodies (refer to manufacturer’s recommendations). In our lab, we routinely use saponin for most purposes and methanol for the detection of phosphorylated signaling molecules. 1. Pellet the cells at 500  g for 5 min remove the supernatant by aspiration or by flicking off the excess fluid and resuspend them in 50 μL of PBS.

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2. Add 500 μL of ice-cold 100% methanol drop by drop. Mix by gentle vortexing. Volumes can be modified—the final concentration of methanol needs to be 90%. 3. Incubate on ice for 30–40 min. Alternatively, this is a convenient stopping point if you cannot proceed. Store samples in the 20
C fridge. 4. Pellet the cells at 500  g for 5 min and wash two times with ample amounts of FACS medium. 5. Continue staining your intracellular targets in FACS medium in a manner identical as described for surface staining. Permeabilization of Cells Using Saponin and Staining of Intracellular Targets

1. During fixation prepare solution of primary antibodies against intracellular targets. Antibodies should be resuspended in 0.1% saponin in PBS solution (see Note 17). Add antibody solution, resuspend the cells and incubate the samples at room temperature for 20–30 min protected from light. 2. Wash thoroughly with 0.05% saponin in PBS (or FACS medium) then pellet the cells at 500  g for 5 min and remove the supernatant. 3. Add secondary antibody solution (if applicable) in 0.1% saponin in PBS, resuspend the cells and incubate the samples at room temperature for 20–30 min protected from light. 4. Wash thoroughly with 0.05% saponin in PBS (or FACS medium) then pellet the cells at 500  g for 5 min and remove the supernatant. 5. Resuspend the cells in 200 μL FACS medium and analyze on flow cytometer.

3.4.8 Measuring Cytokines in Flow Cytometry

Cytokines may be measured inside the cell that produced them by performing intracellular staining in flow cytometry (or immunohistochemistry) or in cell culture supernatants, tissue fluids and tissue homogenates. In addition to ELISA that has been classically used to detect cytokines, a very handy method is cytometric bead array or Luminex technology that utilizes beads. Beads come in different sizes and can be easily discriminated in flow cytometer thus enabling simultaneous detection of multiple cytokines in the same, usually small (50–100 μL) sample volume. These procedures are very well described by the producer and will thus not be described here. Measuring secreted cytokines in the supernatant or organ homogenates will not give information which cells secreted them. If you wish to know which immune cell secreted which cytokine, intracellular staining has to be performed. Since cytokines are secreted out of the cell quite rapidly, immune cells need to be incubated in Brefeldin A for a few hours to prevent cytokine secretion. Additionally, cells usually need a stimulus either

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from antibodies against activating immune cell receptors or via a broader stimulus (LPS, PMA and ionomycin, CpG, etc.) to induce cytokine secretion in culture. 3.4.9 Mononuclear Cell Adoptive Transfer (Via Cell Sorting)

Adoptive cell transfer into MCMV-infected immunodeficient mice is a frequently used method to analyze the functional capacity of these cells to control MCMV infection in vivo. Blood, spleen, and liver are the most accessible sources of mononuclear cells; however, any organ can be used as source of leukocytes. Cell adoptive transfer can be done either by cell sorting or by transferring total number of cells from a cell sample normalized to the desired number of target cell subpopulation. In both approaches, care should be taken to make sure that donor and acceptor mice are syngeneic to avoid graft-versus-host disease. Prior to adoptive transfer, donor mice may be treated with monoclonal antibodies to deplete the unwanted cell subpopulations that could influence the results of the experiment. Acceptor mice may also be treated with depleting antibodies or γ-irradiation to deplete their innate immune cells. Adoptive transfer of all cells normalized to target cell numbers is a simple and fast method with very little manipulation of transferred cells. However, all cells are transferred and in many cases that includes unwanted cell populations that can have a significant impact on the outcome of the experiment. Nowadays there exists many inbred mouse strains lacking different cell type, which can be used as donors in adoptive transfer experiments (e.g., SCID mice as NK cell donors to avoid contamination with B and T cells). There are distinct benefits to cell sorting—the use of sorted cells allows only the transfer of a defined cell population with no interfering cell subpopulations (e.g., CD8 T cells in NK cell adoptive transfer). However, direct labeling of cells with monoclonal antibodies can cause their activation. Additionally, transfer of cells marked with antibodies can lead to rejection of transferred cells via antibody-mediated cellular cytotoxicity (ADCC). Stripping of bound antibodies post sorting from the surface of the cells is not very efficient and leads to loss of viable cells. ADCC and activation of cells can be avoided by utilizing negative sorting—that is, by labeling all the unwanted cell subpopulation or culturing sorted cells for 24–48 h before transferring them into animals. The purity of negative selection sorts is much lower in comparison to directly labeled and sorted cells since committed cell precursors may not express markers used for sorting out a particular cell subpopulation. Additionally, negative cell sorting requires a significant amount of monoclonal antibodies. Cell sorting presents a stress for the cells, which can influence their expression patterns and behavior posttransfer. When in doubt, use lower sorting speed and pressure and try to shorten sorting

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time. Sorting time depends greatly on the amount of the target cell population. The cells can be enriched for sorting beforehand by using either negative or positive cell selection on commercially available columns (e.g., Miltenyi MACS) followed by sorting on cell sorter. Another, often overlooked, parameter is the vessel the cells are sorted into by the cell sorter. Cells are sorted by electrically charging the droplets containing cells and directing them into acceptor vessel via electric field. The tiny droplets with cells hit the acceptor vessels at high speed and may remain stuck there owing to the charged plastic of the vessel. The tiny droplet then dries out quickly, which results in the death of the sorted cell. In order to prevent that issue, it is advisable to coat your acceptor vessel by filling it with medium optimal for sorted cell type and 10–20% FCS. The proteins in the medium coat the plastic and allow the tiny drops to slide to the bottom of the tube. It is also recommended to sort into medium and not into empty vessels. Finally, many cell media are formulated for optimal pH at atmosphere with 5% CO2, and thus do not maintain the optimal pH in regular atmosphere during sorting. Media with HEPES are very good at maintaining buffer for these purposes. Adoptive Cell Transfer by Cell Sorting

1. A day in advance prepare tubes into which the cells will be sorted (acceptor tubes) by coating their sides with media rich in proteins (10–20% FCS, incubated overnight at +4
C with rotation). Alternatively, coat at 37
C for a few hours prior to sorting. 2. Remove the organ–source of transferred cells from the mouse aseptically in laminar flow cabinet. 3. Obtain single cell suspension by cutting the organ to smaller pieces with surgical scissors and then passing it through stainless-steel wire mesh or cell strainer in 10 mL of RPMI. 4. Pellet the cells by centrifuging at 500  g for 5 min. 5. Lyse erythrocytes by addition of 5 mL sterile lysing buffer and incubate on ice for 5 min. 6. Stop the lysis by addition of 10 mL sterile RPMI. 7. Pellet the cells by centrifuging at 500  g for 5 min. 8. Resuspend in 10 mL of sterile RPMI and count the cells. 9. Stain the cells with desired antibodies making sure that the antibodies and/or FACS media do not contain any azide (see Note 18). 10. Remove media from acceptor tubes leaving only 1–2 mL of cell culture media supplied with 10–20% of fetal calf serum (see Note 19). 11. Sort the cells into acceptor tubes. If possible, cool the sorted cells during the procedure.

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12. Retrieve an aliquot of sorted cells, add 100 μL of 1:100 diluted propidium iodide solution (not needed if amine-staining viability dye was used prior to sorting) in and check purity and viability of sorted cells on FACS analyzer or sorter. 13. Pellet the cells by centrifugation at 800  g for 10 min and resuspend in 150–500 μL of pure RPMI or PBS. 14. Inject the sorted cells into tail vein. Adoptive Cell Transfer of all Cells Normalized to Target Cell Numbers

1. Start from step 7 of the previous protocol “Adoptive cell transfer by cell sorting.” 2. Collect an aliquot of cells (typically 1–2 million cells is enough) for flow cytometric analysis and stain them for specific markers following protocols for surface or intracellular staining of cells for flow cytometry. Include a viability dye for your studies. 3. Analyze the cells on the flow cytometer and determine percentages of live, target cell population. 4. Calculate number of all cells needed to transfer selected number of target cells. Example: if you want to transfer 5000 CD8 T cells from spleen, which contains 10% live CD8 T cells, then to transfer that number of CD8 T cells you need to transfer 5000/ 0.10 ¼ 50,000 total splenocytes. 5. Resuspend the number of total cells in maximal total volume of 500 μL of RPMI and inject the cells into tail vein.

3.4.10 Depletion of Immune Cell Subsets in the Brain

The depletion of immune cell subsets is very commonly used to assess the impact of a particular type of immune or other types of cells for virus pathogenesis or analyzed process. Unlike in transgenic knock-out mice, depletion with antibodies is transient and depleted animals have little time to develop compensatory mechanisms like those born with the deficiency. Unfortunately, it is impossible to deplete cell subsets in only one organ. In addition, the depletion of cell subsets in the brain is made even more difficult by the blood-brain barrier necessitating a very long treatment with depleting antibodies [44, 45]. In general, we recommend using mouse-derived antibodies whenever possible. In cases when mouse antibodies are not available, mouse strains lacking antibodies and B cells (e.g., JHT (B6.129P2-Igh-Jtm1Cgn/J) or μMT/μMT (B6.129S2-Ighmtm1Cgn/J)) can be used to prevent the development of mouse antibodies directed against depleting alloantibodies. The efficiency of the depletion depends on the age of the treated mice and state of the blood-brain barrier. We have managed to significantly reduce the amount of tissue resident CD4 and CD8 T cells in 2–4-month-old mice by injecting depleting antibodies every week for a month [44, 45]. Optimal doses of depleting

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antibodies should be determined experimentally for each depleting antibody. In general, usual doses for newborns are 50 μg/mouse and 150–300 μg/mouse for adults. 1. Prepare depleting antibodies in 50 μL of PBS or pure DMEM for newborn mice or 100–500 μL of PBS or pure DMEM for adult mice. 2. Inject the mice with the prepared antibody-solution via i.p. route. 3.5 Assessment of Antiviral Antibody-Mediated Protection of MCMV Infection of Developing Brain 3.5.1 Preparation of Hyperimmune Serum

1. Infect mice with 2  105 PFU of w.t. MCMV via i.v. or i.p. route. 2. Sacrifice mice 3–4 months postinfection. 3. Use insulin syringe and perform cardiac puncture to aspirate the blood. 4. Leave the blood at room temperature for 10 min to allow it to clot. 5. Centrifuge the blood at 500  g for 5 min to separate the cells from serum. 6. Pool the serum from several animals and store it at 20
C.

3.5.2 In vivo Administration of Immune Serum or Monoclonal Antibodies to Newborn Mice

4

1. Infect newborn mice within the first 24 h after birth with 200–500 PFU of w.t. MCMV (see Subheading 3.1.2). 2. Inject 100 μL of immune serum intra peritoneally using insulin syringe at 5 or 9 days postinfection by inserting the syringe subcutaneously in upper thoracic region. Slide the needle up to peritoneal cavity carefully not to injure liver or urinary bladder.

Notes 1. Take care not to allow cells to dry during the procedure. We recommend preparing all media and virus stocks in advance (see Subheading 3 for all reagents needed). 2. Host cell components may be present in virus preparations; therefore, in vivo experiments should be conducted with virus grown on cells syngeneic to the animals used in experiments. 3. During the incubation period, cells should be monitored on a daily basis for the appearance of cytopathic effects, whereby infected cells become rounded (cytomegalic) and start detaching from the surface of the dish. Ideally, all cells in a dish should exhibit cytopathic effect before continuing with the procedure. 4. Most mouse organs are sufficiently small that they can be homogenized whole and their titers expressed as PFU (plaque-forming units) per whole organ. Adult liver, brain,

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and skin are notable exceptions. For these organs prepare sterile microtubes prefilled with media and weigh them in advance. Then cut a small portion of brain, liver or skin in laminar flow hood and place it into the preweighed tube and weigh it again. The difference between empty and filled microtube is the organ weight. Obtained virus titers are divided by the weight of the organ and thus expressed as PFU/g of tissue. 5. Centrifugal enhancement (30 min, 800  g at room temperature following addition of the virus to the cell layer) facilitates synchronous entry of the virus into the cells and enhances infection by 10–25 fold. When determining virus titer in virus stocks centrifugal enhancement is not used. 6. The choice largely depends on the primary antibodies used, as some antibodies do not work on paraffin embedded samples. 7. In some cases, tissue pretreatment may mask antigen epitopes and will require antigen retrieval as an additional step. This is especially true for formalin-fixed tissue. Antigen retrieval is usually performed by boiling slides in various buffers (e.g., citrate, TRIS, or EDTA buffers) or by enzymatic digestion. The type of antigenic retrieval depends on the antibody used and many reputable antibody suppliers offer recommendations for their particular antibodies. 8. Various blocking solutions are commercially available. Alternatively, you may use 3% BSA in TBS or immune serum with irrelevant antibodies from animal species which are the source of secondary antibodies you will use (e.g., if your secondary antibody is derived from rat, use rat serum to block). 9. Blocking serum, antibodies, and enzyme–avidin complex are diluted in 1% BSA-TBS. 10. Although IHC detection can be done with primary antibodies coupled with peroxidase or fluorochrome, usage of secondary or even tertiary antibody increases signal intensity. This is especially true when using secondary biotinylated antibodies detected with streptavidin-POD. Multiple secondary antibodies can bind to the Fc-fragment of the primary antibody. Each secondary antibody can have multiple biotin molecules attached to it, which can further bind multiple streptavidins. 11. Fixing in acetone preserves the tissue structure better but is more damaging to the antigens; on the other hand, PFA conserves antigens but damages the tissue. 12. For slides thicker than 20 μm, cresyl violet solution should be heated to 37–50
C. 13. If the mouse falls from the beam it should be excluded from further analysis and additional beam test should be performed 5 min later.

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14. For isolation of mononuclear cells from brain, the upper suspension should contain 30% Percoll, while the lower should contain 70% Percoll. For liver, upper solution should contain 40% Percoll, while the lower should contain 80% Percoll. 100% Percoll is prepared by mixing 9 mL of Percoll and 1 mL of 10 PBS. 70% Percoll is made by mixing 7 mL of 100% Percoll and 3 mL of media. 15. If perfusion or density gradient centrifugation was not efficient and coagulated blood is still visible after the step 8, add 1 mL of erythrocyte lysis buffer onto pellet and incubate in ice for 1–2 min. Stop the lysis by addition of 10 mL media. (Note: this step will lower your yield). 16. Fc blocking serves to minimize unspecific antibody binding due to the Fc-receptors present on various immune cells (e.g., NK cells, monocytes, macrophages, granulocytes, B lymphocytes). Purified antibodies against murine CD16/CD32 are commercially available. Alternatively, immune serum or irrelevant antibodies may be used but take care that your secondary antibodies (if used) do not recognize your blocking antibodies. MCMV encodes its own Fc receptor [69] and there are currently no blocking antibodies available. In this case, if you are analyzing MCMV-infected cells, use of isotype control antibodies is highly recommended to control for unspecific MCMV Fc-receptor binding-mediated signals. 17. Saponin is a powdery substance that can irritate one’s respiratory system so handle it carefully. It is usually easier to make 1% saponin solution in PBS (1 g of saponin in 100 mL of PBS) and then use this stock solution to generate further dilutions. 1% saponin can also be aliquoted and stored at 20
C. 18. Sodium azide, a commonly used preservative in FACS media or buffers for long-term antibody storage, is an irreversible inhibitor of cellular metabolism. High concentrations will inhibit metabolism of immune cells during antibody-labeling procedure and will result in inactive transferred cells. Amounts found in most commercially available antibody preparations are usually too low and are diluted during the procedure, however make sure to omit it from your FACS media preparation for cells intended for sorting. 19. The high percentage of fetal calf serum is needed since cells are sorted in a small drop and thus the media gets diluted.

Acknowledgments This work has been supported by the Croatian Science Foundation under the project IP-2016-06-5980 (VJL), University of Rijeka

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under the project uniri-biomed-18-234 (BL) and the grant KK.01.1.1.01.0006, awarded to the Scientific Centre of Excellence for Virus Immunology and Vaccines and cofinanced by the European Regional Development Fund (SJ). References 1. Zuhair M et al (2019) Estimation of the worldwide seroprevalence of cytomegalovirus: a systematic review and meta-analysis. Rev Med Virol 29(3):e2034 2. Tastad KJ et al (2019) Awareness of congenital cytomegalovirus and acceptance of maternal and newborn screening. PLoS One 14(8): e0221725 3. Jeon J et al (2006) Knowledge and awareness of congenital cytomegalovirus among women. Infect Dis Obstet Gynecol 2006:80383 4. Gerna G, Kabanova A, Lilleri D (2019) Human cytomegalovirus cell tropism and host cell receptors. Vaccines (Basel) 7(3) 5. Sinzger C, Digel M, Jahn G (2008) Cytomegalovirus cell tropism. Curr Top Microbiol Immunol 325:63–83 6. Britt W (2007) Virus entry into host, establishment of infection, spread in host, mechanisms of tissue damage. In: Arvin A et al (eds) Human herpesviruses: biology, therapy, and immunoprophylaxis. Cambridge University Press Copyright (c) Cambridge University Press 2007, Cambridge 7. Britt W (2011) Chapter 23—Cytomegalovirus. In: Remington JS et al (eds) Infectious Diseases of the Fetus and Newborn (Seventh Edition). W.B. Saunders, Philadelphia, pp 706–755 8. Cannon MJ (2009) Congenital cytomegalovirus (CMV) epidemiology and awareness. J Clin Virol 46(Suppl 4):S6–S10 9. Davis NL, King CC, Kourtis AP (2017) Cytomegalovirus infection in pregnancy. Birth Defects Res 109(5):336–346 10. Wu CA et al (2011) Transmission of murine cytomegalovirus in breast milk: a model of natural infection in neonates. J Virol 85 (10):5115–5124 11. Manicklal S et al (2013) The "silent" global burden of congenital cytomegalovirus. Clin Microbiol Rev 26(1):86–102 12. Dietrich ML, Schieffelin JS (2019) Congenital cytomegalovirus infection. Ochsner J 19 (2):123–130 13. Pass RF (2002) Cytomegalovirus infection. Pediatr Rev 23(5):163–170

14. Morton CC, Nance WE (2006) Newborn hearing screening--a silent revolution. N Engl J Med 354(20):2151–2164 15. Cheeran MC, Lokensgard JR, Schleiss MR, Neuropathogenesis of congenital cytomegalovirus infection: disease mechanisms and prospects for intervention (2009) Clin Microbiol Rev 22(1):99–126, Table of Contents 16. Griffith BP, Lucia HL, Hsiung GD (1982) Brain and visceral involvement during congenital cytomegalovirus infection of Guinea pigs. Pediatr Res 16(6):455–459 17. Keithley EM, Woolf NK, Harris JP (1989) Development of morphological and physiological changes in the cochlea induced by cytomegalovirus. Laryngoscope 99(4):409–414 18. Streblow DN et al (2008) Mechanisms of cytomegalovirus-accelerated vascular disease: induction of paracrine factors that promote angiogenesis and wound healing. Curr Top Microbiol Immunol 325:397–415 19. Priscott PK, Tyrrell DA (1982) The isolation and partial characterisation of a cytomegalovirus from the brown rat, Rattus norvegicus. Arch Virol 73(2):145–160 20. Loh HS et al (2003) Characterization of a novel rat cytomegalovirus (RCMV) infecting placenta-uterus of Rattus rattus diardii. Arch Virol 148(12):2353–2367 21. Loh HS et al (2006) Pathogenesis and vertical transmission of a transplacental rat cytomegalovirus. Virol J 3:42 22. Cloarec R et al (2016) Cytomegalovirus infection of the rat developing brain in utero prominently targets immune cells and promotes early microglial activation. PLoS One 11(7): e0160176 23. Li L et al (2008) Induction of cytomegalovirus-infected labyrinthitis in newborn mice by lipopolysaccharide: a model for hearing loss in congenital CMV infection. Lab Investig 88(7):722–730 24. Ishiwata M et al (2006) Differential expression of the immediate-early 2 and 3 proteins in developing mouse brains infected with murine cytomegalovirus. Arch Virol 151 (11):2181–2196

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Berislav Lisnic´ et al.

25. Tsutsui Y (2009) Effects of cytomegalovirus infection on embryogenesis and brain development. Congenit Anom (Kyoto) 49(2):47–55 26. Koontz T et al (2008) Altered development of the brain after focal herpesvirus infection of the central nervous system. J Exp Med 205 (2):423–435 27. Kosugi I et al (2002) Innate immune responses to cytomegalovirus infection in the developing mouse brain and their evasion by virus-infected neurons. Am J Pathol 161(3):919–928 28. van Den Pol AN et al (1999) Cytomegalovirus cell tropism, replication, and gene transfer in brain. J Neurosci 19(24):10948–10965 29. Stagno S, Britt WJ (2006) Cytomegalovirus. In: Remington JS, Klein JO (eds) Diseases of the fetus and newborn infant. Saunders, W.B, Philadelphia, PA 30. Clancy B, Darlington RB, Finlay BL (2001) Translating developmental time across mammalian species. Neuroscience 105(1):7–17 31. Reddehase MJ, Lemmermann NAW (2018) Mouse model of cytomegalovirus disease and immunotherapy in the immunocompromised host: predictions for medical translation that survived the "Test of Time". Viruses 10 (12):693 32. de Vries LS et al (2004) The spectrum of cranial ultrasound and magnetic resonance imaging abnormalities in congenital cytomegalovirus infection. Neuropediatrics 35 (2):113–119 33. Bradford RD et al (2015) Murine CMV-induced hearing loss is associated with inner ear inflammation and loss of spiral ganglia neurons. PLoS Pathog 11(4):e1004774 34. Carraro M et al (2017) Cytomegalovirus (CMV) infection causes degeneration of Cochlear vasculature and hearing loss in a mouse model. J Assoc Res Otolaryngol 18 (2):263–273 35. Zhuang W et al (2018) MCMV triggers ROS/ NLRP3-associated inflammasome activation in the inner ear of mice and cultured spiral ganglion neurons, contributing to sensorineural hearing loss. Int J Mol Med 41(6):3448–3456 36. Kosmac K et al (2013) Glucocorticoid treatment of MCMV infected newborn mice attenuates CNS inflammation and limits deficits in cerebellar development. PLoS Pathog 9 (3):e1003200 37. Seleme MC et al (2017) Tumor necrosis factor alpha-induced recruitment of inflammatory mononuclear cells leads to inflammation and altered brain development in murine cytomegalovirus-infected newborn mice. J Virol 91:e01983–e16(8)

38. Trgovcich J et al (1998) In: Scholz M et al (eds) Pathogenesis of murine cytomegalovirus infection in neonatal mice. CMV-related immunopathology Monographs in Virology. Karger, Basel, Switzerland, p 12 39. Slavuljica I et al (2010) Recombinant mouse cytomegalovirus expressing a ligand for the NKG2D receptor is attenuated and has improved vaccine properties. J Clin Invest 120(12):4532–4545 40. Hirsl L et al (2018) Murine CMV expressing the high affinity NKG2D ligand MULT-1: a model for the development of cytomegalovirus-based vaccines. Front Immunol 9:991 41. Almishaal AA et al (2017) Natural killer cells attenuate cytomegalovirus-induced hearing loss in mice. PLoS Pathog 13(8):e1006599 42. Zeleznjak J et al (2017) Mouse cytomegalovirus encoded immunoevasins and evolution of Ly49 receptors—sidekicks or enemies? Immunol Lett 189:40–47 43. Bantug GR et al (2008) CD8+ T lymphocytes control murine cytomegalovirus replication in the central nervous system of newborn animals. J Immunol 181(3):2111–2123 44. Brizic I et al (2018) Brain-resident memory CD8(+) T cells induced by congenital CMV infection prevent brain pathology and virus reactivation. Eur J Immunol 48(6):950–964 45. Brizic I et al (2019) CD4 T cells are required for maintenance of CD8 TRM cells and virus control in the brain of MCMV-infected newborn mice. Med Microbiol Immunol 208 (3-4):487–494 46. Jonjic S et al (1994) Antibodies are not essential for the resolution of primary cytomegalovirus infection but limit dissemination of recurrent virus. J Exp Med 179(5):1713–1717 47. Polic B et al (1998) Hierarchical and redundant lymphocyte subset control precludes cytomegalovirus replication during latent infection. J Exp Med 188(6):1047–1054 48. Fowler KB et al (1992) The outcome of congenital cytomegalovirus infection in relation to maternal antibody status. N Engl J Med 326 (10):663–667 49. Adler SP et al (1995) Immunity induced by primary human cytomegalovirus infection protects against secondary infection among women of childbearing age. J Infect Dis 171 (1):26–32 50. Ross SA et al (2011) Mixed infection and strain diversity in congenital cytomegalovirus infection. J Infect Dis 204(7):1003–1007 51. Boppana SB et al (2001) Intrauterine transmission of cytomegalovirus to infants of women

Rodent Models of CMV Infection with preconceptional immunity. N Engl J Med 344(18):1366–1371 52. Cekinovic D et al (2008) Passive immunization reduces murine cytomegalovirus-induced brain pathology in newborn mice. J Virol 82 (24):12172–12180 53. Roncador G et al (2016) The European antibody network’s practical guide to finding and validating suitable antibodies for research. MAbs 8(1):27–36 54. Becroft DM (1981) Prenatal cytomegalovirus infection: epidemiology, pathology and pathogenesis. Perspect Pediatr Pathol 6:203–241 55. Osborn JE, Walker DL (1971) Virulence and attenuation of murine cytomegalovirus. Infect Immun 3(2):228–236 56. Chong KT, Gould JJ, Mims CA (1981) Neutralization of different strains of murine cytomegalovirus (MCMV)-effect of in vitro passage. Arch Virol 69(2):95–104 57. Brizic´ I et al (2018) Cytomegalovirus infection: mouse model. Curr Protoc Immunol:e51 58. Le-Trilling VT, Trilling M (2017) Mouse newborn cells allow highly productive mouse cytomegalovirus replication, constituting a novel convenient primary cell culture system. PLoS One 12(3):e0174695 59. Tan CS, Frederico B, Stevenson PG (2014) Herpesvirus delivery to the murine respiratory tract. J Virol Methods 206:105–114 60. Hsu KM et al (2009) Murine cytomegalovirus displays selective infection of cells within hours after systemic administration. J Gen Virol 90 (Pt 1):33–43

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61. Slavuljica I et al (2015) Immunobiology of congenital cytomegalovirus infection of the central nervous system-the murine cytomegalovirus model. Cell Mol Immunol 12 (2):180–191 62. Kurz S et al (1997) Latency versus persistence or intermittent recurrences: evidence for a latent state of murine cytomegalovirus in the lungs. J Virol 71(4):2980–2987 63. Weiland F et al (1986) Studies on the morphogenesis of murine cytomegalovirus. Intervirology 26(4):192–201 64. Lemmermann NAW et al (2010) CD8 T-cell immunotherapy of cytomegalovirus disease in the murine model. Method Microbiol 37:369–420 65. Suvarna KS, Layton C, Bancroft JD (2018) Bancroft’s Theory and Practice of Histological Techniques. Elsevier, Amsterdam 66. Ross DD et al (1989) Estimation of cell survival by flow cytometric quantification of fluorescein diacetate/propidium iodide viable cell number. Cancer Res 49(14):3776–3782 67. Perfetto SP et al (2006) Amine reactive dyes: an effective tool to discriminate live and dead cells in polychromatic flow cytometry. J Immunol Methods 313(1-2):199–208 68. Perfetto SP et al (2010) Amine-reactive dyes for dead cell discrimination in fixed samples. Curr Protoc Cytom. Chapter 9: p. Unitas 9 34 69. Thale R et al (1994) Identification and expression of a murine cytomegalovirus early gene coding for an fc receptor. J Virol 68 (12):7757–7765

Chapter 19 Recent Approaches and Strategies in the Generation of Anti-human Cytomegalovirus Vaccines Suresh B. Boppana and William J. Britt Abstract Human cytomegalovirus is the largest human herpesvirus and shares many core features of other herpesviruses such as tightly regulated gene expression during genome replication and latency as well as the establishment of lifelong persistence following infection. In contrast to stereotypic clinical syndromes associated with alpha-herpesvirus infections, almost all primary HCMV infections are asymptomatic and acquired early in life in most populations in the world. Although asymptomatic in most individuals, HCMV is a major cause of disease in hosts with deficits in adaptive and innate immunity such as infants who are infected in utero and allograft recipients following transplantation. Congenital HCMV is a commonly acquired infection in the developing fetus that can result in a number of neurodevelopmental abnormalities. Similarly, HCMV is a major cause of disease in allograft recipients in the immediate and late posttransplant period and is thought to be a major contributor to chronic allograft rejection. Even though HCMV induces robust innate and adaptive immune responses, it also encodes a vast array of immune evasion functions that are thought aid in its persistence. Immune correlates of protective immunity that prevent or modify intrauterine HCMV infection remain incompletely defined but are thought to consist primarily of adaptive responses in the pregnant mother, thus making congenital HCMV a potentially vaccine modifiable disease. Similarly, HCMV infection in allograft recipients is often more severe in recipients without preexisting adaptive immunity to HCMV. Thus, there has been a considerable effort to modify HCMV specific immunity in transplant recipient either through active immunization or passive transfer of adaptive effector functions. Although efforts to develop an efficacious vaccine and/or passive immunotherapy to limit HCMV disease have been underway for nearly six decades, most have met with limited success at best. In contrast to previous efforts, current HCMV vaccine development has relied on observations of unique properties of HCMV in hopes of reproducing immune responses that at a minimum will be similar to that following natural infection. However, more recent findings have suggested that immunity following naturally acquired HCMV infection may have limited protective activity and almost certainly, is not sterilizing. Such observations suggest that either the induction of natural immunity must be specifically tailored to generate protective activity or alternatively, that providing targeted passive immunity to susceptible populations could be prove to be more efficacious. Key words HCMV, Vaccination, HCMV transmission, Congenital infection, Pregnancy

Andrew D. Yurochko (ed.), Human Cytomegaloviruses: Methods and Protocols, Methods in Molecular Biology, vol. 2244, https://doi.org/10.1007/978-1-0716-1111-1_19, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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Epidemiology of HCMV infections Human cytomegalovirus (HCMV) infections have been recognized in every human population that has been studied [1, 2]. HCMV acquisition in the population is characterized by an age-dependent rise in seroprevalence, and correlates most closely with socioeconomic level and race [3–7]. About 50% of women of child-bearing age are seronegative in industrialized countries [7, 8]. In this population, HCMV acquisition occurs at a rate of 1–7% per year, and usually follows frequent and prolonged contact with young children (less than 3 years of age) [1, 8–11]. By comparison, in resource-poor communities in industrialized countries and in developing countries, HCMV is usually acquired very early in life owing to breast milk transmission and crowded living conditions and far fewer adult women are seronegative [1, 12–16].

1.1 Transmission of HCMV

Although the exact mode of HCMV acquisition is unknown, it is presumed to be through direct contact with body fluids from an infected person. Breastfeeding, group care of children, overcrowded living conditions, and sexual activity have all been associated with high rates of HCMV infection. Sources of the virus include oropharyngeal, urine, cervical and vaginal secretions, semen, breast milk, blood products, and allografts (Table 1) [17– 20]. Presumably, exposure to saliva and other body fluids containing infectious virus is a primary mode of spread because infected infants typically excrete significant amounts of HCMV for months

Table 1 Sources and routes of transmission of HCMV infection Mode of Exposure and Transmission Community Acquired Age Perinatal

Intrauterine fetal infection (congenital); intrapartum exposure to virus; breast milk acquired; mother-to-infant transmission

Infancy and Childhood

Exposure to saliva and other body fluids; child-to-child transmission

Adolescence and Adulthood

Exposure to young children; sexual transmission; possible occupational exposures

Hospital Acquired Source Blood products

Blood products from seropositive donors; multiple transfusions; white blood cell containing blood products

Allograft recipients

Allograft from seropositive donors

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to years following infection. Even older children and adults can shed virus for prolonged periods (>6 months) following primary HCMV infection. In addition, a significant proportion of seropositive individuals continue to shed virus intermittently. An important determinant of the frequency of congenital and perinatal HCMV infection is the seroprevalence rate in women of child-bearing age (Table 2). Studies from the USA and Europe have shown that the seropositivity rates in young women range from less than 50% to 85% [1, 21–23]. In contrast, the vast majority of women of childbearing age in less well-developed regions are HCMV antibody positive [4, 16, 24–27]. 1.2 Vertical transmission

Perinatal HCMV acquisition, including congenital infection, contributes significantly to the spread of HCMV because infected infants excrete large amounts of virus for prolonged periods. An additional and less well-appreciated mode of virus spread is through breast milk. It is estimated that almost all breastfed infants born to persistently infected mothers will be exposed to HCMV as a result of breast feeding [17, 28, 29]. Rates of congenital CMV infection are higher in developing countries and for low-income groups in developed countries [4, 24, 26, 27, 30, 31]. The prevalence of congenital HCMV infection is directly related to the maternal seroprevalence rates (Table 2). Studies of risk factors for congenital HCMV infection showed that young maternal age, nonwhite race, single marital status, maternal HIV infection, and history of other sexually transmitted diseases have been associated with increased rates of congenital CMV infection [32–39].

Table 2 Rates of maternal HCMV seroprevalence and congenital HCMV infection in various populations

Location

Maternal HCMV seroprevalence (%)

Prevalence of congenital HCMV infection (%)

References

Aarhus-Viborg, Denmark

52

0.4

[398]

Abidjan, Ivory Coast

100

1.4

[24]

Birmingham, USA

[10]

Low income group

77

1.25

Middle income group

36

0.53

Hamilton, Ontario, Canada

44

0.42

[399]

London, UK

56

0.3

[400]

Seoul, South Korea

96

1.2

[30]

New Delhi, India

99

2.1

[27]

Ribeira˜o Preto, Brazil

96

1.1

[26]

Sukuta, The Gambia

96

5.4

[31]

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Similar to congenital infections, infants infected through breast feeding will excrete virus for prolonged periods, making them ideal vectors for the spread of virus. Children continue to acquire HCMV infection throughout childhood and the rate of infection increases during adolescence and early adulthood secondary to sexual exposure. Significant titers of infectious HCMV can be found in semen and cervical secretions, suggesting that exposure to body fluids could result in the transmission of HCMV [40– 43]. The natural history of HCMV infection in adolescents and adults has been shown to parallel sexually transmitted diseases (STDs) [44, 45]. Homosexual men and women attending STD clinics have an increased incidence of HCMV infection [42]. Thus, HCMV should be considered an STD in adults that can effectively spread virus through a sexually active population (Table 1). 1.3 Congenital HCMV Infection Following Maternal Nonprimary Infections

It was recognized shortly after the description of the cytomegalic inclusion disease in infants that preexisting maternal immunity to HCMV does not prevent transmission to the fetus (nonprimary maternal infections) suggesting that the natural history of congenital HCMV was unique among other perinatal infections [46– 48]. The rate of congenital infection in a given population is directly related to the HCMV seroprevalence rates in women of childbearing age, such that populations with high seroprevalence have higher rates of congenital HCMV infections (Table 2) [4, 16, 26, 27, 31, 39]. It was initially thought that this increased rate of congenital HCMV infection in highly seroprevalent maternal populations resulted from an increased likelihood of infection during pregnancy of nonimmune women and intrauterine transmission. However, the increased rates of congenital infection have been reported from equatorial Africa, Brazil, and India where seroimmunity among women of child-bearing age is near-universal. This finding differs dramatically to other causes of congenital viral infections such as rubella [49]. In the case of rubella, the incidence of congenital rubella syndrome fell dramatically when the rate of maternal seroimmunity was above 80–85%, suggesting that a sufficient pool of susceptible seronegative women was necessary to result in maternal infection and transmission to the developing fetus [50]. In contrast, the rates of congenital HCMV infection increase as the rates of maternal seroimmunity increase past 90%. The significance of this finding was not appreciated until recently because it was thought that congenital infections following nonprimary maternal infections are rarely associated with disease and/or sequelae. Since many early studies included patients referred to single centers because of symptomatic infection, the natural history of this infection was not derived from screened populations that would eliminate much of this bias [51, 52]. However, natural history studies in Sweden by Ahlfors et al. in the early 1980s demonstrated the occurrence of symptomatic infections and

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Table 3 HCMV-related hearing loss according to type of maternal infection Maternal HCMV Infection (n ¼ 85)

Hearing Status

Primary (n ¼ 3)

Nonprimary (n ¼ 40)

Moderate to severe unilateral HL Moderate to profound bilateral HL Normal hearing

0 1 2

4 2 34

Indeterminate or samples not available (95% CI) 1 2 39

hearing loss in infants born to women with nonprimary infections [46, 48]. Subsequent studies in screened populations have demonstrated that congenitally infected infants born to women following nonprimary infections account for the majority of HCMV infected newborns [26, 53–58]. Finally, recent studies from populations in which maternal seroimmunity exceeds 96% and almost all infected infants are born to women with nonprimary infection have shown that the prevalence of symptomatic infection and the development of long-term sequelae such as sensorineural hearing loss in infected infants are very similar to those in reported studies of infants infected following primary maternal infections (Table 3) [26, 55, 59]. Therefore, for a vaccine to be effective in preventing or reducing the burden of congenital HCMV infection, the candidate vaccine has to perform better at preventing HCMV reinfection than naturally acquired immunity. Nonprimary maternal infections were initially believed to be secondary to recurrences of preexisting maternal infection, presumably following reactivation of latent infections. Using restriction fragment length polymorphisms of viral DNA, early studies argued for identity between viral isolates, thus suggesting that nonprimary maternal infection could follow reactivation of existing infections and ultimately lead to the infection of the fetus [60]. However, a careful review of the findings from these studies suggests that the identity of viral isolates could be questioned. These studies were carried out with labeled viral DNA and required extensive passage in vitro to adapt the viral isolates for growth in human fibroblasts, a process that would put significant pressure on virus populations present in the initial inoculum. Thus, it is uncertain whether definitive information was indeed derived from these studies. More recent studies utilizing direct sequencing, including next generation sequencing of viruses isolated from mothers and infants, have demonstrated remarkable genetic diversity and the presence of distinct virus strains in specimens from different compartments from the same child [61–63]. The implications of infection with multiple viral strains in the pathogenesis and outcome of maternal

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Table 4 Infection with multiple HCMV strains in mothers according to serologic responses to 2 polymorphic determinants on gH and gB Mothers of infected infants n (%) (n ¼ 40)

Mothers of uninfected infants n (%) (n ¼ 109)

P value

Antibody reactivity against 2 HCMV strains at first prenatal visit

14 (35.0%)

17 (15.6%)

0.009

Seroconversion to new HCMV strain during pregnancy

7 (17.5%)

5 (4.6%)

0.02

Infection with 2 CMV strains before and/or during pregnancy

21 (52.5%)

22 (20.2%)

96% of this maternal population have been infected with HCMV prior to childbearing age [13, 26, 55, 100]. Thus, unless a candidate vaccine can be shown to induce protective immunity that exceeds that afforded by natural immunity, the target population for vaccination could potentially be limited to women of specific racial and socioeconomic backgrounds in North America and Northern Europe [395]. In addition to issues noted above, additional roadblocks exist for the development of an efficacious HCMV vaccine that could be utilized in allograft recipients. The first is the necessity to induce a protective immune response in patients that are often debilitated secondary to chronic disease and organ failure, treatment with immunosuppressive agents, and possibly with an underlying disease that may compromise normal immunity. In addition, the transplant population is aging and recipients of candidate vaccines could have age-related impairments in response to a vaccine and/or adjuvants. Furthermore, in the posttransplant period all solid organ transplant recipients will undergo some form of immunosuppression and many hematopoietic transplant recipients will be severely immunocompromised secondary to conditioning regimens for underlying disease and a poorly functioning graft. Thus, existing immunity induced by a vaccine given in pretransplant period will potentially be eliminated or severely depressed. Moreover, while vaccine immunity could be effective in limiting acquisition of virus following community exposures it could be significantly less effective when virus is acquired following transplantation of an infected organ, including hematopoietic grafts [394]. Under conditions of organ transplantation, HCMV can essentially bypass protective immune responses at host mucosal surfaces, responses that could provide some advantage to the host by slowing the kinetics of virus replication and spread, and thus permitting timely generation of protective immune responses. Lastly, the finding of HCMV in transplanted organs undergoing host vs. graft responses, as well as in hematopoietic transplant recipients undergoing graft vs. host reactions following hematopoietic transplantation suggest that the loss of normal immune regulation in these patients results in further immunosuppression and favors virus replication. Thus, induction

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of lasting protective immunity by vaccines in this population represents a challenging task. In contrast to immunocompromised allograft recipients, women of childbearing age are immunologically competent and can be expected to respond to candidate HCMV vaccines. An efficacious vaccine for HCMV in this population must induce sufficient immunity to limit the transmission of HCMV to the developing offspring and/or modify the infection in the fetus. Although the virologic characteristics of maternal infection that result in fetal infection remain incompletely described, natural history studies of congenital HCMV infections indicate that vaccine immunity must be protective throughout pregnancy as intrauterine transmission and disease in the offspring of pregnant women has been demonstrated following maternal infection in all trimesters of pregnancy [10, 396, 397]. Furthermore, it is well documented that women with preexisting immunity to HCMV can transmit virus to the fetus and that this infection can lead to disease in the newborn infant. Thus, and as noted above, requirements for vaccine-induced protective immunity in normal women could conceivably exceed the immunity induced following naturally acquired infection. A second potential hurdle in the development of a vaccine to limit damaging congenital HCMV infection is the necessity of vaccine immunity to be transferred across the placenta, a requirement that effectively limits potential mechanisms of protective immunity to antiviral antibodies acquired by the developing fetus. In order to prevent damaging intrauterine infection, vaccine-induced antiviral antibodies must restrict virus replication and dissemination in the fetus, particularly dissemination to the CNS. Although antiviral antibodies have been shown to limit virus entry and limit virus replication in models of CMV infection of the CNS, the nature of these antibodies and their mechanism of action in limiting CNS infection have not been defined.

References 1. Krech U, Konjajev Z, Jung M (1971) Congenital cytomegalovirus infection in siblings from consecutive pregnancies. Helv Paediatr Acta 26:355–362 2. Gold E, Nankervis GA (1976) Cytomegalovirus. In: Evans AS (ed) Viral infections of humans: epidemiology and control. Plenum Press, New York, pp 143–161 3. Pass RF, Dworsky ME, Whitley RJ, August AM, Stagno S, Alford CA Jr (1981) Specific lymphocyte blastogenic responses in children with cytomegalovirus and herpes simplex virus infections acquired early in infancy. Infect Immun 34:166–170

4. Stagno S, Pass RF, Dworsky ME, Alford CA (1982) Maternal cytomegalovirus infection and perinatal transmission. Clin Obstet Gynecol 25:563–576 5. Marshall GS, Stout GG (2005) Cytomegalovirus seroprevalence among women of childbearing age during a 10-year period. Am J Perinatol 22:371–376 6. Kenneson A, Cannon MJ (2007) Review and meta-analysis of the epidemiology of congenital cytomegalovirus (CMV) infection. Rev Med Virol 17:253–276 7. Cannon MJ, Schmidt DS, Hyde TB (2010) Review of cytomegalovirus seroprevalence

442

Suresh B. Boppana and William J. Britt

and demographic characteristics associated with infection. Rev Med Virol 20:202–213 8. Hyde TB, Schmidt DS, Cannon MJ (2010) Cytomegalovirus seroconversion rates and risk factors: implications for congenital CMV. Rev Med Virol 20:311–326 9. Hutto C, Ricks R, Garvie M, Pass RF (1985) Epidemiology of cytomegalovirus infections in young children: day care vs. home care. Pediatr Infect Dis 4:149–152 10. Stagno S, Pass RF, Cloud G, Britt WJ, Henderson RE, Walton PD, Veren DA, Page F, Alford CA (1986) Primary Cytomegalovirus Infection in Pregnancy: Incidence, transmission to fetus and clinical outcome. J Am Med Assoc 256:1904–1908 11. Adler SP (1989) Cytomegalovirus and child day care. Evidence for an increased infection rate among day-care workers. N Engl J Med 321:1290–1296 12. Seo S, Cho Y, Park J (2009) Serologic screening of pregnant Korean women for primary human cytomegalovirus infection using IgG avidiyt test. Korean J Lab Med 29:557–562 13. Yamamoto AY, Castellucci RA, Aragon DC, Mussi-Pinhata MM (2012) Early high CMV seroprevalence in pregnant women from a population with a high rate of congenital infection. Epidemiol Infect 141:1–5 14. Saraswathy TS, Az-Ulhusna A, Asshikin RN, Suriani S, Zainah S (2011) Seroprevalence of cytomegalovirus infection in pregnant women and associated role in obstetric complications: a preliminary study. Southeast Asian J Trop Med Public Health 42:320–322 15. Akinbami AA, Rabiu KA, Adewunmi AA, Wright KO, Dosunmu AO, Adeyemo TA, Adediran A, Osunkalu VO (2011) Seroprevalence of cytomegalovirus antibodies amongst normal pregnant women in Nigeria. Int J Women’s Health 3:423–428 16. Arapovic J, Rajic B, Pati S, Brizic I, Azinovic I, Sˇusˇak B, Ostojic M, Tutisˇ B, Raguzˇ AB, Tomic V, Jonjic S, Boppana S (2020) Cytomegalovirus Seroprevalence and Birth Prevalence of Congenital CMV Infection in Bosnia and Herzegovina: A Single-Center Experience. Pediatr Infect Dis J 39:140–144 17. Hayes D, Danks M, Givas H, Jack I (1972) Cytomegalovirus in human milk. N Engl J Med 287:177 18. Lang DJ, Kummer JF (1975) Cytomegalovirus in semen: observations in selected populations. J Infect Dis 132:472–473 19. Reynolds DW, Stagno S, Hosty TS, Tiller M, Alford CA (1973) Maternal cytomegalovirus

excretion and perinatal infection. N Engl J Med 289:1–5 20. Alford CA, Stagno S, Pass RF (1980) Natural history of perinatal cytomegalovirus infection. In: Perinatal Infections. Excerpta Medical, Amsterdam, pp 125–147 21. Gold E, Nankervis GA (1982) Cytomegalovirus. In: Evans AS (ed) Viral infections of humans: epidemiology and control, 2nd edn. Plenum Press, New York, pp 167–186 22. Bate SL, Dollard SC, Cannon MJ (2010) Cytomegalovirus seroprevalence in the United States: the national health and nutrition examination surveys, 1988-2004. Clin Infect Dis 50:1439–1447 23. Puhakka L, Sarvikivi E, Lappalainen M, Surcel HM, Saxen H (2016) Decrease in seroprevalence for herpesviruses among pregnant women in Finland: cross-sectional study of three time points 1992, 2002 and 2012. Infect Dis 48:406–410 24. Schopfer K, Lauber E, Krech U (1978) Congenital cytomegalovirus infection in newborn infants of mothers infected before pregnancy. Arch Dis Child 53:536–539 25. Vial P, Torres J, Stagno S, Gonzalez E, Donoso E, Alford CA, Hirsch T, Rodriguez L (1985) Serological screening for cytomegalovirus, rubella virus, herpes simpelx virus, hepatitis B virus and Toxoplasma gondii in two populations of pregnant women in Chile. Bull Pan Am Health Organ 99:53–61 26. Mussi-Pinhata MM, Yamamoto AY, MouraBritto RM, Lima- Issacs, M., Boppana, S., and Britt, W. J. (2009) Birth prevalence and natural history of congenital cytomegalovirus (CMV) infection in highly seroimmune population. Clin Infect Dis 49:522–528 27. Dar L, Pati SK, Patro AR, Deorari AK, Rai S, Kant S, Broor S, Fowler KB, Britt WJ, Boppana SB (2008) Congenital cytomegalovirus infection in a highly seropositive semi-urban population in India. Pediatr Infect Dis J 27:841–843 28. Stagno S, Reynolds DW, Pass RF, Alford CA (1980) Breast milk and the risk of cytomegalovirus infection. N Engl J Med 302:1073–1076 29. Hamprecht K, Maschmann J, Vochem M, Dietz K, Speer CP, Jahn G (2001) Epidemiology of transmission of cytomegalovirus from mother to preterm infants by breastfeeding. Lancet 357:513–518 30. Sohn YM, Park KI, Lee C, Han DG, Lee WY (1992) Congenital cytomegalovirus infection in Korean population with very high

Recent Approaches and Strategies CMV Vaccines prevalence of maternal immunity. J Kor Med Sci 7:47–51 31. van der Sande MAB, Kaye S, Miles DJC, Waight P, Jeffries DJ, Ojoula OO, Palmero M, Pinder M, Ismaili J, Flanagan KL, Aveika AA, Rowland-Jones S, McConkey SJ, Whittle H, Marchant A (2007) Risk factors for and clinical outcome of congenital cytomegalovirus infection in a peri-urban West-African birth cohort. PLoS One 2:e492 32. Fowler KB, Stagno S, Pass RF (1993) Maternal age and congenital cytomegalovirus infection: screening of two diverse newborn populations, 1980-1990. J Infect Dis 168:552–556 33. Preece PM, Tookey P, Ades A, Peckham CS (1986) Congenital cytomegalovirus infection: predisposing maternal factors. J Epidemiol Community Health 40:205–209 34. Fowler KB, Stagno S, Pass RF (1991) Rates of congenital cytomegalovirus infection based on newborn screening in two populations over an eleven year interval. Pediatr Res 29:90A 35. Gompels UA, Larke N, Sanz-Ramos M, Bates M, Musonda K, Manno D, Siame J, Monze M, Filteau S, Group., a. t. C. S (2012) Human cytomegalovirus infant infection adversely affects growth and development in maternally HIV-exposed and unexposed infants in Zambia. Clin Infect Dis 54:434–442 36. Manicklal S, van Niekerk AM, Kroon SM, Hutto C, Novak Z, Pati SK, Chowdhury N, Hsiao NY, Boppana SB (2014) Birth prevalence of congenital CMV among infants of HIV-infected women on prenatal antiretroviral prophylaxis in South Africa. Clin Infect Dis 58(10):1467–1472 37. Mwaanza N, Chilukutu L, Tembo J, Kabwe M, Musonda K, Kapasa M, Chabala C, Sinyangwe S, Mwaba P, Zulma A, Bates M (2013) High rates of congenital cytomegalovirus infection linked with maternal HIV infection among neonatal admissions at a large referral center in Sub-Saharan Africa. Clin Infect Dis 58 (5):728–735 38. Adachi K, Xu J, Ank B, Watts DH, Camarca M, Mofenson LM, Pilotto JH, Joao E, Gray G, Theron G, Santos B, Fonseca R, Kreitchmann R, Pinto J, MussiPinhata MM, Machado DM, Ceriotto M, Morgado MG, Bryson YJ, Veloso VG, Grinsztejn B, Mirochnick M, Moye J, Nielsen-Saines K, Team, f. t. N. H. S (2018) Congenital Cytomegalovirus and HIV

443

Perinatal Transmission. Pediatr Infect Dis J 37:1016–1021 39. Pathirana J, Groome M, Dorfman J, Kwatra G, Boppana S, Cutland C, Jones S, Madhi SA (2019) Prevalence of congenital cytomegalovirus infection and associated risk of in utero Human Immunodeficiency Virus (HIV) acquisition in a high-HIV prevalence setting, South Africa. Clin Infect Dis 69:1789–1796 40. Jordan MC, Rousseau WE, Noble GR, Stewart JA, Chin TDY (1973) Association of cervical cytomegaloviruses with venereal disease. N Engl J Med 288(18):932–934 41. Willmott FE (1975) Cytomegalovirus in female patients attending a VD clinic. Brit J Vener Dis 51:278–280 42. Drew WL, Mintz L, Miner RC, Sands M, Ketterer B (1981) Prevalence of cytomegalovirus infection in homosexual men. J Infect Dis 143:188–192 43. Chandler SH, Holmes KK, Wentworth BB, Gutman LT, Wiesner PJ, Alexander ER, Handsfield HH (1985) The Epidemiology of cytomegaloviral infection in women attending a sexually transmitted disease clinic. J Infect Dis 152:597–605 44. Sohn YM, Oh MK, Balcarek KB, Cloud GA, Pass RF (1991) Cytomegalovirus infection in sexually active adolescents. J Infect Dis 163:460–463 45. Knox GE, Pass RF, Reynolds DW, Stagno S, Alford CA (1979) Comparative prevalence of subclinical cytomegalovirus and herpes simplex virus infections in the genital and urinary tracts of low income, urban females. J Infect Dis 140:419–422 46. Ahlfors K, Harris S, Ivarsson S, Svanberg L (1981) Secondary maternal cytomegalovirus infection causing symptomatic congenital infection. N Engl J Med 305:284 47. Stagno S, Pass RF, Dworsky ME, Henderson RE, Moore EG, Walton PD, Alford CA (1982) Congenital cytomegalovirus infection: The relative importance of primary and recurrent maternal infection. N Engl J Med 306:945–949 48. Ahlfors K, Forsgren M, Ivarsson SA, Harris S, Svanberg L (1983) Congenital cytomegalovirus infection: on the relation between type and time of maternal infection and infant’s symptoms. Scand J Infect Dis 15:129–138 49. Ross LA (1995) Rubella, Adolescent medicine (Philadelphia, Pa.) 6, pp. 15–26 50. Stagno S, Pass RF, Dworsky ME, Alford CA (1983) Congenital and perinatal cytomegaloviral infections. Semin Perinatol 7:31–42

444

Suresh B. Boppana and William J. Britt

51. Fowler KB, Stagno S, Pass RF, Britt WJ, Boll TJ, Alford CA (1992) The outcome of congenital cytomegalovirus infection in relation to maternal antibody status. N Engl J Med 326:663–667 52. Demmler GJ (1991) Infectious diseases society of america and centers for disease control. Summary of a workshop on surveillance for congenital cytomegalovirus disease. Rev Infect Dis 13:315–329 53. Boppana SB, Fowler KB, Britt WJ, Stagno S, Pass RF (1999) Symptomatic congenital cytomegalovirus infection in infants born to mothers with preexisting immunity to cytomegalovirus. Pediatrics 104:55–60 54. Ahlfors K, Ivarsson S, Harris S (1999) Report on a long-term study of maternal and congenital cytomegalovirus infection in Sweden. Review of prospective studies available in the literature. Scand J Infect Dis 31:443–457 55. Yamamoto AY, Mussi-Pinhata MM, Isaac MDL, Amaral FR, Carvalheiro CG, Aragon DC, da Silva Mafredi AK, Boppana SB, Britt WJ (2011) Congenital cytomegalovirus infection as a cause of sensorineural hearing loss in a highly seropositive population. Ped Infect Dis J 30:1043–4046 56. Townsend CL, Forsgren M, Ahlfors K, Ivarsson S, Tookey P, Peckham CS (2013) Long-term outcomes of congenital cytomegalovirus infection in Sweden and the United Kingdom. Clin Infect Dis 56:1232–1239 57. Puhakka L, Renko M, Helminen M, Peltola V, Heiskanen-Kosma T, Lappalainen M, Surcel H-M, Lo¨nnqvist T, Saxen H (2017) Primary versus non-primary maternal cytomegalovirus infection as a cause of symptomatic congenital infection - register-based study from Finland. Infect Dis 49:445–453 58. Wang C, Zhang X, Bialek S, Cannon MJ (2011) Attribution of congenital cytomegalovirus infection to primary versus non-primary maternal infection. Clin Infect Dis 52: e11–e13 59. Yamamoto AY, Anastasio ART, Massuda ET, Isaac ML, Manfredi AKS, Cavalcante JMS, Carnevale-Silva A, Fowler KB, Boppana S, Britt WJ, Mussi-Pinhata MM (2019) Contribution of congenital cytomegalovirus (cCMV) to permanent hearing loss in a highly seropositive population: "The BraCHS study". Clin Infect Dis 70(7):1379–1384 60. Huang ES, Alford CA, Reynolds DW, Stagno S, Pass RF (1980) Molecular epidemiology of cytomegalovirus infections in women and their infants. N Engl J Med 303:958–962

61. Renzette N, Bhattacharjee B, Jenson JD, Gibson L, Kowalik TF (2011) Extensive genome-wide variability of human cytomegalovirus in congenitally infected infants. PLoS Pathog 7:e1001344 62. Ross SA, Novak Z, Pati S, Patro RK, Blumenthal J, Danthaluri VR, Ahmed A, Michaels MG, Sanchez PJ, Bernstein DI, Tolan RW, Palmer AL, Britt WJ, Fowler KB, Boppana SB (2011) Mixed infection and strain diversity in congenital cytomegalovirus infection. J Infect Dis 204:1003–1007 63. Dobbins GC, Patki A, Chen D, Tiwari HK, Hendrickson C, Britt WJ, Fowler K, Chen JY, Boppana SB, Ross SA (2019) Association of CMV genomic mutations with symptomatic infection and hearing loss in congenital CMV infection. BMC Infect Dis 19:1046–1046 64. Bale JF Jr, Petheram SJ, Robertson M, Murph JR, Demmler G (2001) Human cytomegalovirus a sequence and UL144 variability in strains from infected children. J Med Virol 65:90–96 65. Bale JF, Zimmerman B, Dawson J, Souza I, Petheram B, Murph JR (1999) Cytomegalovirus transmission in child care homes. Arch Pediatr Adolesc Med 153:75–79 66. Chandler SH, McDougall JK (1986) Comparison of restriction site polymorphisms among clinical isolates and laboratory strains of human cytomegalovirus. J Gen Virol 67:2179–2192 67. Ishibashi K, Tokumoto T, Tanabe K, Shirakawa H, Hashimoto K, Kushida N, Yanagida T, Inoue O, Yamaguchi H, Toma T, Suzutami T (2007) Association of the outcome of renal transplantation with antibody response to cytomegalovirus strainspecific glycoprotein H epitopes. J Infect Dis 45:60–67 68. Yamamoto AY, Mussi-Pinhata MM, Boppana SB, Novak Z, Wagatsuma VM, Oliviera PD, Duarte G, Britt WJ (2010) Human cytomegalovirus reinfection is associated with intrauterine transmission in a highly cytomegalovirus-immune maternal population. Am J Obstet Gynecol 202:297. e291–297.e298 69. Novak Z, Ross SA, Patro RK, Pati SK, Kumbla RA, Brice S, Boppana SB (2008) Cytomegalovirus strain diversity in seropositive women. J Clin Microbiol 46:882–886 70. Ross SA, Arora N, Novak Z, Fowler KB, Britt WJ, Boppana SB (2010) Cytomegalovirus reinfections in healthy seroimmune women. J Infect Dis 201:386–389

Recent Approaches and Strategies CMV Vaccines 71. Boppana SB, Rivera LB, Fowler KB, Mach M, Britt WJ (2001) Intrauterine transmission of cytomegalovirus to infants of women with preconceptional immunity. N Engl J Med 344:1366–1371 72. Barbosa NG, Yamamoto AY, Duarte G, Aragon DC, Fowler KB, Boppana S, Britt WJ, Mussi-Pinhata MM (2018) Cytomegalovirus shedding in seropositive pregnant women from a high-seroprevalence population: the brazilian cytomegalovirus hearing and maternal secondary infection study. Clin Infect Dis 67:743–750 73. Rubin RH, Russell PS, Levin M, et, a. l. (1979) Summary of a workshop on cytomegalovirus infections during organ transplantation. J Infect Dis 139:728–734 74. Autran B, Carcelain G, Li TS, Blanc C, Mathez D, Tubiana R, Katlama C, Debre P, Leibowitch J (1997) Positive effects of combined antiretroviral therapy on CD4+ T cell homeostasis and function in advanced HIV disease. Science 277:112–116 75. Komanduri KV, Viswanathan MN, Wieder ED, Schmidt DK, Bredt BM, Jacobson M, McCune JM (1998) Restoration of cytomegalovirus-specific CD4+ T-lymphocyte responses after ganciclovir and highly active antiretroviral therapy in individuals infected with HIV-1. Nat Med 4:953–956 76. Bronke C, Palmer NM, Jansen CA, Westerlaken GH, Polstra AM, Reiss P, Bakker M, Miedema F, Tesselaar K, van Baarle D (2005) Dynamics of cytomegalovirus (CMV)-specific T cells in HIV-1-infected individuals progressing to AIDS with CMV end-organ disease. J Infect Dis 191:873–880 77. Gerna G, Lilleri D, Chiesa A, Zelini P, Furione M, Comolli G, Pellegrini C, Sarchi E, Migotto C, Bonora MR, Meloni F, Arbustini E (2011) Virologic and immunologic monitoring of cytomegalovirus to guide preemptive therapy in solid-organ transplantation. Am J Transplant 11:2463–2471 78. Gratama JW, van Esser JW, Lamers CH, Tournay C, Lo¨wenberg B, Bolhuis RL, Cornelissen JJ (2001) Tetramer-based quantification of cytomegalovirus (CMV)-specific CD8 + T lymphocytes in T-cell-depleted stem cell grafts and after transplantation may identify patients at risk for progressive CMV infection. Blood 98:1358–1364 79. Lu´cia M, Crespo E, Cruzado JM, Grinyo´ JM, Bestard O (2014) Human CMV-specific T-cell responses in kidney transplantation;

445

toward changing current risk-stratification paradigm. Transpl Int 27:643–656 80. Ganepola S, Gentilini C, Hilbers U, Lange T, Rieger K, Hofmann J, Maier M, Liebert UG, Niederwieser D, Engelmann E, Heilbronn R, Thiel E, Uharek L (2007) Patients at high risk for CMV infection and disease show delayed CD8+ T-cell immune recovery after allogeneic stem cell transplantation. Bone Marrow Transplant 39:293–299 81. Reusser P, Riddell SR, Meyers JD, Greenberg PD (1991) Cytotoxic T-lymphocyte response to cytomegalovirus after human allogeneic bone marrow transplantation: pattern of recovery and correlation with cytomegalovirus infection and disease. Blood 78:1373–1380 82. Li CR, Greenberg PD, Gilbert MJ, Goodrich JM, Riddell SR (1994) Recovery of HLA-restricted cytomegalovirus (CMV)specific T-cell responses after allogeneic bone marrow transplant: correlation with CMV disease and effect of ganciclovir prophylaxis. Blood 83:1971–1979 83. Egli A, Humar A, Kumar D (2012) State-ofthe-art monitoring of cytomegalovirusspecific cell-mediated immunity after organ transplant: a primer for the clinician. Clin Infect Dis 55:1678–1689 84. Gabanti E, Lilleri D, Scaramuzzi L, Zelini P, Rampino T, Gerna G (2018) Comparison of the T-cell response to human cytomegalovirus (HCMV) as detected by cytokine flow cytometry and QuantiFERON-CMV assay in HCMV-seropositive kidney transplant recipients. New Microbiol 41:195–202 85. Walter EA, Greenberg PD, Gilbert MJ, Finch RJ, Watanabe KS, Thomas ED, Riddell SR (1995) Reconstitution of cellular immunity against cytomegalovirus in recipients of allogeneic bone marrow by transfer of T-cell clones from the donor. N Engl J Med 333:1038–1044 86. Biron CA, Byron KS, Sullivan JL (1989) Severe herpesvirus infections in an adolescent without natural killer cells. N Engl J Med 320:1731–1735 87. Starr SE, Tolpin MD, Friedman HM, Paucker K, Plotkin SA (1979) Impaired cellular immunity to cytomegalovirus in congenitally infected children and their mothers. J Infect Dis 140:500–505 88. Gehrz RC, Marker SC, Knorr SO, Kalis JM, Balfour HH Jr (1977) Specific cell-mediated immune defect in active cytomegalovirus infection of young children and their mothers. Lancet 2:844–847

446

Suresh B. Boppana and William J. Britt

89. Miles DJ, Sande M, Kaye S, Crozier S, Ojuola O, Palmero MS, Sanneh M, Touray ES, Waight P, Rowland-Jones S, Whittle H, Marchant A (2008) CD4(+) T cell responses to cytomegalovirus in early life: a prospective birth cohort study. J Infect Dis 197:658–662 90. Tu W, Chen S, Sharp M, Dekker C, Manganello AM, Tongson EC, Maecker HT, Holmes TH, Wang Z, Kemble G, Adler S, Arvin A, Lewis DB (2004) Persistent and selective deficiency of CD4+ T cell immunity to cytomegalovirus in immunocompetent young children. J Immunol 172:3260–3267 91. Marchant A, Appay V, Van Der Sande M, Dulphy N, Liesnard C, Kidd M, Kaye S, Ojuola O, Gillespie GM, Vargas Cuero AL, Cerundolo V, Callan M, McAdam KP, Rowland-Jones SL, Donner C, McMichael AJ, Whittle H (2003) Mature CD8(+) T lymphocyte response to viral infection during fetal life. J Clin Invest 111:1747–1755 92. Huygens A, Lecomte S, Tackoen M, Olislagers V, Delmarcelle Y, Burny W, Van Rysselberge M, Liesnard C, Larsen M, Appay V, Donner C, Marchant A (2015) Functional exhaustion limits CD4+ and CD8 + T-cell responses to congenital cytomegalovirus infection. J Infect Dis 212:484–494 93. Griffiths PD, Stagno S, Pass RF, Smith RJ, Alford CA (1982) Infection with cytomegalovirus during pregnancy: specific IgM antibodies as a marker of recent primary infection. J Infect Dis 145:647–653 94. Coppola T, Mangold JF, Cantrell S, Permar SR (2019) Impact of maternal immunity on congenital cytomegalovirus birth prevalence and infant outcomes: a systematic review. Vaccine 7:129 95. Leruez-Ville M, Magny JF, Couderc S, Pichon C, Parodi M, Bussieres L, Guillminot T, Ghout I, Ville Y (2017) Risk factors for congenital cytomegalovirus infection following primary and nonprimary maternal infection: A prospective neonatal screening study using polymerase chain reaction in saliva. Clin Infect Dis 65(3):398–404 96. Britt WJ (2017) Congenital cytomegalovirus infection and the enigma of maternal immunity. J Virol 91:e02392–e02316 97. Brizic´ I, Lisnic´ B, Brune W, Hengel H, Jonjic´ S (2018) Cytomegalovirus infection: mouse model. Curr Protoc Immunol:e51 98. Britt WJ (2018) Maternal immunity and the natural history of congenital human cytomegalovirus infection. Viruses 10:405 99. Krmpotic´ A, Podlech J, Reddehase MJ, Britt WJ, Jonjic´ S (2019) Role of antibodies in

confining cytomegalovirus after reactivation from latency: three decades’ re´sume´. Med Microbiol Immunol 208:415–429 100. Mussi-Pinhata MM, Yamamoto AY, Aragon DC, Duarte G, Fowler KB, Boppana S, Britt WJ (2018) Seroconversion for cytomegalovirus infection during pregnancy and fetal infection in a highly seropositive population: "The BraCHS Study". J Infect Dis 218:1200–1204 101. Dreher MA, Arora N, Fowler KB, Novak Z, Britt WJ, Boppana SB, Ross SA (2014) Spectrum of disease and outcome in children with symptomatic congenital cytomegalovirus infection. J Pediatr 164:855–859 102. Tanimura K, Tairaku S, Ebina Y, Morioka I, Nagamata S, Deguchi K, Morizane M, Deguchi M, Minematsu T, Yamada H (2017) Prediction of congenital cytomegalovirus infection in high-risk pregnant women. Clin Infect Dis 64:159–165 103. Ahlfors K, Ivarsson SA, Harris S, Svanberg L, Holmqvist R, Lernmark B, Theander G (1984) Congenital cytomegalovirus infection and disease in Sweden and the relative importance of primary and secondary maternal infections. Scand J Infect Dis 16:129–137 104. Yeager AS (1983) Transmission of cytomegalovirus to mothers by infected infants: another reason to prevent transfusionacquired infections. Pediatr Infect Dis 2:295 105. Bootz A, Karbach A, Spindler J, Kropff B, Reuter N, Sticht H, Winkler TH, Britt WJ, Mach M (2017) Protective capacity of neutralizing and non-neutralizing antibodies against glycoprotein B of cytomegalovirus. PLoS Pathog 13:e1006601 106. Nelson CS, Cruz DV, Tran D, Bialas KM, Stamper L, Wu H, Gilbert M, Blair R, Alvarez X, Itell H, Chen M, Deshpande A, Chiuppesi F, Wussow F, Diamond DJ, Vandergrift N, Walter MR, Barry PA, Cohen-Wolkowiez M, Koelle K, Kaur A, Permar SR (2017) Preexisting antibodies can protect against congenital cytomegalovirus infection in monkeys. JCI Insight 2:e94002 107. Bravo FJ, Bourne N, Schleiss MR, Bernstein DI (2003) An animal model of neonatal cytomegalovirus infection. Antivir Res 60:41–49 108. Bratcher DF, Bourne N, Bravo FJ, Schleiss MR, Slaoui M, Myers MG, Bernstein DI (1995) Effect of passive antibody on congenital cytomegalovirus infection in guinea pigs. J Infect Dis 172:944–950 109. Farrell HE, Shellam GR (1991) Protection against murine cytomegalovirus infection by passive transfer of neutralizing and

Recent Approaches and Strategies CMV Vaccines non-neutralizing monoclonal antibodies. J Gen Virol 72:149–156 110. Chatterjee A, Harrison CJ, Britt WJ, Bewtra C (2001) Modification of maternal and congenital cytomegalovirus infection by antiglycoprotein b antibody transfer in guinea pigs. J Infect Dis 183:1547–1553 111. Alford CA, Hayes K, Britt WJ (1988) Primary cytomegalovirus infection in pregnancy: comparison of antibody responses to virus encoded proteins between women with and without intrauterine infection. J Infect Dis 158:917–924 112. Lilleri D, Kabanova A, Revello MG, Percivalle E, Sarasini A, Genini E, Sallusto F, Lanzavecchia A, Corti D, Gerna G (2013) Fetal human cytomegalovirus transmission correlates with delayed maternal antibodies to gH/gL/pUL128-130-131 complex during primary infection. PLoS One 8:e59863 113. Forner G, Saldan A, Mengoli C, Gussetti N, Palu` G, Abate D (2016) Cytomegalovirus (CMV) enzyme-linked immunosorbent spot assay but not CMV QuantiFERON assay is a novel biomarker to determine risk of congenital CMV infection in pregnant women. J Clin Microbiol 54:2149–2154 114. Saldan A, Forner G, Mengoli C, Gussetti N, Palu` G, Abate D (2015) Strong cell-mediated immune response to human cytomegalovirus is associated with increased risk of fetal infection in primarily infected pregnant women. Clin Infect Dis 61:1228–1234 115. Nigro G, Adler SP, La Torre R, Best AM (2005) Passive immunization during pregnancy for congenital cytomegalovirus infection. N Engl J Med 353:1350–1362 116. Visentin S, Manara R, Milanese L, Da Roit A, Forner G, Salviato E, Citton V, Magno FM, Orzan E, Morando C, Cusinato R, Mengoli C, Palu G, Ermani M, Rinalid R, Cosmi E, Gussetti N (2012) Early pregnancy cytomegalovirus infection in pregnancy: Maternal hyperimmunoglobulin therapy improves outcomes among infants at 1 year of age. Clin Infect Dis 55:497–503 117. Revello MG, Furione M, Rognoni V, Arossa A, Gerna G (2014) Cytomegalovirus DNAemia in pregnant women. J Clin Virol 61:590–592 118. Kagan KO, Enders M, Schampera MS, Baeumel E, Hoopmann M, Geipel A, Berg C, Goelz R, De Catte L, Wallwiener D, Brucker S, Adler SP, Jahn G, Hamprecht K (2019) Prevention of maternal-fetal transmission of cytomegalovirus after primary maternal infection in the first trimester by biweekly

447

hyperimmunoglobulin administration. Ultrasound Obstet Gynecol 53:383–389 119. Maidji E, McDonagh S, Genbacev O, Tabata T, Pereira L (2006) Maternal antibodies enhance or prevent cytomegalovirus infection in the placenta by neonatal Fc receptor-mediated transcytosis. Am J Pathol 168:1210–1226 120. Pereira L, Maidji E, McDonagh S, Genbacev O, Fisher S (2003) Human cytomegalovirus transmission from the uterus to the placenta correlates with the presence of pathogenenic bacteria and maternal immunity. J Virol 77:13301–13314 121. Shimamura M, Mach M, Britt WJ (2006) Human cytomegalovirus infection elicits a glycoprotein M (gM)/gN-specific virus-neutralizing antibody response. J Virol 80:4591–4600 122. Macagno A, Bernasconi NL, Vanzetta F, Dander E, Sarasini A, Revello MG, Gerna G, Sallusto F, Lanzavecchia A (2010) Isolation of human monoclonal antibodies that potently neutralize human cytomegalovirus infection by targeting different epitopes on the gH/ gL/UL128-131A complex. J Virol 84:1005–1013 123. Vanarsdall AL, Chin AL, Liu J, Jardetzky TS, Mudd JO, Orloff SL, Streblow D, MussiPinhata MM, Yamamoto AY, Duarte G, Britt WJ, Johnson DC (2019) HCMV trimerand pentamer-specific antibodies synergize for virus neutralization but do not correlate with congenital transmission. Proc Natl Acad Sci U S A 116:3728–3733 124. Britt WJ, Fay J, Kneiss N, Mach M (1990) An immunodominant linear epitope on the major envelope glcoprotein complex (gB) of human cytomegalovirus, In VIIIth International Congress of Virology, Berlin 125. Boppana SB, Britt WJ (1995) Antiviral antibody responses and intrauterine transmission after primary maternal cytomegalovirus infection. J Infect Dis 171:1115–1121 126. Saccoccio FM, Jenks JA, Itell HL, Li SH, Berry M, Pollara J, Casper C, Gantt S, Permar SR (2019) Humoral immune correlates for prevention of postnatal cytomegalovirus acquisition. J Infect Dis 220:772–780 127. Nelson CS, Huffman T, Jenks JA, Cisneros de la Rosa E, Xie G, Vandergrift N, Pass RF, Pollara J, Permar SR (2018) HCMV glycoprotein B subunit vaccine efficacy mediated by nonneutralizing antibody effector functions. Proc Natl Acad Sci U S A 115:6267–6272 128. Ross SA, Ahmed A, Palmer AL, Michaels MG, Sa´nchez PJ, Stewart A, Bernstein DI, Feja K,

448

Suresh B. Boppana and William J. Britt

Fowler KB, Boppana SB (2017) Newborn dried blood spot polymerase chain reaction to identify infants with congenital cytomegalovirus-associated senfsorineural hearing loss. J Pediatr 184:57–61.e51 129. Vanarsdall AL, Ryckman BJ, Chase MC, Johnson DC (2008) Human cytomegalovirus glycoproteins gB and gH/gL mediate epithelial cell-cell fusion when expressed either in cis or in trans. J Virol 82:11837–11850 130. Vanarsdall AL, Howard PW, Wisner TW, Johnson DC (2016) Human cytomegalovirus gH/gL forms a stable complex with the fusion protein gB in virions. PLoS Pathog 12:e1005564 131. Gerna G, Sarasini A, Patrone M, Percivalle E, Fiorina L, Campanini G, Gallina A, Baldanti F, Revello MG (2008) Human cytomegalovirus serum neutralizing antibodies block virus infection of endothelial/epithelial cells, but not fibroblasts, early during primary infection. J Gen Virol 89:853–865 132. Chiuppesi F, Wussow F, Johnson E, Bian C, Zhuo M, Rajakumar A, Barry PA, Britt WJ, Chakraborty R, Diamond DJ (2015) Vaccinederived neutralizing antibodies to the human cytomegalovirus gH/gL pentamer potently block primary cytotrophoblast infection. J Virol 89:11884–11898 133. Gerna G, Percivalle E, Perez L, Lanzavecchia A, Lilleri D (2016) Monoclonal antibodies to different components of the human cytomegalovirus (HCMV) pentamer gH/gL/pUL128L and trimer gH/gL/gO as well as antibodies elicited during primary HCMV infection prevent epithelial cell syncytium formation. J Virol 90:6216–6223 134. Fouts AE, Chan P, Stephan JP, Vandlen R, Feirbach B (2012) Antibodies against the gH/gL/UL128/UL130/UL131 complex comprise the majority of the anticytomegalovirus (anti-CMV) neutralizing antibodie response in CMV hyperimmune globulin. J Virol 86:7444–7447 135. Lilleri D, Gerna G (2017) Maternal immune correlates of protection from human cytomegalovirus transmission to the fetus after primary infection in pregnancy. Rev Med Virol 27. https://doi.org/10.1002/rmv. 1921 136. Murrell I, Bedford C, Ladell K, Miners KL, Price DA, Tomasec P, Wilkinson GWG, Stanton RJ (2017) The pentameric complex drives immunologically covert cell-cell transmission of wild-type human cytomegalovirus. Proc Natl Acad Sci U S A 114:6104–6109 137. Jenks JA, Goodwin ML, Permar SR (2019) The roles of host and viral antibody Fc

receptors in herpes simplex virus (HSV) and human cytomegalovirus (HCMV) infections and immunity. Front Immunol 10:1–9 138. Lilleri D, Fornara C, Furione M, Zavattoni M, Revello MG, Gerna G (2007) Development of human cytomegalovirusspecific T cell immunity during primary infection of pregnant women and its correlation with virus transmission to the fetus. J Infect Dis 195:1062–1070 139. Dauby N, Kummert C, Lecomte S, Liesnard C, Delforge M-L, Donner C, Marchant A (2014) Primary human cytomegalovirus infection induces the expansion of virus-specific activated and atypical memory B cells. J Infect Dis 210:1275–1285 140. Dauby N, Sartori D, Kummert C, Lecomte S, Haelterman E, Delforge M-L, Donner C, Mach M, Marchant A (2016) Limited effector memory B-cell response to envelope Glycoprotein B during primary human cytomegalovirus infection. J Infect Dis 213:1642–1650 141. Zanghellini F, Boppana SB, Emery VC, Griffiths PD, Pass RF (1999) Asymptomatic primary cytomegalovirus infection: virologic and immunologic features. J Infect Dis 180:702–707 142. Baldanti F, Paolucci S, Campanini A, Sarasini A, Percivalle E, Revello MG, Gerna G (2006) Human cytomegalovirus UL131A, UL130, and UL128 genes are highly conserved among field isolates. Arch Virol 151:1225–1233 143. Fornara C, Cassaniti I, Zavattoni M, Furione M, Adzasehoun KMG, De Silvestri A, Comolli G, Baldanti F (2017) Human cytomegalovirus-specific memory CD4+ T-cell response and its correlation with virus transmission to the fetus in pregnant women with primary infection. Clin Infect Dis 65:1659–1665 144. Mele, F., Fornara, C., Jarrossay, D., Furione, M., Arossa, A., Spinillo, A., Lanzavecchia, A., Gerna, G., Sallusto, F., and Lilleri, D. (2017) Phenotype and specificity of T cells in primary human cytomegalovirus infection during pregnancy: IL-7Rpos long-term memory phenotype is associated with protection from vertical transmission, PLoS One 12, e0187731 145. Sester M, Sester U, Gartner B, Girndt M, Meyerhans A, Kohler H (2002) Dominance of virus-specific CD8 T cells in human primary cytomegalovirus infection. J Am Soc Nephrol 13:2577–2584 146. Gamadia LE, Remmerswaal EB, Weel JF, Bemelman F, van Lier RA, Ten Berge IJ (2003) Primary immune responses to human

Recent Approaches and Strategies CMV Vaccines CMV: a critical role for IFN-gamma-producing CD4+ T cells in protection against CMV disease. Blood 101:2686–2692 147. Moss P, Khan N (2004) CD8(+) T-cell immunity to cytomegalovirus. Hum Immunol 65:456–464 148. Sylwester AW, Mitchell BL, Edgar JB, Taormina C, Pelte C, Ruchti F, Sleath PR, Grabstein KH, Hosken NA, Kern F, Nelson JA, Picker LJ (2005) Broadly targeted human cytomegalovirus-specific CD4+ and CD8+ T cells dominate the memory compartments of exposed subjects. J Exp Med 202:673–685 149. Gyulai Z, Endresz V, Burian K, Pincus S, Toldy J, Cox WI, Meric C, Plotkin S, Gonczol E, Berencsci K (2000) Cytotoxic T lymphocyte (CTL) resposnes to human cytomegalovirus pp65, IE1-Exaon4, gB, pp150, and pp28 in healthy individuals: reevaluation of prevalence of IE1-specific CTSs. J Infect Dis 181:1537–1546 150. Atkins JT, Demmler GJ, Williamson WD, McDonald JM, Istas AS, Buffone GJ (1994) Polymerase chain reaction to detect cytomegalovirus DNA in the cerebrospinal fluid of neonates with congenital infection. J Infect Dis 169:1334–1337 151. La Rosa C, Limaye AP, Krishnan A, Longmate J, Diamond DJ (2007) Longitudinal assessment of cytomegalovirus (CMV)specific immune responses in liver transplant recipients at high risk for late CMV disease. J Infect Dis 195:633–644 152. Schooley RT, Hirsch MS, Colvin RB, Cosimi AB, Tolkoff-Rubin NE, McCluskey RT, Burton RC, Russell PS, Herrin JT, Delmonico FL, Giorgi JV, Henle W, Rubin RH (1983) Association of herpes virus infection with T-lymphocyte subset alterations, glomerulopathy, and opportunistic infections after renal transplantation. N Engl J Med 308:307–313 153. Sester M, Sester U, Gartner B, Heine G, Girndt M, Mueller-Lantzsch N, Meyerhans A, Kohler H (2001) Levels of virus-specific CD4 T cells correlate with cytomegalovirus control and predict virusinduced disease after renal transplantation. Transplantation 71:1287–1294 154. Reusser P, Cathomas G, Attenhofer R, Tamm M, Thiel G (1999) Cytomegalovirus (CMV)-specific T cell immunity after renal transplantation mediates protection from CMV disease by limiting the systemic virus load. J Infect Dis 180:247–253 155. Pourgheysari B, Piper KP, McLarnon A, Arrazi J, Bruton R, Clark F, Cook M, Mahendra P, Craddock C, Moss PA (2009) Early reconstitution of effector memory CD4

449

+ CMV-specific T cells protects against CMV reactivation following allogeneic SCT. Bone Marrow Transplant 43:853–861 156. Razonable RR (2008) Cytomegalovirus infection after liver transplantation: current concepts and challenges. World J Gastroenterol 14:4849–4860 157. Gallina A, Percivalle E, Simoncini L, Revello MG, Gerna G, Milanesi G (1996) Human cytomegalovirus protein pp65 lower matrix phosphoprotein harbours two transplantable nuclear localization signals. J Gen Virol 77:1151–1157 158. Gerna G, Lilleri D, Fornara C, Comolli G, Lozza L, Campana C, Pellegrini C, Meloni F, Rampino T (2006) Monitoring of human cytomegalovirus-specific CD4 and CD8 T-cell immunity in patients receiving solid organ transplantation. Am J Transplant 6:2356–2364 159. Lilleri D, Gerna G, Fornara C, Lozza L, Maccario R, Locatelli F (2006) Prospective simultaneous quantification of human cytomegalovirus-specific CD4+ and CD8+ T-cell reconstitution in young recipients of allogeneic hematopoietic stem cell transplants. Blood 108:1406–1412 160. Alford CA, Stagno S, Pass RF, Huang ES (1981) Epidemiology of cytomegalovirus. In: Nahmais A, Dowdle W, Schinazi R (eds) The human herpesviruses: an interdisciplinary perspective. Elsevier, New York, pp 159–171 161. Riddell SR, Greenberg PD (1994) Therapeutic reconstruction of human viral immunity by adoptive transfer of cytotoxic T lymphocyte clones. Curr Top Microbiol Immunol 189:9–34 162. Riddell SR, Greenberg PD (2000) T-cell therapy of cytomegalovirus and human immunodeficiency virus infection. J Antimicrobl Chemother 45:35–43 163. Boppana SB, Polis MA, Kramer AA, Britt WJ, Koenig S (1995) Virus specific antibody responses to human cytomegalovirus (HCMV) in human immunodeficiency virus type 1-infected individuals with HCMV retinitis. J Infect Dis 171:182–185 164. Dylewski J, Chou S, Merigan TC (1983) Absence of detectable IgM antibody during cytomegalovirus disease in patients with AIDS [letter]. N Engl J Med 309:493 165. Rasmussen L, Morris S, Wolitz R, Dowling A, Fessell J, Holodniy M, Merigan TC (1994) Deficiency in antibody response to human cytomegalovirus glycoprotein gH in human immunodeficiency virus-infected patients at

450

Suresh B. Boppana and William J. Britt

risk for cytomegalovirus retinitis. J Infect Dis 170:673–677 166. Emery VC, Asher K, Sanjuan Cde J (2012) Importance of the cytomegalovirus seropositive recipient as a contributor to disease burden after solid organ transplantation. J Clin Virol 54:125–129 167. Schachtner T, Stein M, Reinke P (2017) CMV-specific T cell monitoring offers superior risk stratification of CMV-seronegative kidney transplant recipients of a CMV-seropositive donor. Transplantation 101:e315–e325 168. Nesher L, Shah DP, Ariza-Heredia EJ, Azzi JM, Siddiqui HK, Ghantoji SS, Marsh LY, Michailidis L, Makedonas G, Rezvani K, Shpall EJ, Chemaly RF (2016) Utility of the enzyme-linked immunospot interferon-γ-release assay to predict the risk of cytomegalovirus infection in hematopoietic cell transplant recipients. J Infect Dis 213:1701–1707 169. Lee H, Park KH, Ryu JH, Choi A-R, Yu JH, Lim J, Han K, Kim SI, Yang CW, Chung BH, Oh E-J (2017) Cytomegalovirus (CMV) immune monitoring with ELISPOT and QuantiFERON-CMV assay in seropositive kidney transplant recipients. PLoS One 12: e0189488 170. Lochmanova A, Lochman I, Tomaskova H, Marsalkova P, Raszka J, Mrazek J, Dedochova J, Martinek A, Brozmanova H, Grundmann M (2010) Quantiferon-CMV test in prediction of cytomegalovirus infection after kidney transplantation. Transplant Proc 42:3574–3577 171. Chiereghin A, Potena L, Borgese L, Gibertoni D, Squarzoni D, Turello G, Petrisli E, Piccirilli G, Gabrielli L, Grigioni F, Lazzarotto T (2018) Monitoring of cytomegalovirus (CMV)-specific cellmediated immunity in heart transplant recipients: clinical utility of the QuantiFERONCMV assay for management of posttransplant cmv infection. J Clin Microbiol 56: e01040–e01017 172. Favi E, Santangelo R, Iesari S, Morandi M, Marcovecchio GE, Trecarichi EM, Salerno MP, Ferraresso M, Citterio F, Romagnoli J (2017) Enzyme-linked immunospot assay as a complementary method to assess and monitor cytomegalovirus infection in kidney transplant recipients on pre-emptive antiviral therapy: a single-center experience. Transplant Proc 49:1766–1772 173. Kumar D, Mian M, Singer L, Humar A (2017) An interventional study using cellmediated immunity to personalize therapy

for cytomegalovirus infection after transplantation. Am J Transplant 17:2468–2473 174. Manuel O (2013) Clinical experience with immune monitoring for cytomegalovirus in solid-organ transplant recipients. Curr Infect Dis Rep 175. Krawczyk A, Ackermann J, Goitowski B, Trenschel R, Ditschkowski M, Timm J, Ottinger H, Beelen DW, Gru¨ner N, Fiedler M (2018) Assessing the risk of CMV reactivation and reconstitution of antiviral immune response post bone marrow transplantation by the QuantiFERON-CMV-assay and real time PCR. J Clin Virol 99–100:61–66 176. Panagou E, Zakout G, Keshani J, Smith C, Irish D, Mackinnon S, Kottaridis P, Fielding A, Griffiths PD (2016) Cytomegalovirus pre-emptive therapy after hematopoietic stem cell transplantation in the era of realtime quantitative PCR: comparison with recipients of solid organ transplants. Transpl Infect Dis 18:405–414 177. Pergam SA, Xie H, Sandhu R, Pollack M, Smith J, Stevens-Ayers T, Ilieva V, Kimball LE, Huang M-L, Hayes TS, Corey L, Boeckh MJ (2012) Efficiency and risk factors for CMV transmission in seronegative hematopoietic stem cell recipients. Biol Blood Marrow Transplant 18:1391–1400 178. Raanani P, Gafter-Gvili A, Paul M, Ben-Bassat I, Leibovici L, Shpilberg O (2008) Immunoglobulin prophylaxis in hematological malignancies and hematopoietic stem cell transplantation. Cochrane Database Syst Rev 4:CD006501 179. Raanani P, Gafter-Gvili A, Paul M, Ben-Bassat I, Leibovici L, Shpilberg O (2009) Immunoglobulin prophylaxis in hematopoietic stem cell transplantation: systematic review and meta-analysis. J Clin Oncol 27:770–781 180. Slade M, Goldsmith S, Romee R, DiPersio JF, Dubberke ER, Westervelt P, Uy GL, Lawrence SJ (2017) Epidemiology of infections following haploidentical peripheral blood hematopoietic cell transplantation. Transpl Infect Dis 19:e12629. https://doi.org/10. 1111/tid.12629 181. Crocchiolo R, Bramanti S, Vai A, Sarina B, Mineri R, Casari E, Tordato F, Mauro E, Timofeeva I, Lugli E, Mavilio D, Carlo-StellaC, Santoro A, Castagna L (2015) Infections after T-replete haploidentical transplantation and high-dose cyclophosphamide as graftversus-host disease prophylaxis. Transpl Infect Dis 17:242–249 182. Marek A, Stern M, Chalandon Y, Ansari M, Ozsahin H, Gu¨ngo¨r T, Gerber B, Ku¨hne T,

Recent Approaches and Strategies CMV Vaccines Passweg JR, Gratwohl A, Tichelli A, Seger R, Schanz U, Halter J, Stussi G, Swiss Blood Stem Cell, T (2014) The impact of T-cell depletion techniques on the outcome after haploidentical hematopoietic SCT. Bone Marrow Transplant 49:55–61 183. Mulanovich VE, Jiang Y, de Lima M, Shpall EJ, Champlin RE, Ciurea SO (2011) Infectious complications in cord blood and T-cell depleted haploidentical stem cell transplantation. Am J Blood Res 1:98–105 184. Marty FM, Ljungman P, Chemaly RF, Maertens J, Dadwal SS, Duarte RF, Haider S, Ullmann AJ, Katayama Y, Brown J, Mullane KM, Boeckh M, Blumberg EA, Einsele H, Snydman DR, Kanda Y, DiNubile MJ, Teal VL, Wan H, Murata Y, Kartsonis NA, Leavitt RY, Badshah C (2017) Letermovir prophylaxis for cytomegalovirus in hematopoietic-cell transplantation. N Engl J Med 377:2433–2444 185. Roddie C, Peggs KS (2017) Immunotherapy for transplantation-associated viral infections. J Clin Invest 127:2513–2522 186. Pass RF, Griffiths PD, August AM (1983) Antibody response to cytomegalovirus after renal transplantation: comparison of patients with primary and recurrent infection. J Infect Dis 147:40–46 187. Kropff B, Landini MP, Mach M (1993) An ELISA using recombinant proteins for the detection of neutralizing antibodies against human cytomegalovirus. J Med Virol 39:187–195 188. Falagas ME, Snydman DR, Ruthazer R, Werner BG, Griffith J (1997) Surveillance cultures of blood, urine, and throat specimens are not valuable for predicting cytomegalovirus disease in liver transplant recipients. Boston Center for Liver Transplantation Cytomegalovirus Immune Globulin Study Group. Clin Infect Dis 24:824–829 189. Snydman DR (1990) Cytomegalovirus immunoglobulins in the prevention and treatment of cytomegalovirus disease. Rev Infect Dis 12(S):839–848 190. Snydman DR, Werner BG, Heinze-Lacey B, Berardi VP, Tilney NL, Kirkman RL, Milford EL, Cho SI, Bush HL, Levey AS, Strom TB, Carpenter CE, Levey RH, Harmon WE, Zimmerman CE, Shaprio ME, Steinman T, LoGerfo F, Idelson B, Schroter GPJ, Levin MJ, McIver J, Leszczynski J, Grady GF (1987) Use of cytomegalovirus immune globulin to prevent cytomegalovirus disease in renal transplant recipients. N Engl J Med 317:1049–1054

451

191. Rea F, Potena L, Yonan N, Wagner F, Calabrese F (2016) Cytomegalovirus hyper immunoglobulin for CMV prophylaxis in thoracic transplantation. Transplantation 100(Suppl 3):S19–S26 192. Grossi P, Mohacsi P, Szabolcs Z, Potena L (2016) Cytomegalovirus immunoglobulin after thoracic transplantation: an overview. Transplantation 100(Suppl 3):S1–S4 193. Schulz U, Solidoro P, Mu¨ller V, Szabo A, Gottlieb J, Wilkens H, Enseleit F (2016) CMV Immunoglobulins for the treatment of CMV infections in thoracic transplant recipients. Transplantation 100(Suppl 3):S5–S10 ˜o 194. Garcia-Gallo CL, Gil PU, Laporta R, Carren MC, de Pablo A, Ferreiro MJ (2005) Is gammaglobulin anti-CMV warranted in lung transplantation? Transplant Proc 37:4043–4045 195. Ranganathan K, Worley S, Michaels MG, Arrigan S, Aurora P, Ballmann M, Boyer D, Conrad C, Eichler I, Elidemir O, Goldfarb S, Mallory GB Jr, Mogayzel PJ, Parakininkas D, Solomon M, Visner G, Sweet SC, Faro A, Danziger-Isakov L (2009) Cytomegalovirus immunoglobulin decreases the risk of cytomegalovirus infection but not disease after pediatric lung transplantation. J Heart Lung Transplant 28:1050–1056 196. Valantine HA, Luikart H, Doyle R, Theodore J, Hunt S, Oyer P, Robbins R, Berry G, Reitz B (2001) Impact of cytomegalovirus hyperimmune globulin on outcome after cardiothoracic transplantation: a comparative study of combined prophylaxis with CMV hyperimmune globulin plus ganciclovir versus ganciclovir alone. Transplantation 72:1647–1652 197. Ljungman P, Cordonnier C, Einsele H, Bender-Go¨tze C, Bosi A, Dekker A, De la Camara R, Gmu¨r J, Newland AC, Prentice HG, Robinson AJ, Rovira M, Ro¨sler W, Veil D (1998) Use of intravenous immune globulin in addition to antiviral therapy in the treatment of CMV gastrointestinal disease in allogeneic bone marrow transplant patients: a report from the European Group for Blood and Marrow Transplantation (EBMT). Infectious Diseases Working Party of the EBMT. Bone Marrow Transplant 21:473–476 198. Winston DJ, Ho WG, Bartoni K, Champlin RE (1993) Intravenous immunoglobulin and CMV-seronegative blood products for prevention of CMV infection and disease in bone marrow transplant recipients. Bone Marrow Transplant 12:283–288

452

Suresh B. Boppana and William J. Britt

199. Ahn H, Tay J, Shea B, Hutton B, Shorr R, Knoll GA, Cameron DW, Cowan J (2018) Effectiveness of immunoglobulin prophylaxis in reducing clinical complications of hematopoietic stem cell transplantation: a systematic review and meta-analysis. Transfusion 58:2437–2452 200. Abdel-Azim H, Elshoury A, Mahadeo KM, Parkman R, Kapoor N (2017) Humoral immune reconstitution kinetics after allogeneic hematopoietic stem cell transplantation in children: a maturation block of IgM Memory B cells may lead to impaired antibody immune reconstitution. Biology Blood Marrow Transplant 23:1437–1446 201. Bourassa-Blanchette S, Knoll GA, Hutton B, Fergusson N, Bennett A, Tay J, Cameron DW, Cowan J (2019) Clinical outcomes of polyvalent immunoglobulin use in solid organ transplant recipients: A systematic review and meta-analysis. Clin Transpl 33: e13560 202. D’Orsogna LJ, Wright MP, Krueger RG, McKinnon EJ, Buffery SI, Witt CS, Staples N, Loh R, Cannell PK, Christiansen FT, French MA (2009) Allogeneic hematopoietic stem cell transplantation recipients have defects of both switched and igm memory B cells. Biol Blood Marrow Transplant 15:795–803 203. Heimall J, Puck J, Buckley R, Fleisher TA, Gennery AR, Neven B, Slatter M, Haddad E, Notarangelo LD, Baker KS, Dietz AC, Duncan C, Pulsipher MA, Cowan MJ (2017) Current knowledge and priorities for future research in late effects after hematopoietic stem cell transplantation (HCT) for severe combined immunodeficiency patients: a consensus statement from the second pediatric blood and marrow transplant consortium International Conference on Late Effects after Pediatric HCT. Biol Blood Marrow Transplant 23:379–387 204. Boeckh M, Bowden RA, Storer B, Chao NJ, Spielberger R, Tierney DK, Gallez-HawkinsG, Cunningham T, Blume KG, Levitt D, Zaia JA (2001) Randomized, placebo-controlled, double-blind study of a cytomegalovirusspecific monoclonal antibody (MSL-109) for prevention of cytomegalovirus infection after allogeneic hematopoietic stem cell transplantation. Biol Blood Marrow Transplant 7:343–351 205. Anonymous (1997) MSL-109 adjuvant therapy for cytomegalovirus retinitis in patients with acquired immunodeficiency syndrome: the Monoclonal Antibody Cytomegalovirus Retinitis Trial. The Studies of Ocular

Complications of AIDS Research Group. AIDS Clinical Trials Group. Arch Ophthalmol 115:1528–1536 206. Ishida JH, Patel A, Mehta AK, Gatault P, McBride JM, Burgess T, Derby MA, Snydman DR, Emu B, Feierbach B, Fouts AE, Maia M, Deng R, Rosenberger CM, Gennaro LA, Striano NS, Liao XC, Tavel JA (2017) Phase 2 randomized, double-blind, placebocontrolled trial of RG7667, a combination monoclonal antibody, for prevention of cytomegalovirus infection in high-risk kidney transplant recipients. Antimicrob Agents Chemother 61:e01794–e01716 207. Schleiss MR (2006) Nonprimate models of congenital cytomegalovirus (CMV) infection: gaining insight into pathogenesis and prevention of disease in newborns. ILAR J 47:65–72 208. Britt WJ, Boppana S (2004) Human cytomegalviurs virion proteins. Hum Immunol 65:395–402 209. Itell HL, Nelson CS, Martinez DR, Permar SR (2017) Maternal immune correlates of protection against placental transmission of cytomegalovirus. Placenta 60(Suppl 1): S73–S79 210. Reddehase MJ, Lemmermann NAW (2018) Mouse model of cytomegalovirus disease and immunotherapy in the immunocompromised host: predictions for medical translation that survived the "Test of Time". Viruses 10:693 211. Jonjic S, Babic M, Polic B, Krmpotic A (2008) Immune evasion of natural killer cells by viruses. Curr Opin Immunol 20:30–38 212. Jonjic S, Pavic I, Lucin P, Rukavina D, Koszinowski UH (1990) Efficacious control of cytomegalovirus infection after long-term depletion of CD8+ T lymphocytes. J Virol 64:5457–5464 213. Jonjic S, Pavic I, Polic B, Crnkovic I, Lucin P, Koszinowski UH (1994) Antibodies are not essential for the resolution of primary cytomegalovirus infection but limit dissemination of recurrent virus. J Exp Med 179:1713–1717 214. Koszinowski UH, Reddehase MJ, Jonjic S (1991) The role of CD4 and CD8 T cells in viral infections. Curr Opin Immunol 3:471–475 215. Koszinowski UH, Reddehase MJ, Keil GM, Volkmer H, Jonjic S, Messerle M, del Val M, Mutter W, Munch K, Buhler B (1987) Molecular analysis of herpesviral gene products recognized by protective cytolytic T lymphocytes. Immunol Lett 16:185–192 216. Krmpotic A, Bubic I, Polic B, Lucin P, Jonjic S (2003) Pathogenesis of murine

Recent Approaches and Strategies CMV Vaccines cytomegalovirus infection. Microbes Infect 5:1263–1277 217. Polic B, Hengel H, Krmpotic A, Trgovcich J, Pavic I, Luccaronin P, Jonjic S, Koszinowski UH (1998) Hierarchical and redundant lymphocyte subset control precludes cytomegalovirus replication during latent infection. J Exp Med 188:1047–1054 218. Shanley JD (1991) Murine models of cytomegalovirus associated pneumonitis. Transplant Proc 23:S12–S16 219. Shanley JD, Biczak L, Formon SJ (1993) Acute murine cytomegalovirus infection induces lethal hepatitis. J Infect Dis 167:264–269 220. Koszinowski UH, Del Val M, Reddehase MJ (1990) Cellular and molecular basis of the protective immune response to cytomegalovirus infection. Curr Top Microbiol Immuno 154:189–220 221. Martins JP, Andoniou CE, Fleming P, Kuns RD, Schuster IS, Voigt V, Daly S, Varelias A, Tey S-K, Degli-Esposti MA, Hill GR (2019) Strain-specific antibody therapy prevents cytomegalovirus reactivation after transplantation. Science 363:288–293 222. Alcami A, Koszinowski UH (2000) Viral mechanisms of immune evasion. Immunol Today 21:447–455 223. Krmpotic A, Hasan M, Loewendorf A, Saulig T, Halenius A, Lenac T, Polic B, Bubic I, Kriegeskorte A, Pernjak-Pugel E, Messerle M, Hengel H, Busch DH, Koszinowski UH, Jonjic S (2005) NK cell activation through the NKG2D ligand MULT-1 is selectively prevented by the glycoprotein encoded by mouse cytomegalovirus gene m145. J Exp Med 201:211–220 224. Zimmermann A, Trilling M, Wagner M, Wilborn M, Bubic I, Jonjic S, Koszinowski U, Hengel H (2005) A cytomegaloviral protein reveals a dual role for STAT2 in IFN-{gamma} signaling and antiviral responses. J Exp Med 201:1543–1553 225. LoPiccolo DM, Gold MC, Kavanagh DG, Wagner M, Koszinowski UH, Hill AB (2003) Effective inhibition of K(b)- and D (b)-restricted antigen presentation in primary macrophages by murine cytomegalovirus. J Virol 77:301–308 226. Lenac T, Budt M, Arapovic J, Hasan M, Zimmermann A, Simic H, Krmpotic A, Messerle M, Ruzsics Z, Koszinowski UH, Hengel H, Jonjic S (2006) The herpesviral Fc receptor fcr-1 down-regulates the NKG2D ligands MULT-1 and H60. J Exp Med 203:1843–1850

453

227. Krmpotic A, Messerle M, Crnkovic-Mertens I, Polic B, Jonjic S, Koszinowski UH (2001) The immunoevasive function encoded by the mouse cytomegalovirus gene m152 protects the virus against T cell control in vivo. J Exp Med 190:1285–1296 228. Krmpotic A, Busch DH, Bubic I, Gebhardt F, Hengel H, Hasan M, Scalzo AA, Koszinowski UH, Jonjic S (2002) MCMV glycoprotein gp40 confers virus resistance to CD8+ T cells and NK cells in vivo. Nat Immunol 3:529–535 229. Kavanagh DG, Gold MC, Wagner M, Koszinowski UH, Hill AB (2001) The multiple immune-evasion genes of murine cytomegalovirus are not redundant: m4 and m152 inhibit antigen presentation in a complementary and cooperative fashion. J Exp Med 194:967–978 230. Hasan M, Krmpotic A, Ruzsics Z, Bubic I, Lenac T, Halenius A, Loewendorf A, Messerle M, Hengel H, Jonjic S, Koszinowski UH (2005) Selective down-regulation of the NKG2D ligand H60 by mouse cytomegalovirus m155 glycoprotein. J Virol 79:2920–2930 231. Hengel H, Reusch U, Gutermann A, Ziegler H, Jonjic S, Lucin P, Koszinowski UH (1999) Cytomegaloviral control of MHC class I function in the mouse. Immunol Rev 168:167–176 232. Smith HR, Heusel JW, Mehta IK, Kim S, Dorner BG, Naidenko OV, Iizuka K, Furukawa H, Beckman DL, Pingel JT, Scalzo AA, Fremont DH, Yokoyama WM (2002) Recognition of a virus-encoded ligand by a natural killer cell activation receptor. Proc Natl Acad Sci U S A 99:8826–8831 233. Upton JW, Kaiser WJ, Mocarski ES (2010) Virus inhibition of RIP3-dependent necrosis. Cell Host Microbe 7:302–313 234. Lisnic VJ, Krmpotic A, Jonjic S (2010) Modulation of natural killer cell activity by viruses. Curr Opin Microbiol 13(4):530–539 235. Handke W, Luig C, Popovic B, Krmpotic A, Jonjic S, Brune W (2013) Viral inhibition of BAK promotes murine cytomegalovirus dissemination to salivary glands. J Virol 87:3592–3596 236. Brune W, Andoniou CE (2017) Die another day: inhibition of cell death pathways by cytomegalovirus. Viruses 9:249 237. Lio C-WJ, McDonald B, Takahashi M, Dhanwani R, Sharma N, Huang J, Pham E, Benedict CA, Sharma S (2016) cGAS-STING signaling regulates initial innate control of

454

Suresh B. Boppana and William J. Britt

cytomegalovirus infection. J Virol 90:7789–7797 238. Deutschmann J, Schneider A, Gruska I, Vetter B, Thomas D, Kießling M, Wittmann S, Herrmann A, Schindler M, Milbradt J, Ferreiro´s N, Winkler TH, Wiebusch L, Gramberg T (2019) A viral kinase counteracts in vivo restriction of murine cytomegalovirus by SAMHD1. Nat Microbiol 4:2273–2284 239. Stempel M, Chan B, Juranic´ Lisnic´ V, Krmpotic´ A, Hartung J, Paludan SR, Fu¨llbrunn N, Lemmermann NAW, Brinkmann MM (2019) The herpesviral antagonist m152 reveals differential activation of STING-dependent IRF and NF-κB signaling and STING’s dual role during MCMV infection. EMBO J 38:e100983 240. Feng L, Sheng J, Vu G-P, Liu Y, Foo C, Wu S, Trang P, Paliza-Carre M, Ran Y, Yang X, Sun X, Deng Z, Zhou T, Lu S, Li H, Liu F (2018) Human cytomegalovirus UL23 inhibits transcription of interferon-γ stimulated genes and blocks antiviral interferon-γ responses by interacting with human N-myc interactor protein. PLoS Pathog 14: e1006867 241. Daley-Bauer LP, Roback L, Crosby LN, McCormick AL, Feng Y, Kaiser WJ, Mocarski ES (2017) Mouse cytomegalovirus M36 and M45 death suppressors cooperate to prevent inflammation resulting from antiviral programmed cell death pathways. Proc Natl Acad Sci 114:E2786–E2795 242. Kaiser WJ, Upton JW, Mocarski ES (2013) Viral modulation of programmed necrosis. Curr Opin Virol 3:296–306 ˇ eleznjak J, Lisnic´ VJ, Popovic´ B, Lisnic´ B, 243. Z Babic´ M, Halenius A, L’Hernault A, Rovisˇ TL, Hengel H, Erhard F, Redwood AJ, Vidal SM, Do¨lken L, Krmpotic´ A, Jonjic´ S (2019) The complex of MCMV proteins and MHC class I evades NK cell control and drives the evolution of virus-specific activating Ly49 receptors. J Exp Med 216:1809–1827 244. Humphreys IR, de Trez C, Kinkade A, Benedict CA, Croft M, Ware CF (2007) Cytomegalovirus exploits IL-10-mediated immune regulation in the salivary glands. J Exp Med 204:1217–1225 245. Reddehase MJ, Balthesen M, Rapp M, Jonjic S, Pavic I, Koszinowski UH (1994) The conditions of primary infection define the load of latent viral genome in organs and the risk of recurrent cytomegalovirus disease. J Exp Med 179:185–193 246. Mathys S, Schroeder T, Ellwart J, Koszinowski UH, Messerle M, Just U (2003)

Dendritic cells under influence of mouse cytomegalovirus have a physiologic dual role: to initiate and to restrict T cell activation. J Infect Dis 187:988–999 247. Spencer JV, Lockridge KM, Barry PA, Lin G, Tsang M, Penfold ME, Schall TJ (2002) Potent immunosuppressive activities of cytomegalovirus-encoded interleukin-10. J Virol 76:1285–1292 248. Chang WL, Barry PA (2009) Attenuation of innate immunity by cytomegalovirus IL-10 establishes a long-term deficit of adaptive antiviral immunity. Proc Natl Acad Sci U S A 107:22647–22652 249. Mandaric S, Walton SM, Rulicke T, Richter K, Girard-Madoux MJ, Clausen BE, Zurunic A, Kamanaka M, Flavell RA, Jonjic S, Oxenius A (2012) IL-10 suppression of NK/DC crosstalk leads to poor priming of MCMV-specific CD4 T cells and prolonged MCMV persistence. PLoS Pathog 8:e1002846 250. Scalzo AA, Corbett AJ, Rawlinson WD, Scott GM, Degli-Esposti MA, Pyzik M, GendronPontbriand EM, Fodil-Cornu N, Vidal SM, Lemmermann NA, Bohm V, Holtappels R, Reddehase MJ, Babic M, Krmpotic A, Jonjic S, Engel P, Angulo A (2007) The interplay between host and viral factors in shaping the outcome of cytomegalovirus infection. Immunol Cell Biol 85:46–54 251. Deere JD, Chang WLW, Villalobos A, Schmidt KA, Deshpande A, Castillo LD, Fike J, Walter MR, Barry PA, HartiganO’Connor DJ (2019) Neutralization of rhesus cytomegalovirus IL-10 reduces horizontal transmission and alters long-term immunity. Proc Natl Acad Sci U S A 116:13036–13041 252. Geary CD, Sun JC (2017) Memory responses of natural killer cells. Semin Immunol 31:11–19 253. O’Sullivan TE, Sun JC (2015) Generation of natural killer cell memory during viral infection. J Innate Immun 7:557–562 254. Scalzo AA, Yokoyama WM (2008) Cmv1 and natural killer cell responses to murine cytomegalovirus infection. Curr Top Microbiol Immunol 321:101–122 255. Vivier E, Raulet DH, Moretta A, Caligiuri MA, Zitvogel L, Lanier LL, Yokoyama WM, Ugolini S (2011) Innate or adaptive immunity? The example of natural killer cells. Science 331:44–49 256. Rapp M, Messerle M, Lucin P, Koszinowski UH (1993) In vivo protection studies with MCMV glycoproteins gB and gH expressed by vaccinia virus. In: Michelson S, Plotkin SA (eds) Multidisciplinary approach to

Recent Approaches and Strategies CMV Vaccines understanding cytomegalovirus disease. Excerpta Medica, Amsterdam, pp 327–332 257. Reddehase MJ, Mutter W, Munch K, Buhring HJ, Koszinowski UH (1987) CD8-positive T lymphocytes specific for murine cytomegalovirus immediate-early antigens mediate protective immunity. J Virol 61:3102–3108 258. Del Val M, Schlicht HJ, Volkmer H, Messerle M, Reddehase MJ, Koszinowski UH (1991) Protection against lethal cytomegalovirus infection by a recombinant vaccine containing a single nonameric T-cell epitope. J Virol 65:3641–3646 259. Mohr CA, Arapovic J, Muhlbach H, Panzer M, Weyn A, Dolken L, Krompotic A, Voehringer D, Ruzsics Z, Koszinowski U, Sacher T (2010) A spread-deficient cytomegalovirus for assessment of first-target cells in vaccination. J Virol 84:7730–7742 260. Schlub TE, Sun JC, Walton SM, Robbins SH, Pinto AK, Munks MW, Hill AB, Brossay L, Oxenius A, Davenport MP (2011) Comparing the kinetics of NK cells, CD4, and CD8 T cells in murine cytomegalovirus infection. J Immunol 187:1385–1392 261. Gonzalez Armas JC, Morello CS, Cranmer LD, Spector DH (1996) DNA immunization confers protection against murine cytomegalovirus infection. J Virol 70:7921–7928 262. Morello CS, Cranmer LD, Spector DH (2000) Suppression of murine cytomegalovirus (MCMV) replication with a DNA vaccine encoding MCMV M84 (a homolog of human cytomegalovirus pp65). J Virol 74:3696–3708 263. Hakki M, Riddell SR, Storek J, Carter RA, Stevens-Ayers T, Sudour P, White K, Corey L, Boeckh M (2003) Immune reconstitution to cytomegalovirus after allogeneic hematopoietic stem cell transplantation: impact of host factors, drug therapy, and subclinical reactivation. Blood 102:3060–3067 264. Slavuljica I, Busche A, Babic M, Mitrovic M, Gasparovic I, Cekinovic D, Markova Car E, Pernjak Pugel E, Cikovic A, Lisnic VJ, Britt WJ, Koszinowski U, Messerle M, Krmpotic A, Jonjic S (2010) Recombinant mouse cytomegalovirus expressing a ligand for the NKG2D receptor is attenuated and has improved vaccine properties. J Clin Invest 120:4532–4545 265. Cekinovic D, Golemac M, Pugel EP, Tomac J, Cicin-Sain L, Slavuljica I, Bradford R, Misch S, Winkler TH, Mach M, Britt WJ, Jonjic S (2008) Passive immunization reduces murine cytomegalovirus-induced brain pathology in newborn mice. J Virol 82:12172–12180

455

266. Klenovsek K, Weisel F, Schneider A, Appelt U, Jonjic S, Messerle M, BradelTretheway B, Winkler TH, Mach M (2007) Protection from CMV infection in immunodeficient hosts by adoptive transfer of memory B cells. Blood 110:3472–3479 267. Farroway LN, Gorman S, Lawson MA, Harvey NL, Jones DA, Shellam GR, Singleton GR (2005) Transmission of two Australian strains of murine cytomegalovirus (MCMV) in enclosure populations of house mice (Mus domesticus). Epidemiol Infect 133:701–710 268. Gorman S, Harvey NL, Moro D, Lloyd ML, Voigt V, Smith LM, Lawson MA, Shellam GR (2006) Mixed infection with multiple strains of murine cytomegalovirus occurs following simultaneous or sequential infection of immunocompetent mice. J Gen Virol 87:1123–1132 269. Barry PA, Lockridge KM, Salamat S, Tinling SP, Yue Y, Zhou SS, Gospe SM Jr, Britt WJ, Tarantal AF (2006) Nonhuman primate models of intrauterine cytomegalovirus infection. ILAR J 47:49–64 270. Assaf BT, Mansfield KG, Westmoreland SV, Kaur A, Oxford KL, Diamond DJ, Barry PA (2012) Patterns of acute rhesus cytomegalovirus (RhCMV) infection predict long-term RhCMV infection. J Virol 86:6354–6357 271. Lockridge KM, Sequar G, Zhou SS, Yue Y, Mandell CP, Barry PA (1999) Pathogenesis of experimental rhesus cytomegalovirus infection. J Virol 73:9576–9583 272. Kaur A, Daniel MD, Hempel D, Lee-Parritz D, Hirsch MS, Johnson RP (1996) Cytotoxic T-lymphocyte responses to cytomegalovirus in normal and simian immunodeficiency virus-infected rhesus macaques. J Virol 70:7725–7733 273. Yue Y, Zhou SS, Barry PA (2003) Antibody responses to rhesus cytomegalovirus glycoprotein B in naturally infected rhesus macaques. J Gen Virol 84:3371–3379 274. Kaur A, Hale CL, Noren B, Kassis N, Simon MA, Johnson RP (2002) Decreased frequency of cytomegalovirus (CMV)-specific CD4+ T lymphocytes in simian immunodeficiency virus-infected rhesus macaques: inverse relationship with CMV viremia. J Virol 76:3646–3658 275. Kaur A, Kassis N, Hale CL, Simon M, Elliott M, Gomez-Yafal A, Lifson JD, Desrosiers RC, Wang F, Barry P, Mach M, Johnson RP (2003) Direct relationship between suppression of virus-specific immunity and emergence of cytomegalovirus disease in simian AIDS. J Virol 77:5749–5758

456

Suresh B. Boppana and William J. Britt

276. Yue Y, Wang Z, Abel K, Li J, Strelow L, Mandarino A, Eberhardt MK, Schmidt KA, Diamond DJ, Barry PA (2008) Evaluation of recombinant modified vaccinia Ankara virusbased rhesus cytomegalovirus vaccines in rhesus macaques. Med Microbiol Immunol 197:117–123 277. Yue Y, Kaur A, Eberhardt MK, Kassis N, Zhou SS, Tarantal AF, Barry PA (2007) Immunogenicity and protective efficacy of DNA vaccines expressing rhesus cytomegalovirus glycoprotein B, phosphoprotein 65-2, and viral interleukin-10 in rhesus macaques. J Virol 81:1095–1109 278. Wussow F, Yue Y, Martinez J, Deere JD, Longmate J, Herrmann A, Barry PA, Diamond DJ (2013) A vaccine based on rhesus cytomegalovirus UL128 complex induces broadly neutralizing antibodies in rhesus macaques. J Virol 87:1322–1332 279. Valencia S, Gill RB, Dowdell KC, Wang Y, Hornung R, Bowman JJ, Lacayo JC, Cohen JI (2019) Comparison of vaccination with rhesus CMV (RhCMV) soluble gB with a RhCMV replication-defective virus deleted for MHC class I immune evasion genes in a RhCMV challenge model. Vaccine 37:333–342 280. Sturgill ER, Malouli D, Hansen SG, Burwitz BJ, Seo S, Schneider CL, Womack JL, Verweij MC, Ventura AB, Bhusari A, Jeffries KM, Legasse AW, Axthelm MK, Hudson AW, Sacha JB, Picker LJ, Fru¨h K (2016) Natural killer cell evasion is essential for infection by Rhesus cytomegalovirus. PLoS Pathog 12: e1005868 281. Hansen SG, Vieville C, Whizin N, CoyneJohnson L, Siess DC, Drummond DD, Legasse AW, Axthelm MK, Oswald K, Trubey CM, Piatak M Jr, Lifson JD, Nelson JA, Jarvis MA, Picker LJ (2009) Effector memory T cell responses are associated with protection of rhesus monkeys from mucosal simian immunodeficiency virus challenge. Nat Med 15:293–299 282. Hansen SG, Zak DE, Xu G, Ford JC, Marshall EE, Malouli D, Gilbride RM, Hughes CM, Ventura AB, Ainslie E, Randall KT, Selseth AN, Rundstrom P, Herlache L, Lewis MS, Park H, Planer SL, Turner JM, Fischer M, Armstrong C, Zweig RC, Valvo J, Braun JM, Shankar S, Lu L, Sylwester AW, Legasse AW, Messerle M, Jarvis MA, Amon LM, Aderem A, Alter G, Laddy DJ, Stone M, Bonavia A, Evans TG, Axthelm MK, Fru¨h K, Edlefsen PT, Picker LJ (2018) Prevention of tuberculosis in rhesus macaques

by a cytomegalovirus-based vaccine. Nat Med 24:130–143 283. Hansen SG, Powers CJ, Richards R, Ventura AB, Ford JC, Siess D, Axthelm MK, Nelson JA, Jarvis MA, Picker LJ, Fruh K (2010) Evasion of CD8+ T cells is critical for superinfection by cytomegalovirus. Science 328:102–106 284. Holtappels R, Podlech J, Grzimek NK, Thomas D, Pahl-Seibert MF, Reddehase MJ (2001) Experimental preemptive immunotherapy of murine cytomegalovirus disease with CD8 T-cell lines specific for ppM83 and pM84, the two homologs of human cytomegalovirus tegument protein ppUL83 (pp65). J Virol 75:6584–6600 285. Podlech J, Holtappels R, Pahl-Seibert MF, Steffens HP, Reddehase MJ (2000) Murine model of interstitial cytomegalovirus pneumona in syngenic bone marrow transplantation: persistence of protective pulmonary CD8-T-cell infiltrates after clearance of acute infection. J Virol 74:7496–7507 286. Holtappels R, Munks MW, Podlech J, Reddehase MJ (2006) CD8 T-cell-based immunotherapy of cytomegalovirus diseae in the mouse model of the immunocompromised bone marrow transplantation recipient. In: Reddehase MJ (ed) Cytomegaloviruses: molecular biology and immunology. Caister, Norfolk, pp 383–419 287. Hill AB (2018) The immune response to CMV infection and vaccination in mice, monkeys and humans: recent developments. Curr Opin Virol 28:161–166 288. Scalzo AA, Fitzgerald NA, Wallace CR, Gibbons AE, Smart YC, Burton RC, Shellam GR (1992) The effect of the Cmv-1 resistance gene, which is linked to the natural killer cell gene complex, is mediated by natural killer cells. J Immunol 149:581–589 289. Scalzo AA, Fitzgerald NA, Simmons A, La Vista AB, Shellam GR (1990) Cmv-1, a genetic locus that controls murine cytomegalovirus replication in the spleen. J Exp Med 171:1469–1483 290. Brown MG, Zhang J, Du Y, Stoll J, Yokoyama WM, Scalzo AA (1999) Localization on a physical map of the NKC-linked Cmv1 locus between Ly49b and the Prp gene cluster on mouse chromosome 6. J Immunol 163:1991–1999 291. Webb JR, Lee SH, Vidal SM (2002) Genetic control of innate immune responses against cytomegalovirus: MCMV meets its match. Genes Immun 3:250–262

Recent Approaches and Strategies CMV Vaccines 292. Lee SH, Girard S, Macina D, Busa M, Zafer A, Belouchi A, Gros P, Vidal SM (2001) Susceptibility to mouse cytomegalovirus is associated with deletion of an activating natural killer cell receptor of the C-type lectin superfamily. Nat Genet 28:42–45 293. Depatie C, Chalifour A, Pare C, Lee SH, Vidal SM, Lemieux S (1999) Assessment of Cmv1 candidates by genetic mapping and in vivo antibody depletion of NK cell subsets. Int Immunol 11:1541–1551 294. Cheng TP, French AR, Plougastel BF, Pingel JT, Orihuela MM, Buller ML, Yokoyama WM (2008) Ly49h is necessary for genetic resistance to murine cytomegalovirus. Immunogenetics 60:565–573 295. Cooper MA, Yokoyama WM (2010) Memory-like responses of natural killer cells. Immunol Rev 235:297–305 296. Elliott JM, Yokoyama WM (2011) Unifying concepts of MHC-dependent natural killer cell education. Trends Immunol 32:364–372 297. Fodil-Cornu N, Lee SH, Belanger S, Makrigiannis AP, Biron CA, Buller RM, Vidal SM (2008) Ly49h-deficient C57BL/6 mice: a new mouse cytomegalovirus-susceptible model remains resistant to unrelated pathogens controlled by the NK gene complex. J Immunol 181:6394–6405 298. Zeleznjak J, Popovic B, Krmpotic A, Jonjic S, Lisnic VJ (2017) Mouse cytomegalovirus encoded immunoevasins and evolution of Ly49 receptors—Sidekicks or enemies? Immunol Lett 189:40–47 299. Hadaya K, de Rham C, Bandelier C, Bandelier C, Ferrari-Lacraz S, Jendly S, Berney T, Buhler L, Kaiser L, Seebach JD, Tiercy JM, Martin PY, Villard J (2008) Natural killer cell receptor repertoire and their ligands, and the risk of CMV infection after kidney transplantation. Am J Transplant 8:2674–2683 300. Bacon L, Eagle RA, Meyer M, Easom N, Young NT, Trowsdale J (2004) Two human ULBP/RAET1 molecules with transmembrane regions are ligands for NKG2D. J Immunol 173:1078–1084 301. Bauer S, Groh V, Wu J, Steinle A, Phillips JH, Lanier LL, Spies T (1999) Activation of NK cells and T cells by NKG2D, a receptor for stress-inducible MICA. Science 285:727–729 302. Chalupny NJ, Sutherland CL, Lawrence WA, Rein-Weston A, Cosman D (2003) ULBP4 is a novel ligand for human NKG2D. Biochem Biophys Res Commun 305:129–135 303. Eagle RA, Traherne JA, Ashiru O, Wills MR, Trowsdale J (2006) Regulation of NKG2D

457

ligand gene expression. Hum Immunol 67:159–169 304. Eagle RA, Traherne JA, Hair JR, Jafferji I, Trowsdale J (2009) ULBP6/RAET1L is an additional human NKG2D ligand. Eur J Immunol 39:3207–3216 305. Rolle A, Mousavi-Jazi M, Eriksson M, Odeberg J, Soderberg-Naucler C, Cosman D, Karre K, Cerboni C (2003) Effects of human cytomegalovirus infection on ligands for the activating NKG2D receptor of NK cells: up-regulation of UL16-binding protein (ULBP)1 and ULBP2 is counteracted by the viral UL16 protein. J Immunol 171:902–908 306. Orange JS (2013) Natural killer cell deficiency. J Allergy Clin Immunol 132:515–525 307. Walton SM, Wyrsch P, Munks MW, Zimmermann A, Hengel H, Hill AB, Oxenius A (2008) The dynamics of mouse cytomegalovirus-specific CD4 T cell responses during acute and latent infection. J Immunol 181:1128–1134 308. Jonjic S, Mutter W, Weiland F, Reddehase MJ, Koszinowski UH (1989) Site-restricted persistent cytomegalovirus infection after selective long-term depletion of CD4+ T lymphocytes. J Exp Med 169:1199–1212 309. Lucin P, Pavic I, Polic B, Jonjic S, Koszinowski UH (1992) Gamma interferondependent clearance of cytomegalovirus infection in salivary glands. J Virol 66:1977–1984 310. Brizic´ I, Hirsˇl L, Sˇustic´ M, Golemac M, Britt WJ, Krmpotic´ A, Jonjic´ S (2019) CD4 T cells are required for maintenance of CD8 T (RM) cells and virus control in the brain of MCMV-infected newborn mice. Med Microbiol Immunol 208:487–494 311. Holtappels R, Podlech J, Geginat G, Steffens HP, Thomas D, Reddehase MJ (1998) Control of murine cytomegalovirus in the lungs: relative but not absolute immunodominance of the immediate-early 1 nonapeptide during the antiviral cytolytic T-lymphocyte response in pulmonary infiltrates. J Virol 72:7201–7212 312. Reddehase MJ, Weiland F, Munch K, Jonjic S, Luske A, Koszinowski UH (1985) Interstitial murine cytomegalovirus pneumonia after irradiation: characterization of cells that limit viral replication during established infection of the lungs. J Virol 55:264–273 313. Holtappels R, Bohm V, Podlech J, Reddehase MJ (2008) CD8 T-cell-based immunotherapy of cytomegalovirus infection: "proof of

458

Suresh B. Boppana and William J. Britt

concept" provided by the murine model. Med Microbiol Immunol 197:125–134 314. Kurz S, Steffens HP, Mayer A, Harris JR, Reddehase MJ (1997) Latency versus persistence or intermittent recurrences: evidence for a latent state of murine cytomegalovirus in the lungs. J Virol 71:2980–2987 315. Sell S, Dietz M, Schneider A, Holtappels R, Mach M, Winkler TH (2015) Control of murine cytomegalovirus infection by γδ T cells. PLoS Pathog 11:e1004481 316. Baroncelli S, Barry PA, Capitanio JP, Lerche NW, Otsyula M, Mendoza SP (1997) Cytomegalovirus and simian immunodeficiency virus coinfection: longitudinal study of antibody responses and disease progression. J Acquir Immune Defic Syndr Hum Retrovirol 15:5–15 317. Sequar G, Britt WJ, Lakeman FD, Lockridge KM, Tarara RP, Canfield DR, Zhou SS, Gardner MB, Barry PA (2002) Experimental coinfection of rhesus macaques with rhesus cytomegalovirus and simian immunodeficiency virus: pathogenesis. J Virol 76:7661–7671 318. Bialas KM, Permar SR (2016) The march towards a vaccine for congenital CMV: rationale and models. PLoS Pathog 12:e1005355 319. Reuter JD, Gomez DL, Wilson JH, Van Den Pol AN (2004) Systemic immune deficiency necessary for cytomegalovirus invasion of the mature brain. J Virol 78:1473–1487 320. Woolf NK, Jaquish DV, Koehrn FJ (2007) Transplacental murine cytomegalovirus infection in the brain of SCID mice. Virol J 4:26 321. Bia FJ, Griffith BP, Fong CK, Hsiung GD (1983) Cytomegaloviral infections in the guinea pig: experimental models for human disease. Rev Infect Dis 5:177–195 322. Bia FJ, Miller SA, Davidson KH (1984) The guinea pig cytomegalovirus model of congenital human cytomegalovirus infection. Birth Defects 20:233–241 323. Griffith BP, McCormick SR, Booss J, Hsiung GD (1986) Inbred guinea pig model of intrauterine infection with cytomegalovirus. Am J Pathol 122:112–119 324. Borune N, Schleiss M, Bravo F, Bernstein D (2001) Preconception immunization with a cytomegalovirus (CMV) glycoprotein vaccine improves pregnancy outcome in a guniea pig model of congenital CMV infection. J Infect Dis 183:59–64 325. Schleiss MR (2007) Comparison of vaccine strategies against congenital CMV infection in the guinea pig model. J Clin Virol 41:224–230

326. Harrison CJ, Britt WJ, Chapan NM, Mullican J, Tracy S (1995) Reduced congenital cytomegalovirus (CMV) infection after maternal immunization with a guinea pig CMV glycoprotein before gestational primary CMV infection in the guinea pig model. J Infect Dis 172:1212–1220 327. Choi K, Root M, McGregor A (2016) A novel non-replication-competent cytomegalovirus capsid mutant vaccine strategy is effective in reducing congenital infection. J Virol 90:7902–7919 328. Schleiss MR, Permar SR, Plotkin SA (2017) Progress toward development of a vaccine against congenital cytomegalovirus infection. Clin Vaccine Immunol 24:e00268–e00217 329. Swanson EC, Gillis P, Hernandez-Alvarado N, Ferna´ndez-Alarco´n C, Schmit M, Zabeli JC, Wussow F, Diamond DJ, Schleiss MR (2015) Comparison of monovalent glycoprotein B with bivalent gB/pp65 (GP83) vaccine for congenital cytomegalovirus infection in a guinea pig model: Inclusion of GP83 reduces gB antibody response but both vaccine approaches provide equivalent protection against pup mortality. Vaccine 33:4013–4018 330. Schleiss MR, Bourne N, Stroup G, Bravo FJ, Jensen NJ, Bernstein DI (2004) Protection against congenital cytomegalovirus infection and disease in guinea pigs, conferred by a purified recombinant glycoprotein B vaccine. J Infect Dis 189:1374–1381 331. Auerbach MR, Yan D, Vij R, Hongo J-A, Nakamura G, Vernes J-M, Meng YG, Lein S, Chan P, Ross J, Carano R, Deng R, LewinKoh N, Xu M, Feierbach B (2014) A neutralizing anti-gH/gL monoclonal antibody is protective in the guinea pig model of congenital CMV infection. PLoS Pathog 10: e1004060 332. Chatterjee A, Harrison CJ, Britt WJ, Li O, Bewtra C (1999) Immunization with hyperimmune anti-gB serum prevents congenital CMV infection in guinea pigs, In APS/SPR Annual Meeting, San Francisco, CA 333. Hashimoto K, Yamada S, Katano H, Fukuchi S, Sato Y, Kato M, Yamaguchi T, Moriishi K, Inoue N (2013) Effects of immunization of pregnant guinea pigs with guinea pig cytomegalovirus glycoprotein B on viral spread in the placenta. Vaccine 31:3199–3205 334. Woolf NK (1991) Guinea pig model of congenital CMV-induced hearing loss: a review. Transplant Proc 23:32–34 335. Park AH, Gifford T, Schleiss MR, Dahlstrom L, Chase S, McGill L, Li W, Alder SC (2010) Development of

Recent Approaches and Strategies CMV Vaccines cytomegalovirus-mediated sensorineural hearing loss in a Guinea pig model. Arch Otolaryngol Head Neck Surg 136:48–53 336. Bialas KM, Tanaka T, Tran D, Varner V, Cisneros De La Rosa E, Chiuppesi F, Wussow F, Kattenhorn L, Macri S, Kunz EL, Estroff JA, Kirchherr J, Yue Y, Fan Q, Lauck M, O’Connor DH, Hall AHS, Xavier A, Diamond DJ, Barry PA, Kaur A, Permar SR (2015) Maternal CD4+ T cells protect against severe congenital cytomegalovirus disease in a novel nonhuman primate model of placental cytomegalovirus transmission. Proc Natl Acad Sci U S A 112:13645–13650 337. Revello MG, Lazzarotto T, Guerra B, Spinillo A, Ferrazi E, Kustermann A, Guaschino S, Vergani P, Todros T, Frusca T, Arossa A, Furione M, Rognoni V, Rizzo N, Gabrielli L, Klersy C, Gerna G, Group., f. t. C. S (2014) A randomized trial of hyperimmune globulin to prevent congenital cytomegalovirus. N Engl J Med 370:1316–1326 338. Bla´zquez-Gamero D, Galindo Izquierdo A, Del Rosal T, Baquero-Artigao F, Izquierdo Me´ndez N, Soriano-Ramos M, Rojo Conejo P, Gonza´lez-Tome´ MI, Garcı´aBurguillo A, Pe´rez Pe´rez N, Sa´nchez V, Ramos-Amador JT, De la Calle M (2019) Prevention and treatment of fetal cytomegalovirus infection with cytomegalovirus hyperimmune globulin: a multicenter study in Madrid. J Matern Retal Neonatal Med 32:617–625 339. Britt WJ, Cekinovic D, Jonjic S (2013) Murine model of neonatal cytomegalovirus infection. In: Reddehasse M (ed) Cytomegaloviruses. Cassister, London 340. Koontz T, Bralic M, Tomac J, Pernjak-PugelE, Bantug G, Jonjic S, Britt WJ (2008) Altered development of the brain after focal herpesvirus infection of the central nervous system. J Exp Med 205:423–435 341. Bradford RD, Yoo Y-G, Golemac M, Pugel EP, Jonjic S, Britt WJ (2015) Murine CMV-induced hearing loss is associated with inner ear inflammation and loss of spiral ganglia neurons. PLoS Pathog 11:e1004774 342. Kosmac K, Bantug GR, Pugel EP, Cekinovic D, Jonjic S, Britt WJ (2013) Glucocorticoid treatment of MCMV infected newborn mice attenuates CNS inflammation and limits deficits in cerebellar development. PLoS Pathog 9:e1003200 343. Seleme MC, Kosmac K, Jonjic S, Britt WJ (2017) Tumor necrosis factor alpha-induced recruitment of inflammatory mononuclear cells leads to inflammation and altered brain development in murine cytomegalovirus-

459

infected newborn mice. J Virol 91: e01983–e01916 344. Sung CYW, Seleme MC, Payne S, Jonjic S, Hirose K, Britt W (2019) Virus-induced cochlear inflammation in newborn mice alters auditory function. JCI Insight 4:e128878 345. Stagno S, Reynolds DW, Huang E-S, Thames SD, Smith RJ, Alford CA (1977) Congenital cytomegalovirus infection: occurrence in an immune population. N Engl J Med 296:1254–1258 346. Adler SP, Starr SE, Plotkin SA, Hempfling SH, Buis J, Manning ML, Best AM (1995) Immunity induced by primary human cytomegalovirus infection protects against secondary infection among women of childbearing age. J Infect Dis 171:26–32 347. Pass RF, Zhang C, Evans A, Simpson T, Andrews W, Huang ML, Corey L, Hill J, Davis E, Flanagan C, Cloud G (2009) Vaccine prevention of maternal cytomegalovirus infection. N Engl J Med 360:1191–1199 348. Bernstein DI, Munoz FM, Callahan ST, Rupp R, Wootton SH, Edwards KM, Turley CB, Stanberry LR, Patel SM, Mcneal MM, Pichon S, Amegashie C, Bellamy AR (2016) Safety and efficacy of a cytomegalovirus glycoprotein B (gB) vaccine in adolescent girls: A randomized clinical trial. Vaccine 34:313–319 349. Hansen SG, Ford JC, Lewis MS, Ventura AB, Hughes CM, Coyne-Johnson L, Whizin N, Oswald K, Shoemaker R, Swanson T, Legasse AW, Chiuchiolo MJ, Parks CL, Axthelm MK, Nelson JA, Jarvis MA, Piatak M Jr, Lifson JD, Picker LJ (2011) Profound early control of highly pathogenic SIV by an effector memory T-cell vaccine. Nature 473:523–527 350. Wloch MK, Smith LR, Boutsaboualoy S, Reyes L, Han C, Kehler J, Smith HD, Selk L, Nakamura R, Brown JM, Marbury T, Wald A, Rolland A, Kaslow D, Evans T, Boeckh M (2008) Safety and immunogenicity of a bivalent cytomegalovirus DNA vaccine in healthy adult subjects. J Infect Dis 197:1634–1642 351. Griffiths PD, Stanton A, McCarrell E, Smith C, Osman M, Harber M, Davenport A, Jones G, Wheeler DC, O’Beirne J, Thorburn D, Patch D, Atkinson CE, Pichon S, Sweny P, Lanzman M, Woodford E, Rothwell E, Old N, Kinyanjui R, Haque T, Atabani S, Luck S, Prideaux S, Milne RS, Emery VC, Burroughs AK (2011) Cytomegalovirus glycoprotein-B vaccine with MF59 adjuvant in transplant recipients: a phase 2 randomised placebocontrolled trial. Lancet 377:1256–1263

460

Suresh B. Boppana and William J. Britt

352. Bernstein DI, Reap EA, Katen K, Watson A, Smith K, Norberg P, Olmsted RA, Hoeper A, Morris J, Negri S, Maughan MF, Chulay JD (2009) Randomized, double-blind, phase I trial of alphavirus replicon vaccine for cytomegalovirus in CMV seronegative adult volunteers. Vaccine 28:484–493 353. Heineman TC, Schleiss M, Bernstein DI, Spaete RR, Yan L, Duke G, Prichard M, Wang Z, Yan Q, Sharp MA, Klein N, Arvin AM, Kemble G (2006) A phase 1 study of 4 live, recombinant human cytomegalovirus Towne/Toledo chimeric vaccines. J Infect Dis 193:1350–1360 354. La Rosa C, Longmate J, Lacey SF, Kaltcheva T, Sharan R, Marsano D, Kwon P, Drake J, Williams B, Denison S, Broyer S, Couture L, Nakamura R, Kelsey MI, Krieg AM, Diamond DJ, Zaia JA (2012) Clinical evaluation of safety and immunogenicity of PADRE-cytomegalovirus (CMV) and tetanus-CMV fusion peptide vaccines with or without PF03512676 adjuvant. J Infect Dis 205:1294–1304 355. Snydman DR (1993) Review of the efficacy of cytomegalovirus immune globulin in the prophylaxis of CMV disease in renal transplant recipients. Transplant Proc 25:25–26 356. Yeager AS, Grumet FC, Hafleigh EB, Arvin AM, Bradley JS, Prober CG (1981) Prevention of transfusion-acquired cytomegalovirus infections in newborn infants. J Pediatr 98:281–287 357. Rubin R (2002) Clinical approach to infection in the compromised host. In: Rubin R, Young LS (eds) Infection in the organ transplant recipient. Kluwer Academic Press, New York, pp 573–679 358. Nigro G, Adler SP (2011) Cytomegalovirus infections during pregnancy. Curr Opin Obstet Gynecol 23:123–128 359. Hedlund M, Stenqvist AC, Nagaeva O, Kjellberg L, Wulff M, Baranov V, MinchevaNilsson L (2009) Human placenta expresses and secretes NKG2D ligands via exosomes that down-modulate the cognate receptor expression: evidence for immunosuppressive function. J Immunol 183:340–351 360. Iwasenko JM, Howard J, Arbuckle S, Graf N, Hall B, Craig ME, Rawlinson WD (2011) Human cytomegalovirus infection is detected frequently in stillbirths and is associated with fetal thrombotic vasculopathy. J Infect Dis 203:1526–1533 361. La Torre R, Nigro G, Mazzocco M, Best AM, Adler SP (2006) Placental enlargement in women with primary maternal

cytomegalovirus infection is associated with fetal and neonatal disease. Clin Infect Dis 43:994–1000 362. Pereira L, Maidji E (2008) Cytomegalovirus infection in the human placenta: maternal immunity and developmentally regulated receptors on trophoblasts converge. Curr Top Microbiol Immunol 325:383–395 363. Scott GM, Chow SS, Craig ME, Pang CN, Hall B, Wilkins MR, Jones CA, Lloyd AR, Rawlinson WD (2012) Cytomegalovirus infection during pregnancy with maternofetal transmission induces a proinflammatory cytokine bias in placenta and amniotic fluid. J Infect Dis 205:1305–1310 364. Cui X, Meza BP, Adler SP, McVoy MA (2008) Cytomegalovirus vaccines fail to induce epithelial entry neutralzing antibodies comparable to natural infection. Vaccine 26:5760–5766 365. Gerna G, Sarasini A, Partone M, Percivalle E, Fiorina L, Campanini G, Gallina A, Baldanti F, Revello MG (2008) Human cytomegalvirus serum neutralizing antibodies block virus infection of endothelial/epithelial cells but not fibroblasts, early during primary infection. J Gen Virol 89:853–865 366. Lilleri D, Kabanova A, Lanzavecchia A, Gerna G (2012) Antibodies against neutralization epitopes of human cytomegalovirus gH/gL/pUL128-130-131 complex and virus spreading may correlate with virus control in vivo. J Clin Immunol 32 (6):1324–1331 367. Jacobson MA, Adler SP, Sinclair E, Black D, Smith A, Chu A, Moss RB, Wloch MK (2009) A CMV DNA vaccine primes for memory immune responses to live-attenuated CMV (Towne strain). Vaccine 27:1540–1548 368. Boppana SB, Miller J, Britt WJ (1996) Transplacentally acquired antiviral antibodies and outcome in congenital human cytomegalovirus infection. Viral Immunol 9:211–218 369. Borysiewicz LK, Graham KS, Hickling JK, Mason PD, Sissons JCP (1988) Human cytomegalovirus-specific cytotoxic T cells: their precursor frequency and stage specificity. Eur J Immunol 18:269–275 370. Wills MR, Carmichael AJ, Mynard K, Jin X, Weekes MP, Plachter B, Sissons JG (1996) The human cytotoxic T lymphocyte (CTL) response to cytomegalovirus is dominated by structural protein pp65: frequency, specificity and T-cell receptor usage of pp65-specific CTL. J Virol 70:7569–7579 371. Pass RF, Duliege AM, Boppana SB, Britt WJ, Granoff DM, Sekulovich R, Burke RL (1995)

Recent Approaches and Strategies CMV Vaccines A phase I trial of BIOCINE CMV gB vaccine in seronegative adults, In Fifth International Cytomegalovirus Conference, Stockholm, Sweden 372. Sabbaj S, Pass RF, Goepfert PA, Pichon S (2011) Glycoprotein B vaccine is capable of boosting both antibody and CD4 T-cell responses to cytomegalovirus in chronically infected women. J Infect Dis 203:1534–1541 373. Bubic I, Wagner M, Krmpoti A, Saulig T, Kim S, Yokoyama WM, Jonji S, Koszinowski UH (2004) Gain of virulence caused by loss of a gene in murine cytomegalovirus. J Virol 78:7536–7544 374. Crnkovic-Mertens I, Messerle M, Milotic I, Szepan U, Kucic N, Krmpotic A, Jonjic S, Koszinowski UH (1998) Virus attenuation after deletion of the cytomegalovirus Fc receptor gene is not due to antibody control. J Virol 72:1377–1382 375. Chang WL, Barry PA, Szubin R, Wang D, Baumgarth N (2009) Human cytomegalovirus suppresses type I interferon secretion by plasmacytoid dendritic cells through its interleukin 10 homolog. Virology 390:330–337 376. Logsdon NJ, Eberhardt MK, Allen CE, Barry PA, Walter MR (2011) Design and analysis of rhesus cytomegalovirus IL-10 mutants as a model for novel vaccines against human cytomegalovirus. PLoS One 6:e28127 377. Jones BC, Logsdon NJ, Josephson K, Cook J, Barry PA, Walter MR (2002) Crystal structure of human cytomegalovirus IL-10 bound to soluble human IL-10R1. Proc Natl Acad Sci U S A 99:9404–9409 378. Hughes BL (2019) Randomized trial to prevent congenital cytomegalovirus, In IDWeek 2019, Washington, DC 379. Stagno S, Dworsky M, Torres J, Mesa T, Hirsh T (1982) Prevalence and importance of congenital cytomegalovirus infection in three different populations. J Pediatr 101:897–900 380. Adler SP, Finney JW, Manganello AM, Best AM (1996) Prevention of child-to-mother transmission of cytomegalovirus by changing behaviors: a randomized controlled trial. Ped Infect Dis J 15:240–246 381. Picone O, Vauloup-Fellous C, Cordier AG, Parent Du Chatelet I, Senat MV, Frydman R, Grangeot-Kaeros L (2009) A 2-year study on cytomegalovirus infection during pregnancy in a French hospital. BJOC 116:818–823 382. Revello MG, Tibaldi G, Masuelli G, Frisina V, Sacchi A, Furione M, Arossa A, Spinillo A, Klersy C, Ceccarelli M, Gerna G, Todros T

461

(2015) Prevention of primary cytomegalovirus infection in pregnancy. EBioMedicine 2:1205–1210 383. Paya CV, Humar A, Dominguez E, Washburn K, Blumberg E, Alexander B, Freeman R, Heaton N, Pescovitz MD, Group, V. S. O. T. S (2004) Efficacy and safety of valganciclovir vs oral ganciclovir for prevention of cytomegalovirus disease in solid organ transplant recipients. Am J Transplant 4:611–620 384. Kotton CN, Kumar D, Caliendo AM, Asberg A, Chou S, Snydman DR, Allen U, Humar A (2010) International consensus guidelines on the management of cytomegalovirus in solid organ transplantation. Transplantation 89:779–795 385. Asberg A, Humar A, Jardine AG, Rollag H, Pescovitz MD, Mouas H, Bignamini A, Toz H, Dittmer I, Montejo M, Hartmann A, Group, V. S (2009) Longterm outcomes of CMV disease treatment with valganciclovir versus IV ganciclovir in solid organ transplant recipients. Am J Transplant 9:1205–1213 386. Ljungman P, de la Camara R, Cordonnier C, Einsele H, Englehard D, Reusser P, Styczynski J, Ward K, Leukemia, E. C. o. I. i (2008) Manamagement of CMV, HHV-6, HHV-7 and Kaposi-sarcoma associated herpesvirus (HHV-8) infections in patients with hematological malignancies and after SCT. Bone Marrow Transplant 42:227–240 387. Tomblyn M, Chiller T, Einsele H, Gress R, Sepkowitz K, Storek J, Wingard JR, Young JA, Boeckh MJ, Center for International Blood and Marrow Transplant Research (CIBMTR); National Marrow Donor Program (NMDP); European Blood and Marrow Transplant Group (EBMT); American Society of Blood and Marrow Transplantation (ASBMT); Canadian Blood and Marrow Transplant Group (CBMTG); Infectious Disease Society of America (IDSA); Society for Healthcare Epidemiology of America (SHEA); Association of Medical Microbiology and Infectious Diseases Canada (AMMI); Centers for Disease Control and Prevention (CDC) (2009) Guidelines for preventing infectious complications among hematopoietic cell transplant recipients: a global perspective. Biol Blood Marrow Transplant 15:1143–1238 388. El Helou G, Razonable RR (2019) Letermovir for the prevention of cytomegalovirus infection and disease in transplant recipients: an evidence-based review. Infect Drug Resist 12:1481–1491

462

Suresh B. Boppana and William J. Britt

389. Drew WL, Sweet ES, Miner RC, Mocarski ES (1984) Multiple infections by cytomegalovirus in patients with acquired immune deficiency syndrome: documentation by Southern blot hybridization. J Infect Dis 150:952–953 390. Burkhardt C, Himmelein S, Britt W, Winkler T, Mach M (2009) Glycoprotein N subtypes of human cytomegalovirus induce a strain-specific antibody response during natural infection. J Gen Virol 90:1951–1961 391. Dal Monte P, Pignatelli S, Mach M, Landini MP (2001) The product of human cytomegalovirus UL73 is a new polymorphic structural glycoprotein (gpUL73). J Hum Virol 4:26–34 392. Rasmussen L, Geissler A, Cowan C, Chase A, Winters M (2002) The genes encoding the gCIII complex of human cytomegalovirus exist in highly diverse combinations in clinical isolates. J Virol 76:10841–10848 393. Cudini J, Roy S, Houldcroft CJ, Bryant JM, Depledge DP, Tutill H, Veys P, Williams R, Worth AJJ, Tamuri AU, Goldstein RA, Breuer J (2019) Human cytomegalovirus haplotype reconstruction reveals high diversity due to superinfection and evidence of within-host recombination. Proc Natl Acad Sci U S A 116:5693–5698 394. Hodson EM, Barclay PG, Craig JC, Jones C, Kable K, Strippoli GF, Vimalachandra D, Webster AC (2005) Antiviral medications for preventing cytomegalovirus disease in solid organ transplant recipients. Cochrane Database Syst Rev 2:CD003774 395. Mussi-Pinhata MM, Yamamoto AY (2020) Natural history of congenital cytomegalovirus infeciton in highly seropositive populations. J Infect Dis 221:S15–S22 396. Pass RF, Fowler KB, Boppana SB, Britt WJ, Stagno S (2006) Congenital cytomegalovirus infection following first trimester maternal infection: symptoms at birth and outcome. J Clin Virol 35:216–220 397. Enders G, Daiminger A, Bader U, Exler S, Enders M (2011) Intrauterine transmission and clinical outcome of 248 pregnancies with primary cytomegalovirus infection in relation to gestational age. J Clin Virol 52:244–246 398. Andersen HK, Brostrom K, Hansen KB, Leerhoy J, Pedersen M, Osterballe O, Felsager U, Mogensen S (1979) A prospective study on the incidence and significance of congenital cytomegalovirus infection. Acta Paediatr Scand 68:329–336

399. Larke RBP, Wheatley E, Saigal S, Chernesky MA (1980) Congenital cytomegalovirus infection in an urban Canadian community. J Infect Dis 142:647–653 400. Peckham CS, Chin KS, Coleman JC, Henderson K, Hurley R, Preece PM (1983) Cytomegalovirus infection in pregnancy: preliminary findings from a prospective study. Lancet 1:1352–1355 401. Child SJ, Hakki M, De Niro KL, Geballe AP (2004) Evasion of cellular antiviral responses by human cytomegalovirus TRS1 and IRS1. J Virol 78:197–205 402. Brune W, Nevels M, Shenk T (2003) Murine cytomegalovirus m41 open reading frame encodes a Golgi-localized antiapoptotic progein. J Virol 77:11633–11643 403. Jurak I, Schumacher U, Simic H, Voigt S, Brune W (2008) Murine cytomegalovirus m38.5 protein inhibits Bax-mediated cell death. J Virol 82:4812–4822 404. Cam M, Handke W, Picard-Maureau M, Brune W (2010) Cytomegaloviruses inhibit Bak- and Bax-mediated apoptosis with two separate viral proteins. Cell Death Differ 17:655–665 405. Skaletskaya A, Bartle LM, Chittenden T, McCormick AL, Mocarski E, Goldmacher VS (2001) A cytomegalovirus-encoded inhibitor of apoptosis that suppresses caspase-8activation. Proc Natl Acad Sci U S A 98:7829–7834 406. Brune W, Hasan M, Krych M, Bubic I, Jonjic S, Koszinowski UH (2001) Secreted virus-encoded proteins reflect murinr cytomegalovirus productivity in organs. J Infect Dis 184:1320–1324 407. Lenac T, Arapavic J, Traven L, Krmpotic A, Jonjic S (2008) Murine cytomegalovirus regulation of NKG2D ligands. Med Microbiol Immunol 197:159–166 408. Lodoen MB, Abenes G, Umamoto S, Houchins JP, Liu F, Lanier LL (2004) The cytomegalovirus m155 gene product subverts natural killer cell antiviral protection by disruption of H60-NKG2D interactions. J Exp Med 200:1075–1081 409. McCormick AL, Skaletskaya A, Barry PA, Mocarski ES, Goldmacher VS (2003) Differential function and expression of the viral inhibitor of caspase 8-induced apoptosis (vICA) and the viral mitochondria-localized inhibitor of apoptosis (vMIA) cell death suppressors conserved in primate and rodent cytomegaloviruses. Virology 316:221–233 410. Chalupny NJ, Rein-Weston A, Dosch S, Cosman D (2006) Down-regulation of the

Recent Approaches and Strategies CMV Vaccines NKG2D ligand MICA by the human cytomegalovirus glycoprotein UL142. Biochem Biophys Res Commun 346:175–181 411. Fruh K, Yang Y (1999) Antigen presentation by MHC class I and its regulation by interferon gamma. Curr Opin Immunol 11:76–81 412. Bubeck A, Reusch U, Wagner M, Ruppert T, Muranyi W, Kloetzel PM, Koszinowski U (2002) The glycoprotein gp48 of murine cytomegalovirusL proteasome-dependent cytosolic dislocation and degradation. J Biol Chem 277:2216–2224 413. Vidal SM, Lanier LL (2006) NK cell recognition of moouse cytomegalovirus-infected cells. Curr Top Microbiol Immunol 298:183–206 414. Babic M, Pyzik M, Zafirova B, Mitrovic M, Butorac V, Lanier LL, Krompotic A, Vidal SM, Jonjic S (2010) Cytomegalovirus immunoevasion reveals the physiological role of "missing self" recognition in natural killer cell dependent virus control in vivo. J Exp Med 207:2663–2673 415. Tomazin R, Boname J, Hegde NR, Lewinsohn DM, Altschuler Y, Jones TR, Cresswell P, Nelson JA, Riddell SR (1999) Cytomegalovirus US2 destroys two components of the MHC class II pathway, preventing recognition by CD+ T cells. Nat Med 5:1039–1043 416. Jackson SE, Mason GM, Wills MR (2011) Human cytomegalovirus immunity and immune evasion. Virus Res 157:151–160 417. Kharfan-Dabaja MA, Boeckh M, Wilck MB, Langston AA, Chu AH, Wloch MK, Guterwill DF, Smith LR, Rolland AP, Kenney RT (2012) A novel therapeutic cytomegalovirus DNA vaccine in allogeneic haemopoietic

463

stem-cell transplantation: a randomised, double-blind, placebo-controlled, phase 2 trial. Lancet Infect Dis 12:290–299 418. Adler SP, Lewis N, Conlon A, Christiansen MP, Al-Ibrahim M, Rupp R, Fu T-M, Bautista O, Tang H, Wang D, Fisher A, Culp T, Das R, Beck K, Tamms G, Musey L, V160-001 Study Group (2019) Phase 1 clinical trial of a conditionally replication-defective human cytomegalovirus (CMV) vaccine in CMV-seronegative subjects. J Infect Dis 220:411–419 419. La Rosa C, Longmate J, Martinez J, Zhou Q, Kaltcheva TI, Tsai W, Drake J, Carroll M, Wussow F, Chiuppesi F, Hardwick N, Dadwal S, Aldoss I, Nakamura R, Zaia JA, Diamond DJ (2017) MVA vaccine encoding CMV antigens safely induces durable expansion of CMV-specific T cells in healthy adults. Blood 129:114–125 420. Arora N, Novak Z, Fowler KB, Boppana SB, Ross SA (2010) Cytomegalovirus viruria and DNAemia in healthy seropositive women. J Infect Dis 202:1800–1803 421. Rivera LB, Boppana SB, Fowler KB, Britt WJ, Stagno S, Pass RF (2002) Predictors of hearing loss in children with symptomatic congenital cytomegalovirus infection. Pediatrics 110:762–767 422. Ross SA, Novak Z, Fowler KB, Arora N, Britt WJ, Boppana SB (2009) Cytomegalovirus blood viral load and hearing loss in young children with congenital infection. Pediatr Infect Dis J 28:588–592 423. Gnann JW, Ahlmen J, Svalander C et al (1988) Inflammatory cells in transplanted kidneys are infected by human cytomegalovirus. Am J Pathol 132:239–248

INDEX A Acute infection ................................................................ 21 AIDS patients ....................................................... 8, 9, 103 Allografts .................................................... 8, 9, 404, 409, 417–419, 438–441 Antibiotic selection ..................................... 149, 155, 205 Antivirals ........................................ 8, 9, 29, 32, 267, 269, 396, 409–416, 418–420, 422, 424–433, 437–439, 441 Apoptosis ............................... 5, 110, 111, 301, 420, 421 Atherosclerosis .............................................................. 2, 3

Cyclooxygenase-2 (COX-2) ......................................... 161 Cytokines ..............................................54, 69, 84, 89, 95, 96, 98, 104, 292, 350, 358, 359, 366, 370, 387, 392, 393 Cytomegalovirus (CMV) .............................. 1–11, 19–36, 51–80, 83–99, 103–111, 115–131, 133–156, 159–193, 199–209, 213–232, 247–263, 265, 270, 292, 302, 303, 309, 316, 331, 333, 343, 351, 359, 365–398, 403–441 Cytomegalovirus interleukin 10 (cmvIL-10) ..... 291–298 Cytoplasms .........................................116, 233, 266–269, 278, 279, 302

B

D

BAC purification ........................................................... 165 Bacterial artificial chromosomes (BAC)..................25–28, 31–34, 133–156, 165, 173, 176–179, 189, 193, 201, 204, 205, 215, 219–222, 231, 248, 304, 313, 357 B-cell responses ............................................................. 346 Blood cells .......................................................52, 60, 237, 352, 368, 376, 404 Blood products................................................... 6, 52, 404 Brain.................................................... 366–368, 370–372, 374, 376, 386–398, 425 Breast milk ............................................... 6, 366, 404, 405 Buffy coat cells ..................................................... 106, 389

Density gradients ....................................... 35, 53, 60, 77, 103–111, 223, 237, 398 Differentiation............................................ 23, 51, 54–56, 67–71, 78, 79, 84–89, 104, 160, 301, 338 Difficult-to-transfect ............................................ 310, 312

C Cancer........................................................... 2, 3, 5, 8, 11, 103, 118, 161, 210, 301 Carboxymethyl cellulose................................................. 49 Cardiovascular diseases ............................. 2, 3, 9, 11, 214 Cell cycle....................................................................5, 161 CD34+ stem cells ............................................................ 89 CD4+T cells................................................................... 416 CD8+T cells.......................................................... 371, 425 Clinical isolates .......................21, 22, 26, 29, 31–34, 134 CMV promoter .................................................... 118, 128 Coimmunoprecipitation ............................................... 200 Congenital infections ........................................2, 6, 9, 39, 103, 365–369, 405, 406, 408, 410–414, 428, 437 Cre/LoxP ........................................................................ 28 CRISPR/Cas9 screening..................................... 247–263 Culture of primary fibroblasts ..................................41–50

E Early genes....................................................................... 68 Electron microscopy ............................................ 265–285 Endothelial cells ..................................... 3, 20, 23–26, 30, 31, 33, 34, 45, 55, 79, 83, 160, 162, 180–182, 213, 248, 352, 365 Enzyme-linked immunosorbent assay (ELISA) ........... 75, 188, 234, 291–298, 346, 351, 359, 392 Epithelial cells............................................... 3, 24, 25, 30, 31, 33, 55, 66, 67, 83, 160, 162, 213, 244, 247–263, 365, 433 Extreme limiting dilution analysis (ELDA)................... 97

F Fetal infections ...................................368, 404, 411–414, 436, 441 Fibroblasts .....................................................3, 20–26, 28, 31–34, 39–43, 45, 47, 48, 52, 54, 55, 65–68, 70, 75, 77, 83–86, 89, 95, 96, 98, 115–131, 135, 139, 151, 152, 160–162, 166, 167, 179–182, 184, 192, 200, 208, 209, 213, 214, 217, 222, 235, 237, 248, 252, 253, 261, 263, 270, 274, 297, 316, 338, 352, 355, 365, 374, 377, 407, 419

Andrew D. Yurochko (ed.), Human Cytomegaloviruses: Methods and Protocols, Methods in Molecular Biology, vol. 2244, https://doi.org/10.1007/978-1-0716-1111-1, © Springer Science+Business Media, LLC, part of Springer Nature 2021

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466 Index

AND

PROTOCOLS

Flow cytometry ............................. 55, 56, 62, 63, 65, 70, 71, 80, 85, 87, 88, 95, 168, 311, 349, 350, 355, 357–359, 361, 387, 390–393, 395 FLP/frt................................................................. 142, 175

G Gene expression ......................................... 22, 73, 75, 84, 115, 116, 128, 167, 170, 174, 302, 309, 329 Gibson cloning ............................................ 145, 146, 153 Glioblastomas ............................................................3, 161 Gliomas.............................................................................. 3 Glycoprotein B (gB) ............................................. 57, 224, 413, 414, 423, 427, 431, 432 G-protein coupled receptors ........................................ 159 Gradient purification....................................................... 36 Granulocyte-colony stimulating factor (G-CSF) ......... 52, 60, 89, 96, 344, 346, 348, 356 Green fluorescence protein (GFP) ..........................84–87, 91, 93–98, 156, 222, 248, 253–256, 311, 314, 331, 333, 337 Growth of virus in culture ........................................21–24 Guinea pig CMV (GPCMV) .............................. 173, 366, 367, 427, 428

H Hematopoietic cells...........................................83, 84, 89, 91, 163, 344, 345 Hematopoietic progenitor cells (HPCs).................83–88, 90, 95–97, 104, 161, 162, 180, 312, 314, 333, 344, 346, 351, 352, 354, 360, 366 Hematopoietic stem cells (HSCs)............................86, 87 HiBiT .................................................................... 213–232 High-pressure freezing (HPF) .................. 266, 270, 271, 275, 280, 282 Homologous recombination ....................... 28, 139, 141, 143, 154, 173, 176–178, 219, 220 huBLT .................................................................. 343–362 Human cytomegalovirus (HCMV) antibodies ................................................................ 292 latency ............................................................. 343–362 Humanized mice ..................................... 51, 88, 343–345 Human peripheral blood monocytes ......... 110, 236, 243

I Immediate early gene promoter ................................... 219 Immediate early genes .................................................... 68 Immune control ................................................................ 8 Immune evasions.......................................... 32, 420, 421, 423, 432, 434 Immunofluorescence ................................. 25, 26, 30, 35, 75, 181–184, 188 Immunofluorescence analysis ......................................... 45 Immunohistochemistry ................................................ 392

Immunosuppression ........................................... 409, 413, 418, 424, 438, 440 In vitro ....................................................3, 40, 55, 66, 71, 79, 83, 88, 138, 161, 174, 183, 188, 189, 291, 330, 417 In vivo .....................................................3, 32, 39, 40, 83, 88, 160, 161, 180, 183, 188, 189, 191, 193, 330, 344, 355–356, 360, 371, 393, 396 Inducible..................................................... 115–131, 141, 167, 170 Industrialized countries ................................................ 404 Infection in utero ................................................ 366, 410, 411, 439 Intrapartum transmission ................................................. 6 Isolates .................................................. 19, 21–23, 25, 29, 33, 34, 48, 52, 55, 60, 65, 67, 73, 87, 94, 96, 105, 106, 111, 134, 135, 137–139, 141, 144, 146, 153, 201, 205, 206, 320, 358, 359, 383, 384, 407

K Knockdown .......................................................... 115–131

L Lab adapted strain........................................................... 46 Late genes........................................................... 68, 73, 78 Latency .................................................51, 52, 67, 68, 72, 75, 83–88, 91, 93, 95, 96, 98, 99, 104, 161, 186, 187, 234, 343–362, 372, 381, 383, 425 Latency and reactivation ...................................52, 61, 66, 72, 75–76, 83–99, 161, 346, 359 Lentiviral............................................. 116–125, 128–130, 166–168, 170, 249–252, 332, 333 Leukocytes ................................................ 6, 21, 106, 108, 109, 190, 237, 346, 366, 387, 393 Liver .................................................................2, 6, 9, 160, 190, 193, 344–346, 351–353, 356, 358, 360–362, 370, 371, 380, 384, 388, 393, 396–398 Locked nucleic acids (LNA) ...............309, 330, 338, 339 Luciferase assays ......................................... 173, 193, 303, 309, 329–331, 337, 339 Luciferases .................................................. 164, 171, 173, 214, 309, 329, 330, 338 Lytic replication ............................................................ 180

M Macrophages ............................................ 3, 4, 24, 25, 52, 54, 66, 69–72, 79, 83, 104, 105, 180, 213, 344, 366, 398, 421 mCherry protein ............................................................. 91 Membrane fusion ................................................. 247, 416 Membrane receptors ................................... 200, 207, 247 Microarray analyses .............................................. 174, 336 MicroRNAs (miRNAs) ..................................88, 301–339

HUMAN CYTOMEGALOVIRUSES: METHODS Monocytes .......................................................3, 4, 24, 52, 53, 55, 57, 61, 66, 69–72, 79, 83, 103–111, 161, 180, 200, 233–245, 333, 344, 366, 370, 371, 376, 398, 433 Mononuclear cells ...................................... 53, 57, 60–65, 70, 75, 77, 90, 106–108, 111, 237, 370, 371, 388–390, 393–395, 398, 416 Mononucleosis-like syndrome.......................................... 1 Morphogenesis ..................................................... 265–285 Murine cytomegalovirus (MCMV) .................... 160, 170, 171, 173, 181, 189, 190, 193, 302, 366–368, 370–384, 393, 396, 420–422, 424, 425, 429, 432, 434 Myeloid differentiation ................................56, 71–72, 79 Myeloid progenitors .......................................... 52, 86, 87

N NanoLuciferase (NanoLuc)................................. 214, 215 Neutralizing antibodies ................................29, 413–415, 418, 422–424, 427, 432, 433 NK cells............................................................32, 79, 344, 345, 370, 371, 393, 398, 409, 420–425, 430, 434 Northern blot...................................... 304–306, 316–318 Novel antivirals.............................................................. 437 Nucleoside analogues.................................................... 320 Nucleus ...............................................160, 233, 267–269, 277, 278, 282, 302

O Oligonucleotide primers ............................................... 118 Oncogenesis ...................................................................... 3 Oncomodulatory........................................................... 161 Oropharyngeal secretions ............................................. 404

P Paraffin embedded tissue specimens ...........................375, 384–385 Pathogenesis ..............................................1–11, 103–111, 160, 180, 183, 188, 189, 214, 366, 367, 376–384, 395, 407, 420, 425, 427–430 Pentameric complexes.........................415, 416, 423, 433 Permissive cells ..........................................................34, 48 Phosphoproteome......................................................... 234 Phosphoproteomic screen ................................... 233–245 Photoactivatable ribonucleoside-enhanced crosslinking and immunoprecipitation (PAR CLIP) ..........302, 306–307, 320–321, 324, 325, 332 Plaque assay .................................................. 99, 181, 190, 193, 253, 254, 383 Plasmids ...................................................... 25, 28, 31, 57, 117–121, 123, 124, 128, 129, 135–139, 141, 142, 144–146, 150, 152–154, 165, 166, 168, 170, 173, 176, 179, 189, 193, 202, 204, 205, 215, 217, 247–249, 297, 309–313, 315, 338

AND

PROTOCOLS Index 467

Polymerase chain reaction (PCR) ....................52, 56, 57, 72–76, 121, 123, 126–128, 131, 135–137, 139, 141, 143–147, 149, 150, 153–156, 165, 176–178, 202, 205, 215, 217, 219–223, 227, 229, 231, 232, 249, 258–262, 303, 304, 306, 307, 310, 312, 313, 319, 324, 333, 336, 337, 383, 384, 437 Propidium iodide staining ............................................ 395 Protein-protein interactions ................................ 214, 269 Purification ................................................. 22, 23, 34, 57, 63, 67, 77, 78, 90, 127, 134, 135, 139, 144, 146, 150, 154, 165, 179, 201–207, 217, 218, 223, 231, 249, 302–304, 310, 312, 313, 315, 320, 324, 325, 328, 333, 336

Q qRT-PCR.............................................118, 125–127, 333

R Rat CMV (RCMV) ............................................. 160, 173, 214–231, 302, 367, 375 Real-time polymerase chain reaction (PCR) ..............128, 219, 227, 261, 306, 337, 437 Recombination ............................................. 26, 134, 135, 137–139, 141, 143, 144, 148–149, 152, 154–156, 176–178, 209, 216, 219–222, 230, 231, 431 Reconstitution .......................................... 26, 28, 31, 134, 137, 150, 179, 344, 345, 355, 424 Retinitis................................................................... 1, 9, 10 Retroviral vector...........................................202, 205–206 Resource-poor communities ........................................ 404 Rhesus CMV (RhCMV) ............................ 173, 366, 423, 426, 428, 432–435 RISC-immunoprecipitation (RISC-IP) ......................302, 315, 320, 325, 332, 334, 336, 337 RNA-immunoprecipitation ....................... 302, 308–309, 325–326 Rodent models .................................................3, 367, 424

S Saliva ....................................................404, 408, 422, 423 Salivary glands ............................................ 160, 188–190, 193, 377, 422, 425, 434 SCID..................................................................... 344, 393 SCID-hu mouse model ................................................ 344 Secondary envelopment...................................... 267, 269, 270, 277, 279, 283–285 Semen ................................................................... 404, 406 Sensorineural hearing loss (SNHL) ....................... 6, 367, 371, 407 Short hairpin RNAs (shRNAs)..................................... 116 Signal transduction ..................................... 159, 160, 193

HUMAN CYTOMEGALOVIRUSES: METHODS

468 Index

AND

PROTOCOLS

Smooth muscle cellss ......................................3, 4, 24, 83, 161, 248, 366 Soluble glycoproteins................. 201–202, 207, 209, 423 Species specificity......................................... 214, 343, 366 Spleens ............................................... 160, 190, 193, 356, 358, 361, 371, 381, 383, 384, 387, 389, 393, 395 Stem-loop real-time PCR ........................... 306, 316, 319 Strains ......................................................... 3, 5, 8, 19–36, 40, 44–46, 49, 67, 84, 86, 91, 93–95, 98, 134, 135, 137, 138, 141, 153, 155, 167, 173, 177, 179, 180, 189, 193, 199, 200, 208, 209, 215, 219, 222, 244, 261, 271, 284, 303, 304, 344, 348, 351, 362, 367, 371, 374, 377, 378, 380, 381, 393, 395, 407, 408, 411, 422, 423, 431, 437, 438 Surgical manipulation ................................................... 353

T T-cell response...................................................... 409, 413 Tegument proteins............................................... 269, 427 Thymus ....................................... 344–346, 351, 353, 361 Titers .................................................. 2, 3, 22, 23, 25, 26, 28–33, 39–41, 43–47, 49, 50, 98, 99, 129, 160, 179–182, 190, 191, 230, 253, 254, 314, 333, 355, 359, 371, 377, 379–382, 396, 397, 406, 433 Tomography ............................................... 270–273, 276, 277, 279–281, 284, 285 Transplacental transfer .................................................. 373 Transactivation .................................................................. 9 Transductions ...................................................... 119, 120, 122, 124, 125, 128, 129, 166–168, 170, 171, 250, 252, 261 Transfections ............................................ 28, 31, 34, 116, 119, 121, 124, 128, 129, 134, 137, 139, 150–152, 160, 163, 166–168, 170, 173, 174, 179, 180, 188, 189, 193, 205, 220, 222, 223, 231, 248–250, 309–311, 313, 315, 329–333, 337–339

Transplant recipients .......................................2, 8, 9, 103, 343, 408, 417–419, 432, 437–440 Trophoblasts........................................................... 24, 366 Tropism................................................. 21–27, 30–34, 67, 83, 103, 134, 180, 189, 199, 200, 247

U Urine................................................................21, 26, 404, 408, 422, 423

V Vaccinations .........................................427, 435, 437, 440 Vaginal secretions .......................................................... 404 Vascular diseases .......................................... 3, 4, 103, 367 Viral DNA ..............................................25, 72, 150, 189, 217, 223, 228, 229, 303, 310, 312, 333, 356, 357, 383, 384, 407 Viral entry...................................199, 200, 234, 242, 243 Viral gene expression ................................ 45, 75, 84, 134 Viral genetics ....................................................21, 33, 437 Viral GPCR (vGPCR)......................................... 160–162, 165–168, 171, 173, 174, 181, 185, 187, 189, 192 Viral pathogenesis ................................................. 32, 160, 162, 192, 214 Viral replication ........................................ 9, 34, 104, 115, 116, 160, 180, 181, 183, 191, 192, 267, 359, 371, 414, 420 Viremia ................................................367, 369, 378, 422 Virus containing supernatant ................................ 29, 250 Virus reconstitution ........................................................ 28

W Western blots.............................................. 125, 126, 129, 172, 181, 185, 188, 203, 207, 208, 234, 238, 244, 251, 322, 323, 328, 335–337