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Oral inoculation of chickens with a candidate fowl adenovirus 9 vector and functional studies of fowl adenovirus 9 ORF1 by Li Deng A Thesis presented to The University of Guelph In partial fulfilment of requirements for the degree of Doctor of Philosophy in Pathobiology Guelph, Ontario, Canada © Li Deng, May, 2015 ABSTRACT ORAL INOCULATION OF CHICKENS WITH A CANDIDATE FOWL ADENOVIRUS 9 VECTOR AND FUNCTIONAL STUDIES OF FOWL ADENOVIRUS 9 ORF1 Li Deng University of Guelph, 2015 Advisor: Dr. Éva Nagy FAdV-9Δ4, lacking six open reading frames (ORFs) 0, 1, 1A, 1B, 1C and 2 at the left end of FAdV-9 genome, shows potential as a vaccine vector. In vivo studies demonstrated that FAdV-9Δ4 does not replicate as efficiently as wild type (wt) virus in intramuscularly inoculated chickens, suggesting the important roles of the left-end ORFs. The fecal-oral route is the natural route of FAdV infection, and the oral administration confers several advantages compared to administration through other routes when developing a vaccine. Therefore, the objectives of this study were to investigate the effects of oral inoculation with both FAdV-9Δ4 and wtFAdV-9 in chickens and to explore the functions of one of the left-end ORFs, ORF1. Chickens were orally inoculated with FAdV-9Δ4 or wtFAdV-9. Compared to wtFAdV-9 group, reduced virus shedding in feces, lower viral loads in tissues, and lower antibody response were found in FAdV-9Δ4 group. There were also significant differences between FAdV-9Δ4 and wtFAdV-9 in terms of the induction of cytokine mRNA expression, including interferon α (IFN-α), IFN-γ, and interleukin-12 (IL-12). This indicated the important roles of the six ORFs in modulation of the host immune response. ORF1, a homolog of deoxyuridine 5′-triphosphate pyrophosphatase (dUTPase), was hypothesized to be important in virus replication and regulation of host immune response. In this project, FAdV-9 ORF1 was verified as a functional dUTPase and its molecular features were characterized, including transcription and protein expression patterns and cellular localization. A dUTPase knockout virus (ORF1stop) and its rescued revertant (resORF1) were generated. Functional studies showed that FAdV-9 dUTPase was not required for virus replication in vitro, but played a role in virus replication in vivo. FAdV-9 dUTPase also contributed to the regulation of the expression of IFN-α, IFN-β, and IFN-γ both in vitro and in vivo, and the host antibody response as well. This is the first study to functionally identify an early gene of FAdV-9. The data presented are helpful for better understanding of the molecular biology of FAdVs and for exploring the mechanism of the host immune response against FAdV infections. ACKNOWLEDGEMENTS I wish to express my sincere gratitude and thanks to my dear advisor Dr. Éva Nagy, for her continued guidance and support in all aspects of my research in the past years, and for her kindness and patience throughout my PhD program. I am also very grateful for her financial support. I would also like to sincerely thank Dr. Peter Krell, Dr. Shayan Sharif, and Dr. Ray Lu, who served on my advisory committee and offered valuable advice, suggestions, ideas and comments throughout this study. I am also thankful for Dr. Davor Ojkić and Dr. Byram Bridle, who served on my qualifying examination committee and motivated me to make achievements in science. I would like to thank Bryan Griffin and all other members of the Nagy lab, both past and present, including James Ackford, David Leishman, Betty-Anne McBey, Dr. Yanlong Pei, Dr. Robert Kozak, Dr. Helena Grgić, Dr. Andres Diaz, Dexter Endozo, Javier Hernandez, Nathan Orr, and Dr. Xiaobing Qin. You are all good colleagues and made the Nagy lab a wonderful environment. I am also thankful for Dr. Sarah Wootton and her post-docs and students, including Dr. Jondavid de Jong, Dr. Nicolle Petrik, Dr. Darrick Yu, Dr. Scott Walsh, and Lisa Santry, for all the assistance in the past years. I would also like to thank Leah Read and Tony Cengija for their help during my experimental periods. I would also like to acknowledge China Scholarship Council, University of Guelph and Northwest Agriculture and Forestry University (Yangling, China) for providing me the financial support to conduct my graduate studies. iv Last but never be the least, I am very grateful to my family in China, and my wife Ying (Yvonne) He, for all the love, support, assistance, encouragement and all the other things they have offered to me. v TABLE OF CONTENTS ABSTRACT........................................................................................................................ ii ACKNOWLEDGEMENTS ............................................................................................... iv TABLE OF CONTENTS ................................................................................................... vi LIST OF TABLES ............................................................................................................. ix LIST OF FIGURES ............................................................................................................ x LIST OF ABBREVIATIONS ............................................................................................ xi Chapter 1 ............................................................................................................................. 1 GENERAL INTRODUCTION ................................................................................................ 1 LITERATURE REVIEW ....................................................................................................... 2 1. Adenovirus taxonomy ............................................................................................. 2 2. Virion structure ....................................................................................................... 4 3. Genome organization of mastadenoviruses and early gene functions .................... 7 3.1 The E1A region ................................................................................................. 8 3.2 The E1B region ................................................................................................. 9 3.3 E2 proteins ...................................................................................................... 14 3.4 The E3 region .................................................................................................. 15 3.4 E4 proteins ...................................................................................................... 16 3.5 Intermediate and late genes ............................................................................. 16 4. Molecular biology of aviadenoviruses .................................................................. 17 4.1 Genome organization of aviadenoviruses ....................................................... 17 4.2 Functionally identified early genes in FAdV genome .................................... 19 4.3 Study of FAdV-9 genome ............................................................................... 23 5. Disease associated with aviadenoviruses .............................................................. 24 6. Diagnosis of fowl adenovirus infection ................................................................ 25 7. Immune response to human adenovirus ................................................................ 28 7.1 Innate immune responses ................................................................................ 28 7.2 Adaptive immune responses ........................................................................... 31 8. Immune responses to fowl adenoviruses .............................................................. 31 HYPOTHESES AND OBJECTIVES ...................................................................................... 34 Chapter 2. Oral inoculation of chickens with a candidate fowl adenovirus 9 vector ....... 35 vi ABSTRACT ..................................................................................................................... 36 INTRODUCTION .............................................................................................................. 38 MATERIALS AND METHODS ........................................................................................... 40 Viruses and cells ....................................................................................................... 40 Animal experiment.................................................................................................... 40 Statistical analysis ..................................................................................................... 41 RESULTS ........................................................................................................................ 41 Virus shedding .......................................................................................................... 42 Viral genome copy number in tissues ....................................................................... 42 Antibody response .................................................................................................... 43 Cytokine gene expression in tissues ......................................................................... 43 DISCUSSION ................................................................................................................... 45 ACKNOWLEDGMENTS .................................................................................................... 50 Chapter 3. Characterization and functional studies of fowl adenovirus 9 dUTPase ........ 58 ABSTRACT ..................................................................................................................... 59 INTRODUCTION .............................................................................................................. 60 MATERIALS AND METHODS ........................................................................................... 62 Viruses and cells ....................................................................................................... 62 dUTPase enzyme activity assay ................................................................................ 63 Generation of mutant viruses .................................................................................... 64 RT-PCR and qRT-PCR ............................................................................................. 65 Western blot .............................................................................................................. 66 Immunofluorescence assay ....................................................................................... 66 Statistical analysis ..................................................................................................... 66 RESULTS ........................................................................................................................ 67 Bioinformatics analysis of FAdV-9 ORF1 as a potential dUTPase ......................... 67 FAdV-9 ORF1 has dUTPase enzymatic activity ...................................................... 68 In vitro characteristics of ORF1 mutant viruses ....................................................... 69 Transcription and protein expression profiles of ORF1 ........................................... 69 Cellular localization of ORF1 protein....................................................................... 70 Cytokine gene mRNA expression induced by wild type and mutant viruses ........... 71 DISCUSSION ................................................................................................................... 73 ACKNOWLEDGEMENTS................................................................................................... 77 vii Chapter 4. Fowl adenovirus 9 dUTPase plays a role in virus replication in vivo and in the regulation of the host immune response ........................................................................... 89 ABSTRACT ..................................................................................................................... 90 INTRODUCTION .............................................................................................................. 91 MATERIALS AND METHODS ........................................................................................... 93 Viruses and cells ....................................................................................................... 93 Experimental animals................................................................................................ 93 Animal experimental design ..................................................................................... 93 DNA and RNA extraction ......................................................................................... 94 qPCR and qRT-PCR ................................................................................................. 94 Enzyme-linked immunosorbent assay (ELISA) ....................................................... 95 Statistical analysis ..................................................................................................... 95 RESULTS ........................................................................................................................ 95 Virus shedding .......................................................................................................... 95 Viral loads in tissues ................................................................................................. 96 Antibody response .................................................................................................... 97 Cytokine gene expression in tissues ......................................................................... 97 DISCUSSION ................................................................................................................. 100 ACKNOWLEDGEMENT .................................................................................................. 103 Chapter 5. General discussion......................................................................................... 111 References ....................................................................................................................... 119 viii LIST OF TABLES TABLE 1.1 Summary of 13 adenoviral proteins and their functions ................................. 6 TABLE 1.2 Adenovirus gene products and their functions .............................................. 11 TABLE 2.1 Virus titers (pfu/ml) in the feces of chickens orally inoculated with FAdV9Δ4 and wtFAdV-9. .................................................................................................. 51 TABLE 2.2 Viral genome copy number in tissues ........................................................... 52 TABLE 3.1 Primers for generating mutant viruses ......................................................... 78 TABLE 3.2 Primers used for cytokines for qRT-PCR .................................................... 79 SUPPLEMENTARY TABLE S1 Pairwise identities of dUTPase amino acid sequences of adenoviruses. ............................................................................................................. 80 TABLE 4.1 Virus titers in the feces of infected chickens .............................................. 105 TABLE 4.2 Viral genome copy number in tissues ......................................................... 106 ix LIST OF FIGURES FIGURE 1.1 Phylogenetic tree of adenovirus genomes. .................................................... 3 FIGURE 1.2 Structure of adenovirus.................................................................................. 5 FIGURE 1.3 Transcription map of the mastadenovirus genome. ..................................... 10 FIGURE 1.4 ORFs in the left end of FAdV-9 genome. ................................................... 27 FIGURE 2.1 Antibody response in orally inoculated chickens ........................................ 53 FIGURE 2.2 Cytokine mRNA expression in spleen samples........................................... 54 FIGURE 2.3 Cytokine mRNA expression in liver samples ............................................. 55 FIGURE 2.4 Cytokine mRNA expression in bursa of Fabricius samples ........................ 56 FIGURE 2.5 Cytokine mRNA expression in cecal tonsil samples ................................... 57 SUPPLEMENTARY FIGURE S1 Generation of the mutant viruses. ............................. 81 FIGURE 3.1 Multiple alignment of amino acid sequences of dUTPase homologs. ........ 82 FIGURE 3.2 PCR-based dUTPase enzyme activity assay. .............................................. 83 FIGURE 3.3 Viral DNA accumulation curve and one-step growth curve. ...................... 84 FIGURE 3.4 Transcription and protein expression profiles of ORF1. ............................. 85 FIGURE 3.5 Cellular localization of ORF1 protein ......................................................... 86 FIGURE 3.6 Cytokine mRNA expressions in CH-SAH cells .......................................... 87 FIGURE 3.7 Cytokine mRNA expressions in Celi cells .................................................. 88 FIGURE 4.1 FAdV-specific IgG antibody response in chickens. .................................. 107 FIGURE 4.2 Cytokine mRNA expression in spleen samples of chickens. .................... 108 FIGURE 4.3 Cytokine mRNA expression in liver samples of chickens. ....................... 109 FIGURE 4.4 Cytokine mRNA expression in cecal tonsil samples of chickens. ............ 110 x LIST OF ABBREVIATIONS aa Ab Ad ADP AdV AGPT ANOVA APCs BAdV bp CAV CBP Celi CFIA CH-SAH CPE CTLs DBP d.p.i. DCs DMEM-F12 dsDNA dUTPase E. coli E1A EDSV EDTA EGFP ELISA ER FAdV FBS GAM-1 GoAdV gp19k h.p.i. HAdV HAPI HAT HDAC1 HHV HI amino acid antibody adenoviral adenovirus death protein adenovirus agar gel precipitation test analysis of variance antigen presenting cells bovine adenovirus base pair chicken anemia virus CREB-binding protein chicken embryo liver Canadian Food Inspection Agency chicken hepatoma cytopathic effect cytotoxic T lymphocytes DNA binding protein days post-inoculation dendritic cells Dulbecco's modified Eagle's medium and nutrient mixture Ham's F-12 medium double-strand DNA deoxyuridine 5′-triphosphate pyrophosphatase Escherichia coli early region 1A egg drop syndrome virus ethylenediamine tetraacetic acid enhanced green fluorescence protein enzyme-linked immunosorbent assay endoplasmic reticulum fowl adenovirus fetal bovine serum Gallus anti morte protein 1 goose adenovirus glycoprotein 19k hours post-infection human adenovirus highly pathogenic avian influenza histone-acetyltransferase histone deacetylase 1 human herpesvirus hemagglutination inhibition xi hMDM HPS hsp40 HSV-1 i.m. IBDV IBH IBV IFN Ig IL IRF ITR kDa mDCs MHC-I MHV-68 ml MLP MLTU MOI MTOC NBs NF NF-κB NK nt ORF PAdV PAMP PBMCs PBS PCR pDCs pfu PHA-P PKR PML pol PPi pRb PRRs pTP RFLP RNA RT human monocyte-derived macrophage hydropericardium syndrome heat-shock proteins 40 herpes simplex virus type 1 intramuscularly infectious bursal disease virus inclusion body hepatitis infectious bronchitis virus interferon immunoglobulin interleukin interferon regulatory factor inverted terminal repeat kilodalton myeloid DCs major histocompatability complex class I murine gammaherpesvirus 68 mililiter major late promoter major late transcription unit multiplicity of infection microtubule organizing center nuclear bodies nuclear factor nuclear factor kappa B natural killer nucleotide open reading frame porcine adenovirus pathogen-associated molecular pattern peripheral blood mononuclear cells phosphate buffered saline polymerase chain reaction plasmacytoid dendritic cells plaque forming unit phytohemagglutinin-P protein kinase R promyelocytic leukemia polymerase pyrophosphate retinoblastoma protein pattern recognition receptors precursor terminal protein restriction fragment length polymorphisms ribonucleic acid reverse transcription xii SAdV SEM SPF SRBC ssDNA TAdV Th1 TLRs TNF TP TPL TR TSA TSAdV VA-RNAs VZV wt μl simian adenovirus standard error of the mean specific-pathogen-free sheep red blood cells single-strand DNA turkey adenovirus T helper 1 toll-like receptors tumor necrosis factor terminal protein tripartite leader sequence tandem repeat region trichostatin A tree shrew adenovirus virus-associated RNAs varicella zoster virus wild type microliter xiii Chapter 1 General introduction Adenoviruses (AdVs) have been extensively developed as vaccine vectors. Numerous studies have proven that adenoviral (Ad) vectors meet the most important criteria of an ideal vaccine vector in terms of efficacy, safety and stability (Tatsis and Ertl, 2004). Compared to other viral vectors, Ad vectors offer a number of advantages: a broad range of natural host, infection of a variety of both dividing and quiescent cell types, robust transgene expression, easily grown to high titers, easy manipulation, lack of serious threat of horizontal transmission, induction of potent innate and adaptive immune responses, and transduction of antigen presenting cells (Ahi et al., 2011; Bangari and Mittal, 2006). Fowl adenoviruses (FAdVs) can be isolated from both sick and healthy birds in poultry farms and have a worldwide distribution (Hess, 2013). In recent years, a number of researchers have focussed on developing FAdVs, such as FAdV-1, FAdV-8, FAdV-9 and FAdV-10, as vaccine vectors and have shown promising results (Francois et al., 2004; Sheppard et al., 1998; Johnson et al., 2003; Corredor and Nagy, 2010b; 2011; Ojkić and Nagy, 2001). However, compared to the extensive studies in human adenoviruses (HAdVs), the study of the molecular biology of FAdVs is far behind. The genome of FAdV-9 has been fully sequenced (Ojkić and Nagy, 2000). Based on this, a deleted virus FAdV-9Δ4, lacking six open reading frames (ORFs), was generated in our laboratory and has shown potential as a vaccine vector (Corredor and Nagy, 2010b; 2011; Ojkić and Nagy, 2001). However, to better understand the molecular biology of FAdV-9, research on exploring the functions of viral genes is required. 1 Literature review 1. Adenovirus taxonomy Adenoviruses, first isolated as respiratory pathogens and characterized in 1953 (Rowe et al., 1953), belong to the family Adenoviridae and are classified into five genera (Harrach et al., 2011): Mastadenovirus, isolated from mammals; Aviadenovirus, isolated from birds; Atadenovirus, isolated from mammals, birds and reptiles and named due to their high adenine and thymine contents in the genomes; Siadenovirus, isolated from a frog and birds and named because they encode a putative sialidase (Davison and Harrach, 2002); and Ichtadenovirus, isolated from a sturgeon (Berk, 2013; Harrach et al., 2011). Specifically, each genus has one or more species (Fig. 1.1). To date, 56 human adenovirus (HAdVs) serotypes have been identified and classified into 7 species, Human adenovirus A to Human adenovirus G, based on serology, genome sequencing, phylogenetic distance, hemagglutination, and oncogenicity in rodents (Berk, 2013). Other species in the genus Mastadenovirus include Murine adenovirus A to Murine adenovirus C, Ovine adenovirus A, Ovine adenovirus B, Porcine adenovirus A to Porcine adenovirus C, Simian adenovirus A, and Tree shrew adenovirus A (Harrach et al., 2011). There are 12 fowl adenovirus (FAdV) serotypes. They are grouped into 5 species, Fowl adenovirus A to Fowl adenovirus E, on the basis of phylogeny, genome organization, restriction fragment length polymorphism (RFLP) profiles and the lack of significant cross-neutralization (Zsak and Kisary, 1984; Harrach et al., 2011). Other species in the genus Aviadenovirus are Falcon adenovirus A, Goose adenovirus A, and Turkey adenovirus B. Recently, several potential species have been proposed to belong to the 2 Figure 1. 1 Phylogenetic tree of adenovirus genomes. Adenoviruses, belonging to family Adenoviridae, are classified into five genera, Mastadenovirus, Aviadenovirus, Atadenovirus, Siadenovirus, and Ichtadenovirus. Each genus has one or more species. Taken from Harrach et al., 2011. 3 genus Aviadenovirus but have not been approved yet, including Pigeon aviadenovirus A, Duck aviadenovirus B, Turkey adenovirus C and Turkey adenovirus D (Marek et al., 2014a, 2014b). Atadenovirus includes Ovine adenovirus D, Duck adenovirus A (egg drop syndrome virus, EDSV), Bovine adenovirus D, Possum adenovirus and Snake adenovirus A. Viruses in the genus Siadenovirus have a putative sialidase gene present at the left end of the genome and include three species: Frog adenovirus (Davison et al., 2000), Raptor adenovirus A and Turkey adenovirus A. Ichtadenovirus has been newly established and contains one single speices Sturgeon adenovirus A (Doszpoly et al., 2009), which has been found in white sturgeon (Benkö et al., 2002). 2. Virion structure Adenoviruses are non-enveloped, icosahedral particles with a diameter from 70 to 100 nm. Each virion consists of a protein shell called a capsid that surrounds a viral core containing the DNA genome. The capsid has two roles: it protects the core genome and mediates the entry of the virus into cells. There are at least 13 proteins in the adenovirus virion (Fig. 1.2, Table 1.1), including II-IX, IIIa, Iva2, terminal protein (TP), μ, and p23, the viral protease (Russell, 2009). They are all structural proteins encoded by late regions L1 to L5 of the viral DNA genome. Historically, these proteins were numbered (II–IX) in order due to their molecular weight (smallest to greatest) when they were first identified (Vellinga et al., 2005). The capsid is composed of 252 subunits called capsomeres, which are collectively made up of 240 hexons and 12 pentons. Each capsid consists of at least 7 proteins, three of 4 Figure 1. 2 Structure of Adenovirus Taken from Russell, 2009 5 Table 1.1 Summary of 13 adenoviral proteins and their functions Name Location Known Functions II Hexon monomer Structural III Penton base Penetration IIIa Associated with penton base Penetration IV Fiber Receptor binding; haemagglutination V Core: associated with DNA and penton base Histone-like; packaging? VI Hexon minor polypeptide Stabilization/assembly of particle? VII Core Histone-like VIII Hexon minor polypeptide Stabilization/assembly of particle? IX Hexon minor polypeptide Stabilization/assembly of particle? TP Genome - Terminal Protein Genome replication μ Nucleoprotein Genome replication? IVa2 Nucleoprotein Genome packaging Protease Associated with pentons? Maturation Taken from Russell, 2009 6 which are termed major proteins, hexon (protein II), penton base (pIII) and fiber (pIV); and the other four are termed minor capsid proteins, IIIa, VI, VIII and IX (Vellinga et al., 2005). The hexon protein comprises the majority of the outer shell of the capsid, which is made up of 240 homotrimers that encapsidate most parts of the virus, including the viral genome and associated proteins. The trimeric fiber projects from each of the 12 vertices of the icosahedron, while the penton base lies at the base of each fiber. These three capsid proteins contribute to the majority of activities required for the early stages of Ad infection (Russell, 2009; Medina-Kauwe, 2013; Berk, 2013). The remaining six structural components are situated in the virus core, five of which are associated with the double-stranded DNA genome (V, VII, μ, IVa2 and TP), while the 23 K virion protease plays a vital role in the assembly of the virion. The TP is covalently attached to the 5’ ends of the adenoviral DNA, circularising it. Newly synthesized TP acts as a primer for DNA replication in the cell nucleus (Berk, 2013; Russell, 2009). 3. Genome organization of mastadenoviruses and early gene functions Adenoviruses have a linear, double-stranded DNA (dsDNA) genome with a size range of 26-48 kb (Harrach et al., 2011). At the 5’ end of each strand, a terminal protein is covalently linked which functions as a protein primer at the onset of viral DNA replication. The genomes of all adenoviruses have inverted terminal repeat (ITR) sequences, ranging in size from 36 to over 200 base pairs (bp), which function as origins of DNA replication at each end of the viral genome. DNA sequences, which encode most virion structural proteins and proteins involved in viral DNA replication and virion assembly, are well conserved in all adenovirus genomes. In contrast, DNA sequences encoding regulatory proteins, usually located at both ends of the genome, are not 7 conserved among the genera (Berk, 2013). According to the time of expression during the viral life cycle, human adenovirus genomes contain 5 early transcription units (E1A, E1B, E2, E3 and E4), 4 intermediate transcription units at the onset of viral DNA replication (IX, IVa2, L4 intermediate, and E2 late), and one late transcription unit (major late) that is processed to generate 5 families of late mRNAs (L1-L5) (Fig. 1.3, Table 1.2). All of these transcripts are transcribed by RNA polymerase II. In addition, the human adenovirus genome also carries one or two small virus-associated RNA (VA-RNA), which varies in the serotypes. The VA-RNA is transcribed by RNA polymerase III. The early genes are involved in regulation and virus replication (E1); viral DNA replication (E2); escape from the immune system (E3) and cell cycle control (E4). The late genes are transcribed later than the early genes and encode the structural proteins described above. 3.1 The E1A region During adenovirus infection, E1A is the first viral transcription unit to be transcribed once the viral DNA reaches the nucleus. It is essential for adenovirus mediated cell transformation (Mymryk et al., 1994; Gallimore and Turnell, 2001; Zhang et al., 2004). E1A has been extensively studied in adenovirus serotype 5, and most of the literature relate to this serotype. The E1A gene encodes, using alternative splicing, two mRNAs during the early phase of the infection and three other mRNAs (11S, 10S and 9S) during the late phase. The two early E1A proteins are called 13S and 12S, which are composed of 298 and 243 amino acids (aa), respectively. The 13S has a 46 aa internal region that is not found in the 12S form. Based on sequence comparison of different serotypes, these two proteins both contain two conserved regions, CR1 and CR2, while 13S contains the 8 CR3 internal 46 aa region. The various functions of E1A are mediated mainly through interactions with many key cellular proteins via these conserved regions. The CR1 conserved region induces host cell cycling through interaction with many chromatin-remodelling proteins for gene activation and repression. E1A interacts with p300 and its paralog CREB-binding protein (CBP), which are co-activators for transcription. It has been reported that E1A can inhibit the histone-acetyltransferase (HAT) activity of p300 (Chakravarti et al., 1999; Hamamori et al., 1999) and stimulate transcription through p300/CBP. Other E1A-CR1 interacting HAT complexes include PCAF (Lang and Hearing, 2003), p400, a member of the SWI/SNF family (Fuchs et al., 2001), and TRAPP, a component of HAT complexes (Deleu et al., 2001; Nikiforov et al., 2002). However, the exact mechanism for how chromosome remodelling induces cell cycling by E1A is still unclear. E1A CR3 region is involved in the activation of other early viral transcription units. Its transcription activation function is dependent on the interaction of E1A-CR3 with MED23 (formally called SUR-2), a component of the Mediator complex (Ablack et al., 2010). Cell transformation induced by expression of E1A alone is unstable. Creation of stable transformed cells requires both E1A and E1B products. E1A induces the activation of p53 and thus E1A-transformed cells die rapidly from apoptosis (Debbas and White, 1993; Grand et al., 1994). It has been demonstrated that the function of both E1B products in transformation is to block p53-activated apoptosis in order to establish stably transformed cells (Grand et al., 1994). 3.2 The E1B region The E1B transcription unit encodes five products by alternative splicing. The two major 9 polypeptides E1B19K and E1B55K are transcribed using two different ORFs. In addition, Figure 1.3 Transcription map of the mastadenovirus genome. The mastadenovirus genome is organized in early (red), intermediate (blue) and late (green) transcriptional units. Taken from Weitzman, 2005. 10 Table 1.2 Adenovirus gene products and their functions Phase Gene E1A E1B E2A Early E2B E3 E4 IVa2 IX Intermediate VAI VAII Late L1-L5 Products and their function Inactivates pRB to release E2F - cell cycle deregulation Transactivates viral promoters 55K targets p53 and participate in transport of late viral mRNA 19K is a Bcl-2 homologue - anti-apoptotic Preterminal protein (pTP) and DNA polymerase (pol) - DNA replication Single-strand DNA binding protein (DBP) - DNA replication gp19K inhibits MHC I expression 10.4K/14.5K (RID complex) inhibits tumor necrosis factor (TNF) apoptosis, internalizes TNF receptor and degrades Fas ligand 14.7K inhibits TNF apoptosis, stabilized NFκB 11.6K (ADP) induces cell lysis Products: orf1, orf2, orf3, orf4, orf6 and orf6/7 modulate viral mRNA metabolism, promote virus DNA replication and block host protein synthesis Initiate the major late promoter (MLP), which regulates late genes Non-coding RNA that stimulates translation of viral genes and blocks double stranded RNA activated protein kinase R (PKR) during interferon response Non-coding RNA that blocks PKR during interferon response Structural proteins: L1 (IIIa); L2 (penton base, V, VII); L3 (hexon, VI, virus protease); L4 (VIII); L5 (fiber) Taken from Weitzman, 2005. 11 three other minor products E1B-156R, E1B-93R and E1B-84R are transcribed from the ORF for E1B55K, (Virtanen and Pettersson, 1985). It is well known that the major function of the E1B proteins, at least when functioning on their own, is to counteract E1A-induced apoptosis in order to prevent premature lysis of the host cell, thus ensuring an efficient viral replication (Berk, 2013). E1B19K protein prevents p53 independent apoptosis. Upon activation of p53, the proapoptotic BCL-2 family member BAK forms oligomers with another proapoptotic protein BAX to form pore-like structures in the mitochondrial membrane. This process leads to the release of cytochrome C and Smac/DIABOLO which induce the caspasedependent pathway, resulting in apoptosis. This effect can be suppressed by high levels of expression of BCL-2 in uninfected cells. E1B19K protein is a viral mimic of BCL-2, which sequesters BAK, thus preventing its association with BAX to prevent apoptosis. E1B19K alone is sufficient to support E1A-mediated cell transformation; however, E1B55K is also required for viral replication. E1B55K is involved in many functions during both early and late phases of mastadenovirus infection, most of which are dependent on the formation and the activity of the E4orf6/E1B55K E3 ligase complex. The most studied and key early function of E1B55K is to counteract p53-induced cell cycle arrest and early apoptosis which is caused by the interaction of E1A protein interaction with either retinoblastoma protein (pRB) family proteins or p300/CBP/p400, resulting in activation and accumulation of p53 (Chiou and White, 1997). The p53 is a well-known tumour suppressor that functions mainly as a sequence specific transcription factor, activating or repressing transcription and leading to ultimate cell cycle arrest or apoptosis (Woods and Vousden, 2001). 12 E1B55K antagonises the effect of p53 by at least two separate mechanisms: proteasomal degradation of p53 by the E4orf6/E1B55K E3 ligase and E1B55K-mediated inhibition of p53 transcriptional activity. The localization of E1B55K is largely dependent on two other adenoviral proteins, E4orf3 and E4orf6. In the context of expression of E1B55K alone, HAdV-5 E1B55K adopts a predominantly cytoplasmic localization, exhibiting in immunofluorescence studies bright staining of perinuclear structures generally believed to be aggresomes (Goodrum et al., 1996). Aggresomes are perinuclear bodies formed at the microtubule organizing center (MTOC) in response to accumulation of misfolded and aggregated proteins. E4orf6 is localized in the nucleus when expressed alone. When E1B55K is coexpressed with E4orf6, E1B55K co-localizes with E4orf6 in the nucleus and the aggresome structures remain unaffected (Goodrum et al., 1996). E4orf3 can also bind to E1B55K. E4orf3 has been found to modify promyelocytic leukemia (PML) nuclear bodies (NBs) into track-like structures in the nucleus. Co-expression of these two proteins results in the translocation of E1B55K into the PML NBs (Leppard and Everett, 1999). The interactions of E1B55K with E4orf6 and E4orf3 are mutually exclusive but when these three proteins are co-expressed, E1B55K co-localizes with E4orf6 (Konig et al., 1999). In the context of infection, E1B55K is re-localized by E4orf6 mainly to nuclear viral replication centers where DNA replication and subsequent transcription take place. E1B55K is also found in the nuclear filamentous structures of PML and the aggresomes (Ornelles and Shenk, 1991). E4orf6 and E1B55K nuclear co-localization requires a primate-specific factor, RUNX1 (Marshall et al., 2008). Apart from the direct effects described above on p53, E1B55K is believed to inactivate 13 p53 by sequestration to the perinuclear aggresome structures (Zantema et al., 1985). During infection, as E4orf3 is believed to be expressed earlier than E4orf6, E1B55K is first observed to associate with PML alone with p53 (Konig et al., 1999). 3.3 E2 proteins The E2 transcription unit encodes three proteins, all of which are directly involved in DNA replication. From the same promoter, two different transcripts, E2A and E2B, are produced by alternative splicing. E2A transcripts encode a DNA binding protein (DBP) and E2B encodes both the adenovirus DNA polymerase (pol) and the precursor terminal protein (pTP). E2A expression is mediated by the E1A region. Viral DNA replication requires all three E2 proteins, plus two cellular proteins, nuclear factor I (NF-I) and nuclear factor III (NF-III) (Challberg and Kelly, 1981). DBP is the first non-structural adenoviral protein identified due to its abundance in infected cells (van der Vliet and Levine, 1973). It is a 72 kDa protein that binds to singlestrand DNA (ssDNA) with high affinity and to dsDNA and RNA with lower affinity. It has many roles in both the initiation and elongation steps of viral DNA replication, especially to unwind DNA templates. Ad pol is a 140 kDa viral DNA polymerase, a major component of the initiation complex of viral DNA replication. Unlike cellular polymerase, it possesses 5’ to 3’ polymerase activity as well as the 3’ to 5’ exonuclease activity. The pTP is initially produced as an 80 kDa protein, which is responsible for protein priming for the initiation of DNA replication. At the late phase of infection, pTP is processed by proteolysis into a smaller fragment of 55 kDa during the assembly of virions involving the adenovirus protease. TP acts as a primer for viral DNA replication and hence ends up being covalently attached to the 5’ termini of each end of the viral 14 DNA (Challberg and Kelly, 1981). 3.4 The E3 region The E3 region, which is dispensable for the replication of adenovirus, encodes a range of proteins that subvert the host immune response. The E3 promoter contains nuclear factor kappa B (NF-κB) -binding sites that can be induced by cytokines such as tumour necrosis factors (TNFs) (Deryckere et al., 1995). The adenovirus E3-19K glycoprotein (gp19k) is localized in the membrane of the endoplasmic reticulum (ER), where it binds to major histocompatability complex (MHC) class I antigens, preventing their export from the ER to the cell surface. It is the presence of a lysine based motif (KKXX) that is responsible for retention in the ER. The E3-10.4K/14.5K complex (RID α/β) also inhibits TNF-α and Fas ligand-induced cell death by internalising their receptors (TNFR1, Fas and TNF-related apoptosis-inducing ligand) and promoting their degradation in lysosomes (Elsing and Burgert, 1998; Tollefson et al., 1998). RID α/β localize to the plasma membrane, Golgi, ER and vesicles within the cell (Tollefson et al., 1998). The E3-14.7K protein inhibits apoptosis through the TNFR, Fas and TRAIL pathways, with the effect on Fas being less pronounced. The E3-14.7 kDa protein is localized in the cytosol and nucleus (Li et al., 1999) and functions by binding to cellular proteins that mediate apoptosis including NEMO/IKK-γ. Apoptosis induced by transfecting cells with NEMO/IKK-γ, is reversed by 70% in the presence of E3-14.7K protein (Li et al., 1999). The E3-11.6K, adenovirus death protein (ADP), is the only E3 protein not involved in subverting the host cell immune response. It is produced late in infection and induces cell 15 death, resulting in the release of progeny virus from the cell. 3.4 E4 proteins Transcripts from the E4 region are subject to alternate splicing events, leading to the production of approximately 18 distinct mRNAs, which are predicted to encode 7 different proteins named Orf1, Orf2, Orf3, Orf4, Orf3/4, Orf6, Orf6/7 (Virtanen et al., 1985). However, one of these polypeptides (E4orf3/4) has never been detected in infected cell. The E4 region has a similar genomic organisation throughout all serotypes, except that E4orf1 is not conserved in HAd40 (Davison et al., 2003). The E4 region is required for efficient viral DNA replication, late gene expression and shutoff of the synthesis of host cell proteins. This is confirmed by Halbert et al. (1985) through generating mutant HAdV-5 viruses carrying defined lesions in the E4 region. A mutant virus dl366, lacking the majority of the E4 region, is severely defective and can only be propagated in stable Vero cells expressing the E4 region (Halbert et al., 1985). Another mutant virus dl355, lacking 14 bp within the segment encoding the E4Orf6 protein, shows a delayed onset of viral DNA synthesis. Expression of late viral proteins is reduced in both dl355 and dl366 viruses, although more severely in dl366. Shutoff of the host protein synthesis is also less efficient with the mutant viruses (Halbert et al., 1985). 3.5 Intermediate and late genes After the onset of virus DNA replication, the intermediate transcription units IVa2 and IX genes are expressed at high levels, activating transcription of the MLP and resulting in expression of the late genes (Lutz et al., 1997; Tribouley et al., 1994). The major late transcription unit (MLTU) encodes multiple proteins from five regions, 16 L1 to L5, by differential splicing and polyadenylation. It is the primary transcript that encodes most of the structural proteins and some non-structural proteins needed for the assembly of progeny virions. All late mRNAs contain a 5’ tripartite leader sequence (TPL), which is important for their nuclear export, translation and stability. The major structural proteins produced by MLTU include L2-III (penton), L3-II (hexon) and L5-IV (fiber), and non-structural proteins include L1-52/55K, L4-22K, L4-33K and L4-100K. Expression from the MLTU is temporally regulated. During the early phase of infection, the MLP is active at low levels and transcription does not proceed to the end of the MLTU. This results in the production of one major product, the L1-52/55K protein. After viral DNA replication, the MLP becomes fully activated and its activation depends on many known activators including the E1A protein, the product of the intermediate transcription unit IVa2 and transcription factors MAZ and Sp1. Viral DNA replication is also required prior to MLP activation, suggesting that a cis-acting type of regulation is involved (Berk, 2013). 4. Molecular biology of aviadenoviruses 4.1 Genome organization of aviadenoviruses Compared to mastadenoviruses, the molecular biology of aviadenoviruses is less studied. The genome size of aviadenoviruses is considerably larger compared to that of mastadenoviruses, representing the largest adenovirus DNA after that of white sturgeon adenovirus (Harrach et al., 2011). To date, the complete genome sequences have been determined for members of all FAdV species: Fowl adenovirus A (FAdV-1, Chiocca et al., 1996), Fowl adenovirus B (FAdV-5, Marek et al., 2013), Fowl adenovirus C (FAdV4, Griffin and Nagy, 2011; Marek et al., 2012), Fowl adenovirus D (FAdV-9, Ojkić and 17 Nagy, 2000) and Fowl adenovirus E (FAdV-8, Grgić et al., 2011). In addition to FAdVs, the genomes of two other aviadenovirus species have been fully sequenced: Turkey adenovirus B (TAdV-1, Kaján et al., 2010) and Goose adenovirus A (GoAdV-4, Kaján et al., 2012). Recently, more complete genomes of several other avian adenoviruses have become available, including pigeon adenovirus 1 (PiAdV-1), duck adenovirus 2 (DAdV2), TAdV-4 and TAdV-5 (Marek et al., 2014a, 2014b). They represent the potential species Pigeon aviadenovirus A, Duck aviadenovirus B, Turkey adenovirus C and Turkey adenovirus D, respectively. All of these are suggested to belong to the genus Aviadenovirus but have not been approved yet by the international committee on taxonomy of viruses (Marek et al., 2014a, 2014b). The organization of the central part of the aviadenovirus genomes (from IVa2 to fiber gene) is similar to that of mastadenoviruses, including E2 proteins, IVa2, fiber, hexon and other late proteins. However, no regions homologous to the E1, E3 and E4 of mastadenoviruses are recognized in aviadenovirus genomes. Despite the lack of genetic similarity to other genera, the ends of the aviadenovirus genomes are termed E1 and E4 (Davison et al., 2003; Marek et al., 2013). In the left-end region (E1 region) of aviadenovirus genomes, there are few variations in ORF constitution. The E4 region, larger in aviadenoviruses than in members of other genera, contains several transcription units that are unique for aviadenoviruses. ORFs 22, 20A, 20, 19, 8 (GAM-1, for Gallus anti morte), and 17 of this region seem to be well conserved in all currently sequenced aviadenoviruses except that ORF17 is absent in GoAdV-4 genome (Kaján et al., 2012). On the other hand, the order and orientation of shared ORFs are also well conserved among different FAdVs (Marek et al., 2013). 18 The fiber of FAdVs has been implicated to play an important role in the infectivity and pathogenicity of FAdVs (Pallister et al., 1996; Grgić et al., 2014; Schachner et al., 2014). Contrary to most mastadenoviruses, all FAdVs that have been examined have two fibers protruding from each penton base (Gelderblom and Maichle-Lauppe, 1982). Two fiber genes are identified in members of Fowl adenovirus A and Fowl adenovirus C (Chiocca et al., 1996; Griffin and Nagy, 2011; Marek et al., 2012), while only a single fiber gene is found in sequenced members of Fowl adenovirus B, Fowl adenovirus D and Fowl adenovirus E (Grgić et al., 2011; Marek et al., 2013; Ojkić and Nagy, 2000). Although it is still unknown how many fibers per penton base they possess, two fiber genes are identified in the genomes of TAdV-1, TAdV-5, GoAdV-4 and PiAdV-1 (Kaján et al., 2010, 2012; Marek et al., 2014a, 2014b), whereas one fiber gene was found in the genome of TAdV-4 and DAdV-2 (Marek et al., 2014a, 2014b). Nevertheless, it is predicted that, based on the phylogenetic analysis of aviadenoviruses, the ancestor strain of all current aviadenoviruses probably had two fiber genes, of which fiber-1 was lost in the branch leading to FAdV-B, FAdV-D, FAdV-E and TAdV-C (Marek et al., 2014a). 4.2 Functionally identified early genes in FAdV genome At least one complete genome from members of all FAdV species has been sequenced, and the FAdV genomes encode numerous genes. However, unlike the early genes of human adenoviruses that are extensively defined, the majority of early genes of FAdVs have not yet been characterized in detail. To date, there are only 3 early genes that have been characterized beyond their sequences for FAdVs: ORF8 (GAM-1), ORF22, and ORF1 (dUTPase). In addition, yet not characterized, some genes are found to show homology with proteins of other viruses, for example, ORF2, a homolog of nonstructural 19 protein NS1 (Rep protein) of parvoviruses, and ORF19, a homolog of lipase of Marek's disease virus (Corredor et al., 2006, 2008; Washietl and Eisenhaber, 2003). ORF8 ORF8, also named GAM-1, is a conserved early gene located at the right-end of the genomes of all FAdVs. Encoding a 31 kDa nuclear protein, GAM-1 was originally discovered upon screening for viral proteins that regulate cellular apoptosis (Chiocca et al., 1997). Though sharing no sequence homology, it mimics the functions of Bcl-2 and E1B-19K of human adenoviruses and blocks apoptosis (Chiocca et al., 1997). Glotzer et al. (2000) have demonstrated that GAM-1 of FAdV-1 is required for virus replication. They showed that the GAM-1 protein increases the cellular levels of heatshock proteins 40 (hsp40) and hsp70 and relocates these proteins to the nucleus. GAM-1 activates host heat-shock responses with hsp40. Heat shock or overexpression of hsp40 can partially replace the roles of GAM-1 in viral replication. GAM-1 influences the expression of cellular genes. Chiocca and co-workers (2002) have shown that expression of GAM-1 increases the level of transcription from a variety of eukaryotic promoters, probably by effectively inactivating histone deacetylase 1 (HDAC1) both in vitro and in vivo. They also demonstrated that a FAdV-1 lacking GAM-1 is replication defective, but the defect can be overcome by either expressing an interfering HDAC1 mutant or by treating infected cells with HDAC inhibitor trichostatin A (TSA). Further evidence showed that GAM-1 interferes with SUMOylation of HDAC1, through destroying promyelocytic leukemia nuclear bodies and delocalizing SUMO-1 into the cytoplasm (Colombo et al., 2002). The SUMOylation inhibitory function of the purified GAM-1 protein has been recently confirmed in an in vitro assay (Avila et al., 2015). 20 Hacker et al. (2005) also demonstrated that the FAdV-1 GAM-1 protein enhances transient and stable recombinant protein expression in Chinese hamster ovary cells. Everett et al. (2014) showed that GAM-1 increases the infection efficiency of a regulatory protein ICP0-deficient mutant of herpes simplex virus 1 (HSV-1). ORF22 ORF22 of FAdV-1 is identified as an early transcription product that is detectable from 2 hours post-infection (h.p.i.) (Payet et al., 1998). The ORF22 protein is expressed as early as 6 h p.i. and is still accumulating at 30 h.p.i. (Lehrmann and Cotten, 1999). In terms of its function, ORF22 is found to interact with the pRb to activate the E2F pathway, in cooperation with GAM-1 (Lehrmann and Cotten, 1999). ORF22 binds to the pocket domain of pRb, similar to other DNA tumor virus proteins, while GAM-1 interacts with pRb regions outside the pocket domain. ORF1 Localized in the left end of FAdV genomes, ORF1 is a homolog of deoxyuridine 5′triphosphate pyrophosphatase (dUTPase). Corredor et al. (2006) showed that ORF1 is well conserved in all genomes of FAdVs, with an amino acid identity ranging from 56 to 100% among FAdV genomes. Cao et al. (1998) showed that FAdV-9 ORF1 (formerly named ORF LTR1) is transcribed at 2 h.p.i., while no data are available regarding its protein expression. ORF1 of FAdV-1 has been demonstrated to possess dUTPase enzyma activity (Weiss et al., 1997); however, its function in the virus life cycle is unknown. dUTPase is a ubiquitous enzyme which exists widely in eukaryotic and prokaryotic cells, viruses and some other biological organisms. This enzyme catalyzes the hydrolysis of 21 dUTP to dUMP and pyrophosphate (PPi), thereby preventing incorporation of uracil into DNA and reducing dUTP/dTTP ratio in cells. To date, the dUTPase gene has been characterized among a number of viruses, including herpesvirus (Fisher and Preston, 1986; Zhao et al., 2008), retrovirus (Payne and Elder, 2001), lentiviruses (Threadgill et al., 1993), African swine fever virus (Oliveros et al., 1999), parapoxvirus (Cottone et al., 2002), white spot syndrome virus (Liu and Yang, 2005), chlorella virus (Zhang et al., 2005), Rana grylio virus (Zhao et al., 2007). According to a previous report (McGeoch, 1990), dUTPases from different species are classified into two types: class 1 and class 2, based on the arrangements of five conserved motifs. The class 1 dUTPases have a chain length of around 150 aa residues, and the arrangement of the motifs is 1-2-3-4-5, which is known from bacteria, fungi, plants, metazoans, and a range of viruses. The class 2 dUTPases are about twice as long as those in class 1 with the 3-1-2-4-5 motifs, which are found only in the alpha- and gamma- herpesviruses (McGeehan et al., 2001). Based on the molecular assembly, dUTPases are grouped into three families: the monomeric form of the enzyme encoded by mammalian and avian herpesviruses; the dimeric form of dUTPase encoded by protozoan parasites and the bacterium; and the trimeric form, which is the most studied family, discovered in eukaryotes, prokaryotes and many viruses. The monomeric form is believed to have evolved via gene duplication from a standard dUTPase-coding sequence of the trimeric form followed by a subsequent loss of one copy of each motif from the double-length chain (McGeehan et al., 2001). These two forms have similar enzymatic properties. In contrast, the dimeric enzymes possess no similarity to members of other classes in sequence, structure or enzymatic characteristics. 22 4.3 Study of FAdV-9 genome FAdV-9 genome has been fully sequenced, with the size of 45,063 bp (Ojkić and Nagy, 2000). Transcriptional organization of FAdV-9 has been further analyzed as well (Ojkić et al., 2002). The genome does not contain identifiable mammalian E1, E3, E4 regions, or protein IX sequences, while sequences for E2 region, delayed-early protein IVa2, and late proteins are relatively well conserved. There are two regions of tandemly repeated sequences, TR-1 (nt 37,648–37,812) and TR-2 (nt 38,807–40,561), although their functions are unknown. In previous studies in our laboratory, FAdV-9 has been used as a recombinant virus vector to express the enhanced-green fluorescent protein (EGFP) with the insertion of an EGFP gene in the TR-2, located at the right end of the genome, and its growth in vitro has been characterized (Ojkić and Nagy, 2001). In addition, it has been demonstrated that a 2.4 kb region at the left end of the FAdV-9 genome containing two putative motifs of the packaging signal domain and six ORFs is dispensable for virus replication in vitro. However, a mutant virus FAdV-9Δ4, which is devoid of 6 ORFs (0, 1, 1A, 1B, 1C and 2) (Fig. 1.4) but contains all packaging motifs, cannot replicate at wildtype level in vivo (Corredor and Nagy, 2010a). In another study of this group (Corredor and Nagy, 2010b), an EGFP gene is inserted into the left end of genome of FAdV-9 at the tenth codon of ORF1B and the genome of FAdV-9Δ4 to replace the deleted ORFs, respectively. They showed that the left end region of the FAdV-9 genome is suitable as an insertion/replacement site for foreign gene expression. All these data suggest the importance of the left and right end genes of the FAdV-9 genome during their replication cycle, vector design, and pathogenesis. However, further research about the function of each ORF of FAdV-9 genome still needs to be conducted. 23 5. Disease associated with aviadenoviruses Despite mild or no apparent clinical signs in birds, most members of Aviadenovirus have been associated with a number of diseases including inclusion body hepatitis (IBH), hydropericardium syndrome (HPS), gizzard erosions, proventriculitis and tenosynovitis (Hess, 2013). Of these diseases, IBH and HPS are the most important ones that have been reported from different parts of the world (Hess, 2013; Mittal et al., 2014). IBH IBH was first reported by Helmboldt and Frazier in 1963 in the United States (Helmboldt and Frazier, 1963). IBH is prevalent in broiler chickens at 3-7 weeks of age, but case studies have reported that it could be found in birds as young as 7 days old and as old as 20 weeks (Hess, 2013). IBH is characterized by sudden onset of mortality with a peak after 3-4 days. It usually stops on day 5 but occasionally continues for 2-3 weeks. Morbidity is low, and mortality may reach 10%. Outbreaks of IBH in chickens less than 3-weeks of age could cause the mortality to rise to 30%. Dar et al. (2012) reported that the experimental infection of 2-day-old and 2-week-old chickens with FAdV-8b results in 83% and 43% mortalities, respectively. In the past several decades, IBH has been reported in broiler flocks in numerous countries, including Canada (Nakamura et al., 2011; Kim et al., 2008; Ojkić et al., 2008b; Philippe et al., 2005; Gomis et al., 2006; Zadravec et al., 2013). Almost all serotypes of FAdV have been reported to cause IBH in broiler chickens (Hess, 2013; Mittal et al., 2014). In Canada, the most commonly isolated serotypes are FAdVs-2, -7, -8, and -11 (Ojkić et al., 2008b), of which FAdV-8 had become the dominant serotype in Ontario since 2001 (Ojkić et al., 2008a). 24 Immunosuppressive virus agents such as infectious bursal disease virus (IBDV) and chicken anemia virus (CAV) are thought to be involved in the outbreak of IBH (Fadly et al., 1976; Hess, 2013). However, it is reported that IBH occurs in chickens before IBD is present in New Zealand (Christensen and Saifuddin, 1989). HPS HPS is a emerged disease, which has caused huge economic losses to the poultry industry in Pakistan since 1987 when it was first recognized. Compared to IBH, the mortality rate and incidence of hydropericardium is higher for HPS, resulting in between 20% and 80% mortality, though with low morbidity (Cowen, 1992; Hess, 2013). In addition to Pakistan, the disease has been recognized in India, Middle East, Japan, Mexico, and South America (Abe et al., 1998; Hess et al., 1999; Toro et al., 1999). Unlike IBH that could be caused by any of 12 serotypes, HPS has been solely associated with FAdV-4 from Fowl adenovirus C (Kim et al., 2008; Mittal et al., 2014). HPS primarily occurs in broilers of 3-6 weeks of age, and it also occurs in breeding and layer flocks as well, with lower mortality rates (Hess, 2013). 6. Diagnosis of fowl adenovirus infection Diagnosis of FAdV infections can be carried out by the observation of gross and histopathological changes in the liver, electron microscopic detection of viral particles, various serological tests, such as enzyme-linked immunosorbent assay (ELISA), agar gel precipitation test (AGPT), indirect hemagglutination, immunofluorescence, PCR, nested PCR and real-time PCR (Hess, 2000, 2013; Ganesh et al., 2002; Philippe et al., 2007; Romanova et al., 2009). 25 Several methods have been described for isolation and identification of FAdV. Cell lines of chicken liver-origin such as primary chicken embryo liver (Celi) and hepatoma cell lines (CH-SAH) can be used for the recovery of FAdV from specimens (Alexander et al., 1998; Ojkić et al., 2008a). They produce typical CPE characterized by rounding, syncytium formation (which resembles a bunch of grapes), and cell detachment. Negative stain electron microscopy and thin-section electron microscopy have proven to be useful for rapid detection of FAdV in suspected materials and tissues based on their characteristic morphology (Hess, 2013). Immunocytochemistry can also be used to detect the virus in infected cells by immunofluorescence staining with avian adenovirus antiserum (Hess, 2013). Several researchers have employed PCR alone (Jiang et al., 1999), or in combination with RFLP of different regions of the hexon gene as a very sensitive and specific method for detection, differentiation and phylogenetic analysis of FAdV (Meulemans et al., 2004; Raue et al., 2005; Ojkić et al., 2008b). For these molecular methods, PCR primers are frequently chosen in the region of the hexon gene and fiber gene because they have hypervariable regions among serotypes (Ojkić et al., 2008b). Recently, a few research groups developed a genotyping technique for FAdVs based on real-time PCR and highresolution melting-curve analysis (Marek et al., 2010; Steer et al., 2009, 2011), which is demonstrated to be accurate, rapid, and robust for the identification of FAdV serotypes. Serologic methods are also employed to identify FAdV infection. The AGPT is a widely used serologic test for the detection of FAdV antibodies, since it is fast and economical. However, its use is limited when detecting FAdV infections in SPF flocks, due to its lack of sensitivity (Hess, 2013). ELISA for the detection of group-specific or type-specific 26 Figure 1.4 ORFs in the left end of FAdV-9 genome. Taken from Corredor and Nagy, 2010a 27 antibodies to FAdVs has been described (Corredor and Nagy, 2011; Ojkić and Nagy, 2003; Junnu et al., 2014). Philippe et al. (2007) compared AGPT offered by diagnostic laboratories that uses FAdV-1 as the antigen with a FAdV group-specific ELISA. They demonstrated that ELISA is considerably more sensitive than the AGPT in early stages of infection and shows good potential for practical application to monitor for the presence of adenovirus antibodies in commercial flocks. Recently, Xie and co-workers (2013) developed an ELISA for the diagnosis of FAdVs, based on two non-structural proteins, 100K and 33K of FAdV-1, and demonstrated that the 100K-33K-ELISA method is sensitive, specific and can distinguish an acute FAdV infection from an inactivated virusbased vaccination response. In addition, it is reported that monoclonal antibody against FAdV has been also used to improve diagnostic assays, study pathogenesis and identify strains (Ahmad and Burgess, 2001). 7. Immune response to human adenovirus 7.1 Innate immune responses Adenoviruses are highly immunogenic and elicit potent innate and adaptive immune responses. Since adenoviral infections are common, the majority of adults have acquired immunity, often against multiple Ad serotypes (Nayak and Herzog, 2009). The innate response is comprised of various cells, including macrophages, dendritic cells (DCs), neutrophils and natural killer (NK) cells as well as serum proteins such as chemokines, cytokines and complement. Structural components of AdV, both DNA and capsid proteins, play roles in triggering innate responses and the outcomes can vary depending on Ad species and infected cell type (Liu and Muruve, 2003; Muruve, 2004). 28 Many of these pathways lead to activation of NF-κB and interferon regulatory factor 3 (IRF3), followed by production of inflammatory mediators and interferons (IFNs) (Nociari et al., 2009; Randall and Goodbourn, 2008). Adenoviral components are detected by pattern recognition receptors (PRRs) such as tolllike receptors (TLRs) (Nociari et al., 2009). In mammals, there are at least 13 TLRs with unique specificities. Both TLR2, TLR9 have been implicated in AdV recognition (Appledorn et al., 2008). TLR2 is a cell surface receptor that is known to bind peptidoglycan and zymosan moieties present in bacteria. Studies have shown that TLR2 is a key mediator in responses to some dsDNA viruses as well; however, the exact ligand has not been characterized. TLR9 is an endosomal receptor recognizing DNA with unmethylated CpGs motifs (Appledorn et al., 2008; Compton et al., 2003). The complement system is an important defense mechanism, which consists of plasma proteins that are important in the defense against pathogens and induces rapid destruction and phagocytosis of pathogens (Cichon et al., 2001). The main functions of the complement components contain lysis of pathogens, opsonization, activation of inflammatory response, and clearance of immune complexes (Kiang, et al., 2006). In addition to complement activation, another early defense against pathogens is the recognition of conserved microbial structures known as pathogen-associated molecular patterns (PAMPs). Members of the TLR family are transmembrane proteins that recognize PAMPs (Takeda and Akira, 2005). Adenoviral DNA is sensed by TLR9. Through the adaptor protein myeloid differentiation factor 88 (MyD88), a signalling cascade is initiated, whereupon production of type I interferons in plasmacytoid dendritic cells (pDCs) is induced (Yamaguchi et al., 2007). Type I IFNs (IFN-α and IFN-β) as well 29 as the pro-inflammatory cytokines interleukin-6 (IL-6), IL-12 and tumor necrosis factor α (TNF-α) are also secreted by other antigen presenting cells (APCs) such as myeloid DCs (mDCs), macrophages and Kupffer cells. These cells recognize adenoviral DNA through a TLR-independent pathway that is not fully characterized (Zhu et al., 2007a). One proposed pathway is the induction of interferon regulatory factor 3 (IRF3) by double stranded viral DNA that leads to transcription of type I IFNs (Nociari et al., 2007). The IFNs activate a positive feedback loop, which results in maturation of the APCs (Nociari et al., 2009). It is believed that DCs are activated by binding of the RGD motif on the adenoviral penton base to αVβ integrins on DCs, which leads to TNF-α secretion and maturation by autocrine TNF-α stimulation (Philpott et al., 2004). Type I IFN represents one of the most important antiviral defense mechanisms (O'Neill and Bowie, 2010). First of all, they promote innate immune responses by the activation of NF-κB that induces production of pro-inflammatory cytokines and chemokines. Effector cells like neutrophils, natural killer (NK) cells and monocytes are hence recruited to the site of infection. Type I IFNs also induce adaptive immune response by promoting T and B cell responses. How the adaptive immunity is regulated by type I IFNs is not completely known but they do induce production of IL-15, which stimulates NK and T cell proliferation (Bonjardim et al., 2009). In addition, type I IFNs upregulate IFN-γ production in NK cells, which induces T helper 1 (Th1) response that will activate cytotoxic T lymphocytes (CTLs) against virus-infected cells (Zhu et al., 2007b). B-cell activation is characterized by IL-10, which peaks 72-96 hours after intravenous infusion of a replication-selective adenovirus in cancer patients (Nemunaitis et al., 2001). Reid et al. (2002) reported that patients who receive HAdV-5 systemically are positive for 30 anti-HAdV-5 antibodies after the first viral dose, which complicates re-administration. 7.2 Adaptive immune responses Cell-mediated immune responses to HAdVs have been observed both for CD8+ cytotoxic T cells (CTLs) and for CD4+ T helper cells. Multiple MHC class I and class II-restricted epitopes have been mapped within the conserved region of the hexon (Leen et al., 2004, 2008). Only minimal cell-mediated immune responses to other capsid proteins, fiber or penton base, are detected. CTLs kill infected cells by multiple mechanisms including perforin, Fas-L and TNF-α, and thereby the life cycle of HAdV is disrupted before progeny viruses are released (Leen et al., 2008). Specific CD4+ T cells play a critical role in driving B-cell activation and differentiation. Activated B-cells undergo formation into plasma cells, which produce antibodies directed against adenoviral epitopes located on the major capsid proteins, i.e., hexon, penton and fiber (MacLennan et al., 1997). Binding of antibodies induces effective clearance of viruses from the circulation and enhances the interaction of AdVs with leukocytes through the Fcγ- and complement receptors (Spear et al., 2001). Adenoviral infection generates serotype-specific neutralizing antibodies (NAbs). Anti-hexon NAbs were initially proposed as the most important antibodies in the neutralization process (Roberts et al., 2006). However, recent data indicate that NAbs against fiber and penton proteins may also have great relevance (Myhre et al., 2007; Sarkioja et al., 2008). It seems that antibodies directed against various Ad capsid components synergize in the neutralization process (Gahery-Segard et al., 1998). 8. Immune responses to fowl adenoviruses Despite an increasing interest from researchers, the immune response against fowl adenovirus infection is less well studied, compared to that of human adenoviruses. 31 Recently, it has been demonstrated that FAdVs regulate a variety of cytokine gene expression in infected birds. Grgić et al. (2013b) demonstrated that chickens intramuscularly inoculated with FAdV-8 have significantly higher IFN-γ mRNA expression and significantly lower IL-8 mRNA expression in spleen and liver, compared to mock-infected chickens. FAdV-8 also upregulates, although not significantly, the mRNA expression of IL-18 and IL-10 in spleen and liver. In another study by this group (Grgić et al., 2013a), they showed that FAdV-4 induces significantly higher mRNA expression of IFN-γ and IL-10, compared to the uninfected group, while no significant difference was found in terms of the mRNA expression of IL-8 and IL-18. Several studies have shown that FAdVs infection is involved in inducing immunosuppression. For example, Shivachandra et al (2003) demonstrated that chickens infected with serotype FAdV-4 have a significant decrease in the percentage of IgMproducing B cells in bursa of Fabricius. Schonewille et al. (2008) showed that the virulent FAdV-4 infection results in a severe reduction in CD3+, CD4+ (helper T cells) and CD8+ cells (cytotoxic T cells) in spleen, and a decrease of CD4+ and CD8+ T-lymphocytes in the thymus. More recently, Hussain et al. (2012) showed that broilers inoculated with a FAdV-4 field strain have immune system dysfunction in terms of lymphoid organ integrity, antibodies, and cell-mediated immune responses, as demonstrated by atrophy of bursa of Fabricius, thymus, spleen, lower antibody titers against sheep red blood cells (SRBC) and significantly reduced phytohemagglutinin-P (PHA-P), compared to the uninfected group. Similarly, chickens orally-inoculated with a FAdV-1 field strain, isolated from field outbreaks of IBH, have a significant decrease in the antibody response to Brucella abortus (T-cell-independent antigen) and a significant decrease in 32 blastogenesis response of peripheral blood lymphocytes to PHA-P (Singh, et al., 2006). These data indicate that suppression of the antibody and cell-mediated immune responses might be a universal phenomenon of virulent fowl adenoviruses. Fowl adenoviruses elicit a strong humoral response in the hosts. FAdV-specific Abs are commonly detected in breeder and layer chickens (Hess, 2013). Following infection, birds rapidly developed detectable NAbs (type-specific) at 1 week p.i., and the peak of the Ab level is around 3 week p.i. (Ojkić and Nagy, 2003; Hess, 2013). Kim et al. (2014) demonstrated that chickens vaccinated with an inactivated FAdV-4 still remain high Ab level at 7 week p.i.. However, it is reported that the Ab response to FAdV-8 appears at 2 week p.i., followed by a slight decrease at 3 week p.i., and increases again at 4 week p.i. (Grgić et al., 2011). Corredor and Nagy (2010a) showed that the Ab response to FAdV-9 (A-2A) appears at 1 week p.i. and continues to increase until the end of the experiment at 4 week p.i.. Schachner et al. (2014) reported that 1-day-old chickens intramuscularly injected with fiber 2 of FAdV-4 induced anti-fiber 2 Abs which peak at the age of 4 weeks and protect chicken from the virulent FAdV-4 challenge. Several studies have demonstrated that the host Ab response depends on the dosage of the inoculum and the route of inoculation. For example, Ojkić and Nagy (2003) showed that the chickens intramuscularly inoculated with a higher dose of FAdV-9 have significantly higher Ab level than those inoculated with a lower dose from 1 to 4 weeks p.i.. They also demonstrated that intramuscularly inoculated chickens have significantly higher Ab level than chickens that receive virus through water or feed. Similarly, Grgić et al. (2011) showed that the level of Ab response to FAdV-8 is significantly higher in intramuscularly inoculated chickens than in orally inoculated chickens. 33 Hypotheses and Objectives Hypotheses 1. FAdV-9 ORF1 is a genuine dUTPase, which plays important roles in virus replication in vitro and in vivo. 2. FAdV-9 ORF1 modulates the host immune response against the virus infection. Objectives 1. To explore the effects of the left-end genes on virus replication and modulation of immune response on orally inoculated chickens. 2. To generate ORF1 knockout virus and HA-tagged ORF1 recombinant FAdV-9 virus. 3. To characterize the molecular features of FAdV-9 ORF1. 4. To explore the roles of FAdV-9 ORF1 on virus replication and modulation of the host response in vitro and in vivo, through comparing the ORF1 knockout virus and wild type virus. 34 Chapter 2. Oral inoculation of chickens with a candidate fowl adenovirus 9 vector Li Deng, Shayan Sharif, Éva Nagy* Department of Pathobiology, Ontario Veterinary College, University of Guelph, Guelph, Ontario, Canada *Corresponding author: Éva Nagy Tel.: +1-(519) 824-4120 Ext. 54783 Fax: +1-(519) 824-5930 E-mail address: [email protected] Mail address: Department of Pathobiology, Ontario Veterinary College, University of Guelph, 50 Stone Road East, Guelph, ON, Canada, N1G 2W1 Author’s contributions LD designed and performed all experiments, conducted the data analysis and wrote the first draft of the manuscript. SS and ÉN provided guidance during the experiments and critical review for the manuscript. Published in Clinical and Vaccine Immunology. 2013. 20(8):1189-1196. 35 Abstract Fowl adenoviruses (FAdVs) are a potential alternative to human adenovirus-based vaccine vectors. Our previous studies demonstrated that a 2.4-kb region at the left end of the FAdV-9 genome is nonessential for virus replication and is suitable for the insertion or replacement of transgenes. Our in vivo study showed that the virus FAdV-9Δ4, lacking six open reading frames (ORFs) at the left end of its genome, replicates less efficiently than wild-type FAdV-9 (wtFAdV-9) in chickens that were infected intramuscularly. However, the fecal-oral route is the natural route of FAdV infection, and the oral administration of a vaccine confers some advantages compared to administration through other routes, especially when developing an adenovirus as a vaccine vector. Therefore, we sought to investigate the effects of FAdV-9 in orally inoculated chickens. In the present study, we orally inoculated specific-pathogen-free (SPF) chickens with FAdV-9 and FAdV-9Δ4 and assessed virus shedding, antibody response, and viral genome copy number and cytokine gene expression in tissues. Our data showed that FAdV-9Δ4 replicated less efficiently than did wtFAdV-9, as evidenced by reduced virus shedding in feces, lower viral genome copy number in tissues, and lower antibody response, which are consistent with the results of the intramuscular route of immunization. Furthermore, we found that both wtFAdV-9 and FAdV-9Δ4 upregulated the mRNA expression of alpha interferon (IFN-α), IFN-γ, and interleukin-12 (IL-12). In addition, there was a trend toward downregulation of IL-10 gene expression caused by both viruses. These findings indicate that one or more of the six deleted ORFs contribute to modulating the host response against virus infection as well as virus replication in vivo. 36 Key words fowl adenovirus, oral inoculation, virus replication, host response 37 Introduction Fowl adenoviruses (FAdVs), of the genus Aviadenovirus and the family Adenoviridae (Harrach et al., 2011), have a worldwide distribution and can be isolated from both sick and healthy birds (Adair and Fitzgerald, 2008). Infection with pathogenic FAdVs can lead to inclusion body hepatitis (IBH) in broiler chickens, causing very significant losses to the poultry industry worldwide, including in Canada (Dar et al., 2012; Ojkić et al., 2008). FAdVs are transmitted horizontally and vertically, can cause persistent infections, and are excreted through feces and the respiratory tract (Adair and Fitzgerald, 2008; Grgić et al., 2006). To date, the genomes of four fowl adenoviruses (those of FAdV-1, FAdV-9, FAdV-8, and FAdV-4) have been fully sequenced (Chiocca et al., 1996; Grgić et al., 2011; Griffin and Nagy, 2011; Ojkić and Nagy, 2000), and they are about 10 kb larger than those of mastadenoviruses. Human adenoviruses (HAdVs) and other mammalian adenoviruses are used both as oncolytic viruses (Cody and Douglas, 2009; Gallo et al., 2005; Shashkova et al., 2005) and vaccine vectors (Lasaro and Ertl, 2009; Sharma et al., 2010). FAdVs are also suitable vectors; for example, FAdV-1- and FAdV-8-based recombinant viruses have induced protective immune responses against infectious bursal disease virus and infectious bronchitis virus, respectively (Francois et al., 2004; Johnson et al., 2003). The nonpathogenic FAdV-9 has also been developed as a virus vector. We demonstrated that the tandem repeat region 2 (TR-2) at the right end of the genome is dispensable and is suitable for foreign gene insertion (Ojkić and Nagy, 2001). More recently, a 2.4-kb 38 region at the left end of the FAdV-9 genome, containing two putative motifs of the packaging signal domain and six open reading frames (ORFs), was shown to be nonessential for virus replication in vitro. However, a deletion virus (FAdV-9Δ4) that lacks the six ORFs (0, 1, 1A, 1B, 1C, and 2) replicated less efficiently than the wild-type (unmodified) FAdV-9 (wtFAdV-9) in chickens inoculated intramuscularly, and the antibody (Ab) level was lower in the FAdV-9Δ4-inoculated birds (Corredor and Nagy, 2010a). We have also demonstrated that the left end of the FAdV-9 genome is a suitable site for the insertion and replacement of foreign genes (Corredor and Nagy, 2010b). Moreover, in chickens immunized with a recombinant virus containing the enhanced green fluorescence protein (EGFP) gene, antibodies were detected against the foreign protein (Corredor and Nagy, 2011). All these studies suggest the importance of the leftend genes of the FAdV-9 genome in virus replication, immune response modulation, and vector design. Moreover, the optimization of delivery routes and regimens is important for overcoming the potential limitations of AdV-based vaccines for both human and animal applications (Thacker et al., 2009). FAdVs are normally transmitted by the fecal-oral route, so we wanted to learn more about the replication of our vector virus and its effect on the chicken immune system after oral administration. Therefore, the aims of the present work were to study virus replication and host response in chickens that were inoculated orally with an FAdV vector virus (FAdV-9Δ4) and a wild-type virus (wtFAdV-9). Specifically, virus shedding in feces, viral genome copy number in tissues, antibody response, and expression levels in tissues of selected cytokine genes, alpha interferon (IFN-α), IFN-γ, interleukin-10 (IL10), and IL-12 were determined. 39 Materials and Methods Viruses and cells FAdV-9 (strain A-2A) and FAdV-9Δ4 were propagated and titrated in chicken hepatoma cells (CH-SAH) as described previously (Alexander et al., 1998). The cells were maintained in Dulbecco's modified Eagle's medium and nutrient mixture Ham's F-12 medium (DMEM-F12) supplemented with 10% non-heat-inactivated fetal bovine serum (FBS), 2 mM l-glutamine, 100 U/ml penicillin, and 100 μg/ml streptomycin. Animal experiment The experiment was reviewed and approved by the Animal Care Committee of the University of Guelph in accordance with the Guide to the Care and Use of Experimental Animals of the Canadian Council on Animal Care. One hundred thirty-five 1-day-old specific-pathogen-free (SPF) White Leghorn chickens were obtained from the Canadian Food Inspection Agency (CFIA) (Ottawa, Canada) and were housed in the isolation unit of the University of Guelph. At 7 days of age, the chickens were wing tagged and randomly divided into three groups (groups I, II, and III). On day 10, the chickens were inoculated orally with 1.5 × 107 PFU/chick with wtFAdV-9 (group I), FAdV-9Δ4 (group II), or PBS (group III). The chickens were observed daily for clinical signs of infection. To detect virus in the feces, cloacal swabs were collected in 1 ml PBS with antibiotics at 0, 1, 3, 5, 7, 10, 14, 21, and 28 days postinoculation (d.p.i.) and were stored at −80°C until processing. Sample preparation and virus titration were performed as described previously (Corredor and Nagy, 2010a). A sample was regarded as negative if it tested negative at least twice and at two different times. Blood samples were collected from all chickens on 0, 7, 14, 21, and 28 d.p.i., and sera were tested for antibodies by 40 enzyme-linked immunosorbent assay (ELISA) using purified FAdV-9 as an antigen and following the method described previously (Ojkić and Nagy, 2003). Five chickens from each group were randomly drawn for euthanasia and necropsy on 1, 2, 3, 4, 5, 7, 14, 21, and 28 d.p.i.. Liver, cecal tonsil, spleen, and bursa of Fabricius samples were collected and sectioned into two portions: one was placed in a sample bag and stored at -80°C for viral genome copy number determination, as described previously (Romanova et al., 2009), and the other was collected in a 1.5-ml sterile Eppendorf tube containing RNAlater (Invitrogen Canada, Inc., Burlington, Ontario, Canada) and stored at -80°C. The expression levels of the IFN-α, IFN-γ, IL-10, and IL-12 p40 cytokine genes were evaluated by real-time quantitative PCR (RT-qPCR), with β-actin as a reference gene, as described previously (Abdul-Careem et al., 2006, 2007; Grgić et al., 2013). Statistical analysis Statistical analyses were performed using GraphPad Prism 5.0 software (San Diego, CA). A one-way analysis of variance (ANOVA) was used to determine significant differences between the groups. The critical level for significance was set at a P value of <0.05. The data were expressed as mean ± standard error of the mean (SEM), determined from five individual birds at the designated days. Results Throughout the experiment, no clinical signs of infection were seen in any groups of chickens, and there were no pathological lesions at necropsy. 41 Virus shedding Virus titers in cloacal swabs were determined by the plaque assay. No virus was detected in any groups of chickens before inoculation and in the mock-infected group throughout the study. The virus titers in groups inoculated with wtFAdV-9 and FAdV-9Δ4 are shown in Table 2.1. For FAdV-9Δ4, virus was detected only at days 1 and 7 p.i., and the titers were significantly lower than those of wtFAdV-9-infected chickens. In the wtFAdV-9infected group, virus was detected with high titers at 1 to 14 d.p.i., but virus was not detected at the later days (21 and 28 d.p.i.). The highest titer appeared at 5 d.p.i. with 4.0 ×103 PFU/ml. Viral genome copy number in tissues Viral genome copy numbers in liver, cecal tonsil, bursa of Fabricius, and spleen samples were determined by quantitative PCR (qPCR), and the results are summarized in Table 2.2. No viral DNA was detected in the mock-infected chickens. Throughout the study, viral DNA was detected in cecal tonsil and spleen samples from both virus-infected groups from 1 d.p.i. until 21 d.p.i. and also in liver samples collected at 1, 3, 5, and 7 d.p.i. At day 14 p.i., 40% and 60% of the liver samples had detectable virus levels for the FAdV-9Δ4 and wtFAdV-9 groups, respectively. At 21 d.p.i., only 20% of the samples, and only from the wtFAdV-9 group, were positive for virus. Viral DNA was also detected in some samples of the bursa of Fabricius until day 14 p.i.; however, the genome copy numbers were low compared to those in other tissue samples. The viral genome copy number was highest in cecal tonsil samples, and it was higher in the wtFAdV-9 group than in FAdV-9Δ4-infected chickens. 42 Antibody response The presence of FAdV-9-specific Ab was determined by ELISA as described previously (Ojkić and Nagy, 2003) and is shown in Fig. 2.1. No antibodies were detected in any groups before inoculation or in the mock-infected group at any time. Over the study, antibody levels increased in both the wtFAdV-9- and FAdV-9Δ4-infected groups from week 1 p.i. until the end of the experiment at week 4 p.i.. The antibody response to wtFAdV-9 was significantly higher (P<0.001) than that to FAdV-9Δ4 throughout the experiment. Cytokine gene expression in tissues The expression of mRNA of cytokines in the spleen, liver, bursa of Fabricius, and cecal tonsil samples was measured by RT-qPCR. The expression of IFN-α, IFN-γ, IL-10, and IL-12 genes in spleen samples is shown in Fig. 2.2. There was a statistically significant upregulation (P<0.05) in the expression of IFN-α in the spleen samples from wtFAdV-9-infected chickens at 7 d.p.i. compared to that in the mock-infected group. In addition, the expression of IFN-γ was significantly upregulated (P<0.05) at both 5 and 7 d.p.i. upon wtFAdV-9 infection compared to that in both the FAdV-9Δ4-infected and mock-infected groups. IL-12, similar to the pattern of IFN-γ, was also significantly upregulated (P<0.05) in the spleen samples from wtFAdV9-infected chickens at 3, 5, and 7 d.p.i. compared to that in the other two groups. Moreover, there was also a significant upregulation (P<0.05) of the expression of IL-12 in the spleen samples from FAdV-9Δ4-infected chickens. The expression of IL-10 showed some variations, including both upregulation and downregulation, upon wtFAdV-9 or FAdV-9Δ4 infection, although they were not significant (P>0.05). It should 43 be noted that IL-10 was downregulated, although not significantly, by wtFAdV-9 at 5 and 7 d.p.i., while IFN-γ was significantly upregulated. The expression of IFN-α, IFN-γ, IL-10, and IL-12 genes in liver samples is presented in Fig. 2.3. Similar to the cytokine patterns in spleen samples, upregulation in the expression of IFN-α, IFN-γ, and IL-12 was found in the wtFAdV-9-infected group at certain d.p.i. not seen in the mock-infected group. For example, IFN-α was significantly upregulated (P<0.05) at 3, 5 and 7 d.p.i., as was the case for IFN-γ at 3, 5, and 14 d.p.i. and for IL-12 at all designated time points except 1 d.p.i. Additionally, compared to FAdV-9Δ4 infection, the wtFAdV-9 caused a greater level (P<0.05) of induction of IFNα, IFN-γ, and IL-12 at 7, 3, and 7 d.p.i., respectively. In FAdV-9Δ4 infection, upregulation was noted for only IFN-γ at 3 and 5 d.p.i. (P<0.05). The expression of IL-10 was downregulated (P<0.05) in wtFAdV-9-infected chickens at 5 d.p.i, while IFN-γ was significantly upregulated (0.001<P<0.05) at that time. The expression of IFN-α, IFN-γ, IL-10, and IL-12 genes in bursa of Fabricius samples is illustrated in Fig. 2.4. There was a statistically significant upregulation (P<0.05) of the expression of IFN-α in bursa samples from wtFAdV-9-infected chickens at 5, 7, and 14 d.p.i. and in bursa samples from FAdV-9Δ4-infected chickens at 14 d.p.i. compared to that in the mock-infected group. For the expression of IFN-γ, significant upregulation (P<0.05) was found at 5 and 14 d.p.i. in only the wtFAdV-9-infected chickens. IL-12 was upregulated significantly by both wtFAdV-9 at 7 and 14 d.p.i. and by FAdV-9Δ4 at 14 d.p.i. On the other hand, the expression of IL-10 was noted for downregulation at 14 d.p.i., while IFN-γ was significantly upregulated (0.001<P<0.05) in the wtFAdV-9 group at that time. 44 The expression of the IFN-α, IFN-γ, IL-10, and IL-12 genes in cecal tonsil samples is presented in Fig. 2.5. There was no significant difference (P>0.05) in the expression of IFN-α, IFN-γ, and IL-10 between any two groups. However, there might be some downregulation of IL-10 expression at 5 and 7 d.p.i. Unlike other cytokines, IL-12 was significantly upregulated (P<0.05) by wtFAdV-9 infection at 7 and 14 d.p.i. compared to the mock-infected group. Discussion In the present study, we investigated virus replication and host responses in chickens that were orally inoculated with our adenovirus vector candidate, FAdV-9Δ4, which lacks six ORFs at the left end of the viral genome. FAdV-9Δ4, although it replicated less efficiently in vivo than did wtFAdV-9, induced an antibody response, albeit at a lower level than in wtFAdV-9-inoculated birds. The cytokine gene expression profiles upon virus infection showed that wtFAdV-9 significantly upregulated the mRNA expression of IFN-α, IFN-γ, and IL-12 in all tested tissues except cecal tonsils at least at one tested time point throughout the experiment, while FAdV-9Δ4 did not. Human adenoviruses, such as HAdV-5, have been extensively investigated for vectored vaccine and gene therapy due to their aptitude for inducing potent innate and adaptive immune responses (Cody and Douglas, 2009; Lasaro and Ertl, 2009; Yamamoto and Curiel, 2010). However, the use of HAdV-based vectors is hampered by the widespread preexisting immunity in humans (Lasaro and Ertl, 2009). This initiated interest in the development of nonhuman AdVs, including FAdVs, which are an attractive choice both as vaccine vectors for poultry (Francois et al., 2004; Johnson et al., 2003) and as gene therapy vectors. The optimization of delivery routes and application regimens of AdV 45 vectors are also needed to counteract the limitations of HAdV-based vaccines (Thacker et al., 2009). Moreover, oral administration of AdV vectors is better able to avoid systemic neutralizing antibodies than are other routes of administration (Tucker et al., 2008; Xiang et al., 2003). Nonpathogenic FAdV-9 is being studied and developed as a vector in our laboratory. Earlier, we employed both oral and intramuscular administration routes to evaluate the Ab response to FAdV-9 (Ojkić and Nagy, 2003). However, in that study, virus was given through water and feed, which means the amount of virus dose taken up by the chickens was unknown. In more recent studies (Corredor and Nagy, 2010a, 2011), we evaluated the FAdV-9Δ4 vector virus administered intramuscularly (i.m.), and in the present work, the chickens were inoculated orally. Similar to with i.m. administration, virus was rarely detected (only at days 1 and 7 p.i.) in the feces of the orally inoculated FAdV-9Δ4 group and with titers very significantly lower than those of the wtFAdV-9 group. The route of inoculation did not alter the period of virus shedding for wtFAdV-9-infected chickens. After oral inoculation, virus was detected by both plaque assay and quantitative PCR at 1 d.p.i., which showed the highest viral genome copy number throughout the study. One explanation is that the detected viruses were from the initial inoculum, i.e., the parental viruses. At 3 d.p.i., the viral genome copy number dropped markedly in all tissue samples. A second peak of viral genome copy numbers in the liver and bursa of Fabricius samples from the wtFAdV-9 group occurred at 5 d.p.i., which was well in accord with the highest titer detected at that time point. Similar trends have been seen for FAdV-8 (Grgić et al., 2011). Viral genome copy numbers, indicating the virus load in different tissues, were significantly higher in the wtFAdV-9 group than in the FAdV-9Δ4 group tissue samples 46 except in spleen and were shown to be the highest in the cecal tonsil samples. Similar results have been obtained for both FAdV-8 (Grgić et al., 2011) and FAdV-4 (unpublished data). The antibody response after oral inoculation in the wtFAdV-9 group was significantly higher (P<0.001) than in the FAdV-9Δ4 group throughout the study (Fig. 2.1), which is similar to the i.m. inoculation results of Corredor and Nagy (Corredor and Nagy, 2010a). The fact that the mutant virus elicits a less-robust antibody response than the wild-type virus might be advantageous when the same vector virus is considered in a secondary treatment or vaccination. The i.m. inoculation induced a higher Ab level not only for wFAdV-9 but for FAdV-8 (Grgić et al., 2011) and FAdV-4 as well (our unpublished data). In addition to the Ab response, we investigated the expression of IFN-α, IFN-γ, IL-12, and IL-10 genes at different days after oral inoculation. Type I IFNs are essential for the mediation of potent antiviral responses, and they also upregulate IFN-γ production in natural killer (NK) cells, which induces a T helper 1 (Th1) response that will activate cytotoxic T lymphocytes (CTLs) against virus-infected cells. One of the major roles of IL-10 is to counteract the effects of Th1 responses by inhibiting IFN-γ synthesis (Endharti et al., 2005). Chicken IL-10 also possesses a similar function (Rothwell et al., 2004). In the present study, a trend of downregulation of IL-10 gene expression in both wtFAdV-9 and FAdV-9Δ4 groups was found, which was not surprising considering that IFN-γ expression was upregulated by both viruses. Likewise, it was also apparent that IL10 gene expression, similar to expression of other cytokine genes, was downregulated to a larger extent in wtFAdV-9-infected birds than in FAdV-9Δ4 birds. Previous studies 47 (Liu et al., 2003; Muruve et al., 2004) showed that the inflammatory response against an AdV vector in mice was transient and did not extend beyond 24 h, followed by a somewhat resting period of inflammatory gene expression that occurred in the liver samples lasting until 72 h.p.i. At days 4 to 5 p.i. a second dominant peak of inflammatory gene expression appeared in the liver samples, which is consistent with the adaptive immune response (Lieber et al., 1997). The cytokine gene expression in our study was investigated from only day 1 p.i., and thus, the first peak of inflammatory gene expression might have been missed, although this is unlikely. Nevertheless, the second peak beginning about 5 d.p.i. was confirmed. IFN-α and IFN-γ mRNA expression was upregulated in all tissues, except in the cecal tonsils, of the wtFAdV-9-infected group. The upregulation of these two cytokines was not remarkable soon after infection (day 1 p.i.), but it became statistically significant at days 3, 5, and 7 p.i. These data were similar to the results of our study on FAdV-8 (Grgić et al., 2013). FAdV-9Δ4 infection also upregulated the expression of IFN-α and IFN-γ mostly in the liver and bursa of Fabricius samples. However, the upregulation by FAdV9Δ4 was less than that by wtFAdV-9 and was statistically significant in the liver samples only at days 3 and 5 p.i. A significant difference was also noted for the expression of IFN-α and IFN-γ between wtFAdV-9 and FAdV-9Δ4 groups, which might be due to the less-efficient replication of FAdV-9Δ4 in inoculated chickens. However, it might also be due to the deleted ORFs (0, 1, 1A, 1B, 1C, and 2) that potentially have roles in modulating the host immune response against FAdV infection, as wtFAdV-9 induced a significantly higher IFN-γ expression than FAdV-9Δ4 at 3 and 5 d.p.i. in spleen samples, where no significant difference was found in terms of the viral genome copy number. 48 IL-12 is a pleiotropic heterodimeric cytokine comprising two subunits (p35 and p40) and is secreted by monocytes, macrophages, and dendritic cells (Kato et al., 1996). In mammals, the key role of IL-12 is the initiation and progression of the Th1-type immune response that is typically associated with IFN-γ induction by resting and activated T and NK cells, through inducing the proliferation of the activated T and NK cells (Cho et al., 1996; Trinchieri, 2003). Both the p40 and p35 genes of chicken interleukin-12 (chIL-12) are cloned and characterized (Degen et al., 2004), and similar to the mammalian IL-12, chIL-12 also induces IFN-γ synthesis and proliferative activity of freshly exposed chicken spleen cells. We found that the mRNA expression of IL-12 (p40) was upregulated throughout the study in all selected tissues from both the wtFAdV-9 and FAdV-9Δ4 groups. We found that the mRNA expression of IL-12 (p40) was upregulated in all selected tissues of wtFAdV-9-infected chickens. Moreover, we also noted that on the days when IL-12 was significantly upregulated, IFN-γ was also upregulated, which is in agreement with the results of Degen and coworkers (Degen et al., 2004). In conclusion, we investigated virus replication and host responses of orally inoculated chickens with a candidate FAdV vector virus, FAdV-9Δ4. Based on virus shedding and the number of viral genome copies in selected tissues, virus replication in FAdV-9Δ4 was less efficient than that in wtFAdV-9, which was similar to that of intramuscular inoculation. We also demonstrated that both wtFAdV-9 and FAdV-9Δ4 generally upregulated the mRNA expression of IFN-α, IFN-γ, and IL-12 and had a trend of downregulation of IL-10 gene expression in vivo. wtFAdV-9 normally caused a larger extent of regulation than FAdV-9Δ4. Our data suggest that the six deleted ORFs of FAdV-9Δ4 play an important role not only in virus replication in vivo but also in 49 modulating the host response against FAdV infection, the areas we are currently studying. Acknowledgments Li Deng is a recipient of a China Scholarship Council Ph.D. fellowship. This work was supported by the Natural Sciences and Engineering Research Council of Canada, the Canadian Poultry Research Council, and the Ontario Ministry of Agriculture and Food. We thank Sara Languay and Betty-Anne McBey for their technical assistance and the personnel in the Isolation Unit for their animal care. 50 Table 2.1 Virus titers (pfu/ml) in the feces of chickens orally inoculated with FAdV9Δ4 and wtFAdV-9. Days p.i. FAdV-9Δ4 Titer wtFAdV-9 %a Titer % 0 NDb 0 ND 0 1 1.04 × 102 ± 1.50 × 102 88.9 8.26 × 102 ± 5.81 × 102 100 3 ND 0 2.14 × 103 ± 1.46 × 103 100 5 ND 0 3.97 × 103 ± 4.07 × 103 100 7 1.60 × 101 ± 3.51 × 101 25 4.16 × 102 ± 3.89 × 102 100 10 ND 0 1.94 × 102 ± 2.65 × 102 80 14 ND 0 3.21 × 101 ± 3.12 × 101 53.3 21 ND 0 ND 0 28 ND 0 ND 0 a b The percentage of chickens shedding the virus ND, not detected. 51 Table 2.2 Viral genome copy number in tissues of chickens orally inoculated with FAdV-9Δ4 and wtFAdV-9. Days p.i. 1 3 5 Tissuesa FAdV-9Δ4 copy number % wtFAdV-9 copy number % L 4.27×102±1.84×101 100 1.72×105±1.76×104 100 CT 1.68×104±1.58×103 100 1.88×106±1.32×105 100 B 1.15×102±4.84×101 100 2.17×103±1.71×102 100 S 1 8.77×10 ±6.43×10 L 2.08×101±6.42×100 CT 14 21 2 1 0 0 2.98×10 ±1.71×10 100 2 2.29×10 ±4.62×10 100 100 4.12×102±1.18×101 100 100 3 6 3 100 0 0 1.21×10 ±3.08×10 B 6.52×10 ±3.76×10 40 9.65×10 ±2.71×10 60 S 3.75×102±2.05×101 100 6.03×102±2.36×101 100 L 3.39×101±3.88×100 100 8.92×102±5.64×101 100 CT 2.65×103±7.46×101 100 9.80×105±9.35×102 100 B 7 2 b 1 0 1.86×10 ±2.68×10 1 60 0 2 1 60 2 1 3.42×10 ±7.14×10 S 9.54×10 ±7.1×10 100 1.75×10 ±2.01×10 100 L 4.55×101±4.56×100 100 2.12×102±2.46×101 100 CT 2 1 0 0 6.54×10 ±2.79×10 100 3 1 100 1 0 7.53×10 ±5.86×10 B 8.90×10 ±2.76×10 40 8.28×10 ±2.93×10 80 S 8.15×101±3.71×100 100 1.46×102±1.71×101 100 L 1.62×101±3.24×100 40 7.64×101±2.72×100 60 CT 7.30×101±5.45×100 100 1.77×103±1.63×101 100 B 6.73×100±2.26×100 40 3.22×101±5.50×100 40 S 2.65×101±2.35×100 100 1.27×102±5.21×101 100 L NDc 0 1.56×101±4.35×100 20 CT 4.35×101±1.15×100 100 1.44×103±2.93×101 100 B ND 0 ND 0 S 1 0 1.52×10 ±5.76×10 100 a 52 1 1.15×10 ±2.71×10 L, liver; CT, cecal tonsil; B, bursa of Fabricius; S, spleen. The percentage of tissues in which the viral genome was detected c ND, not detected. b 2 100 Figure 2.1 Antibody response in orally inoculated chickens Antibody (IgG) response to FAdV-9 in chickens orally inoculated with FAdV-9Δ4 (checkered bars) or wtFAdV-9 (striped bars) and in mock-inoculated chickens (white bars), as measured by ELISA, shown as S/P (sample-to-positive) ratios. *, statistical significance (P<0.05) compared to the mock-infected group. Brackets above the bars indicate comparison between wtFAdV-9- and FAdV-9Δ4-infected chickens. 53 Figure 2.2 Cytokine mRNA expression in spleen samples Cytokine mRNA expression in spleen samples from wtFAdV-9 (striped bars), FAdV-9Δ4 (checkered bars), and mock-infected (white bars) chickens. Target and reference gene expression levels were quantified by RT-qPCR, and levels are presented relative to βactin expression and normalized to a calibrator. Error bars represent standard error of the means. The significance of the regulation level between any two groups was analyzed. *, significant (P<0.05) upregulation compared to the mock-infected group; **, very significant (0.001<P<0.01) upregulation compared to the mock-infected group. Brackets above the bars indicate comparison between wtFAdV-9- and FAdV-9Δ4-infected chickens. 54 Figure 2.3 Cytokine mRNA expression in liver samples Cytokine mRNA expression in liver samples from wtFAdV-9 (striped bars), FAdV-9Δ4 (checkered bars), and mock-infected (white bars) chickens. Target and reference gene expression levels were quantified by RT-qPCR and are presented relative to β-actin expression and normalized to a calibrator. Error bars represent the standard error of the means. The significance of the regulation level between any two groups was analyzed. *, significant (P<0.05) up- or downregulation compared to the mock-infected group; **, very significant (0.001<P<0.01) up- or downregulation compared to the mock-infected group. Brackets above the bars indicate comparison between wtFAdV-9- and FAdV-9Δ4infected chickens. 55 Figure 2.4 Cytokine mRNA expression in bursa of Fabricius samples Cytokine mRNA expression in bursa of Fabricius samples from wtFAdV-9 (striped bars), FAdV-9Δ4 (checkered bars), and mock-infected (white bars) chickens. Target and reference gene expression levels were quantified by RT-qPCR and are presented relative to β-actin expression and normalized to a calibrator. Error bars represent the standard error of the means. The significance of the regulation level between any two groups was analyzed. *, significant (P<0.05) up- or downregulation compared to the mock-infected group; **, very significant (0.001<P<0.01) up- or downregulation compared to the mock-infected group. Brackets above the bars indicate comparison between wtFAdV-9and FAdV-9Δ4-infected chickens. 56 Figure 2.5 Cytokine mRNA expression in cecal tonsil samples Cytokine mRNA expression in cecal tonsil samples from wtFAdV-9 (striped bars), FAdV-9Δ4 (checkered bars), and mock-infected (white bars) chickens. Target and reference gene expression levels were quantified by RT-qPCR and are presented relative to β-actin expression and normalized to a calibrator. Error bars represent the standard error of the means. The significance of the regulation level between any two groups was analyzed. *, significant (P<0.05) up- or downregulation compared to the mock-infected group; **, very significant (0.001<P<0.01) up- or downregulation compared to the mock-infected group. Brackets above the bars indicate comparison between wtFAdV-9and FAdV-9Δ4-infected chickens. 57 Chapter 3. Characterization and functional studies of fowl adenovirus 9 dUTPase Li Deng1, Xiaobing Qin2, Peter Krell3, Ray Lu3, Shayan Sharif1, Éva Nagy1* 1. Department of Pathobiology, University of Guelph, Guelph, Canada 2. College of Animal Science and Veterinary Medicine, Qingdao Agricultural University, Qingdao, China 3. Department of Molecular and Cellular Biology, University of Guelph, Guelph, Canada * Corresponding author: Éva Nagy Tel.: +1-(519) 824-4120 Ext. 54783 Fax: +1-(519) 824-5930 E-mail address: [email protected] Mail address: Department of Pathobiology, Ontario Veterinary College, University of Guelph, 50 Stone Road East, Guelph, ON, Canada, N1G 2W1 Author’s contributions LD designed all experiments and performed the majority of them, conducted the data analysis and wrote the first draft of the manuscript. XQ made an intermediate construct for the FAdV-9HA-ORF1 virus. RL, SS and ÉN provided guidance during the experiments. PK and ÉN provided critical review for the manuscript. To be submitted to Virology. 58 Abstract Fowl adenoviruses (FAdVs) are being developed as recombinant vaccine vectors and cancer therapy tools due to their large genomes that allow for a large transgene capacity. dUTPase, a ubiquitous enzyme that catalyzes the hydrolysis of dUTP to dUMP, found in many viruses including adenoviruses, has yet to be identified in FAdV-9. A multiple alignment of dUTPase amino acid sequences suggested that dUTPase exists in most AdVs including FAdVs. FAdV-9 ORF1 was very similar to other viral dUTPases and contained the five conserved motifs that define the protein family. Indeed, ORF1 was verified as a functional dUTPase through a PCR-based dUTPase enzymatic assay. Moreover, the transcription and protein expression patterns were characterized by RTPCR and Western blot, which showed that FAdV-9 dUTPase was transcribed from 2 hours post-infection (h.p.i.) and the protein was translated from 6 h.p.i., and both continued to the late phase of virus infection. By immunofluorescence microscopy with a HA-tagged dUTPase the recombinant virus FAdV-9 dUTPase was localized in both the cytoplasm and nucleus. Using a dUTPase knockout virus generated through site-directed mutagenesis and homologous recombination, we found that FAdV-9 dUTPase was not required for virus replication in vitro, but upregulated the expression of type I interferons. This is the first study that functionally characterized an early gene in FAdV-9. Our findings on the FAdV-9 dUTPase have shed new light on the mechanism of host immune response against FAdV infection. Key words fowl adenovirus 9, dUTPase, virus replication, type I interferon 59 Introduction Fowl adenoviruses (FAdVs) are distributed worldwide in poultry farms. Some FAdVs are important causative agents of inclusion body hepatitis (IBH) and hydropericardium syndrome (HPS) that result in significant economic losses to the poultry industry (Hess, 2013). FAdVs belong to the genus Aviadenovirus in the family Adenoviridae. The 12 serotypes are classified into five species: Fowl adenovirus A to Fowl adenovirus E. Due to the large DNA genome that allows for a large transgene capacity and no or low pathogenicity, FAdVs are considered as attractive alternatives to human adenoviruses (HAdVs) as recombinant vaccine vectors and gene therapy tools (Thacker et al., 2009; Corredor and Nagy, 2010b). To date, five FAdV genomes have been completely sequenced, including FAdV-1, -4, -5, -8 and -9 (Chiocca et al., 1996; Griffin and Nagy, 2011; Grgić et al., 2011; Marek et al., 2012; Marek et al., 2013; Ojkić and Nagy, 2000). Unlike HAdVs, however, very limited data are available in terms of the molecular biology of FAdVs, as most of the putative FAdV ORFs share no homology to any known genes, especially for the genus-specific open reading frames (ORFs) at both ends of the genome. To date, only a few genes have been functionally characterized and functional studies of them lag behind those of HAdVs. GAM-1 and ORF22 of FAdV-1 (CELO virus) show the ability of activating E2F-dependent transcription through interacting with the retinoblastoma protein (pRb), although neither exhibits any significant sequence homology to E1A proteins of HAdVs (Lehrmann and Cotten, 1999). In addition, FAdV-1 GAM-1 is verified as an antiapoptotic protein that functions similarly to Bcl-2 and human adenovirus E1B-19K (Chiocca et al., 1997). Genomic sequence analysis (Cao et al., 1998; Davison et al., 2003) 60 showed that FAdVs encode a deoxyuridine 5′-triphosphate pyrophosphatase (dUTPase) homolog, which has already been functionally verified in FAdV-1 (Weiss et al., 1997) but not yet in other FAdVs. However, its function has not been demonstrated. dUTPase is a ubiquitous enzyme that catalyzes the hydrolysis of dUTP to dUMP and pyrophosphate (PPi), thereby reducing the dUTP/dTTP ratio in cells and preventing the incorporation of uracil into DNA. The hydrolysis of dUTP generates dUMP that serves as a precursor for the biosynthesis of thymidine nucleotides (Harris et al., 1999). All freeliving organisms, as well as many viruses encode dUTPase, including herpesvirus (Glaser et al., 2006), retrovirus (Payne and Elder, 2001), African swine fever virus (Oliveros et al., 1999), poxvirus (Cottone et al., 2002), and adenovirus (Weiss et al., 1997). Based on the oligomerization state, there are three forms of dUTPase representing three distinct families. The first family contains the monomeric dUTPase encoded exclusively by mammalian and avian herpesviruses. The second family contains the homodimeric dUTPases, which are present in protozoa and bacteria. The largest family forms homotrimers, which are found in prokaryotes, most eukaryotes, some DNA viruses such as poxvirus, and a number of retroviruses (Tarbouriech et al., 2005). It has been demonstrated that viral dUTPase is important for virus replication (Threadgill et al., 1993; Voronin et al., 2014). Other studies showed that the dUTPase of EpsteinBarr virus (EBV) is able to upregulate pro-inflammatory cytokines in unstimulated peripheral blood mononuclear cells (PBMCs) (Glaser et al., 2006) and the dUTPase of the murine gammaherpesvirus 68 (MHV-68) can inhibit type I interferon signaling (Leang et al., 2011). Interferons (IFNs) are characterized by their potent antiviral properties (Takaoka and 61 Yanai, 2006). They are grouped into two classes: type I IFNs (majorly IFN-α and IFN-β), which are produced in virally infected cells to confer an antiviral state on uninfected cells, and type II IFN, which comprises a sole member, IFN-γ, and is strongly produced by activated T cells or NK cells and implicated in macrophage activation and antiviral response (Vilcek, 2003; Takaoka and Yanai, 2006). The objectives of this study were to characterize FAdV-9 ORF1 as a dUTPase enzyme and to investigate its molecular features including transcription and protein expression patterns; subcellular localization and its potential role in virus replication in vitro and modulating cytokine gene expression, including IFN-α, IFN-β, IFN-γ, and IL-10, a pleiotropic cytokine that inhibits the synthesis of proinflammatory cytokines (Rothwell et al., 2004). Materials and Methods Viruses and cells The wild type FAdV-9 (wtFAdV-9; strain A-2A) and all mutant viruses including ORF1stop, resORF1, and FAdV-9HA-ORF1 were propagated and titrated in chicken hepatoma cells (CH-SAH) as described previously (Alexander et al., 1998). The one-step growth curves in CH-SAH cells were also done as described (Alexander et al., 1998). Cytokine gene expression, ORF1 transcription and protein translation, were determined in Celi cells that were collected from the livers of 15-day-old chicken embryos as described by Schat and Purchase (1989) and in CH-SAH cells. All cells were grown in Dulbecco’s modified Eagle’s medium/nutrient mixture F-12 Ham (DMEM-F12) supplemented with 10% fetal bovine serum (FBS), 2 mM L-glutamine, 100 U/ml penicillin, and 100 µg/ml streptomycin. 62 dUTPase enzyme activity assay The assay for testing the dUTPase enzyme activity was adapted from a protocol (Leang et at., 2011). This is based on the ability of dUTP to be incorporated into a PCR product during amplification in lieu of dTTP. If presented with active dUTPase, the dUTP would be hydrolyzed to dUMP which cannot be incorporated and will prevent forming a PCR product. Briefly, to produce ORF1 protein, ORF1 was cloned into pET-28a first, and the His-tagged ORF1 protein was expressed in BL21(DE3) E. coli following the induction with isopropyl-beta-D-thiogalactopyranoside, followed by purification on a Ni-column. To test the dUTPase enzyme activity, five microliters of purified His-tagged ORF1 protein was incubated with 5 μl of 5 mM dUTP (Roche U1191) for 24 hours in 10 μl of dUTPase reaction buffer (100 mM Tris pH 7.5, 20 mM MgCl2, 20 mM DTT, and 0.2 mg/ml BSA) at 37 °C. Reactions were terminated by freezing. Instead of a dNTP mix, dUTP, or His-ORF1-treated dUTP, together with a mixture of individual dATP, dCTP, and dGTP, were used for PCR amplification of ORF1 as a template (ORF1-For: 5’ACTGAAACCTCGCAGAGGTCC-3’; ORF1-Rev: 5’- ACTGAAACCTCGCAGAGGTCC-3’). PCR was conducted using Taq polymerase with a high concentration of MgCl2 (5 mM). In combination with individual dATP, dCTP, and dGTP, dTTP served as a positive control, while DNase and RNase-free H2O served as the negative control. Heat-inactivated His-tagged ORF1 protein (95 °C, 5 mins) was also used as a control. Cycle conditions were 95°C 10 min; 95 °C 30 seconds, 57 °C 30 seconds, 72 °C 30 seconds for 35 cycles; 72°C 10 minutes; hold at 4°C. PCR products were run in a 1.0 % agarose gel. 63 Generation of mutant viruses The FAdmid clone pPacFAdV-9 containing the whole FAdV-9 genome was constructed previously in our laboratory to manipulate the FAdV-9 genome (Ojkić and Nagy, 2001). First, the left-end 3.1 kb (nt 210–3316) of FAdV-9 genome was cloned to the pGEM-T vector by primers pleft-For and pleft-Rev (Table 3.1) to generate the intermediate construct pGEM-T-pleft (Step 2 in Supplementary Fig. S1). Primers (Table 3.1) were designed to introduce three stop codons (TAA at nt 850, TGA at nt 865, and TAG at nt 871) and one SwaI site (ATTTAAAT at nt 852; no other SwaI cutting site would be present in the final construct) within ORF1 (nt 847-1338) for knocking out ORF1, through two rounds of PCR-based site-directed mutagenesis (SDM) (Step 3 in Fig. S1). The PCR product was digested with DpnI to get rid of the original methylated pGEM-Tpleft template plasmid, followed by transformation into DH5α. The ORF1-mutated pGEM-T-pleft was screened by SwaI digestion and sequencing, and the positive plasmid clone was named ORF1-SDM-pleft. Secondly, the ORF1-mutated left-end FAdV-9 genome was amplified from ORF1-SDM-pleft with pleft-F and pleft-R (Table 3.1), followed by gel purification (Step 4 in Fig. S1). The FAdmid pPacFAdV-9 was then digested with SgfI and purified through ethanol precipitation (Step 1 in Fig. S1). Homologous recombination was performed by co-transforming 500-800 ng purified mutated PCR product and 50-80 ng SgfI-digested pPacFAdV-9 into E. coli BJ5183 strain (Step 5 in Fig. S1), as described (Corredor and Nagy, 2010a). The positive mutant FAdmid was screened by SwaI digestion and sequencing; followed by PacI digestion and transfection into CH-SAH cells with 2-4 µg DNA. Upon the appearance of cytopathic effect (CPE) at around 5-6 days post-transfection, cell cultures were frozen and thawed 64 three times, followed by virus propagation in CH-SAH cells in a 150 mm dish. The virus was obtained through ultra-centrifugation and viral DNA extraction was conducted as described (Ojkić and Nagy, 2001). The positive mutant virus was confirmed by sequencing the DNA and named ORF1stop. Similarly, the ORF1-rescued virus resORF1 and HA-tagged virus FAdV-9HA-ORF1 were obtained with primers listed in Table 3.1. RT-PCR and qRT-PCR CH-SAH cells were infected with FAdV-9HA-ORF1 at a multiplicity of infection (MOI) of 5. At 0, 1, 2, 4, 6, 8, 10, 12, 16, 20, and 24 h.p.i., cells were collected for RNA extraction with Tri Reagent (Molecular Research Center), followed by DNase I treatment (Fermentas), cDNA synthesis with SuperScript® II Reverse Transcriptase (Invitrogen) and random primer. The cDNA was amplified by primers HA-tag-For and ORF1-Rev (Table 3.1). The pcDNA-HA-ORF1 plasmid and RNA of mock-infected cells served as positive and negative controls, respectively. PCR products were run in a 1.0 % agarose gel. RNAs were obtained from CH-SAH cells at 4, 8, 12, 16, 24 h.p.i. and primary chicken embryo liver cells at 8, 12, and 16 h.p.i. mock-infected or infected with wtFAdV-9, ORF1stop, resORF1FAdV-9 at a MOI of 5, followed by cDNA synthesis as described above. The 1/10 diluted cDNA was used for real-time PCR with the gene specific primers for different cytokines (Table 3.2) and LightCycler® 480 SYBR Green I Master as described (Grgić et at., 2011). β-actin was used as a housekeeping gene. All data were normalized to a positive plasmid, which contains the same sequence as the amplicon of the gene as a calibrator. 65 Western blot CH-SAH cells infected with FAdV-9HA-ORF1 at an MOI of 5 were washed with PBS and harvested at 2, 4, 6, 8, 12, 16, 20 and 24 h.p.i. by removing the monolayer with a cell scraper in PBS, followed by centrifugation at 6000 rpm at 4°C for 10 mins. The cell pellets were then lysed with RIPA buffer containing a protease inhibitor cocktail on ice for 20 mins. The supernatant was harvested by centrifugation at 6000 rpm at 4°C for 20 mins. HA-ORF1 fusion protein was probed with mouse anti-HA monoclonal antibody (Sigma-Aldrich, 1:750) and horseradish peroxidase-conjugated goat anti-mouse IgG antibody (Invitrogen, 1:10, 000). Immunofluorescence assay CH-SAH cells were infected with FAdV-9HA-ORF1 at an MOI of 5. At 6, 12, 18, and 24 h.p.i., cells were fixed with 3.7% paraformaldehyde for 30 mins and permeabilized with 0.1% NP-40 for 20 mins at room temperature. Afterwards, cells were incubated with mouse anti-HA monoclonal antibody (Sigma-Aldrich, 1:750) for 1 hour, and DyLight 549-conjugated goat anti-mouse IgG antibody (Jackson ImmunoResearch, 1:10, 000) for 1 hour. The ProLong® Gold antifade reagents with DPAI (Life Technologies) was added and incubated for 10 minutes prior to analysis by confocal microscopy. Statistical analysis Statistical analyses were performed with GraphPad Prism 6.0 software (San Diego, CA). Significant differences of the cytokine gene expressions among the groups were determined through the two-way ANOVA analysis. The critical level for significance was set at P<0.05. The data were expressed as mean ± S.E.M., determined from three 66 independent experiments. Results Bioinformatics analysis of FAdV-9 ORF1 as a potential dUTPase The 489 bp nucleotides of FAdV-9 ORF1, located at the left end of the genome (nt 8471335), encodes a polypeptide of 163 amino acids (aa) with a calculated molecular mass of 17.4 kDa. Previous studies suggested that ORF1 of FAdVs is a dUTPase homolog (Davison et al., 2003; Corredor and Nagy, 2006). The amino acid sequences of 20 identified or putative viral dUTPases, including 11 mastadenoviruses and 9 aviadenoviruses, were obtained from GeneBank (dUTPase homolog in HAdV-F genome is not available). The pairwise identities of dUTPase sequences between any two species were calculated and summarized in the Supplementary Table S1. The data showed that all HAdV dUTPases share high amino acid sequence identities (40.8% - 68.8%) with each other, but low (less than 29.1%) with other members in the genus Mastadenovirus, with the exception of Simian adenovirus A (SAdV-A) (39% - 51.3%). However, dUTPases from all aviadenoviruses show high amino acid sequence identities (44.3% - 85.0%) of dUTPase with each other. Low identities were found between the two genera Mastadenovirus and Aviadenovirus (less than 33.3%) except for that between aviadenoviruses and Porcine adenovirus C (PAdV-C) (46.5 to 59.9%), Bovine adenovirus A (BAdV-A) (45.8 to 54.9%), and Tree shrew adenovirus A (TSAdV-A) (43.0 to 53.0%). These results indicat that the dUTPase genes, while conserved, are highly variable within the family Adenoviridae, though more conserved within the genus Aviadenovirus. In addition, 30 dUTPase amino acid sequences from several viruses and different organisms were aligned to identify the conserved regions. As illustrated in Fig. 67 3.1, the 5 motifs forming the active site are well conserved in all sequences except in human adenoviruses, SAdV-A and PAdV-A, in which motif 5 is missing. Due to the high identities of amino acid sequences between FAdV-9 ORF1 and other dUTPases and the conserved motifs found among them, we concluded that FAdV-9 ORF1 very likely encodes a dUTPase. FAdV-9 ORF1 has dUTPase enzymatic activity To determine if FAdV-9 ORF1 indeed encodes a functional dUTPase, a PCR-based enzymatic assay was employed. The rationale of the assay is explained in Materials and Methods. Initially, to express the dUTPase, ORF1 was cloned into pET-28a. The Histagged ORF1 protein was expressed in Escherichia coli (E. coli) BL21 (DE3) following induction with isopropyl-beta-D-thiogalactopyranoside at 37 ºC. The fusion protein HisORF1, calculated to have a molecular mass of approximately 22.0 kDa, was detected by SDS-PAGE. The His-ORF1 protein was further confirmed by Western blot, using mouse anti-His monoclonal antibody (Sigma-Aldrich) and horseradish peroxidase-conjugated goat anti-mouse IgG antibody (Invitrogen). The His-ORF1 protein was purified through a Ni-column for the dUTPase enzymatic assay. As shown in Fig. 3.2, PCR products were detected when using a mixture of dCTP, dGTP, and dATP with either dTTP (lane 1) or dUTP (lane 2), which served as positive controls. In lanes 3 and 4, PCR products were detected in the presence of dUTPase reaction buffer itself, indicating that the buffer did not interfere with the PCR reaction. In lanes 5 and 7, in which dTTP or dUTP was preincubated with His-tagged ORF1 protein before use, PCR product was present in lane 5 (dTTP) but not in lane 7 (dUTP), suggesting that the dUTP was hydrolyzed in the presence of His-tagged ORF1 protein and the hydrolysis was specific to dUTP (lane 7) 68 but not dTTP (lane 5). To demonstrate that it was the enzymatic activity and not just the ORF1 protein itself that was responsible for the lack of PCR product in lane 7, the HisORF1 protein was heat-inactivated prior to the incubation with dUTP in dUTPase reaction buffer in lane 6, in which the PCR product was present. Together, it is concluded that the ORF1 protein possesses dUTPase enzymatic activity. In vitro characteristics of ORF1 mutant viruses Many studies have shown that the viral dUTPase is important for virus replication in nondividing cells (Oliveros et al., 1999; Threadgill et al., 1993). However, the role of FAdV9 ORF1 during the virus life cycle is still unknown. To determine if ORF1 affects virus replication, we generated an ORF1 knockout virus, ORF1stop, and its rescued revertant resORF1, as depicted in Supplementary Fig. 3.1. Viral DNA accumulation and one-step growth curves were analyzed among ORF1stop, resORF1, and wild type FAdV-9 viruses. As shown in Fig. 3.3, both viral DNA accumulation and virus growth kinetics of ORF1stop were similar to the wild type and rescued virus, indicating that ORF1 deletion did not affect viral DNA accumulation or virus replication in vitro. Transcription and protein expression profiles of ORF1 To study protein expression of FAdV-9 ORF1, an HA-tagged recombinant FAdV-9 (FAdV-9HA-ORF1) was constructed by inserting an HA tag into the N-terminus of ORF1 in the genome through site-directed mutagenesis and homologous recombination. CH-SAH cells were infected with FAdV-9HA-ORF1, and the transcription profile of ORF1 was determined by RT-PCR, which was expected to generate a 516 bp PCR fragment with primers ORF1-For and ORF1-Rev (Table 3.1). As shown in Fig. 3.4, ORF1 was transcribed as early as 2 h.p.i. and it continued through the entire time-course. 69 This is consistent with previous results in our laboratory (Cao et al., 1998). CH-SAH cells were infected with FAdV-9HA-ORF1, and protein expression was evaluated by Western blot using mouse anti-HA monoclonal antibody (Sigma-Aldrich) and horseradish peroxidase-conjugated goat anti-mouse IgG antibody (Invitrogen). As shown in Fig. 3.4, no band was detected at 2 and 4 h.p.i.. However, two bands were observed from 6 h.p.i. to 24 h.p.i.. Based on the predicted size of HA-ORF1 (approximately 18.5 kDa), the faster migrating band of the two presumably represents HA-ORF1 protein. The slower migrating band (at ~21 kDa) was present with the faster migrating one in all lanes from 6 to 24 h.p.i. but not detected at earlier time (2 and 4 h.p.i.). This suggests that the higher molecular weight protein detected by the anti-HA antibody was at least virus infection dependent and could represent an alternative form of HA-ORF1. To determine if expression of the larger protein is cell-dependent, DF-1 cells, primary chicken embryo liver cells (Celi), and primary duck fibroblast cells were also infected and analyzed by Western blot. The slower migrating band (~21 kDa) and the band of HA-ORF1 protein appeared for all cells (data not shown). Cellular localization of ORF1 protein Studies have shown that other viral dUTPases localize in both the nucleus and cytoplasm; however, no data is available for any adenoviral dUTPase. In order to determine the cellular localization of FAdV-9 ORF1, immunofluorescence assay was conducted in CHSAH cells infected with FAdV-9HA-ORF1. As shown in Fig. 3.5, the red fluorescence representing HA-ORF1 protein first appeared at 6 h.p.i. in the cytoplasm, though fluorescence in some cells was weak. Starting from 12 h.p.i., some of the red fluorescence was also seen in the nucleus, and continued to accumulate up to 18 h.p.i.. By 70 24 h.p.i., the red fluorescence was close to be evenly distributed in both the cytoplasm and nucleus and was quite strong. Cytokine gene mRNA expression induced by wild type and mutant viruses The dUTPases of some gammaherpesviruses are demonstrated to modulate the host immune response against virus infection (Glaser et al., 2006; Leang et al., 2011).To investigate if FAdV-9 ORF1 has similar functions, CH-SAH cells were infected with ORF1stop, resORF1 and wtFAdV-9, respectively. Mock-infected cells were used as a negative control. The mRNA expression of IFN-α, IFN-β, IFN-γ, and IL-10 was determined by qRT-PCR in each group, relative to β-actin. As illustrated in Fig. 3.6, the resORF1 group showed very similar results with wtFAdV-9 for all cytokines and all time points. This demonstrated that the only change made in the genome of ORF1stop was the knockout of ORF1, to which any difference found between ORF1stop and wtFAdV-9 could be ascribed. Hence the comparisons are reported only for wtFAdV-9. As shown in Fig. 3.6, despite a slight increase, transcript levels of IFN-α in ORF1stop group were not statistically different from the mock group from 8 to 24 h.p.i.. However, IFN-α transcripts were upregulated by wtFAdV-9 at 12 (2.7 fold, 0.001<P<0.01) and 16 h.p.i. (1.9 fold, 0.001<P<0.01), compared to mock control. In addition, there was a statistically significant difference of IFN-α transcripts between wtFAdV-9 group and ORF1stop group at 12 (1.8 fold, 0.001<P<0.01) and 16 h.p.i. (1.6 fold, 0.001<P<0.01). IFN-β transcripts in both ORF1stop and wtFAdV-9 groups were significantly upregulated at 12 (2.0 fold, P<0.05; 3.5 fold, 0.001<P<0.01) and 16 h.p.i. (1.5 fold, P<0.05; 2.3 fold, 0.001<P<0.01), compared to mock control. However, wtFAdV-9 upregulated the IFN-β transcripts to a larger extent than ORF1stop did at 12 (1.7 fold, 0.001<P<0.01) and 16 71 h.p.i. (1.5 fold, 0.001<P<0.01). This may suggest that, in addition to ORF1, there is another FAdV gene that is responsible for the up-regulation of IFN-β. wtFAdV-9 generally seemed to have induced higher IFN-γ mRNA expression than ORF1stop at all time points (except at 4 h.p.i.), although the differences were not statistically significant. Transcript levels of IL-10 were significantly downregulated by ORF1stop (2.0 fold, P<0.05), resORF1 (2.0 fold, P<0.05), and wtFAdV-9 (1.8 fold, P<0.05) at 8 h.p.i.. At 12 and 16 h.p.i., despite a downregulation by all viruses compared to mock control, no significant difference (P>0.05) was found between any two groups. To investigate if any of the differences were cell line-dependent in modulating cytokine transcript levels, primary chicken embryo liver cells (Celi) were also tested. Only 8, 12, 16 h.p.i. were selected, since any major differences were found at these time points for CH-SAH cells. As shown in Fig. 3.7, all viruses upregulated transcript levels of IFN-α, IFN-β, and IFN-γ, compared to the mock control. The results were very similar to those of CH-SAH cells. Specifically, despite an increase, IFN-α transcripts in ORF1stop group were not statistically different with the mock control at all time points, while it was significantly up-regulated by wtFAdV-9 at 12 h.p.i. (3.6 fold, 0.001<P<0.05) and 16 h.p.i. (5.6 fold, 0.001<P<0.05). There was a significant difference of the IFN-α transcripts between ORF1stop and wtFAdV-9 groups at 12 (2.7 fold, 0.001<P<0.05) and 16 h.p.i. (3.4 fold, 0.001<P<0.05), which was similar to that in CH-SAH cells as well. While there was some minor increase of transcript level of IFN-β and IFN-γ compared to mock control, no statistically significant differences were found for any two groups. Downregulation of IL-10 transcripts was found by all viruses at 16 h.p.i., though the differences between any two groups were not statistically significant. 72 Discussion In contrast to human adenoviruses, the molecular biology of fowl adenoviruses remains poorly understood. In the present study, we showed that FAdV-9 ORF1 encodes a genuine and active dUTPase enzyme that could catalyze the hydrolysis of dUTP. This dUTPase would be the first early gene functionally characterized in FAdV-9. FAdV-9 dUTPase was first transcribed at 2 h.p.i. and its protein was expressed at 6 h.p.i. in the cytoplasm, which was localized in both the nucleus and cytoplasm later. We also found that the presence of FAdV-9 dUTPase did not affect virus replication in vitro but contributed to the up-regulation of type I interferons. To date, a number of viral dUTPases have been characterized (Cottone et al., 2002; Glaser et al., 2006; Oliveros et al., 1999; Payne and Elder, 2001; Weiss et al., 1997). In this study, we aligned 20 dUTPase amino acid sequences from different adenoviral species and showed that the dUTPase, though conserved among aviadenoviruses, is highly variable within the family Adenoviridae. The multiple alignment of 30 dUTPase amino acid sequences from different viruses and organisms demonstrated that FAdV-9 ORF1 shares high dUTPase amino acid identity with many other viruses and organisms, with the 5 conserved motifs forming the active site well conserved. On the other hand, HAdVs dUTPases less well conserved with others, in which of the 5 conserved motifs, motif 5 is not present and motifs 1, 2 and 3 are not well conserved. This is probably the reason why HAdV dUTPases lack dUTPase enzymatic activity (Weiss et al., 1997), as the motifs 3 and 5 were demonstrated to be critical for the catalytic activity in the dUTPases of herpesvirus and equine infectious anemia virus (Shao et al., 1997; Harris et al., 1999; Freeman et al., 2009). It is speculated that the HAdVs-encoded dUTPases have 73 evolved from an ancestral dUTPase enzyme to an oncogene leading to cell transformation (Weiss et al., 1997). Viral dUTPase is dispensable in dividing cells but is required for efficient virus replication in non-dividing macrophages, for example, for African swine fever virus (Oliveros et al., 1999) and equine infectious anemia virus (Threadgill et al., 1993). However, Voronin and co-workers (2014) have recently showed that even in dividing cells (Cf2Th cells), the dUTPase of bovine immunodeficiency virus is critical for the production of progeny virus, although both viral cDNA synthesis and integration into the host cell DNA were unaltered. In the present study, viral DNA accumulation and onestep growth curves were identical between ORF1stop and wtFAdV-9 in both CH-SAH cells and primary chicken embryo liver cells, although, it could not be ruled out that FAdV-9 dUTPase may be important for virus replication in vivo. The dUTPase of herpes simplex virus does not affect the replication of the virus in vitro, but severely reduces the replication in the central nervous system of mice (Pyles et al., 1992). Similarly, MHV-68encoded dUTPase is necessary for efficient virus replication in the lungs of infected mice (Leang et al., 2011). Previous studies showed that FAdV-9 ORF1 (formerly named ORF LTR1) was transcribed at 2 h.p.i. (Cao et al., 1998). In this study, we generated a recombinant virus FAdV-9HA-ORF1 so that the HA-ORF1 protein could be detectable by Western blot with an anti-HA antibody. An 18.5 kDa HA-ORF1 protein band and a slower migrating band (~21 kDa) were detected as early as 6 h.p.i., and continued to the late phase of the virus life cycle and only in virus-infected cells. Other viral dUTPases are also found to be transcribed and translated at the early stage of virus replication. Zhao et al. (2008) 74 reported that duck enteritis virus dUTPase is transcribed as early as 30 min p.i. and its protein expression first occurred at 8 h.p.i., whereas the Rana grylio virus-encoded dUTPase is both transcribed and expressed at 4 h.p.i. (Zhao et al., 2007). It should be noted that the level of transcription and expression of other viral dUTPase generally peaked at a certain time point, while that of FAdV-9 dUTPase continuously maintained a very high level from the beginning to the late phase. In addition, from 12 h.p.i., ORF1 protein appeared and accumulated in the nucleus, and was distributed evenly in both cytoplasm and nucleus at 24 h.p.i., which is consistent with the results of other viral dUTPase proteins (Zhao et al., 2007, 2008). These data suggest that FAdV-9 dUTPase might play an important role in the nucleus during the virus life cycle, perhaps related to its role in maintaining a low dUTP/dTTP ratio to minimize the incorporation of uracil into the viral DNA. The dUTPases encoded by the gammaherpesviruses have novel functions, which are independent of their enzymatic activity. For instance, Glaser and co-workers (2006) showed that the EBV-encoded dUTPase is able to induce immune dysregulation in vitro as demonstrated by the up-regulation of TNF-α, IL-1β, IL-8, IL-6, and IL-10 in unstimulated PBMCs. In the present study, we compared the transcript level of INFs -α, β, -γ, and one anti-inflammatory cytokine IL-10 in cells infected with ORF1stop, resORF1, and wtFAdV-9. Our results showed that wtFAdV-9 induced 1.8 and 1.7 fold higher levels of IFN-α and IFN-β transcripts than ORF1stop, respectively, indicating a role of FAdV-9 dUTPase in the upregulation of type I IFNs. This is similar to the data on EBV-encoded dUTPase which upregulated IFN-β by 11-fold (Ariza et al., 2013). Other studies have also demonstrated that the CD4 and cytokine receptor interacting region of 75 gp120 evolved from an ancestral dUTPase gene (Abergel et al., 1999) and that gp120 of human immunodeficiency virus (HIV) is involved in higher levels of TNF-α, IL-6, IL-10, INF-α, and IFN-γ (Rychert et al., 2010). Type I IFNs represent the first line of defense for the innate immune system (Kawai and Akira, 2006), which plays an essential role in the host response against viral infections (Taniguchi and Takaoka, 2001). Moreover, many viruses have developed a myriad of strategies to evade the host immune response through targeting the type I IFN system. Viruses encode certain proteins that can inhibit IFN-α and IFN-β synthesis, inactivate secreted IFN molecules, interfere with IFNAR signaling, and/or block the activation of antiviral effector proteins upregulated by IFNs (Weber et al., 2004). The examples include the NS1 protein of influenza A virus (Hale et al., 2008), the NS3/4A of hepatitis C virus (Li et al., 2005), and the V protein of paramyxovirus (Andrejeva et al., 2004). Previously, we found that FAdV-9 grows as well in CH-SAH cells pre-treated with recombinant chicken IFN-α (AbD Serotec®) as in untreated cells (data not shown), suggesting that FAdV-9 can evade the antiviral response of IFN-α. This indicated that there should be some viral protein that can counteract the type I IFN response against FAdV-9 infection. It was hypothesized that dUTPase can perform this function, as is the case for the dUTPase of the MHV-68 (Leang et al., 2011). However, our data showed that FAdV-9 dUTPase was able to induce type I interferons, which was surprising but very interesting as this may indicate that dUTPase might function like a pathogenassociated molecular pattern (PAMP) instead. Ariza and co-workers (2014) have verified the identity of dUTPases of human herpesvirus 6A (HHV-6A), HHV-8, and varicella zoster virus (VZV) as a PAMP, as evidenced by activating NF-κB through ligation of 76 TLR2/TLR1 heterodimers and inducing the secretion of the inflammatory cytokines and IFN-γ. It should be noted, however, that the dUTPase of herpesviruses is monomeric and is normally twice the size of the sequence of the trimeric dUTPase (typically around 150 aa). Moreover, in the herpesvirus dUTPases, the 5 conserved motifs are reshuffled to an order of 3-1-2-4-5 instead of the order 1-2-3-4-5 in the trimeric dUTPase (McGeehan et al., 2001). It is well known that AdVs induce robust innate and adaptive immune responses, although the detailed mechanism remains to be clarified. Yamaguchi and co-workers (2010) have shown that the virus-associated RNAs (VA-RNAs) induce the production of IFN-α and IFN-β. FAdVs do not have VA-RNAs; therefore, if FAdV-9 dUTPase indeed functions like a PAMP, it would help our understanding of the mechanism of the FAdVinduced immune response. Further studies are required to test this hypothesis,. In conclusion, we verified that FAdV-9 ORF1 encodes a functional dUTPase enzyme and characterized its molecular features including transcription and translation patterns and cellular localization. We also demonstrated that, although FAdV-9 dUTPase did not affect virus replication in vitro, it contributed to the up-regulation of type I interferons in vitro. Our data lays the foundation for further exploration on the mechanism of the host immune response against FAdV infection. Acknowledgements Li Deng was a recipient of China Scholarship Council PhD fellowship. This work was supported by the Natural Sciences and Engineering Research Council of Canada, the Canadian Poultry Research Council, and the Ontario Ministry of Agriculture, Food and Rural Affairs. 77 Table 3.1 Primers for generating mutant viruses Letters in bold indicate nucleotides that were changed; the letters in italic represent enzyme sites; the underlined letters indicate stop codons. Length Primer Name Sequence (5’-3’) Functions (bp) pleft-F GCACAGTCCCAATGGCTT 18 Amplify the leftend FAdV-9 pleft-R AAATTCTGGTCCGTTACCGA 20 genome SDMORF1-1st- TCCCGTTTAGGTGAGGAGATGTAATTT Introduce one stop 50 For AAATCCGGTTGCCCCCCGACTCC codon and one SwaI site right SDMORF1-1st- GGAGTCGGGGGGCAACCGGATTTAAA after the start 50 Rev TTACATCTCCTCACCTAAACGGGA codon of ORF1 SDMORF1AGATGTAATTTAAATCCGGTTGACCC Introduce another 50 2nd-For TAGACTCCACCGGTAAAGCTGCTG two stop codons further away from SDMORF1CAGCAGCTTTACCGGTGGAGTCTAGG the start codon of 50 2nd-Rev GTCAACCGGATTTAAATTACATCT ORF1 TCCCGTTTAGGTGAGGAGATGTATCC N-HA-ORF1ATATGATGTTCCAGATTATGCTTCTT 68 For Fuse HA tag at the TCGATTCCGGTTGCCC N-terminus of GGGCAACCGGAATCGAAAGAAGCAT ORF1 N-HA-ORF1AATCTGGAACATCATATGGATACAT 68 Rev CTCCTCACCTAAACGGGA Sequence the leftSeq pleftGACCCTGGTCGGAAACGAT 19 end of FAdV-9 FAdV-9 genome Determine the transcription HA-tag-For TATCCATATGATGTTCCAGATTATGCT 27 profile of HAORF1 CAGGAATTCTCTTTCGATTCCGGTTGC Clone ORF1 into ORF1-For 30 CCC pET-28a plasmid Clone ORF1 into pET-28a plasmid; GCTGCGGCCGCCTAAGAAAAGGAGG determine ORF1-Rev 30 AGGGA transcription profile of HAORF1 78 Table 3.2 Primers used for cytokines for qRT-PCR Name Sequence Length (bp) β-actin For β-actin Rev IFN-α For IFN-α Rev IFN-β For IFN-β Rev IFN-γ For IFN-γ Rev IL-10 For IL-10 Rev CAACACAGTGCTGTCTGGTGGTA ATCGTACTCCTGCTTGCTGATCC ATCCTGCTGCTCACGCTCCTTCT GGTGTTGCTGGTGTCCAGGATG GCCTCCAGCTCCTTCAGAATACG CTGGATCTGGTTGAGGAGGCTGT ACACTGACAAGTCAAAGCCGCACA AGTCGTTCATCGGGAGCTTGGC AGCAGATCAAGGAGACGTTC ATCAGCAGGTACTCCTCGAT 23 23 23 22 23 23 24 22 20 20 79 Mastadenovirus Supplementary Table S1. Pairwise identities of dUTPase amino acid sequences of adenoviruses. Twenty adenovirus dUTPase amino acid sequences were obtained from NCBI and aligned with Genious 8.0 program. The horizontal and vertical lines divides mastadenoviruses and aviadenoviruses. The highest and lowest identity in each group is in bold. HAdV-A HAdV-B HAdV-C HAdV-D HAdV-E HAdV-G SAdV-A PAdV-A PAdV-C BAdV-A Aviadenovirusa TSAdV-A FAdV-A FAdV-B FAdV-C FAdV-D FAdV-E GoAdV-A TAdV-B TAdV-C TAdV-D HA dVA HA dVB HA dVC HA dVD HA dVE HA dVG 50.0 52.0 48.0 45.2 52.4 51.3 14.3 25.8 22.8 17.5 24.2 22.7 25.8 25.0 25.8 22.7 25.8 25.8 21.9 46.0 44.8 68.8 50.0 45.8 16.7 26.0 24.4 19.8 22.8 21.3 22.8 22.8 22.0 19.7 22.8 21.3 21.3 44.8 46.8 46.0 44.2 18.3 23.4 19.7 18.3 22.5 18.6 20.9 20.9 20.2 20.9 21.7 20.2 20.9 43.2 40.8 40.2 22.4 20.6 18.3 18.3 21.4 22.2 26.2 22.2 22.2 21.4 23.8 22.2 19.8 50.0 39.0 17.5 23.6 22.0 22.2 23.6 22.8 24.4 24.4 23.6 18.9 21.3 23.6 21.3 41.5 17.5 29.1 26.0 21.4 27.6 23.6 25.2 26.0 26.0 24.4 26.8 24.4 25.2 SAd V-A PAd V-A PAd V-C 13.4 23.3 23.5 17.8 25.6 23.1 24.8 24.0 26.4 23.1 26.4 25.6 24.0 32.5 36.5 34.4 28.6 29.4 27.0 32.5 30.2 31.0 26.2 33.3 28.6 70.9 49.7 59.9 55.6 58.5 56.3 52.8 46.5 47.9 57.0 55.6 a BA dVA TS Ad V-A 48.6 53.5 51.4 51.4 53.5 50.7 45.8 46.5 54.9 52.8 52.3 47.0 47.0 49.0 47.0 50.0 43.0 45.6 53.0 FAd V-A FAd V-B FAd V-C FAd V-D FAd V-E 56.7 51.2 58.3 60.8 53.5 45.2 59.6 72.7 48.4 68.2 67.5 46.8 51.9 67.5 53.8 55.7 57.3 45.4 56.8 52.2 49.7 85.0 50.7 53.0 73.9 55.7 50.7 59.1 73.9 56.1 Go Ad V-A TA dVB TA dVC 46.8 51.0 53.8 55.4 44.3 55.6 TA dVD Genomes of TAdV-C and TAdV-D have been recently fully sequenced (Marek et al., 2014a). Though not officially accepted yet, TAdV-C and TAdV-D were suggested to belong to the genus Aviadenovirus. Abbreviations: HAdV: human adenovirus; SAdV: Simian adenovirus; PAdV: Porcine adenovirus; BAdV: Bovine adenovirus; TSAdV: Tree shrew adenovirus; FAdV: fowl adenovirus; GoAdV: Goose adenovirus; TAdV: Turkey adenovirus 80 Supplementary Figure S1 Generation of the mutant viruses. (1) FAdmid pPacFAdV-9 was linearized through digestion with SgfI for later use; (2) the intermediate construct pGEM-T-pleft was generated by cloning the left-end of the FAdV9 genome onto the pGEM-T easy vector by primers pleft-For and pleft-Rev; (3) two rounds of site-directed mutagenesis were conducted to introduce the unique SwaI site and 3 stop codons in ORF1 to generate the mutated intermediate construct ORF1-SDM-pleft; (4) linear mutated left-end of FAdV-9 genome was obtained through amplification of ORF1-SDM-pleft with primers pleft-For and pleft-Rev for later use; (5) homologous recombination was carried out with products of (1) and (4) in E.coli BJ5183. Positive recombinant construct was transformed and propagated in E.coli DH5ɑ, followed by PacI digestion and transfection into CH-SAH cells. The mutant virus was harvested following observation of cytopathic effects. 81 Figure 3.1 Multiple alignment of amino acid sequences of dUTPase homologs. Thirty identified or putative dUTPase amino acid sequences were aligned by ClustalW. The alignment was conducted by ClastW through Geneious 8.0 program. Letters in different shadow indicates different similarities among all the sequences as below: black (100%), dark grey (80 to 100%), light grey (60 to 80%), and no shadow (less than 60%). The five conserved motifs 1-5 are indicated on top of the sequences. 82 Figure 3.2 PCR-based dUTPase enzyme activity assay. PCR amplification of ORF1 was conducted with a mixture of individual dATP, dGTP, dCTP, and either dTTP (lanes 1, 3, 5) or dUTP (lanes 2, 4, 6, 7). In the presence of dUTPase reaction buffer, dTTP or dUTP used for the PCR was incubated with water (V/V=1:1, lanes 3 and 4), or His-tagged protein (V/V=1:1, lanes 5 and 7), respectively for 24 hours at 37 °C. Heat-inactivated His-tagged ORF1 protein (95 °C, 5 mins) was also used as a negative control (lane 6). All PCR products were run in a 1.0% agarose gel. 83 Figure 3.3 Viral DNA accumulation curve and one-step growth curve. CH-SAH cells were infected at an MOI of 5 with ORF1stop, resORF1 or wtFAdV-9. At the indicated time points, (A) cells were collected for DNA extraction, followed by qPCR of viral DNA genome; (B) cells with supernatant were collected for virus titration. 84 Figure 3.4 Transcription and protein expression profiles of ORF1. CH-SAH cells were infected with FAdV-9HA-ORF1 at an MOI of 5. At the indicated time points, RNA and cell lysates were collected for RT-PCR (A) and Western blot (B), respectively. The HA-tag-For and ORF1-Rev for amplification of ORF1 template are described in Table 1. PCR products were run in an 1.0% agarose gel. Protein was run in a 12% SDS-PAGE gel and transferred to PVDF membrane (GE Healthcare). Mouse antiHA monoclonal antibody and horseradish peroxidase-conjugated goat anti-mouse IgG antibody were used at 1:750 and 1:10, 000 dilutions, respectively. Anti-β-actin antibody was used to indicate equivalent loading. 85 Figure 3.5 Cellular localization of ORF1 protein Immunofluorescence of HA-ORF1 in CH-SAH cells infected with FAdV-9HA-ORF1 virus at an MOI of 5. At the indicated time points, cells were fixed and permeabilized , followed by incubation with mouse anti-HA monoclonal antibody (1:750, Sigma-Aldrich) and DyLight 549-conjugated goat anti-mouse IgG antibody (1:10 000, Jackson ImmunoResearch). The ProLong® Gold antifade reagents with DAPI (Life Technologies) was added and incubated for 10 minutes prior to analysis with confocal microscopy. 86 Figure 3.6 Cytokine mRNA expressions in CH-SAH cells CH-SAH cells were infected with ORF1stop virus and cytokine mRNA expressions were analyzed at different time points relative to β-actin. Mock-infected cells or cells infected with resORF1 and wtFAdV-9 were used as controls. RNA were collected for qRT-PCT with gene-specific primers as described above. Data was analyzed with Graphpad prism 6.0. Two-way ANOVA was used for the statistical analysis between any of the two groups. * denotes P<0.05 and ** denotes 0.001<P<0.01 when compared to mock control. Comparison between wtFAdV-9 and ORF1stop-infected group is indicated with brackets above the bars. 87 Figure 3.7 Cytokine mRNA expressions in Celi cells Celi cells were infected with ORF1stop virus and cytokine mRNA expressions were analyzed at different time points relative to β-actin. Mock-infected cells or cells infected with resORF1 and wtFAdV-9 were used as controls. RNA were collected for qRT-PCT with gene-specific primers as described for Fig. 6. Data was analyzed with Graphpad prism 6.0. Two-way ANOVA was used for the statistical analysis between any of the two groups. * denotes P<0.05 and ** denotes 0.001<P<0.01 when compared to mock control. Comparison between wtFAdV-9 and ORF1stop-infected group is indicated with brackets above the bars. 88 Chapter 4. Fowl adenovirus 9 dUTPase plays a role in virus replication in vivo and in the regulation of the host immune response Li Deng, Bryan Griffin, Yanlong Pei, David Leishman, Betty-Anne McBey, Shayan Sharif, Éva Nagy* Department of Pathobiology, University of Guelph, Guelph, Canada * Corresponding author: Éva Nagy Tel.: +1-(519) 824-4120 Ext. 54783 Fax: +1-(519) 824-5930 E-mail address: [email protected] Mail address: Department of Pathobiology, Ontario Veterinary College, University of Guelph, 50 Stone Road East, Guelph, ON, Canada, N1G 2W1 Author’s contributions LD designed all experiments and performed the majority of them, conducted the data analysis and wrote the first draft of the manuscript. BG, YP, DL and B-AM helped with the chicken experiment. DL and B-AM provided technical assistance. SS and ÉN provided guidance during the experiments. ÉN provided critical review for the manuscript. To be submitted to Viral Immunology 89 Abstract Fowl adenoviruses (FAdVs) are distributed worldwide in poultry farms. Some of FAdVs are important causative agents of inclusion body hepatitis and hydropericardium syndrome that cause significant economic losses to the poultry industry. Previously, we identified FAdV-9 ORF1 as a deoxyuridine 5′-triphosphate pyrophosphatase (dUTPase) enzyme, which is not required for virus replication in vitro but contributes to the upregulation of type I interferons. In the present study, we compared virus replication in vivo and the host immune response in chickens orally inoculated with ORF1stop (dUTPase knockout virus), resORF1 (the rescued revertant of ORF1stop), and wtFAdV-9, respectively. Our data showed that replication of ORF1stop was delayed on days 1 and 3 post inoculation (p.i.), as evidenced by significantly less virus shedding in feces and lower viral loads in tissues, compared to those of wtFAdV-9 group. However, there was no significant difference between the two groups from 5 d.p.i. to the end of the study. Moreover, we found that there were significant differences between ORF1stop and wtFAdV-9 in terms of the induction of cytokine gene mRNA expression in tissues and IgG antibody responses, suggesting the important roles of dUTPase in modulating the host immune response. Our study clarified the roles of FAdV-9 dUTPase in vivo that are helpful for better understanding the virus-host interaction. Key words fowl adenovirus 9, dUTPase, in vivo, host immune response, cytokine gene expression 90 Introduction Fowl adenoviruses (FAdVs), belonging to the genus Aviadenovirus in the family Adenoviridae, are ubiquitous infectious agents in poultry farms and are distributed worldwide. Although most of them are non-pathogenic or associated with only mild clinical signs, some FAdV strains are associated with a number of diseases including inclusion body hepatitis (IBH) and hydropericardium syndrome (HPS) (Hess, 2013). There are 12 FAdV serotypes, which are subgrouped into five species (Fowl adenovirus A to Fowl adenovirus E), and each consists of one or more serotypes. The genomes of the representatives of all five speices have been fully sequenced, such as FAdV-1, -4, -5, -8 and -9 (Chiocca et al., 1996; Griffin and Nagy, 2011; Grgić et al., 2011; Marek et al., 2013; Ojkić and Nagy, 2000), which are considerably larger than those of mastadenoviruses. FAdVs are promising alternatives to human adenovirus as vaccine vectors (Francois et al., 2004; Johnson et al., 2003; Corredor and Nagy, 2010b). Previously we identified ORF1 of FAdV-9 as a functional dUTPase enzyme (unpublished data), which was the first early gene functionally identified in FAdV-9. dUTPase is a ubiquitous enzyme that catalyzes the cleavage of dUTP to dUMP and pyrophosphate (PPi), maintaining a low dUTP/dTTP ratio in cells and preventing the misincorporation of deoxyuridine into DNA. To date, the dUTPase gene has been characterized in many viruses, such as adenovirus (Weiss et al., 1997), herpesvirus (Glaser et al., 2006), retrovirus (Payne and Elder, 2001), African swine fever virus (Oliveros et al., 1999), and poxvirus (Cottone et al., 2002). In addition, a dUTPase knockout virus ORF1stop and its rescued revertant resORF1 were previously generated in our laboratory for functional studies. Our data showed that FAdV-9 dUTPase was not essential for virus replication in 91 vitro as the knockout of dUTPase did not affect either the viral DNA replication or the production of progeny virus in CH-SAH cells and primary chicken embryo liver (Celi) cells. However, it could be not ruled out that FAdV-9 dUTPase might be important for replication in vivo. For herpes simplex virus (HSV), the deletion of dUTPase did not affect virus replication in vitro but resulted in reduced replication in the central nervous system of mice (Pyles et al., 1992). Leang et al. (2011) also demonstrated that dUTPase of murid herpesvirus 68 (MHV-68) is necessary for efficient virus replication in the lungs of infected mice. Therefore, it is worth investigating the roles of FAdV-9 dUTPase in vivo. It has been demonstrated that the dUTPase of Epstein-Barr virus (EBV) up-regulates several pro-inflammatory cytokines including TNF-α, IL-1β, IL-8, IL-6, and IL-10 in unstimulated peripheral blood mononuclear cells (PBMCs) (Glaser et al., 2006). Another group showed that the dUTPase of the murine gammaherpesvirus 68 (MHV-68) can inhibit type I interferon signaling (Leang et al., 2011). We demonstrated earlier that although FAdV-9 dUTPase is not required for virus replication in vitro, it contributes to the up-regulation of the mRNA expressions of type I interferons (unpublished data). However, how FAdV-9 dUTPase functions in vivo remains unknown. The aims of this study were to explore the functions of FAdV-9 dUTPase in virus replication in orally inoculated chickens and its roles in modulating the host immune response against virus infection. Clocal swabs were collected to test virus shedding among virus-infected groups. Due to the relatively high tropism of the virus in chickens (Ojkić and Nagy, 2003), spleen, liver and cecal tonsil were collected for the determination of viral loads and cytokine gene expression in tissues. Serum samples were collected to compare host antibody responses induced by all viruses. 92 Materials and Methods Viruses and cells ORF1stop, the dUTPase knockout virus, and resORF1, the rescued revertant, were both generated through site-directed mutagenesis and homologous recombination, based on a FAdmid clone pPacFAdV-9 that contains the whole FAdV-9 genome (Ojkić and Nagy, 2001). These two viruses, together with wild type virus, wtFAdV-9, were propagated and titrated in chicken hepatoma cells (CH-SAH) as described previously (Alexander et al., 1998). CH-SAH cells were maintained in Dulbecco’s modified Eagle’s medium/nutrient mixture F-12 Ham (DMEM-F12), supplemented with 10% non-heat-inactivated fetal bovine serum (FBS), 2 mM L-glutamine, 100 U/ml penicillin and 100 μg/ml streptomycin. Experimental animals Specific pathogen free (SPF) white Leghorn chickens were obtained from the Canadian Food Inspection Agency (Ottawa, ON, Canada) and housed in the Isolation Unit of University of Guelph throughout the study. All animal experiments were reviewed and approved by the Animal Care Committee of the University of Guelph according to the Guide to the Care and Use of Experimental Animals of the Canadian Council on Animal Care. Animal experimental design To explore the functions of FAdV-9 dUTPase in virus replication in vivo and modulating the host immune responses, a chicken experiment was carried out. One hundred and sixty four day-old chickens were divided into four groups (n≥40). At the age of 10 days, 93 chickens were orally inoculated with (1) PBS, served as a mock control, (2) 2×107 p.f.u./chick ORF1stop, (3) 2×107 p.f.u./chick wtFAdV-9, and (4) 2×107 p.f.u./chick resORF1, respectively. At 0, 1, 3, 5, 7, 10, 14, 21, 28 and 35 days post inoculation (d.p.i.), cloacal swabs were collected from all birds in 1 ml PBS supplemented with 100 U/ml penicillin and 100 μg/ml streptomycin to determine the virus titer in feces. At 10 hours p.i. (h.p.i.), 1, 3, 5, 7, 14 and 35 d.p.i., liver and cecal tonsil were collected from 5 birds in each group to determine the viral load in tissues; spleen, another portion of liver and cecal tonsil were collected from 5 birds in each group to investigate the cytokine gene expression in tissues, including IFN-α, IFN-β, IFN-γ, IL-10, and IL-8. In addition, serum samples were collected at 0, 7, 14, 21, 28, and 35 d.p.i. to test FAdV-specific IgG antibodies. DNA and RNA extraction Viral DNA was extracted from tissues stored at -80ºC with QIAamp DNA Mini Kit (QIAGEN Inc) following the manufacturer’s instructions. The DNA concentration was measured using a NanoDrop 2000 spectrophotometer. RNA was extracted with Trizol (Invitrogen) from tissues stored in RNAlater as described previously (Deng et al., 2013), prior to digestion with DNase I (Fermentas) and quantification with NanoDrop 2000 spectrophotometer. qPCR and qRT-PCR To determine the viral load in tissues, viral DNA was quantified by qPCR with LightCycler® 480 SYBR Green I Master Kit (Roche Diagnostics) and FAdV-9-specific primers as described previously (Romanova et al., 2009). To determine the cytokine gene expression in tissues, cDNA was first synthesized from total RNA with the 94 SuperScriptTM II Reverse Trancriptase (Invitrogen) and random primer. The 1:10 diluted cDNA was used for qPCR with LightCycler® 480 SYBR Green I Master Kit (Roche Diagnostics) and gene-specific primers as described previously (Abdul-Careem et al., 2006, 2007). Enzyme-linked immunosorbent assay (ELISA) Heat-inactivated serum samples were diluted 1:100 in wash buffer (0.05% Tween 20 in PBS) and used for ELISA as previously described (Ojkić and Nagy, 2003). The sample to positive (S/P) ratio was used to indicate the antibody level (Ojkić and Nagy, 2003). A serum sample from a previous chicken trial, collected from wtFAdV-9-infected chicken at 28 d.p.i, was used as a positive control. Statistical analysis Statistical analyses were performed using GraphPad Prism 6.0 software (San Diego, CA). A two-way analysis of variance (ANOVA) was used to determine significant differences between any two groups. The critical level for significance was set at a P value of <0.05. The data were expressed as mean ± standard error of the mean (SEM). Results Virus shedding To compare virus shedding in chickens among ORF1stop, resORF1 and wtFAdV-9 groups, cloacal swabs were collected for the determination of virus titers in feces by plaque assay. No virus was detected in any samples from chickens prior to inoculation and mock-infected group (data not shown), and the virus titers for other groups are summarized in Table 4.1. Virus was detected in chickens of all three virus-infected 95 groups from day 1 to day 14 p.i., but no virus was detected at days 21, 28 and 35 p.i.. The peak of virus shedding in all three groups occurred at day 5 p.i., with the titer around 5.0×104 p.f.u./ml. At days 1 and 3 p.i., the virus titers in the samples of ORF1stop group were significantly lower (P<0.05) than that of wtFAdV-9 group (approximately 4 and 14 fold at 1 and 3 d.p.i., respectively), while no significant difference was found at later days (5 to 14 d.p.i.). The virus titers of samples from resORF1 group were quite similar to that of wtFAdV-9 throughout the experiment. Viral loads in tissues To determine the viral loads in tissues, the viral genome copy numbers in liver and cecal tonsil were quantified by real-time PCR with specific FAdV-9 primers. No virus was detected in samples from chickens before inoculation and mock-infected groups (data not shown), and the results of other groups are summarized in Table 4.2. As shown in Table 4.2, virus was detected in cecal tonsils of all groups at 10 h.p.i., with the viral genome copy numbers around 3.0×104/μg total tissue DNA. However, no virus was detected in livers of any group at this time point. At days 1 and 3 p.i., the viral genome copy numbers were significantly lower (P<0.05) in both liver and cecal tonsil of ORF1stop group than those in wtFAdV-9 group (4.3 fold in liver and 3.3 fold in cecal tonsil at 1 d.p.i.; 3.8 fold in liver and 2.5 fold in cecal tonsil at 3 d.p.i.). At days 5 and 7 p.i., although wtFAdV-9 group had generally higher viral genome copy numbers in both liver and cecal tonsil than ORF1stop group, there was no significant difference between the two groups. The resORF1 group showed very similar results with wtFAdV-9, and there was no significant difference between these two groups at all time points. 96 Antibody response Antibody levels against FAdV-9 were determined to investigate the role of ORF1 in modulating the immune response. No antibodies were detected in samples from any group prior to inoculation (0 d.p.i.). The results of other day points, based on the sample to positive (S/P) ratio, are shown in Fig. 4.1. Throughout the experiment, no antibodies were detected in the mock group, while antibody levels increased in all virus-infected groups from 1 week p.i. until the end of the experiment at 35 d.p.i.. Except at 7 d.p.i., the antibody level (S/P ratio) in ORF1stop group was significantly lower than that of wtFAdV-9, while the resORF1 group had very similar antibody response with wtFAdV-9 throughout the study. Cytokine gene expression in tissues In order to determine whether FAdV-9 dUTPase could modulate the innate immune response as it does in vitro, the mRNA expression of IFN-α, IFN-β, IFN-γ, and IL-10 was investigated in the spleen, liver, and cecal tonsil by real-time PCR. The mRNA expression of IL-8 was also followed. In spleens, as is shown in Fig. 4.2, compared to mock control, all viruses resulted in an increase of the mRNA expression of IFN-α at all time points except for 10 h.p.i.. At 1 and 3 d.p.i. the increase was significantly different with mock control for both resORF1 and wtFAdV-9 but not for ORF1stop. At these time points, there were significant differences between ORF1stop and wtFAdV-9. No significant difference was found between any two groups at 5 and 7 d.p.i.. For the mRNA expression of IFN-β, in spite of some increase in all virus-infected groups, no significant difference was found between any two groups at all time points. Similar to IFN-α, the mRNA expression of IFN-γ was upregulated by all 97 three viruses at all time points, compared to mock control. Significant increase was found in both resORF1 and wtFAdV-9 groups throughout the study. However, no significant difference was found in the ORF1stop group. There were significant differences between wtFAdV-9 and ORF1stop at 1, 5 and 7 d.p.i. as well. The mRNA expression of IL-10 was relatively stable, although increased expression was found at 3 d.p.i. and decreased expression was found at 5 and 7 d.p.i. However, differences between any two groups were not significant. In terms of the mRNA expression of IL-8, in spite of some upregulation by all three viruses, there was no significant difference between any two groups at all time points. The mRNA expression of IFN-α, IFN-β, IFN-γ, IL-10, and IL-8 in liver samples is presented in Fig. 4.3. All viruses induced higher IFN-α mRNA expression compared to mock control at 1, 3, 5 and 7 d.p.i.. At 5 d.p.i., the increase compared to mock found in both resORF1 and wtFAdV-9 groups but not in ORF1stop was significant. There was no significant difference between any two virus-infected groups at all time points. Similarly, the mRNA expression of IFN-β was upregulated by all viruses throughout the study. The increase in both resORF1 and wtFAdV-9 groups but not in ORF1stop was significant at 5 d.p.i.. Also at this time point, a significant difference was found between ORF1stop and wtFAdV-9. For the mRNA expression of IFN-γ, all viruses upregulated its expression compared to mock control at all time points. Both resORF1 and wtFAdV-9 had significantly higher level of IFN-γ mRNA expression than mock control at 3 and 5 d.p.i.. In addition, there was a significant difference between ORF1stop and wtFAdV-9 at 3 d.p.i.. The production of IL-10 mRNA expression by ORF1stop was very close to that by mock control at all time points. However, compared to mock control, there were 98 significant decreases of IL-10 mRNA expression in both resORF1 and wtFAdV-9 groups at 10 h.p.i., 1, 3, and 5 d.p.i.. At 7 d.p.i., decreased IL-10 mRNA expression was also found in both resORF1 and wtFAdV-9 groups but not in ORF1stop, although there was no significant difference between any two groups. The mRNA expression of IL-8 was upregulated by all viruses at all time points, compared to mock control. Only the increase in both resORF1 and wtFAdV-9 groups but not in ORF1stop group at 1 d.p.i. was found to be significant. The mRNA expression of IFN-α, IFN-β, IFN-γ, IL-10, and IL-8 in cecal tonsil samples is presented in Fig. 4.4. Compared to mock control, there was increased mRNA expression of all cytokines including IFN-α, IFN-β, IFN-γ, IL-10, and IL-8 in all virus-infected groups throughout the study (except for a slight decrease for IFN-γ by ORF1stop at 3 d.p.i.), Although not significant, resORF1 and wtFAdV-9 had higher level of IFN-α mRNA expression at 3, 5, and 7 d.p.i. than ORF1stop. Increased mRNA expression of IFN-β was found in all virus-infected groups. In addition, resORF1 and wtFAdV-9 but not ORF1stop showed significant increase of the mRNA expression of IFN-β at 1 d.p.i., compared to mock control. resORF1 and wtFAdV-9 had generally higher level of IFN-β mRNA expression than ORF1stop at 1, 3, 5, and 7 d.p.i.. Increased IFN-γ mRNA expression was found in all virus-infected groups at all time points (except for ORF1stop at 1 d.p.i.), while significant increase was only found at 7 d.p.i., compared to mock control. There was no significant difference in the mRNA expression of IL-8 between any two groups at all time points, in spite of some up-regulation by all viruses. 99 Discussion The early genes of human adenoviruses have been extensively studied (reviewed in Chakraborty and Tansey, 2009; Horwitz, 2004; Weitzman, 2005), while only three early genes are functionally identified in fowl adenoviruses (Chiocca et al., 1997; Lehrmann and Cotten, 1999; Weiss et al., 1997). We previously characterized FAdV-9 ORF1 as a dUTPase enzyme, which is not required for virus replication but is able to upregulate the expression of type I interferons in CH-SAH cells and Celi cells (unpublished data). In the present study, we demonstrated that ORF1stop, the dUTPase knockout virus, did not replicate as efficiently as the wild type virus at the early phase of infection (up to day 3 p.i.). However, no significant difference was observed between the two groups from 5 d.p.i. to the end of the study. Moreover, ORF1stop induced significantly less expression of IFN-α, IFN-β, IFN-γ and significantly more expression of IL-10 in tissues, and elicited significantly lower antibody response, compared to wtFAdV-9, suggesting that FAdV-9 dUTPase plays important roles in virus replication in vivo and in modulating the host immune response. A number of researchers have reported the roles of viral dUTPase in virus replication both in vitro and in vivo. For the in vitro studies, the most recognized conclusion is that the viral dUTPase is not essential in dividing cells but is required for efficient virus replication in non-dividing host macrophages. This has been demonstrated for the dUTPase of African swine fever virus (Oliveros et al., 1999) and equine infectious anemia virus (Threadgill et al., 1993). Consistent with these results, our data also demonstrated that FAdV-9 dUTPase did not affect virus replication in CH-SAH cells and Celi cells (unpublished data). However, Voronin and co-workers (2014) showed that the 100 dUTPase of bovine immunodeficiency virus is critical for the virus replication even in dividing cells, although synthesis of the viral cDNA was not altered. On the other hand, different results were reported for the roles of viral dUTPase in vivo. Pyles et al. (1992) showed that the deletion of dUTPase of HSV resulted in reduced replication of virus in the central nervous system of mice. In contrast, the dUTPase of vaccinia virus is not required for virus replication in vivo (Prichard et al., 2008). In the present study, we showed that the replication of ORF1stop, the dUTPase knockout virus, was retarded in chickens at the early phase of infection (1 and 3 d.p.i.) but the overall virus level was the same as wild type from 5 d.p.i. to the end of the experiment. This suggests that FAdV-9 dUTPase has an important role in virus replication in vivo, at least at the early phase of infection. Interestingly, Francois et al. (2001) demonstrated that the transfection of the dUTPase-deficient FAdV-1 cosmid did not produce cytopathic effects (CPE) in chicken hepatocarcinoma cells (LMH), and the infection with the transfected cell lysates produced a CPE later than the wild type FAdV-1 cosmid-transfected cell lysates. Along with our data, this suggests that the lack of dUTPase might be able to retard virus replication among all FAdVs. The antibody level in wtFAdV-9 group increased throughout the experiment, which is consistent with the result of our previous study (Deng et al., 2013). The antibody level in ORF1stop group is significantly lower than that in wtFAdV-9 group at all time points except for 7 d.p.i., suggestting that FAdV-9 dUTPase plays significant roles in the regulation of the host antibody response. This is possibly related to the ability of FAdV-9 dUTPase in up-regulating the expression of type I interferons. Theofilopoulos et al. (2005) reported that type I IFNs can promote antibody-mediated immune responses through 101 promoting adaptive T and B cell responses. Zhu et al. (2007b) showed that type I IFNs induced upon adenoviral infection are critical for the formation virus-specific IgG2a and IgG2b isotypes, as they mediate the activation of early B cells, formation of the germinal center, Ig isotype switching and plasma cell differentiation. They also found that Ig isotype switching and the formation of neutralizing Ab rely on type I IFN signaling in both B and CD4 T cells. Other studies have also shown that antibody response to adenoviral vectors is CD4 Th cell dependent (Yang et al., 1995, 1996). Similarly, Swanson and co-workers (2010) demonstrated that type I IFNs promote production of antigen-specific IgG2c by follicular B cells, which therefore enhances the magnitude and quality of the T cell-independent type 2 antibody responses. Previously, we demonstrated that despite inducing a significantly higher antibody response, compared to the mock control, a multiORF-deleted virus FAdV-9Δ4, lacking ORFs 0, 1, 1A, 1B, 1C and 2, induced significantly lower level of antibody than the wild type virus (Corredor and Nagy, 2010a; Deng et al., 2013). Together with our data, this suggests that of the six deleted genes, dUTPase, if not the only, plays significant roles in the mediation of antibody response. Indeed, ORF1C shares some amino acid identities to the E5 oncoprotein of bovine papillomavirus type 1 (Ojkić and Nagy, 2000) and is hypothesized to downregulate major histocompatibility complex class I molecules (Corredor and Nagy, 2010a). However, to determine if there are more genes other than dUTPase (ORF1) involved in the mediation of antibody response, more research has to be done. Among numerous innate immune cytokines, type I interferons are pleiotropic cytokines that are capable of exerting direct antiviral immune response against virus infection. Induction of type I IFNs is part of the innate immune response to adenovirus infection. 102 We previously found that FAdV-9 dUTPase is important for the up-regulation of type I IFNs in CH-SAH cells and Celi cells. In this study, the mRNA expression of IFN-α and IFN-β was investigated in spleen, liver and cecal tonsil. According to our data, the dUTPase knockout virus, ORF1stop, induced significantly less mRNA expression of IFN-α in spleen at 1 and 3 d.p.i., and significantly less mRNA expression of IFN-β in liver at 5 d.p.i., compared to the wild type virus. This indicates that the role of FAdV-9 dUTPase in up-regulating type I IFNs is not limited to in vitro, but also applies to in vivo. IFN-γ is the only type II interferon that plays key roles in modulating immunity to infectious diseases. In our in vitro study, in which both CH-SAH cells and Celi cells were infected with ORF1stop or wtFAdV-9, no significant difference was found in terms of the mRNA expression of IFN-γ between these two groups. However, the in vivo study showed that there was significantly less expression of IFN-γ in livers of ORF1stop group than that of wtFAdV-9 group at 3 d.p.i.; so was the case in spleens at 1, 5, and 7 d.p.i.. This could be due to that organs like spleen and liver have more types of cells capable of producing INF-γ, compared to a single type of cell lines. Overall, our data demonstrated that FAdV-9 dUTPase retarded the virus replication at the early stage of the infection and played important roles in modulating the host immune response against virus infection as well. Our study provides useful data that will shed more lights on the mechanism of the host immune response against the fowl adenovirus infection. Acknowledgement Li Deng was a recipient of China Scholarship Council PhD fellowship. This work was supported by the Natural Sciences and Engineering Research Council of Canada, the 103 Canadian Poultry Research Council, and the Ontario Ministry of Agriculture, Food and Rural Affairs. The authors wish to thank the personnel of the Isolation Unit for their professional animal care and assistance. 104 Table 4.1 Virus titers in the feces of infected chickens Days ORF1stop resORF1 wtFAdV-9 Titer (pfu/ml) %a Titer (pfu/ml) % Titer (pfu/ml) % 1 1.6×103±3.5×102 100 5.7×103±1.0×103 100 6.4×103±1.2×103 100 3 3.6×103±5.5×102 100 4.8×104±2.1×104 100 4.9×104±1.1×104 100 5 5.0×104±1.1×104 100 5.2×104±1.4×104 100 5.5×104±1.1×104 100 7 5.4×103±1.9×103 100 4.9×103±1.4×103 100 5.3×103±1.3×103 100 10 7.3×102±4.4×102 66.7 7.9×102±1.9×102 85.7 7.1×102±1.9×102 80 14 3.4×101±5.8×100 33.3 3.8×101±4.4×100 71.4 3.8×101±7.1×100 66.7 21 NDb 0 ND 0 ND 0 28 ND 0 ND 0 ND 0 35 ND 0 ND 0 ND 0 p.i. a percentage of chickens shedding viruses b ND = not detected 105 Table 4.2 Viral genome copy number in tissues Viral genome copy number in liver (L) and cecal tonsil (CT) of chickens orally inoculated with ORF1stop, resORF1 and wtFAdV-9, expressed as copies/μg total tissue DNA. Days p.i. Tissue ORF1stop 10 hpi L NDa 1 3 5 7 a resORF1 wtFAdV-9 ND 3 3 ND 3 CT 2.74×10 ±1.6×10 2.88×10 ±7.8×10 3.12×103±7.4×102 L 4.91×101±9.8×100 1.90×102±3.7×101 2.10×102±6.2×101 CT 2.18×105±4.1×104 6.37×105±1.3×105 7.16×105±1.9×105 L 1.31×102±4.2×101 5.35×102±2.2×102 4.92×102±1.1×102 CT 3.27×105±1.4×105 8.32×105±3.5×105 8.11×105±2.8×105 L 1.15×103±3.6×102 1.56×103±2.9×102 1.46×103±4.8×102 CT 2.61×105±8.2×104 3.26×105±9.7×104 3.20×105±1.0×105 L 3.89×102±1.1×102 5.75×102±2.3×102 5.29×102±1.8×102 CT 2.0×104±9.6×103 2.76×104±7.1×103 3.19×104±1.7×104 ND = not detected 106 2 Figure 4.1 FAdV-specific IgG antibody response in chickens. Chickens were orally inoculated with ORF1stop, or resORF1, or wtFAdV-9, or mockinfected. Heat-inactivated serum samples were diluted 1:100 in wash buffer (0.05% Tween 20 in PBS) and used for ELISA. The optical density (OD) value was read in a Bio-Tek ELISA microplate reader at 405 nm. The sample to positive (S/P) ratio was used to indicate the antibody level. A serum sample that is collected from wtFAdV-9-infected chicken at 28 d.p.i in a previous chicken trial was used as a positive control. Statistical significance was indicated by * (P<0.05), ** (0.001<P<0.01), *** (P<0.001). Brackets above the bars indicate comparison between ORF1stop and wtFAdV-9. 107 Figure 4.2 Cytokine mRNA expression in spleen samples of chickens. Chickens were orally inoculated with ORF1stop, resORF1, or wtFAdV-9, or mockinfected. The mRNA expression of target and reference genes was quantified by qRTPCR, and is presented relative to β-actin expression and normalized to a positive plasmid calibrator. Error bars represent standard error of the means. Statistical significance was indicated by * (P<0.05) or ** (0.001<P<0.01), compared to mock control. Brackets above the bars indicate comparison between ORF1stop group and wtFAdV-9 group. 108 Figure 4.3 Cytokine mRNA expression in liver samples of chickens. Chickens were orally inoculated with ORF1stop, resORF1, or wtFAdV-9, or mockinfected. The mRNA expression of target and reference genes was quantified by qRTPCR, and is presented relative to β-actin expression and normalized to a positive plasmid calibrator. Error bars represent standard error of the means. Statistical significance was indicated by * (P<0.05) or ** (0.001<P<0.01), compared to mock control. Brackets above the bars indicate comparison between ORF1stop group and wtFAdV-9 group. 109 Figure 4.4 Cytokine mRNA expression in cecal tonsil samples of chickens. Chickens were orally inoculated with ORF1stop, resORF1, or wtFAdV-9, or mockinfected. The mRNA expression of target and reference genes was quantified by qRTPCR, and is presented relative to β-actin expression and normalized to a positive plasmid calibrator. Error bars represent standard error of the means. Statistical significance was indicated by * (P<0.05) or ** (0.001<P<0.01), compared to mock control. 110 Chapter 5. General discussion It is well known that the major limitation of HAdV-based vectors for clinical use is the universal pre-existing immunity in humans (Lasaro and Ertl, 2009). To overcome this, alternative serotypes and non-human adenoviruses have been investigated for the ability to avoid acute toxicity and evade anti-AdV immune clearance. Tordo et al. (2008) and Xiang et al. (2014) used an E1-deleted canine adenoviruses type-2 (CAdV-2) and an experimental chimpanzee adenovirus vector Simian adenovirus serotype 24 (SAdV-24), respectively, to express the rabies virus glycoprotein, both of which induced high levels of anti-rabies virus neutralizing antibody and conferred protection against the rabies virus lethal challenge. Fowl adenoviruses are also investigated as vaccine vectors for avian diseases. For example, FAdV-1- and FAdV-10-based vectors have been demonstrated to induce protective immunity against IBDV (Francois et al., 2004; Sheppard et al., 1998); and a FAdV-8-based vector was studied against infectious bronchitis virus (IBV) (Johnson et al., 2003). FAdV-9, which is being studied in our laboratory, also shows promising results, as is described in more detail below. In addition to searching alternatives of HAdV vectors, the optimization of delivery routes and regimens is considered as another potential solution (Thacker et al., 2009). For example, Steitz et al. (2010) compared intratracheal, conjunctival, subcutaneous, and in ovo routes to evaluate the optimal vaccine administration for the HAdV-5-based H5N1 highly pathogenic avian influenza (HAPI) vaccine in chickens and revealed that the subcutaneous injection induces the highest humoral immune responses measurable by hemagglutination inhibition test (HI). In particular, it is demonstrated that oral administration of AdV vectors is better able to avoid systemic neutralizing antibodies 111 compared with other routes of administration (Xiang et al., 2003) and that the efficacy of oral vaccination with HAdV-5-based vectors is unaffected by pre-existing immunity in BALB/c mice (Tucker, 2008). Previous studies in our laboratory employed intramuscular inoculation as the administration route to develop FAdV-9 as a vaccine vector. However, the fecal-oral route is a more natural transmission route of FAdVs infection. In the study of Chapter 2, chickens were orally inoculated with FAdV-9Δ4 or wtFAdV-9. Results showed that wtFAdV-9 resulted in virus shedding in feces with titers similar to that of intramuscular inoculation of chickens. However, it should be noted that a higher inoculum was used in this study than that in the previous intramuscular inoculation study (1.5×107 pfu/chick versus 2.0×106 pfu/chick), with the consideration of the strong innate mucosal immunity in the gastrointestinal tract. The results showed that the highest titer in feces appeared earlier, compared to that of the intramuscular route study (4.0 ×103 pfu/ml at 5 d.p.i versus 9.6 ×103 pfu/ml at 7 d.p.i), which might indicate that the oral inoculation triggers the host immune response to clear the virus at an earlier stage than the intramuscular inoculation. In addition, the antibody response induced by both wtFAdV-9 and FAdV-9Δ4 through oral administration are similar to those by intramuscular administration, suggesting that the oral administration could be, if not better, at least competitive with the intramuscular route and easier to administer. FAdV-9 has been developed as a vaccine vector in our laboratory. It has been demonstrated that the tandem repeat region 2 (TR-2) at the right end of FAdV-9 genome is dispensable for virus propagation in vitro and suitable for insertion of foreign genes (Ojkić and Nagy, 2001). FAdV-9Δ4, a deleted virus lacking ORFs 0, 1, 1A, 1B, 1C and 2, replicates less efficiently and induces lower FAdV-specific IgG antibody level than wild 112 type virus in intramuscularly inoculated chickens (Corredor and Nagy, 2010a). These data suggest the importance of the left end genes of FAdV-9 genome in terms of virus replication and modulation of immune response. Among the six deleted ORFs, ORF1, a dUTPase homolog, is the most conserved one in all FAdV genomes, and its amino acid identities ranges from 56% to 100% among FAdV dUTPases (Corredor and Nagy, 2006). This implies that ORF1 is very likely to play important roles in the FAdV life cycle. In this study, FAdV-9 ORF1, containing 5 conserved motifs that form the active sites of the enzyme, was verified as a functional dUTPase, as demonstrated by its ability of catalyzing the hydrolysis of dUTP. dUTPase exists in all free-living organisms, as well as in many viruses. In recent years, functional studies of viral dUTPases have demonstrated that this gene is important for virus replication (Pyles et al., 1992; Glaser et al., 2006; Leang et al., 2011; Ariza et al., 2013). In the study of Chapter 3, it was demonstrated that FAdV-9 dUTPase was not required for virus replication in CH-SAH cells and Celi cells, which is consistent with the results of studies of other viral dUTPases (Oliveros et al., 1999; Threadgill et al., 1993). However, in the in vivo study of Chapter 4, ORF1stop, the dUTPase knockout virus, did not replicate as well as wild type virus in orally inoculated chickens at days 1 and 3 p.i., indicating that FAdV-9 dUTPase plays a role in virus replication in vivo. In the study of Chapter 3, the mRNA expression of INFs -α, -β, -γ, and IL-10 was investigated in CH-SAH cells and Celi cells infected with ORF1stop or wtFAdV-9. Results showed that wtFAdV-9 induced significantly higher mRNA expression of IFN-α and IFN-β than ORF1stop, indicating the role of FAdV-9 dUTPase in the upregulation of type I IFNs. This was contrary with the results of Leang et al. (2011), which showed that 113 the MHV-68-encoded dUTPase counteracts the antiviral reponse of type I IFNs. Nevertheless, it is demonstrated that EBV-encoded dUTPase is able to upregulate the expression of IFN-β (~11 fold) (Ariza et al., 2013). Another group also showed that gp120 of HIV is involved in the higher levels of TNF-α, IL-6, IL-10, IFN-α, and IFN-γ (Rychert et al., 2010). It should be noted that cytokine receptor interacting region of gp120 is evolved from an ancestral dUTPase gene (Abergel et al. 1999). In the study of Chapter 4, similar results were found in terms of the mRNA expression of INFs -α, -β, -γ, IL-10 in tissues of chickens inoculated with ORF1stop or wtFAdV-9. Together, these data indicate that FAdV-9 dUTPase plays important roles in the upregulation of type I IFNs both in vitro and in vivo. Recent studies demonstrated that functions of several dUTPases in affecting virus replication and modulating the immune response are not related to their enzymatic activity. For instance, Voronin et al. (2014) showed that despite the lack of dUTPase activity, the dUTPase-related gene of bovine immunodeficiency virus is critical for viral replication. Leang et al. (2011) showed that the inhibition of the type I interferon response by MHV-68-encoded dUTPase is independent of its enzymatic activity. Therefore, it would be interesting to investigate whether the capacity of FAdV-9 dUTPase to affecting virus replication and upregulating type I IFNs is independent of its enzymatic activity. To achieve this, it is necessary to determine the key residues and peptide for its catalysis through site-directed mutagenesis. Several studies have shown that the motifs 3 and 5 of dUTPase are critical for the catalysis. For example, Freeman et al. (2009) showed that deletion of the flexible Cterminal tail carrying motif 5 of EBV-encoded dUTPase resulted in a protein completely 114 devoid of enzymatic activity. Results of site-directed mutagenesis within motif 5 further demonstrated that replacement of arginine at nt 268 with alanine largely affects the enzyme activity, and replacements of phenylalanine at nt 273 with alanine leads to a nondetectable enzymatic activity (Freeman et al., 2009). It is very likely that motif 5 is also critical for the dUTPase of HAdVs, as dUTPase of HAdV-9 lacking the motif 5 does not show detectable enzymatic activity (Weiss et al., 1997). Whether dUTPases of other HAdVs possess the enzymatic activity is unknow, however, it was demonstrated in Chapter 3 that the absence of the motif 5 is universal for dUTPases of all human adenoviruses (Fig 3.1). To test if the motif 5 is also critical for FAdV-9 dUTPase, future research needs to be done. Also, it would be interesting to explore if there is other critical motif and core sequence responsible for the enzymatic activity of FAdV-9 dUTPase. Truncated mutants of FAdV-9 dUTPase could be generated through site-directed mutagenesis to achieve this aim. In the present work, FAdV-9 dUTPase was demonstrated to contribute to the upregulation of type I interferons in vitro and in vivo. However, the detailed mechanism is still obscure. Glaser et al. (2006) showed that purified EBV-encoded dUTPase is able to induce immune dysregulation in vitro by up-regulating the expression of proinflammatory cytokines including TNF-α, IL-1β, IL-8, IL-6, and IL-10 in PBMCs. It was further demonstrated that this increased expression in response to the EBV-encoded dUTPase treatment was dependent on the activation of NF-ĸB (Waldman et al., 2008). Ariza et al. (2009) showed that EBV-encoded dUTPase activates NF-ĸB in a dosedependent manner through TLR2, but not TLR3, TLR4, or TLR4/MD2, and that the activation of NF-ĸB requires the recruitment of the adaptor molecule MyD88, indicating 115 that EBV-encoded dUTPase could potentially modulate the innate immune response in EBV-permissive cells through the TLR2 and MyD88 signal transduction pathway. To determine how FAdV-9 dUTPase up-regulates the cytokine gene expression, future research needs to be done. For human adenoviruses, several TLRs are involved in sensing the virus. TLR-9 was found to sense HAdV-B in peripheral blood mononuclear cells and plasmacytoid dendritic cells (pDCs) (Sirena et al., 2004; Hendrickx et al., 2014). In addition, TLR2 knockout mice showed reduced NF-κB activation and humoral responses to HAdV vectors (Appledorn et al., 2008). In mammals, high TLR9 expression in pDCs serves as a sensor for DNA virus infection (such as HSV-1 and HSV-2) by exerting an effective antiviral immune response by producing type I IFN (Lund et al., 2003; Krug et al., 2004). Chicken TLR2 has been identified, while there is no orthologous gene of mammalian TLR9 idetified in chickens (Juul-Madsen et al., 2011; Chen et al., 2013). It is reported that TLR21, identified in chickens but not in mammals, has a similar function to mammalian TLR9 in recognition of CpG DNA (Keestra et al., 2010). Therefore, it would be worth exploring whether FAdV-9 dUTPase up-regulates the type I interferons through TLR21 or other TLRs, and investigating what up-stream cytokines of type I IFNs pathway are involved and how their gene expressions are regulated. Generally, viruses encode certain proteins that can counteract the antiviral response of type I IFNs, for example, the NS1 protein of influenza A virus (Hale et al., 2008), the NS3/4A of hepatitis C virus (Li et al., 2005), and the V protein of paramyxovirus (Andrejeva et al., 2004). In human adenoviruses, E1A is known to block this type I IFNmediated response, although the exact mechanisms have not been elucidated. This 116 brought up the question that what viral genes of FAdV-9 would function as an antiviral response inhibitor. Initially, ORF1 of FAdV-9 was hypothesized to be one potential target, as is the case for the MHV-68-encoded dUTPase (Leang et al., 2011). However, based on the study of Chapter 3, it seems that dUTPase of FAdV-9 does not possess this function. Previously, it was found that both wtFAdV-9 and FAdV-9Δ4 grow in CH-SAH cells pretreated with recombinant chicken IFN-α as well as in untreated ones (data not shown), suggesting that both viruses can evade the antiviral response of IFN-α. This indicates that there should be some viral proteins in FAdV-9 genome, rather than the six ORFs (0, 1, 1A, 1B, 1C and 2) deleted in FAdV-9Δ4, which are responsible for this inhibition of IFNα antiviral response. The E4 region might be translocated from the right end of human adenovirus genomes to the left end in FAdV genomes. For example, E4orf1, the dUTPase homolog, is located in the right end of HAdV genomes, while dUTPases of FAdVs are located in the left end of the genome (Weiss et al., 1997; Harrach et al., 2011). Therefore, it would be interesting to investigate whether the genes that function like E1A, at the left end of mastadenovirus genome, would be translocated to the right end of FAdV genomes. In conclusion, in this work, FAdV-9 ORF1 was characterized as a functional dUTPase enzyme and its molecular features including transcription and translation patterns, cellular localization were determined. Functional studies of FAdV-9 dUTPase demonstrated that although FAdV-9 dUTPase did not affect virus replication it contributed to the up-regulation of type I interferons in vitro. Moreover, in vivo study showed that FAdV-9 dUTPase plays significant roles in virus replication at the early 117 stage of the infection (up to 3 d.p.i.) and in modulating the host immune response against virus infection. The data presented in this work contribute to the better understanding of molecular biology of FAdVs, and are also helpful for exploring the mechanism of the host immune response against the fowl adenovirus infection. 118 References Abdul-Careem, M.F., Hunter, B.D., Parvizi, P, Haghighi, H.R., Thanthrige-Don, N., Sharif, S., 2007. 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