Rational Design of Human Metapneumovirus Live Attenuated Vaccine Candidates by Inhibiting Viral Messenger RNA Cap Methyltransferase

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Rational design of human metapneumovirus live attenuated vaccine candidates by inhibiting viral messenger RNA cap methyltransferase

DISSERTATION

Presented in Partial Fulfillment of the Requirements for the Degree Doctor of Philosophy in the Graduate School of The Ohio State University

Yu Zhang

The Graduate Program in Food Science and Technology

The Ohio State University

2014

Dissertation Committee:

Dr. Jianrong Li, advisor

Dr. Melvin Pascall

Dr. Stefan Niewiesk

Dr. Tracey Papenfuss

Copyrighted by

Yu Zhang

2014

Abstract

Human metapneumovirus (hMPV) is a newly discovered paramyxovirus, first identified in 2001 in the Netherlands in infants and children with acute respiratory tract infections. Soon after its discovery, hMPV was recognized as a globally prevalent pathogen. Epidemiological studies suggest that 5 to 15% of all respiratory tract infections in infants and young children are caused by hMPV, a proportion second only to that of human respiratory syncytial virus (hRSV). Despite major efforts, there are no therapeutics or vaccines available for hMPV. In the last decade, approaches to generate vaccines employing viral proteins or inactivated vaccines have failed either due to a lack of immunogenicity or the potential for causing enhanced pulmonary disease upon natural infection with the same virus.

In contrast to inactivated vaccines, enhanced lung diseases have not been observed for candidate live attenuated hMPV vaccines. Thus, a living attenuated vaccine is the most promising vaccine candidate for hMPV. However, it has been a challenge to identify an hMPV vaccine strain that has an optimal balance between attenuation and immunogenicity. In addition, hMPV grows poorly in cell culture and the growth is trypsin-dependent. To evaluate the safety and efficacy of a vaccine candidate, a robust small animal model is required.

To enhance the growth of hMPV, the cleavage site (99RQSR102 motif) of fusion (F) protein was mutated to 99RRRR102 in an infectious clone of hMPV. The resultant recombinant hMPV (rhMPV) displayed trypsin-independent growth phenotype and promoted earlier cytopathic effects (CPE) in cell culture. Interestingly, rhMPV formed clear viral plaques in a number of mammalian cell lines (such as Vero-E6 and

LLC-MK2 cells) in an agarose overlay plaque assay, which allows for plaque purification of the viruses. Thus, the trypsin-independent rhMPV is an improved backbone virus for development of live attenuated vaccines.

To identify a small animal model for hMPV, we compared the replication and pathogenesis of rhMPV in BALB/c mice, Syrian golden hamsters, and cotton rats. It was found that BALB/c mice are not permissive for rhMPV infection. In hamsters, rhMPV had an efficient replication in nasal turbinate but was restricted in lungs. In contrast, hMPV replicated efficiently in both nasal turbinate and lung when intranasally administered with three doses (104, 105 and 106 PFU) in cotton rats. Lungs of cotton rats infected by rhMPV developed histological changes including interstitial pneumonia, mononuclear cells infiltrates, and increased lumen exudates.

Immunohistochemistry examination found that viral antigens were expressed at the luminal surfaces of the bronchial epithelium cells in lungs. Thus, we conclude that that cotton rat is a robust small animal model for rhMPV infection.

We hypothesize that viral messenger RNA (mRNA) cap methyltransferase

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(MTase) is a novel target to identify a live attenuated hMPV strain that has a proper balance between attenuation and immunogenicity. The rationale for this hypothesis is that mRNA cap methylation is essential for viral gene expression and, subsequently, viral replication. To test this hypothesis, we performed a mutagenesis analysis in the putative mRNA cap MTase catalytic site and S-adenosyl methionine (SAM) binding site located in the conserved region VI (CR-VI) of large (L) polymerase protein.

Alanine substitutions to the amino acid residues involved in MTase catalysis and SAM binding diminished reporter gene expression in a minigenome replication assay. Using a reverse genetic system, we recovered three recombinant hMPVs carrying mutations in the SAM binding site. Trans-methylation assay showed that these rhMPV mutants were specifically defective in ribose 2’-O, but not guanine N-7 (G-N-7) methylation.

These MTase-defective rhMPVs showed delayed growth kinetics, reduced viral genome replication and mRNA synthesis, and formed smaller plaques in cell culture compared to wildtype rhMPV. Therefore, rhMPVs lacking 2’-O methylation were highly attenuated in cell culture.

To determine whether MTase-defective rhMPVs can be used as live attenuated vaccine candidates, recombinants rhMPV-G1696A, G1700A, and D1755A were inoculated intranasally into cotton rats. It was found that MTase-defective rhMPVs were highly attenuated in viral replication in upper and lower respiratory tracts in cotton rats. In addition, these recombinant viruses caused minimal lung histological changes and viral antigen expression in bronchial epithelium cells. Importantly, cotton iv

rats vaccinated with these MTase-defective rhMPVs triggered a high level of neutralizing antibody and were completely protected from challenge with wildtype rhMPV.

In conclusion, we found that mRNA cap MTase is a novel target to rationally design live attenuated vaccines for hMPV. These MTase-defective rhMPVs were sufficiently attenuated but retained high immunogenicity in cotton rats. Hence,

MTase-defective rhMPVs are excellent vaccine candidates for hMPV.

Dedication

This dissertation is dedicated to my parents Pei-Xin Zhang and Yun-Zhu Wang

for their love and support.

Acknowledgement

First and foremost, I would like express my deepest gratitude to my advisor Dr. Jianrong Li for his guidance and support during my time in his lab. I am always impressed and inspired by his devotion and enthusiasm to science. Without him, I would not have achieved my dissertation.

I also want to thank my committee members Dr. Stefan Niewiesk, Dr. Melvin Pascall, and Dr. Tracey Papenfuss for the mentorship and the intellectual environment they provided.

All members in Dr. Li’s group: Dr. Yuanmei Ma, Fangfei Lou, Erin DiCaprio, Dr. Yongwei Wei, Dr. Hui Cai, Brie Coyle, Jing Sun, Xueya Liang, Dr. Xiaodong Zhang, Yue Duan, Dr. Junan Li, Kurtis Feng, Ashley Predmore, Elbashir Araud, Jiawei Yeap, Anastasia Purgianto, Dr. Ran Zhao, Dr. Xiangjie Yao, and Yang Zhu, Rongzhang Wang, and Dr. Xianjun Dai, I really appreciate and thank you all for the help and friendship over the years.

I would like to thank all my friends from our collaborate labs, Devra Huey, Dr. Dhohyung Kim, Michelle (Gia) Green, Dr. Miyoung Kim, Yaoling Shu, Dr. Supranee Chaiwatpongsakorn, Dr. Heather Costello, Sara Johnson, Dr. Gabriel Sanglay, Dr. Lizanel Feliciano; thank you for your help all these years.

I also would like to thank the entire faculty, staff and students in Department of Food Science.

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Vita

June 2002…………………………………… Huzhou High School

June 2006…………………………………… B.S. Biotechnology, East China

University of Science and

Technology, Shanghai, China

September 2008 –present…………………… Graduate Research Associate,

Department of Food Science and

Technology, The Ohio State

University

Publications

Zhang Y, Li J, Rational design of human metapneumovirus live attenuated vaccine candidates by inhibiting viral messenger RNA cap methyltransferase (Manuscript in preparation)

Wei Y, Zhang Y, Cai H, Mirza MA, Iorio MR, Peeples EM, Niewiesk S, Li J, Roles of the putative integrin-binding motif of the human metapneumovirus fusion (F) protein in

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cell-cell fusion, viral infectivity, and pathogenesis, J Viro., Manuscript Submitted in November, 2013

Kim MY, Ma Y, Zhang Y, Li J, Shu Y, Oglesbee M., hsp70-dependent antiviral immunity against cytopathic neuronal infection by vesicular stomatitis virus. J Virol. 2013 Oct;87(19):10668-78

Ma Y, Wei Y, Zhang X, Zhang Y, Cai H, Zhu Y, Shilo K, Oglesbee M, Krakowka S, Whelan S, Li J, Messenger RNA cap methylation influences pathogenesis of vesicular stomatitis virus in vivo. Manuscript accepted in December, 2013

Zhang Y, Wei Y, Li J, Li J, Development and optimization of a direct plaque assay for human and avian metapneumoviruses, J Virol Methods. 2012 Oct;185(1):61-8.

PATENTS

Jianrong Li, Yongwei Wei, Yu Zhang. Novel live attenuated vaccines for human metapneumovirus. Patent filed Jun 2012

Jianrong Li, Yuanmei MA, Yu Zhang. World Patent Application WO 2012170997 A1, Paramyxovirus immunogens and related materials and methods, Patent publication date: Dec 13, 2012

BOOK CHAPTERS

Li J, Zhang Y. Messenger RNA Cap Methylation in Vesicular Stomatitis Virus, a Prototype of Non‐Segmented Negative‐Sense RNA Virus, Methylation - From DNA, RNA and Histones to Diseases and Treatment, Prof. Anica Dricu (Ed.), ISBN: 978-953-51-0881-8, InTech, DOI: 10.5772/54598. November 28, 2012

Fields of Study

Major Field: Food Science and Technology

Minor Field: Molecular Virology

Table of Contents

Abstract……………… ...... ii

Dedication…………… ...... xi

Acknowledgement ...... vii

Vita………………...... viii

List of Tables……………… ...... xiv

List of Figures……………...... xv

Chapter 1. Literature review: human metapneumovirus ...... 1

1.1. Introduction ...... 1

1.2. Taxonomy and classification ...... 2

1.3. Epidemiology and clinical features ...... 4

1.4. hMPV virion structure and genome organization ...... 5

1.5. hMPV replication cycle ...... 8

1.6. hMPV viral proteins ...... 12

1.7. Mechanism of cap formation ...... 22

1.8. Reverse genetics ...... 32

1.9. Animal models for hMPV ...... 34

1.10. hMPV vaccine development ...... 37

1.11. Treatment and prevention ...... 50 xi

Chapter 2. Development and optimization of a direct plaque assay for human and avian metapneumoviruses ...... 52

2.1. Abstract ...... 52

2.2. Introduction ...... 53

2.3. Materials and methods ...... 56

2.3.7. Statistical analyses ...... 60

2.4. Results ...... 60

2.5. Discussion ...... 73

Chapter 3. Development of a small animal model for human metapneumovirus: cotton rat is more permissive than hamster and mouse ...... 78

3.1. Abstract ...... 78

3.2. Introduction ...... 79

3.3. Materials and Methods ...... 81

3.4. Results ...... 88

3.5. Discussion ...... 96

Chapter 4. Recovery and characterization of methyltransferase-defective recombinant human metapneumovirus ...... 100

4.1. Abstract ...... 100

4.2. Introduction ...... 101

4.3. Materials and Methods ...... 105

4.4. Results ...... 114

4.5. Discussion ...... 131

Chapter 5. Pathogenicity and immunogenicity of methyltransferase-defective human metapneumovirus in cotton rats ...... 138 xii

5.1. Abstract ...... 138

5.2. Introduction ...... 139

5.3. Materials and Methods ...... 142

5.4. Results ...... 152

5.5. Discussion ...... 166

Chapter 6. Future directions ...... 173

6.1. Determine the genetic stability of MTase-defective rhMPV vaccine candidates… ...... 173

6.2. Optimize the vaccination strategy for MTase-defective rhMPV vaccine candidates…...... 174

6.3. Evaluate the cross protection of MTase-defective rhMPV vaccine candidates…...... 174

6.4. Determine the mechanism of protective immune responses induced by MTase-defective rhMPVs...... 175

6.5. Improve the growth of MTase-defective rhMPV vaccine candidates 175

6.6. Use MTase-defective rhMPV as a vector to deliver antigens from other human paramyxoviruses ...... 176

6.7. Recover and characterize rhMPVs carrying mutations in MTase catalytic site and RNA binding site………...... 177

6.8. Facilitate the clinical trials of the MTase-defective rhMPV vaccine candidates in nonhuman primates ...... 178

References………...... 179

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List of Tables

Table 3.1 Experimental design of hMPV replication study in cotton rats ...... 83

Table 3.2 Reaction components for one-step RT-PCR ...... 88

Table 3.3 Replication of rhMPV in upper and lower respiratory tracts of cotton rats92

Table 4.1 Optimization the growth of rhMPV-D1755A ...... 126

Table 5.1 Reaction components for real-time PCR ...... 150

Table 5.2 Reaction components for one-step RT-PCR ...... 151

Table 5.3 Replication of MTase-defective rhMPVs in cotton rats ...... 152

Table 5.4 Pulmonary histological changes in cotton rats vaccinated with MTase

defective rhMPVs and challenged with rhMPV ...... 156

Table 5.5 Immunogenicity of MTase-defective rhMPVs in cotton rats ...... 161

Table 5.6 Pulmonary histological changes in cotton rats infected by MTase-defective

rhMPVs ...... 164

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List of Figures

Figure 1.1 Phylogenetic distribution of viruses belonging to the Paramyxoviridae family ...... 4

Figure 1.2 Electron microscopic image of hMPV viron particles ...... 6

Figure 1.3 hMPV virion structure ...... 7

Figure 1.4 Comparison the genomic organization of hMPV and hRSV ...... 7

Figure 1.5 Schematic representation of the hMPV life cycle ...... 8

Figure 1.6 Schematic representation of hMPV L protein, putative functions of CR III, V, and VI as determined in VSV ...... 19

Figure 1.7 Molecular architecture of VSV L ...... 22

Figure 1.8 mRNA cap structure ...... 22

Figure 1.9 Comparison of mRNA cap formation in eukaryotic cells and VSV ...... 26

Figure 2.1 Plaque formation of aMPV subtype C MN strain in Vero, Vero-E6, and LLC-MK2 cells...... 62

Figure 2.2 Trypsin dependence of rhMPV-F and rhMPV...... 65

Figure 2.3 Plaque formation of rhMPV-F in Vero, Vero-E6, and LLC-MK2 cells...... 66

Figure 2.4 The effect of actinomycin-D on plaque formation of rhMPV-F...... 69

Figure 2.5 Dynamics of plaque formation of rhMPV-F and aMPV in Vero cells...... 70

Figure 2.6 Effect of actinomycin-D on the number and size of viral plaques...... 71

Figure 2.7 Growth kinetics of rhMPV-F as determined by direct plaque assay and immunostaining assay...... 73

Figure 3.1 Body weight changes of BLAB/c mice infected by rhMPV...... 89

Figure 3.2 Body weight changes of hamsters infected by rhMPV...... 90

Figure 3.3 Virus titer in nasal turbinates and lungs of Syrian golden hamsters infected by rhMPV...... 91

Figure 3.4 Histological changes in lungs of hamsters infected by rhMPV...... 91

Figure 3.5 Histological changes in lungs of cotton rats infected by rhMPV...... 94

Figure 3.6 Immunohistochemical staining of lung tissues from cotton rats infected by rhMPV...... 95

Figure 4.1 Conserved regions (CR) in the L proteins of NNS RNA viruses...... 102

Figure 4.2 Schematic representation of hMPV minigenome construct...... 106

Figure 4.3 Schematic diagram of minigenome assay...... 108

Figure 4.4 Amino acid sequence alignment of conserved region VI (CR VI) of NNS RNA viruses L proteins and modeling with two known 2’-O MTase structures, VP39 and RRMJ...... 115

Figure 4.5 Mutations to the CR VI of hMPV L gene resulting diminished GFP expression in a minigenome assay...... 117

Figure 4.6 Recovery of rhMPV from an infectious cDNA clone by reverse genetics ...... 119

Figure 4.7 rhMPVs carrying mutations in the SAM binding site formed smaller plaques in Vero E6 cells...... 120

Figure 4.8 RT-PCR and sequencing analysis of rhMPVs carrying mutations in the SAM binding site...... 121

Figure 4.9 Growth kinetics of rhMPVs carrying mutations in the SAM binding site...... 124

Figure 4.10 rhMPV mutants had delayed cytopathic effects (CPE)...... 125 xvi

Figure 4.12 Quantification of viral N mRNA and genomic RNA by real-time RT-PCR...... 128

Figure 4.13 In vitro trans-methylation assay...... 130

Figure 5.1 Schematic diagram of rhMPV replication and pathogenesis study in cotton rats...... 144

Figure 5.2 Lung histological changes in cotton rats infected by MTase-defective rhMPVs...... 155

Figure 5.3 Immunohistochemical (IHC) staining of lungs from cotton rats infected by MTase-defective rhMPVs...... 157

Figure 5.4 Schematic diagram of immunogenicity study in cotton rats...... 159

Figure 5.5 Serum neutralizing antibody titers in cotton rats vaccinated with MTase-defective rhMPVs...... 160

Figure 5.6 Lung histological changes in cotton rats vaccinated with MTase-defective rhMPVs followed by challenge with rhMPV...... 163

Figure 5.7 Immunohistochemical (IHC) staining of lungs from cotton rats vaccinated by MTase-defective rhMPVs followed by rhMPV challenge...... 165

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Chapter 1.

Literature review: human metapneumovirus

1.1. Introduction

Human metapneumovirus (hMPV) is a relatively newly discovered human pathogen, first described in 2001 in the Netherlands. It was isolated from infants and children experiencing respiratory tract infections similar to the symptoms caused by human respiratory syncytial virus (hRSV) (van den Hoogen, de Jong et al. 2001). The unknown virus isolate replicated poorly and slowly in cell culture and required trypsin for growth. The cytopathic effects (CPE) caused by this agent were indistinguishable from hRSV. Subsequently, a random PCR amplification strategy (RAP-PCR) was used to identify the unknown pathogen. It was found that this virus shared 20-40% amino acid sequence homology with hRSV and 50-80% homology with avian metapneumovirus (aMPV). Thus, this unknown virus was named human metapneumovirus (hMPV) because it shares the highest homology with aMPV within the genus metapneuvmovirus. Soon after its discovery, hMPV was recognized as a globally prevalent pathogen; virtually all children by the age of 5 were seropositive. It is a major causative agent of acute respiratory tract disease in individuals of all ages, especially in infants, children, the elderly, and immunocompromised individuals

(Pelletier, Dery et al. 2002; Williams, Harris et al. 2004). Epidemiological studies suggest that 5 to 15% of all respiratory tract infections in infants and young children are caused by hMPV, a proportion second only to that of hRSV (Boivin, De Serres et al. 2003; Esper, Boucher et al. 2003; van den Hoogen, van Doornum et al. 2003;

Williams, Harris et al. 2004; de Graaf, Osterhaus et al. 2008). Currently, there are no therapeutics or vaccines available for hMPV (Boivin, Abed et al. 2002; Peret, Boivin et al. 2002).

1.2. Taxonomy and classification

HMPV belongs to the order Mononegavirales, in the family Paramyxoviridae. It is further classified as a member of the subfamily Pneumovirinae of the family of

Paramyxoviridae. This subfamily is composed of the genus Pneumovirus, exemplified by hRSV, bovine respiratory syncytial virus (bRSV), ovine and caprine RSVs, and pneumonia virus of mice (PVM) and the genus metapneuvmovirus which only includes two members, hMPV and aMPV (Figure 1.1). Based on the sequence variability of surface glycoproteins, the fusion (F) protein and the attachment glycoprotein G, hMPV can be classified into two antigenic different groups A and B and each of these groups can be further divided into two subgroups A1 and A2, and

B1and B2.

AMPV is the only other member of the genous metapneumovirus. It is also known as avian pneumovirus or turkey rhinotracheitis. Based on antigenicity and

genetic diversity, four subtypes of aMPV, designated A, B, C, and D, have been characterized. Subtypes A, B, and D are found mainly in Africa, Europe, and Asia while subtype C is prevalent in the US. Interestingly, sequence analyses have shown that the US subgroup C of aMPV is more closely related to hMPV than to other three aMPV subtypes (A, B and D) (Skiadopoulos, Biacchesi et al. 2004; van den Hoogen,

Osterhaus et al. 2004; de Graaf, Osterhaus et al. 2008).

The other subfamily within Paramyxoviridae is called Paramyxovirinae, which also includes many highly infectious human pathogens such as measles (MeV), mumps (MuV), and human parainfluenza virus 3 (PIV3), deadly zoonotic viruses such as Hendra (HeV) and Nipah (NiV), and economically important viruses such as

Newcastle disease virus (NDV) and Sendai virus (SeV). Together, paramyxoviruses are responsible for the majority of respiratory tract diseases and inflict significant economic, health, and emotional burdens. For many of these viruses, there are no effective vaccines or anti-viral drugs.

Figure 1.1 Phylogenetic distribution of viruses belonging to the Paramyxoviridae family, adapted from Feuillet et. al, 2011. hRSV (human respiratory syncytial virus), bRSV (bovine respiratory syncytial virus), oRSV (ovine respiratory syncytial virus), PMV (murine pneumovirus), hMPV (human metapneumovirus), APV (avian metapneumovirus), hPIV2 (human parainfluenza virus type 2), hPIV4a and b (human parainfluenza virus types 4a and b), hPIV5 (human parainfluenza virus type 5), SV41(simian virus 41), MuV (mumps virus), NDV (Newcastle disease virus), MeV (measles virus), RPV (Rinderpest virus), DMV (dolphin morbilivirus), PDV (phocine distemper virus), CDV (canine distemper virus), TPMV (Tupaia paramyxovirus), heV (Hendra virus), NiV (Nipah virus), hPIV1 (human parainfluenza virus type 1), hPIV3 (human parainfluenza virus type 3), bPIV3 (bovine parainfluenza virus type 3), SeV(Sendai virus). Genetic analysis was performed using the gene encoding the nucleoprotein N. The scale represents the number of nucleotide changes.

1.3. Epidemiology and clinical features

HMPV is a globally prevalent pathogen. HMPV infections are observed in all age groups, especially in infants and children. Most children by the age of 5 years have already been infected by hMPV. It has been estimated that hMPV accounts for

5-15% of hospitalization for respiratory tract infections in children, second only to 4

that of hRSV. Similar to hRSV, hMPV also displays a seasonal distribution with the peak occurring in winter and spring.

HMPV causes upper and lower respiratory tract infection with a spectrum of illnesses that range from asymptomatic infection to severe bronchiolitis. Infections in young adults usually cause mild flu-like symptoms. However, the infection can cause more severe disease in infants, children, the elderly, and immunocompromised individuals. HMPV infections in the upper respiratory tract are often associated with symptoms such as cough, fever, and rhinorrhea. In rare cases, infection can cause conjunctivitis, diarrhea, vomiting, and rash. HMPV infections in the lower respiratory tract can cause pneumonia, bronchiolitis, croup, and asthma exacerbation (Boivin,

Abed et al. 2002). These clinical signs caused by hMPV are similar and usually indistinguishable from hRSV and PIV3. In addition, co-infection of hRSV has been observed in 5-17% of patients infected with hMPV. However, it is not clear if the co-infection is related to the exacerbated disease.

1.4. hMPV virion structure and genome organization

Under electron microscopy, hMPV virions are pleomorphic particles, measuring between 150 to 600 nm in diameter, with short spikes of 13 to 17 nm on the lipid envelope (Figure 1.2).

Figure 1.2 Electron microscopic image of hMPV viron particles, adapted from Schildgen et. al, 2011

HMPV is a nonsegmented single-stranded negative-sense RNA virus. The viral genome is approximately 13 kb in length, which contains 8 genes and 9 open reading frames (ORFs) (Figure 1.4) (van den Hoogen, Bestebroer et al. 2002; Bastien,

Normand et al. 2003). The leader (Le) sequence, nucleoprotein(N), phosphorprotein

(P), matrix (M), fusion (F), small hydrophobic protein (SH), attachment glycoprotein

(G), M2 (expresses M2-1 and M2-2 proteins), large (L) polymerase, and trailer (Tr) sequence are arranged in the order: 3’-Le-N-P-M-F-M2-SH-G-L-5’. Each gene begins with a highly conserved 9 nucleotide gene-start (GS) transcriptional signal and ends with a less conserved 6 to 9 nucleotide gene end (GE) signal. Similar to other pneumoviruses, hMPV virion contains a lipid envelope surrounding the M protein.

Three surface glycoproteins, G, F, and SH, form the projections on the envelope

(Figure 1.3). Inside the virion, the single-stranded negative-sense RNA genome is encapsidated by N protein to form an N-RNA complex which serves as a template for viral mRNA synthesis and genome replication. The N-RNA complex is associated 6

with P, L, M2-1, and also likely M2-2, which form a helical ribonucleoprotein (RNP) complex. The viral RNA-dependent RNA polymerase (RdRp) complex includes the catalytic subunit L and the accessory proteins P and M2-1.

Figure 1.3 hMPV virion structure

The genome structure of hMPV is identical to that of aMPV. However, compared with hRSV, hMPV genome lacks nonstructural protein 1 and 2 (NS1 and NS2) and also differs in orders of certain genes (Figure 1.4).

Figure 1.4 Comparison the genomic organization of hMPV and hRSV

1.5. hMPV replication cycle

Figure 1.5 Schematic representation of the hMPV life cycle, adapted from Schildgen et. al, 2011

1.5.1. Attachment and entry

The hMPV life cycle begins with the attachment of the virus to host cells.

Although the detailed mechanisms of initial attachment and receptor binding are not clear, evidence suggest that both hMPV G and F proteins interact with cell surface 8

glycosaminoglycans (GAGs), particularly heparin sulfate (HS), in the initial attachment step. In addition, it has been reported that integrinαvβ1, α5β1 and αv may serve as functional receptors for hMPV entry. Indeed, a putative integrin binding motif, Arg-Gly-Asp (RGD), presents in the F proteins of all strains of hMPV isolates.

Mutations to the RGD motif reduced the binding of soluble F protein to host cells.

These results suggest that those integrins may serve as cellular receptors for hMPV F protein. It is possible that hMPV utilizes two receptors for attachment. However, the mechanism of coordination of these two receptors is unknown. It has been proposed that attachment of hMPV to host cells may involve non-specific interaction of both G and F proteins with GAGs, followed by a specific interaction of F with cellular receptors such as integrin αvβ1, α5β1, and αv (Cseke, Maginnis et al. 2009; Chang,

Masante et al. 2012; Cox, Livesay et al. 2012).

Upon receptor binding, hMPV enters host cells through the fusion of the viral envelope with the cellular membrane. For viruses in the Paramyxovirinae subfamily, membrane fusion requires both the attachment protein (G, H, or HN) and the F protein.

In contrast, membrane fusion of pneumoviruses (such as hMPV) is unique among the paramyxoviruses, in that fusion is accomplished by the F protein alone without help from the attachment glycoprotein G. Consistent with this, recombinant hMPV lacking the G protein was found to replicate efficiently in cell culture. Another unique characteristic of hMPV entry is that fusion of some hMPV strains requires low pH whereas fusion of all other paramyxoviruses occurs at neutral pH. Using the hMPV 9

strain CAN 97-83 that requires low pH for fusion, it was found that inhibition of endosomal acidification or endocytosis significantly reduced hMPV infection, suggesting that hMPV utilizes an endocytic entry mechanism, in contrast to what has been proposed for most paramyxoviruses.

1.5.2. Gene expression and genome replication

Upon virus entry, the ribonucleoprotein (RNP) complex, which containsviral replication and transcription machinery, is released to the cytoplasm where viral replication and gene expression occurs. During primary transcription, the RNA dependent RNA polymerase (RdRp) recognizes the leader sequence at the 3’ end of the viral genome and follows a start-stop-restart model guided by the gene start (GS) and gene end (GE) signals flanking each viral gene to transcribe ten discrete RNAs: a leader RNA (Le+), which is neither capped nor polyadenylated, and eight mRNAs that are capped and methylated at the 5’ end and polyadenylated at the 3’ end. A portion of RdRp disassociated at intergenic regions, thus the viral mRNA expression appears as a decreasing gradient: genes at proximal to the 3’ leader are transcribed more abundantly than downstream genes. The capping and methylation at the 5’ end of viral mRNA and polyadenylation at the 3’ end of mRNA are carried out by the L protein. The 3’ end polyadenylation is accomplished though a stuttering mechanism, where the poly U at the GE signal is transcribed repeatedly.

HMPV encodes 8 mRNAs which are further translated into 9 proteins. Except

for M2 mRNA, which was translated into two proteins, M2-1 and M2-2, all other viral mRNA produces one individual protein. The newly synthesized proteins are either released to cytoplasm (N, P, M, M2-1, M2-2, and L) or transported to host cell membrane (F, G, and SH). Among these proteins, the P and L form the polymerase complex. M2-1 may serve as a co-factor of the polymerase. M2-2 acts as a negative regulator of the mRNA synthesis and genome replication.

During genome replication, the RdRp initiates at the extreme 3’ end leader sequence of the genome and synthesizes a full-length complementary antigenome. All transcription signals are omitted by RdRp in the antigenome synthesis, and the antigenome is neither capped nor polyadenylated. Similar to viral genome, the antigenome is also encapsidated by N protein, which subsequently serves as a template for synthesis of full-length progeny genomes. The RdRp utilizes the 3’ trailer sequence on the antigenome as the promoter and the progeny genomes can then be utilized as templates for secondary transcription, or assembled into infectious particles

(Whelan, Barr et al. 2004).

1.5.3. Assembly and budding

Most of our knowledge about hMPV assembly and budding comes from that of hRSV. Similar to all paramyxoviruses, M protein plays a central role in mediating the assembly and budding of the new virion. It has been demonstrated that M interacts with cytoplasmic domain of F and G proteins on the lipid rafts on the cell membrane.

M protein also interacts with the newly synthesized viral genome as well as the associated N, P, L, and possibly M2-1 and M2-2 to initiate the budding. It has been shown that the minimum viral protein requires for assembly of viruslike particles

(VLPs) are F, G, and M protein. It has also been thought that hMPV hijacks the cellular apical recycling endosomes (ACEs) for budding in the polarized cells.

1.6. hMPV viral proteins

1.6.1. The fusion (F) protein

The hMPV F protein is a class I fusion protein, which directs penetration by triggering fusion between virus envelope and cell membrane. In virus-infected cells, F protein is expressed on the cell surface which induces fusion of neighboring cells and forms syncytia.

The hMPV F is first synthesized as a precursor protein, F0, and subsequently cleaved at an Arg-Gln-Ser-Arg site into two disulfide-linked subunits, F1 and F2. The cleavage site of hMPV F does not conform to the consensus furin motif, thus hMPV clinical isolates require trypsin for growth in cell culture. This cleavage generates a hydrophobic fusion peptide (FP), which is directly inserted into the membrane to initiate fusion. Paramyxovirus F proteins contain two conserved heptad repeat (HR) regions, the N-terminal heptad (HRA) and the C-terminal heptad (HRB), which are located downstream of the fusion peptide and upstream of the transmembrane (TM) domain, respectively. Upon triggering, the metastable pre-triggered F protein

undergoes a series of dramatic and irreversible conformational changes to bring two membranes together to initiate fusion. Currently, the mechanism by which fusion is regulated such that it occurs at the proper time and place remains poorly understood.

Similar to hRSV, hMPV fusion is accomplished by the F protein alone without help from the attachment glycoprotein G. This suggests that G protein is dispensable for attachment and fusion. Consistent with this observation, recombinant hMPV lacking the G protein was found to replicate efficiently in cell culture. Another unique characteristic of hMPV entry is that fusion of some hMPV strains requires low pH whereas fusion of all other paramyxoviruses occurs at neutral pH. In addition, fusion of hMPV requires trypsin. Together these results suggest that hMPV fusion possesses many unique features that are distinct from other paramyxoviruses, although the detailed mechanism underlying fusion promotion in hMPV is still poorly understood.

The fact that the F protein of hMPV alone is sufficient for induction of cell-cell fusion and viral entry suggests that the hMPV F protein possesses dual functions, receptor binding and fusion promotion. It has been shown that integrin αvβ1 and αV are essential for cell-cell fusion and hMPV infectivity (Cseke, Maginnis et al. 2009).

Consistent with this, the F proteins of all known hMPV strains contain a putative integrin-binding motif (329RGD331), which has been shown to be essential for cell-cell fusion,viral infectivity, and replication (Cox, Livesay et al. 2012). Most recently, Chang, Masante et al. 2012 found that heparan sulfate proteoglycans function as primary receptors for hMPV, binding to which is mediated by the F 13

protein. It was hypothesized that the interaction between integrins and hMPV occurs after the initial binding of hMPV F to heparan sulfate proteoglycans. Nonetheless, molecular mechanisms underlying the interaction between hMPV F and integrin, and their roles in cell-cell fusion, hMPV life cycle, and viral pathogenesis remain to be further elucidated.

1.6.2. The attachment glycoprotein (G)

The G protein of hMPV is 217 to 236 amino acids in length among different strains (Bastien, Liu et al. 2004; Peret, Abed et al. 2004; Padhi and Verghese 2008). It is a type II mucin-like glycosylated membrane protein, with its signal sequence and membrane anchor proximal to N terminus and ectodomain on its C terminus. The ectodomain has a high content of serine, threonine and proline residues. In addition it is modified by N-linked and O-linked sugars to yield a mature form that migrates in

SDS-PAGE as a heterogeneous smear with an estimated molecular mass of 38 to 80 kDa, with a major band at about 50kDa. However, the molecular mass of the predicted unmodified form of G protein is approximately 25.9 kDa. HMPV G protein is highly variable among different strains, and the amino acid sequence identities between two serotypes range from only 31 to 35% (Liu, Bastien et al. 2007).

hMPV G was originally thought to be the only viral attachment protein, but now it has been shown that F protein also plays a key role. Similar to RSV G protein, it was reported that hMPV G protein binds to cell surface glycosaminoglycans (GAGs)

in immortalized cells (Liu, Bastien et al. 2007). However, recent studies also demonstrated that virus-cell binding is driven primary by F protein with cell surface

GAGs, mainly heparin sulfate (HS), in tissue culture. However, it is likely that hMPV utilizes different receptors during a natural infection since human airway epithelial

(HAE) cells lack cell surface HS expression. Deletion of hMPV G has little effects on viral replication in cell culture, however the G deleted hMPV mutant is highly attenuated in vivo (Biacchesi, Skiadopoulos et al. 2004). The reduced attachment ability may be the reason for the attenuation. Interestingly, a recent study found that infection of HAE cells with G deleted hMPV mutant increased the activation of transcription factor NF-κB and IRF families compared to wildtype hMPV, suggesting that G protein inhibits innate immune response. Further analysis revealed that the inhibitory affect was achieved by the direct interaction between G and retinoic induced gene (RIG)-I and the disruption of mitochondrial signaling pathway (Bao,

Liu et al. 2008; Kolli, Bao et al. 2011; Bao, Kolli et al. 2013).

1.6.3. The small hydrophobic (SH) protein

The hMPV SH protein is of 177 to 183 amino acids in length, depending on the strain of the virus. SH protein is a type II transmembrane glycoprotein. It has a high sequence variability among different strains with only about 60% identity between different hMPV lineages. In hRSV, it has been suggested that SH forms channel-like transmembrane structure, however, the function of hMPV SH is still unknown. It has been suggested that hMPV SH protein inhibits NF-κB transcription (Bao, Kolli et al. 15

2008), however, recent studies showed no differences in the expression of genes involved in TNF-α-induced NF-κB activation (de Graaf, Herfst et al. 2013). In addition, rhMPV lacking SH protein replicated efficiently in cell culture as well as in upper and lower respiratory tracts in hamsters. In addition, it is highly immunogenic in hamsters. This suggests that SH protein is dispensable in vitro and does not affect viral replication and immunogenicity in vivo.

1.6.4. The matrix (M) protein

The M protein of hMPV is 254 amino acids in length. The detail function of hMPV M protein is still poorly understood. In fact, most of the knowledge about hMPV M is inferred from its counterpart in hRSV. It is likely that M protein plays two major roles: (i) it drives viral assembly and budding and organizes virion structure at the cell membrane (Bagnaud-Baule, Reynard et al. 2011; Sabo, Ehrlich et al. 2011) and (ii) it may inhibit viral RNA transcriptase activity before assembly. Upon infection, the synthesized M protein interacts with the cell membrane through electrostatic and hydrophobic interactions, which leads to the formation of a layer beneath the envelope of virion. The M protein interacts with the cytoplasmic tails of surface glycoprotein F and G, and the N protein that drives viral assembly and budding. In addition, it has been reported that co-expression of F, G, and M leads to the assembly of virus like particles (VLPs). Although the detailed mechanism of viral assembly and budding is not clear, recently study has shown that a conserved -YAGL- motif located at a hydrophobic cleft of M protein is an important determinant of viral 16

assembly (Loo, Jumat et al. 2013). It is not clear which step in viral assembly in which the -YAGL- motif plays a role.

1.6.5. The M2-1 and M2-2 proteins

The M2 gene is a unique gene present in all known members of the

Pneumovirinae subfamily. The M2 mRNA is translated into M2-1 and M2-2 proteins by utilizing two overlapping ORFs. The hMPV M2-1 is a protein of 187 amino acids.

The M2-1 proteins of the pneumoviruses possess a conserved -CCCH-

(Cys-Cys-Cys-His) zinc binding motif located at the N-terminus, which is an important structural component for nucleic acid binding. Functions of its counterpart hRSV M2-1 have been extensively studied. The hRSV M2-1 protein is essential for transcription processivity and it is an anti-terminator. The hRSV M2-1 has been shown to directly interact with N, P, and L proteins. In the absence of M2-1 protein, hRSV polymerase terminates prematurely and randomly. Although deletion of M2-1 is lethal for hRSV, hMPV lacking the M2-1 gene can be recovered and was only slightly attenuated in cell culture. Recombinant hMPV lacking M2-1 could not replicate in a hamster model and was unable to elicit a protective immunity. This demonstrated that hMPV M2-1 is important for in vivo replication.

The second ORF of hMPV M2 gene encodes a M2-2 protein of up to 71 amino acids. The M2-2 ORF possibly initiates at position 525 or 537 on M2 mRNA, which overlaps 53 or 41 nt with M2-1 ORF. HRSV M2-2 appears to function as a regulator

of transcription and genome replication, which inhibits mRNA synthesis and favors genome replication. The hMPV M2-2 protein has been shown to interact with the L protein in an immunoprecipitation assay (Schickli, Kaur et al. 2008; Kitagawa, Zhou et al. 2010). In contrast to hRSV M2-2, hMPV M2-2 is shown to inhibit both mRNA transcription and genomic RNA replication in a minigenome system. Recombinant hMPV lacking M2-2 has been recovered and showed increased viral mRNAs and genomic RNAs, demonstrating the regulatory function of M2-2 in RNA synthesis.

Recently, hMPV M2-2 has also been shown to target mitochondrial antiviral-signaling protein (MAVS) and inhibit the MAVS-activated innate cellular signaling, an immune regulation function which hasn’t yet been reported in hRSV M2-2 (Ren, Wang et al.

2012).

1.6.6. The nucleoprotein (N)

The N protein of hMPV is 394 amino acids in length. It tightly encapsidates the entire viral genomic RNA and antigenomic RNA to form an RNase-resistant helical

N-RNA complex, known as nucleocapsid structure. The encapsidation not only protects the genome from degradation, but also likely protects the genome from the recognition by host pattern recognition receptors (PRRs), especially the retinoic aicd-induced gene I (RIG-I), melanoma differentiation-associated protein 5 (MDA-5), and RNA-induced protein kinase R (PKR). The N-RNA template is recognized by

RdRp whose major components are the L, P, and M2-1 proteins. The N-RNA serves as template for both mRNA transcription and genome replication. In addition, N 18

protein has been shown to interact with L, P, M2-1, and M proteins.

1.6.7. The phosphoprotein (P)

The P protein of hMPV is 294 amino acids in length and is highly phosphorylated. Similar to its counterpart in hRSV, hMPV P also forms a stable homotetramer through a central α-helical coiled-coil region. During mRNA synthesis and RNA replication, P protein appears to be responsible for the recruitment of L protein onto the viral neucleocapsid through the direct interaction with N and L, thus

P is an essential co-factor of RdRp.

1.6.8. The large (L) polymerase protein

The 2005 amino acid hMPV L protein is the largest viral protein. Similar to all other NNS RNA viruses, L protein is a multifunctional protein. It contains enzymatic activities for nucleotide polymerization, mRNA cap addition, cap methylation, and polyadenylation.

Figure 1.6 Schematic representation of hMPV L protein, putative functions of CR III, V, and VI as determined in VSV

Amino acid sequence alignment between the L proteins of representative members of each family within NNS RNA viruses has identified six conserved regions numbered I to VI (CRs I–VI) (Poch, Blumberg et al. 1990) (Figure 1.6). Thus, there is a general thought that the enzymatic activities of L protein are located in these conserved regions. VSV L protein has been used as a model to study the enzymatic activities of NNS RNA virus L proteins because robust transcription can be reconstituted in vitro using VSV L protein. In addition, it has been reported that the purified VSV L protein retains all the enzymatic activities that can process short virus-specific mRNA in trans. Recently, with many breakthroughs in the characterization of the function of VSV L protein, the enzymatic activities have been mapped to CRs at single amino acid level. Within the 6 CRs shared among all NNS

RNA virus L proteins (Figure 1.6), the RdRP activity has been identified in CR III, and this region is also required for polyadenylation. Consistent with this, a GDN 20

motif is conserved in CR III of all NNS RNA virus L proteins, and which is functionally equivalent to the GDD polymerization motif characteristic of positive strand RdRPs. The mRNA capping enzyme is mapped to CR V, whereas the mRNA cap methyltransferase (MTase) is mapped to CR VI. Although functions have not yet to be assigned to the other three conserved regions (CRs I, II, and IV), experiments with Sendai virus (SeV) have indicated that CR I binds P protein and CR II binds the

RNA template (Li, Fontaine-Rodriguez et al. 2005; Ogino, Kobayashi et al. 2005;

Ogino and Banerjee 2007; Li, Bai et al. 2008; Murphy and Grdzelishvili 2009;

Rahmeh, Li et al. 2009).

Although to date, the structure of L protein, or L protein fragments, has not been determined for any of the NNS RNA viruses, limited evidence suggest that the Sendai and measles virus L form homo-multimers through an N-terminal self-assembly domain. L protein also interacts with its co-factor (P) though another N-terminal domain to form the active polymerase complex essential for L protein activity.

More recently, the structure of VSV L protein has been revealed using negative stain electron microscopy (EM) in combination with proteolytic digestion and truncation mutation mapping. It was found that VSV L protein is organized into a ring domain containing the RNA polymerase and an appendage of three globular domains containing the cap-forming activities (Figure 1.7). The capping enzyme maps to a globular domain, which is juxtaposed to the ring, and the cap methyltransferase maps

to a more distal and flexibly connected globule. Interestingly, upon binding to P protein, L protein undergoes a significant structural rearrangement that may facilitate the coordination between mRNA synthesis and capping apparatus (Rahmeh, Schenk et al. 2010).

Figure 1.7 Molecular architecture of VSV L (adapted from Rahmeh et al., 2010). (A) L is organized into a core ring structure harboring the RdRP domain and a flexible appendage containing the activities necessary for cap formation (capping + methylation). The arrows depict putative flexible linkers. (B) Upon binding to P, L undergoes a structural rearrangement. The L–P complex exists as a monomer or a dimer in which the L pairs are likely bridged by interaction with an oligomer of P. The arrow represents the variable orientation of L proteins in the dimers.

1.7. Mechanism of cap formation

1.7.1. Introduction to messenger RNA (mRNA) cap

Figure 1.8 mRNA cap structure

The messenger RNA (mRNA) cap was first discovered in mid 1970s by

(Furuichi, Morgan et al. 1975; Furuichi, Muthukrishnan et al. 1975; Filipowicz,

Furuichi et al. 1976). It was first described as a guanine-N-7 (G-N-7) methylated guanosine residue linked to the 5’ end of mRNA though an unusual 5’ to 5’ triphosphate bridge. It was further revealed that the 2’ hydroxyl group of the first one or two ribose moieties of the 5’ end of the mRNA are also methylated (2’-O methylation) (Figure 1.8). The cap structure was found to be essential for efficient translation by directly binding with the eukaryotic translation initiation factor 4E

(eIF-4E). This complex facilitates the recruitment of other subunits, notably, eIF-4G, and also interacts with poly(A)-binding protein to form a circularized mRNA to promote translation. Later it was shown that the mRNA cap structure is also essential for mRNA nuclear exportation, mRNA stability, and it is also involved in microRNA-mediated gene silencing by interacting with other cap-binding proteins. It is firmly established that G-N-7 methylation is essential for mRNA translation

(Furuichi, Morgan et al. 1975; Furuichi, LaFiandra et al. 1977; Furuichi and Shatkin

1977; Nuss and Furuichi 1977). However, the biological role of 2’-O methylation remains elusive although it has been discovered for more than 40 years. Recent studies on West Nile virus (WNV) suggest that the 2’-O methylation of the 5’- cap of viral RNA functions as a molecular signature to distinct self and non-self mRNA to evade innate host antiviral responses through escape of the suppression of interferon-stimulated genes, tetratricopeptide repeats (IFIT). This provides us a new

insight into the evolutionary basis of the cap structure (Chappell, Nall et al. 2005;

Daffis, Szretter et al. 2010; Zust, Cervantes-Barragan et al. 2011; Habjan, Hubel et al.

2013).

1.7.2. mRNA cap formation in eukaryotic cells: a conventional mechanism.

In eukaryotic cells, mRNA cap formation is an early posttranscriptional event that is essential for subsequent processing, nuclear export, stability, and translation of mRNA. Cap formation is mediated by a series of enzymatic reactions. First, the 5’ triphosphate end of the nascent mRNA chain (5’pppN-RNA) is hydrolyzed by an

RNA triphosphatase (RTPase) to yield the diphosphate 5’ ppN-RNA. Second, an RNA guanylyltransferase (GTase) reacts with GTP to form a covalent enzyme-GMP intermediate and transfers GMP to 5’ppN-RNA via a 5′-5′triphosphate linkage to yield 5’ GpppN-RNA. Third, the capping guanylate is methylated by a G-N-7 methyltransferase (MTase) to yield 7mGpppN-RNA (cap 0). Finally, the G-N-7 methylated cap structure can then be further methylated by a ribose-2’-O (2’-O)

MTase to yield 7mGpppNm-RNA (cap 1). This mRNA cap formation is conserved among all eukaryotes. During mRNA cap methylation, S-adenosyl-L-methionine

(SAM) serves as the methyl donor, and the by-product S-adenosyl-homocysteine

(SAH) is the competitive inhibitor of the SAM-dependent MTase. In this conventional methylation reaction, G-N-7 methylation occurs prior to 2’-O methylation and the two methylase activities are carried out by two separate enzymes, each containing its own binding site for the methyl donor, SAM. 24

1.7.3. mRNA cap formation in NNS RNA viruses: an unconventional

mechanism.

Available evidence suggests that the cap structure of NNS RNA viruses is formed by a mechanism which is different from eukaryotic cap formation. For VSV,

RSV, and spring viremia of carp virus, the two italicized phosphates of the 5′Gppp5′

NpNpN triphosphate bridge have been shown to be derived from a GDP donor, rather than GMP. Using VSV as a model, it was found that capping of VSV mRNA was achieved by a novel polyribonucelotidyltransferase (PRNTase) which transferred a monophosphate RNA onto a GDP acceptor through a covalent L-RNA intermediate.

In the first step, a GTPase associated with the VSV L protein removes the γ-phosphate group of GTP to generate GDP, an RNA acceptor. In the second step, the PRNTase activity of the L protein specifically transfers a 5′-monophosphorylated (p-) RNA moiety of pppRNA with the conserved VSV mRNA-start sequence (AACAG) to GDP to yield a GpppA capped mRNA (Li, Fontaine-Rodriguez et al. 2005; Li, Wang et al.

2006).

Like the unconventional capping enzyme, methylation of the VSV mRNA cap structure is also unique: the mRNA cap is modified by a dual specificity MTase activity within CR VI of L protein whereby ribose 2′-O methylation precedes and facilitates subsequent G-N-7 methylation. Specifically, alterations to the MTase catalytic motif in conserved domain VI of VSV L protein abolished both G-N-7 and

2'-O methylations, suggesting that a single active site is essential for both methylase 25

activities. This is in contrast to all other known MTases, in which the two activities are either catalyzed by separate proteins, each having its own SAM binding site, or, in the case of reovirus, catalyzed by separate domains of the same protein. Interestingly, recent studies of Flaviviruses found that the N terminus of the NS5 protein also encodes dual MTase activities (Li, J., Y., Zhang, 2012).

Figure 1.9 Comparison of mRNA cap formation in eukaryotic cells and VSV (adapted from Li et. al, 2012). (A) Cellular mRNA cap formation. (B) VSV mRNA cap formation.

1.7.4. Capping and methylation of the paramyxovirus mRNA

Similar to VSV L protein, paramyxovirus L possesses all the enzymatic activities needed for viral mRNA synthesis and modifications. Interestingly, it has been shown that paramyxoviruses possess diverse methylated cap structure. For example, measles Sendai virus were found to be methylated at both G-N-7 and 2’-O

positions in their cap structures. In contrast, early studies showed that mRNAs of

Newcastle disease virus (NDV) and hRSV are not 2’-O methylated (Colonno and

Stone 1976). However, recently it has been shown that hRSV mRNA could be doubly methylated at higher SAM concentrations (Liuzzi, Mason et al. 2005).

Sequence alignment found that the signature motifs for mRNA capping and methylation are located in CR-V and CR-VI of paramyxovirus L proteins, respectively.

However, further mechanistic study has been severely hampered due to the lack of a robust in vitro mRNA synthesis for paramyxoviruses, and the technical challenge of expression and purification of a functional paramyxovirus L protein. Recently, a fragment of Sendai virus L protein that includes CR-VI was able to methylate short

Sendai virus-specific RNA sequences in vitro at the G-N-7 position (Ogino, et al.

2005). However, the trans-methylation assay used in that study does not allow the detection of 2’-O methylation. Recently, it was shown that expression a fragment containing CR I to V and another fragment containing CR VI of L proteins from measles virus, Nipah virus, or hRSV separately cannot trans-complement bioactivity of L in an in vitro replicon reporter assay. However, when the fragments of L protein containing CR I-V and CR VI were expressed with a dimerization tag (GCN4) at C terminus and N terminus respectively, the activity of L was restored. These results suggest that CR VI of paramyxovirus L protein mostly likely catalyzes mRNA cap methylation, and additionally this function may require cooperation with the other domains within L. Overall, the mechanism of mRNA capping and methylation in 27

paramyxoviruses has not been characterized. In addition, the capping and methylation status is not known for any member of the genus Metapneumovirus (Dochow, Krumm et al. 2012).

1.7.5. mRNA cap formation as an attractive target for antiviral strategies.

It appears that the entire mRNA capping and methylation machinery in NNS

RNA viruses is different from that of their hosts. This difference, coupled with the fact that replication of NNS RNA viruses occurs in the cytoplasm, suggests that mRNA cap formation is an excellent target for anti-viral drug discovery. Inhibition of the viral mRNA cap formation would likely inhibit downstream events such as replication, gene expression, viral spread, and ultimately viral infection. Since the mRNA of all

NNS RNA viruses contains a methylated cap structure, classes of broadly active anti-viral agents may be developed by targeting the viral cap formation. For human

RSV, several compounds were shown to inhibit polymerase activity which resulted in the synthesis of short uncapped transcripts. RSV mutants resistant to these inhibitors were selected and sequenced. It was found that these resistant mutants contained substitutions in CR V of L, specifically at E1269D, I1381S, and L1421F, suggesting that the mechanism of the action of these compounds is the inhibition of viral mRNA cap addition. Interestingly, these compounds showed strong antiviral activity against

RSV infection in cell culture as well as in a mouse model, demonstrating that mRNA cap addition is an attractive antiviral target. It is known that SAH can inhibit viral mRNA cap methylation. Therefore, many adenosine analogues such as 28

3-deazaeplanocin-A are potent antiviral agents which can significantly inhibit VSV replication in cell culture. The mechanism of the action of these adenosine analogues is through the interference with the host enzyme SAH hydrolase that catalyzes the hydrolysis of SAH to adenosine and L-homocysteine. This reaction is reversible, and the products of this reaction are inhibitory to SAH hydrolase. Obviously, compounds that directly inhibit viral mRNA cap methylation are potent antiviral drugs. For example, sinefungin (SIN), a natural S-adenosyl-L-methionine analog produced by

Streptomyces griseolus, is an inhibitor of methyltransferases. SIN is structurally related to SAM, with the exception that the methyl group that is donated from SAM is replaced by an amino group in SIN. Crystal structures of several MTases have been solved in complex with SIN, which binds to a region that overlaps the SAM binding site. It was found that SIN inhibited VSV G-N-7 and 2’-O methylation with a 50% inhibitory concentration (IC50) of 2.5 μM and 40 μM, respectively. In cell culture,

SIN efficiently inhibited VSV replication, gene expression, and diminished the size of viral plaques without having significant effect on cell viability (Li, Chorba et al.

2007). SIN was also shown to inhibit the MTases of other NNS RNA viruses such as

NDV (Pugh, Borchardt et al. 1978). These examples demonstrate that mRNA capping and methylation is an excellent antiviral target for NNS RNA viruses. An important future direction is to develop high-throughput screening (HTS) to systemically screen compounds that can inhibit mRNA capping and/or methylation of VSV and other

NNS RNA viruses. These inhibitors may be broadly active anti-viral agents against

NNS RNA viruses.

1.7.6. The MTase may provide a new target for the development of a stable

and efficacious live vaccine.

Since mRNAs of NNS RNA viruses are capped and methylated, it is generally believed that viral proteins are translated through a cap-dependent translational pathway. Thus, viral mRNAs lacking a cap structure will be lethal to the virus.

However, inhibition of viral mRNA cap methylation would diminish viral protein translation, which in turn leads to attenuation of the virus since viral replication requires ongoing protein synthesis. Remarkably, recombinant virus lacking mRNA cap MTase can be recovered from the cloned full-length viral cDNA by a reverse genetics system. Thus, this strategy will allow us to rationally design attenuated virus strains by inhibiting mRNA cap methylation. In theory, viruses lacking methylation activity would likely not affect immunogenicity since the MTase is located in the L protein, which is not a neutralizing antibody target. By combining multiple substitutions within the MTase region in the L protein, it should be possible to generate an attenuated virus that is genetically stable, because reversion to wild type at any single amino acid should not provide a fitness gain. Thus, ablating viral mRNA cap methylation would provide a new strategy to rationally attenuate these viruses for development of live attenuated vaccines and their exploitation as viral vectors for vaccines, oncolytic therapy, and gene delivery.

Previously, a panel of MTase-defective VSVs has been generated. Based on the status of mRNA methylation, these recombinant VSVs can be classified into three groups. Viruses in the first group are completely defective in both G-N-7 and 2’-O methylation, including mutations in MTase active site (rVSV-K1651A, D1762A,

K1795A, E1833Q, and E1833A). Viruses in the second group are specifically defective in G-N-7, but not 2’-O MTase, including mutants in SAM binding site

(rVSV-G1670A, G1672A, and S1673A). Viruses in third group that require elevated

SAM concentrations to permit full methylation including a mutant in SAM binding site (rVSV-G1674A) and putative RNA binding site (rVSV-Y1835A). With the exception of rVSV-G1674A and Y1835A, all MTase-defective VSVs were attenuated in cell culture as judged by diminished viral plaque size, reduced infectious viral progeny release (in single-step growth curves), and decreased levels of viral genomic

RNA, mRNA, and protein synthesis (Li, Fontaine-Rodriguez et al. 2005; Li, Wang et al. 2006; Zhang, Wei et al. 2010). Recently, Ma et al., (2013) examined the pathogenicity of these MTase-defective VSVs in mice. It was found that VSV mutants, rVSV-K1651A, D1762A, and E1833Q, which have mutations in the MTase catalytic site and are defective in both G-N-7 and 2’O methylation, were highly attenuated in mice. Recombinant rVSV-G1670A and G1672A, which have mutations in the SAM binding site and are defective in G-N-7 but not 2’-O methylation, retained low virulence in mice. Recombinant rVSV-G1674A, which contains a point mutation in the SAM binding site and requires elevated SAM concentrations to permit full

methylation, was still virulent to mice. These results suggest that MTase is a potential target for generating new live attenuated vaccine candidates for VSV, and perhaps other NNS RNA viruses. This new target for virus attenuation is particularly attractive for paramyxoviruses for which live attenuated vaccine is the most promising vaccine strategy.

1.8. Reverse genetics

NNS RNA viruses can be recovered entirely from cloned cDNA by transfecting mammalian cells with plasmids encoding the viral components of a functional nucleocapsid. This technique is called reverse genetics, or infectious cDNA clone, and it allows us to manipulate the viral genome. Specifically, recombinant viruses can be generated by transfection of cultured cells with a plasmid encoding full-length genomic or antigenomic cDNA and plasmids encoding the major proteins involved in replication and transcription, namely the N, P, and L proteins. For pneumoviruses, this system requires an additional protein, the transcription elongation factor M2-1. For viruses in Filoviridae such as Marburg or Ebola virus, it requires the transcription activator factor VP30.

The first reverse genetics system for NNS RNA viruses was described in 1994 for rabies virus, followed by VSV in 1995. Since then, the reverse genetics for representative members in each family within NNS RNA viruses has been established.

In most cases, plasmid expression is driven by bacteriophage T7 RNA polymerase.

Two strategies have been frequently used for recovery of the virus. First, mammalian cells were infected with a recombinant vaccinia virus expressing T7 polymerase, followed by co-transfection of a plasmid encoding the viral antigenome and support plasmids encoding the nucleocapsid complex. Second, cells (such as BSRT7 cells, derived from a baby hamster kidney line) that constitutively express T7 polymerase were co-transfected with all the plasmids. Intracellular expression of these RNA and protein components results in the assembly of a viral nucleocapsid that is competent for gene expression and genome replication, which leads to the production of infectious virus particles.

The reverse genetics system for hMPV was developed in 2004. HMPV infectious clones for both lineage A and B were successfully established (Herfst, de Graaf et al.

2004). Reverse genetics is a powerful tool to study the functions of each viral protein.

This system allows us to genetically manipulate the viral genome by engineering mutations to viral genome, deleting non-essential viral genes, inserting foreign genes, or swabbing genes from a related virus. Recombinant hMPVs lacking G, SH, M2-1, or M2-2 have been recovered (Biacchesi, Skiadopoulos et al. 2004). In addition, hMPV expressing green fluorescence protein (GFP) has been generated. This system also provides a powerful tool to rationally design live attenuated vaccine candidates

(Biacchesi, Skiadopoulos et al. 2004).

1.9. Animal models for hMPV

Humans are the natural hosts for hMPV. It has been reported that chimpanzees are also naturally infected by hMPV, probably transmitted from the human handlers.

Indeed, approximately 60% of chimpanzees were seropositive for hMPV. However, it is not clear if hMPV can transmit between chimpanzees. Upon experimental hMPV infection, chimpanzees developed clinical signs similar to those observed in humans

(Skiadopoulos, Biacchesi et al. 2004). However, despite the clinical manifestations, the replication titers in upper and lower respiratory tracts of chimpanzees only ranged from

1.5 to 3.2 log10TCID50/mL in nasopharyngeal swab samples. Overall, chimpanzees are the only animal model that has developed diseases similar to humans. However, since

2012, chimpanzees were prohibited from use in biomedical research by NIH because of ethical issues.

In addition to chimpanzees, other nonhuman primates were evaluated to study hMPV pathogenesis, including African green monkeys and rhesus macaques.

Skiadopoulos et al. (2004) reported that rhesus macaques did not exhibit any respiratory tract symptoms when they were intranasally (IN) and intratracheally (IT) infected by 5.2 log10TCID50 of hMPV CAN75 (lineage B) and CAN83 (lineage A).

Nasopharyngeal swab sampling showed that only limited viral replication occurred in rhesus monkeys, ranging from 0.9 to 2.6 log10TCID50 unit. The serum neutralizing antibodies were also relatively low. In comparison, African green monkeys shed viral titer ranging from 2.2 to 4.9 log10TCID50 unit and developed a much higher serum 34

neutralizing antibody titer upon hMPV infection. In both animal models, viral replication was detected in the first 6-7 days post infection. In addition, hMPV immunized rhesus macaques or African green monkeys developed protective immunity against challenge with wildtype hMPV. Thus, while rhesus macaque is less permissive,

African green monkey appears to be a permissive non-human primate model to evaluate the efficacy of vaccine candidates.

Because of the ethical issues, high cost, and limited access to nonhuman primate models, research efforts have been devoted to develop small animal models for hMPV.

To date, mice, hamsters, cotton rats, guinea pigs, and ferrets have been tested to study virus-host interaction, viral pathogenesis, and antiviral immunity. Because of the accessibility and availability of reagents and tools, most studies have been focused on mouse, hamster, and cotton rat models. In the past 10 years, many contradictory results have been reported using these animal models. It seems that many factors such as hMPV strains, environmental conditions (humidity and temperature), sampling methods, and viral titration methods affect the permissiveness of an animal model.

Mice are the most widely used animal used in medical and veterinary research because they are easily accessible and cost effective. Two separate groups reported that hMPV infection in BALB/c led to an appearance of signs of illness, including weight loss, ruffled coat, huddling and respiratory clinical symptoms (such as heavy breathing).

The peak titers in lung tissue occurred at days 7 post-infection (108 PFU/g) and lung

viral titer at day 14 post-infection was 107 PFU/g of tissue (Alvarez, Harrod et al. 2004;

Hamelin, Yim et al. 2005; Schildgen, Simon et al. 2007). This unusual high viral replication has not been observed for any other paramyxoviruses (such as RSV). In addition, many other researchers found that mice are not permissive animal models for hMPV although an extremely high inoculation dose (8 log10TCID50) was used. Thus, the debate about the mouse model for hMPV infection is highly controversial.

Syrian golden hamsters (Mesocricetus auratus) were found to be more permissive for hMPV infection than mice (MacPhail et al., 2004). In this study, Syrian golden hamsters were administered intranasally a 0.1 ml volume containing 1.3 ×106 pfu of hMPV. They found that 105.3 and 104.3 pfu/g tissue of virus was detected in nasal turbinate and lung, respectively, at day 4 post-inoculation. However, cotton rats

(Sigmodon hispidus) appear to be the best small animal models for hMPV. Williams et al. (2005) directly compared the permissiveness of mice, hamster, and cotton rats at the same inoculation dose. They found that cotton rats were more permissive for hMPV than hamsters. In their study, cotton rats were inoculated with 105 PFU of hMPV and were sacrificed at 2, 4, 6, 8, 10, or 14 days postinfection. It was found that the replication of hMPV in the lung tissues peaked on day 4 postinfection at a mean titer of

1.8 × 105 PFU/g and declined gradually. Virus was not detected in the lung after day 6.

Similarly, Wyde et al. (2005) found cotton rats permissive for hMPV replication in nose and lungs, with a peak titer of 3.6 log10PFU/g in the nose and 4.4 log10PFU/g in the lung on day 4 post-infection. HMPV infection in cotton rats is also associated with 36

significant pulmonary histopathological changes. Interestingly, immunohistochemistry revealed that, similar to those of human infection, hMPV antigens were found in airway epithelial cells in lung (MacPhail, Schickli et al. 2004; Hamelin, Yim et al. 2005;

Williams, Tollefson et al. 2005; Wyde, Chetty et al. 2005; Schildgen, Simon et al. 2007;

Mok, Tollefson et al. 2008). In addition, cotton rats have already been proven to be more permissive for hRSV. In fact, cotton rats vaccinated with formalin-inactivated

RSV vaccine developed enhanced pulmonary disease upon wildtype challenge, similar to what has been observed in clinical trials of formalin-inactivated RSV vaccine in children in early 1960. Since cotton rats resembled certain similarities upon respiratory tract infection, it was considered a preferred model for pediatric respiratory tract pathogens (Green, Huey et al. 2013).

1.10. hMPV vaccine development

1.10.1. Significance of vaccine development

A vaccine is the most effective strategy against infectious diseases. Despite major effort, there is no FDA approved vaccine for hMPV, although it is recognized as a globally prevalent pathogen. Epidemiological data suggest that hMPV is one of the major respiratory diseases in the pediatric population as well as the elderly and immunocompromised individuals. It is reported that more than 95% of children have been infected with hMPV by age 5, and hMPV infection occurs frequently and repeatedly throughout life. hMPV infection causes a spectrum of respiratory symptoms

ranging from the common cold, pneumonia, bronchiolitis to asthma and death.

Therefore, there is an urgent need to develop a safe and efficacious vaccine for hMPV.

The priority of a vaccine is to establish long term immunity against natural infection and prevent severe disease caused by lower respiratory infections in the most susceptible populations. Since the discovery of hMPV in 2001, several vaccine candidates have been developed and tested in animal models and nonhuman primates, which will be discussed in this section.

1.10.2. Inactivated vaccine: lessons learned from hRSV inactivated vaccine

It has been a challenge to develop vaccines for human paramyxoviruses. Generally, inactivated and live attenuated vaccines are the two most common strategies used in vaccine development. For safety reasons, an inactivated vaccine is often preferred.

However, development of an inactivated vaccine for human paramyxoviruses resulted in serious complications when tested in human clinical trials. HRSV, a member in the

Pneumovirinae family, is the most significant respiratory pathogen in infants and children. In 1960s, a formalin inactivated (FI) RSV vaccine was developed and tested in clinical trial in children. However, this inactivated vaccine not only failed to induce protective immunity, but also caused vaccine-enhanced pulmonary disease in response to natural RSVinfection. The vaccinated children suffered severe bronchiolitis and pneumonia, which led to 80% hospitalization and two deaths. In contrast, the unvaccinated control group had no deaths and 5% hospitalization. Further studies suggested that the mechanism of enhanced lung damage may be associated with the 38

presence of a Th2 bias response, which led to eosinophilia, goblet cell hyperplasia, overproduction of mucus, production of IgE, and airway hypersensitivity. Recent studies also showed that the incomplete protection and enhanced pulmonary disease may due to insufficient activation of toll like receptors (TLR) by the non-replicable vaccine.

In addition, vaccine-enhanced disease was also observed in an FI vaccine of human parainfluenza type 3 (PIV3), another important human paramyxovirus. Since the discovery of hMPV, an FI hMPV vaccine candidate has been tested in an animal model. de Swart, van den Hoogen et al. 2007; Hamelin, Couture et al. 2007; Yim,

Cragin et al. 2007 showed that FI hMPV vaccine caused enhanced pulmonary disease upon reinfection in cotton rats, similar to that observed for RSV and PIV3. Although the lungs of the vaccinated cotton rats were completely protected from viral replication, dramatically increased lung pathology including enhanced interstitial pneumonitis and alveolitis was observed. These results suggested that inactivated vaccine is not the primary choice for RSV, hMPV, and PIV3, all of which causes severe respiratory disease in same population, infants, childrens, the elderly, and immunocompromised individuals. Therefore, inactivated vaccines were not considered for prevention of human paramyxoviruses.

1.10.3. Live vectored vaccine candidate

Vectored vaccine strategy involves expression of foreign viral antigen using an

attenuated viral backbone. With the development of reverse genetics technology, several paramyxoviruses have been tested as potential vaccine vectors. For example, a live attenuated Newcastle disease virus (NDV) LaSota strain expressing the hemagglutinin protein of avian influenza virus was developed by Avimex (Swayne,

Suarez et al. 2003). Animal experiments showed that this vaccine candidate was effective against both NDV and avian influenza virus. In addition, the major surface glycoproteins of paramyxoviruses have been expressed in a variety of live viral vectors.

For example, Ceva Biomune developed a fowlpox virus vector expressing NDV hemagglutinin-neuraminidase (HN) and fusion (F) protein which led to a bivalent vectored vaccine against both fowlpox virus and NDV (Karaca, Sharma et al. 1998).

Vectored vaccine strategy has been shown to be effective in protecting respiratory diseases. MedImmune has developed a chimeric bovine parainfluenza virus type 3

(bPIV3) backbone expressing HN and F proteins of human parainfluenza virus type 3 and F protein of hRSV(MEDI-534). The resultant vaccine candidates are immunogenic and protective in animal models and currently are under phase 1 clinical trial to evaluate the safety and efficacy in infants and children (Schmidt, McAuliffe et al. 2001; Schmidt 2007).

Several vectored vaccine candidates have been tested for hMPV. Roderick S. Tang et. al reported the use of a chimeric live attenuated bPIV3 harboring hPIV3 F and HN genes (b/h PIV3) to express the hMPV F protein. They showed that the hMPV F

protein was efficiently expressed by the chimeric b/h PIV3 vector. Immunization of

Syrian golden hamsters with this vectored vaccine candidate induced high level of neutralizing antibody and provided protective immunity against wildtype hMPV challenge at 28 days post immunization (Tang, Mahmood et al. 2005). However, it is unknown whether the pre-existing immunity against b/hPIV3 vector will affect the efficacy of the vaccine candidate. In 2008, Mok, Tollefson et al. constructed

Venezuelan equine encephalitis virus replicon particles (VRPs) expressing F

(VRP-MPV.F) or G (VRP-MPV.G) protein of hMPV A2 strain. Intranasal immunization of VRP-MPV.F in cotton rats induced serum neutralizing antibody as well as IgA antibody in secretions at the respiratory mucosa. The immunized cotton rats were protected from viral replication in lungs and nasal turbinate after challenge with an hMPV A2 strain. In addition, no enhanced disease was observed for this vaccine candidate. In comparison, VRP-MPV.G was not immunogenic and protective in animal models. Although this vectored vaccine candidate is promising, the safety is a concern since Venezuelan equine encephalitis virus is a biodefense agent.

1.10.4. Subunit vaccine candidate

It is known that the major surface glycoproteins of paramyxoviruses are responsible for inducing neutralizing antibodies and protective immunities. Thus, subunit vaccine development has been focused on the expression and purification of fusion (F) and attachment (HN, H, or G) proteins. . Previously, the postfusion form of F proteins of PIV3 and RSV, HN protein of PIV3, and G protein of RSV have been 41

purified. Animals immunized with these antigens induced variable levels of neutralizing antibody and reduced viral load in respiratory tracts. In fact, a hRSV F

(postfusion form) subunit vaccine incorporated in nanoparticles was developed by

Novavax, which was found to induce a robust protection in animal models. It is currently under phase 2 clinical trial to assess the efficacy and safety in young women of child-bearing potential (Smith, Raghunandan et al. 2012; Glenn, Smith et al. 2013).

Since subunit vaccines are capable of inducing neutralizing antibody, it could be potentially used for immunization of mothers to produce sufficient maternal antibody to protect infants against RSV infection. Most recently, the crystal structure of prefusion and postfusion RSV F proteins was solved and the immunogenicity of these two forms of F proteins were compared. It was found that prefusion RSV F protein triggered significantly higher neutralizing antibody than postfusion F protein

(McLellan, Chen et al. 2013). Thus, prefusion F protein is a better subunit vaccine for

RSV.

Soluble hMPV F and G proteins have also been purified and tested in animal models (Cseke, Wright et al. 2007). Specifically, cotton rats immunized with a purified soluble F protein (lacking the transmembrane domain) triggered neutralizing antibody responses and provided partial protection against virus replication in the upper and lower respiratory tracts. In addition, F-based subunit vaccine did not induce enhanced pulmonary disease upon reinfection. In a separate study, immunization of Syrian golden hamsters with the soluble F proteins with adjuvant induced high neutralizing 42

titers against both homologous and heterologous hMPV strain. Upon challenge, viral titers in the nasal turbinates of immunized animals were significantly reduced compared with those of unimmunized challenge controls. It is likely that these soluble

F proteins are postfusion forms because they do not have stabilizers to stabilize the F protein. Given the fact that prefusion form of RSV F protein is capable of triggering significantly higher neutralizing antibody than the postfusion form of F protein, it will be interesting to determine the immunogenicity of prefusion form of hMPV F protein in animal models. In comparison, the soluble G protein-based subunit vaccine was immunogenic in cotton rats but was incapable of eliciting neutralizing antibodies and protection against hMPV infection (Ryder, Tollefson et al. 2010). Future experiments to determine whether combination of F and G proteins can enhance the immunogenicity are needed.

1.10.5. DNA vaccine candidate

Similar to the subunit vaccine candidates, a DNA vaccine expressing hMPV F protein (lacking transmembrane domain) was constructed and administered intramuscularly to cotton rats. The F-based DNA vaccine was immunogenic and safe

(inducing no vaccine-enhanced disease). However, only 1.5 log reduction of viral replication in lungs was observed (Cseke, Wright et al. 2007; Liu, Zheng et al. 2009). In addition, it did not protect viral replication in the upper respiratory tract. The results showed that a DNA vaccine may be useful for boosting immunization in combination with other vaccine candidates (such as subunit vaccine), although it cannot induce 43

durable immune responses as it is a non-replicating antigen.

1.10.6. Live attenuated vaccine candidate

Live attenuated vaccines generally mimic natural infection and stimulate humoral, mucosal, and cellular responses, and therefore, usually confer long-term immunity. In contrast to inactivated vaccine candidates, enhanced lung damage has not been observed for live attenuated vaccine candidates in RSV, PIV3, and hMPV. Therefore, live attenuated vaccine candidates are one of most promising vaccine candidates for human paramyxoviruses.

(1) Cold passage (cp) temperature sensitive (ts) live attenuated vaccine candidates

Traditionally, a combination of cold passage (cp) and chemical mutagenesis to induce temperature sensitivity (ts) has been widely used for attenuating respiratory viruses. Since the temperature of upper respiratory tract (approximately 32 oC) is much lower than lower respiratory tract (37 oC), the replication of cpts viruses is expected to be restricted in upper respiratory tract, which may be able to trigger sufficient protection against respiratory diseases. This strategy leads to the successful development of live attenuated influenza virus vaccine which is currently used in human population. Using similar strategy, a number of cpts RSV vaccine candidates have been developed and tested clinically (Karron, Wright et al. 1997; Wright, Karron et al. 2000). However, none of them were found to have satisfactory balance between

attenuation and immunogenicity. For example, cpts RSV vaccine candidates cause upper respiratory tract congestion associated with immunization, raising the concern of the safety issue. The current lead vaccine candidate (MEDI-559) encodes a number of cpts mutations as well as a deletion of the SH gene. This candidate was found to have an acceptable safety profile. However, this candidate vaccine induced detectable antibody responses in only 44% of vaccinees, even after two doses. Thus, enhancement of the immunogenicity of cpts RSV vaccine candidates is needed.

Soon after the discovery of hMPV, a number of cpts hMPV strains have been isolated by repetitive passing of the virus in cell culture for 35 passages at temperature as low as 25 oC (Herfst, de Graaf et al. 2008). These cpts-hMPVs were attenuated in replication in cell culture. Infection of hamsters with the vaccine candidates showed significant reduction in viral replication in upper respiratory tract, and no replication in lower respiratory tract. Immunization of hamsters showed increased neutralizing antibody and protective immunity in both upper and lower respiratory tracts against wildtype hMPV challenge. Genomic sequencing identified 20 amino acid mutations in these cpts hMPV strains. Although the detailed functions of these mutations were not clear, these cpts mutations could be combined with “rational” designed vaccine strategy

(such as gene deletion mutations) using the reverse genetics techniques. To date, the safety and immunogenicity of these cpts hMPV vaccine candidate in humans are unknown.

(2) Gene-deleted live attenuated vaccine candidate

More recently, the use of reverse genetics systems, whereby infectious virus is derived from a full-length cDNA clone, has allowed the development of live attenuated vaccine candidates encoding specific mutations, gene deletions and insertions, and chimeric genes. This strategy provides new avenues for developing safe and effective paramyxovirus vaccines. The nonessential proteins in RSV include the nonstructural

NS1 and NS2, the attachment G, the small hydrophobic (SH), and the M2-2. These proteins have been shown to be dispensable for in vitro replication. Deletion of these genes can result in attenuation of RSV. HRSV lacking NS1, NS2, G, SH, and M2-2 have been recovered. These gene-deleted hRSVs were highly attenuated and provided a variable level of protection against hRSV challenge in animal models. However, it has been a challenge to balance the attenuation and immunogenicity. For example, hRSV lacking NS1 or G protein were too attenuated to generate sufficient protective immunity.

In contrast, hRSV lacking SH protein is not sufficiently attenuated. When cpts mutations and SH gene deletion were combined, the resultant vaccine candidate

(MEDI-559) has a satisfactory safety profile but not sufficient immunogenicity.

In hMPV, these nonessential proteins include the attachment G, the SH, the M2-1 and M2-2. Indeed, recombinant hMPV mutants lacking SH, G, M2-1, and M2-2 have been generated using reverse genetics techniques (Biacchesi, Pham et al. 2005;

Buchholz, Biacchesi et al. 2005; Schickli, Kaur et al. 2008). The rhMPV lacking G or

SH protein replicates efficiently in cell culture. When rhMPV-∆G was administered 46

intranasally to hamsters, 40 fold and 600-fold reduction in viral replication in upper and lower respiratory tract, respectively, were observed. The rhMPV-∆G mutant induced a significantly lower level of neutralizing antibody compared to the wildtype rhMPV.

Despite the lower antibody response, hamsters were completely protected from wildtype hMPV challenge. Recent studies have shown that hMPV G modulates the innate immune response, which subsequently affects adaptive immunity. Thus, rhMPV-∆G may impair the immunogenicity. In contrast, rhMPV-∆SH showed a slight increase in viral replication in hamsters. It appears that rhMPV-∆SH is not sufficiently attenuated in vivo. Thus, it may not be feasible for using as a live attenuated vaccine.

In another study, rhMPVs lacking M2-1, M2-2, or both were recovered. This is in contrast to RSV for which deletion of M2-1 is lethal to virus. Both rhMPV-∆M2-1 and rhMPV-∆M2-(1+2) replicate efficiently in cell culture. However, no infectious virus can be recovered from lung and nasal turbinate after inoculation to hamsters, suggesting that they were highly attenuated in vivo. Unfortunately, no neutralizing antibody or protective immunity was observed. It appears that rhMPV-∆M2-1 and rhMPV-∆M2-(1+2) were too attenuated to generate sufficient immunity. Recombinant rhMPV-∆M2-2 also showed efficient replication in cell culture and was highly attenuated in hamsters. Interestingly, hamsters immunized with rhMPV-∆M2-2 developed a high level of neutralizing antibody and were completely protected from wildtype challenge. Thus, rhMPV-∆M2-2 is a potential live attenuated vaccine candidate. 47

In addition, the replication and immunogenicity of rhMPVs lacking SH, G, or M2 have been evaluated in African green monkeys. Similar to the observation in hamster models, rhMPV-∆SH was not attenuated in nonhuman primates either. However, rhMPV-∆G and ∆M2-2 were reduced 6-fold and 160-fold in upper respiratory tract and

3200-fold and 4000-fold in the lower respiratory tract, respectively. These two deletion mutants were highly immunogenic and protective against wildtype challenge

(Biacchesi, Pham et al. 2005).

(3) Chimeric live attenuated vaccine candidate

Another strategy is to generate live attenuated chimeric vaccines, which involves exchange of genes between closely related viruses. Previously, it was shown that replacement of bRSV F and G genes with their counterparts in hRSV yielded a chimeric virus that replicated efficiently in vitro. This chimeric virus is highly attenuated in chimpanzees, but didn’t protect chimpanzees against subsequent challenge with wild-type hRSV. This suggests that the chimeric virus is too attenuated to elicit protective immune responses.

Avian metapneumovirus (aMPV) is the only other member in the pneumovirus genus, and is closely related to hMPV. In 2005, Pham, Biacchesi et al. reported a chimeric vaccine candidate in which the N or P protein of hMPV backbone was replaced with that of aMPV type C. Surprisingly, the N and P chimeric rhMPVs replicated to a peak titer that was 11- and 25- fold higher than that of wildtype hMPV,

respectively. In contrast to the enhanced viral replication in vitro, the N and P chimeras were reduced approximately 100-fold in upper and lower respiratory tracts in hamsters on day 3 postinfection. In African green monkeys, replication of the P chimeric hMPV was reduced 100- and 1000-fold in upper and lower respiratory tract, respectively.

Replication of N chimeric hMPV was reduced 10-fold in lower respiratory tract but no attenuation was observed in upper respiratory tract. Thus, the P chimeric hMPV is more attenuated than the N chimeric hMPV. In hamster and non-human primate models, both chimeric hMPVs induced high levels of neutralizing antibody and protective immunity.

The safety and immunogenicity of P chimeric rhMPV vaccine candidate is currently under phase 1 human clinical trial.

1.10.7. Conclusion:

In recent years, a number of hMPV vaccine candidates including subunit vaccines, vectored vaccines, and live attenuated vaccines have been evaluated in small animal models and nonhuman primates; however, there remains no currently licensed hMPV vaccine. Still, the live attenuated vaccine is the most promising vaccine candidate.

These live vaccines offer several advantages: (i) enhanced lung damage has not been observed either after natural infection or vaccination with live attenuated viruses; (ii) live attenuated hMPV vaccines mimics natural viral infection that can induce balanced and long-term immune responses; and (iii) intranasal vaccination with live attenuated viruses should induce better local immunity compared with intramuscular injection of subunit vaccines. Although live attenuated vaccine is the most attractive vaccine 49

strategy, the major challenge is to identify a vaccine strain that has an optimal balance between attenuation and immunogenicity. Live attenuated RSV vaccines have been in development for several decades, but none were found to have satisfactory balance between attenuation and immunogenicity. Similarly, several live attenuated hMPV vaccines have been developed and tested in animal models. Most of them were either not sufficiently attenuated or lacked a robust immunogenicity. Therefore, there is urgent need to explore new approaches to generate live hMPV vaccines that are highly attenuated but retain high immunogenicity. The goal of this thesis is to explore new live attenuated hMPV vaccines by inhibiting viral mRNA cap methyltransferase.

1.11. Treatment and prevention

Currently, there is no standardized treatment or preventive strategy against hMPV.

Ribavirin is a synthetic guanosine triphosphate (GTP) analog, which acts by inhibiting viral RNA polymerase. It has been successfully used for treatment of RSV infection in lung transplant recipients. Hamellin et al. (2008) showed that ribavirin has a similar inhibitory activity against hMPV in vitro and in a mouse model. Recently, ribavirin has been successfully used for the treatment of immunocompromised patients infected by hMPV. However, the application of ribavirin treatment is limited due to its known teratogenic potential.

Several antibodies specifically targeting F protein have been shown to neutralize all lineages of hMPV in vitro. Passive immunization by intranasal delivery of the antibody to cotton rats significantly reduces viral replication in both lung and nasal tissues. These antibodies showed prophylactic or therapeutic applications against 50

severe hMPV infection.

Chapter 2.

Development and optimization of a direct plaque assay for

human and avian metapneumoviruses

2.1. Abstract

The genus Metapneumovirus within the subfamily Pneumovirinae and family

Paramyxoviridae includes only two viruses, human metapneumovirus (hMPV) and avian metapneumovirus (aMPV), which cause respiratory disease in humans and birds, respectively. These two viruses grow poorly in cell culture and other quantitation methods, such as indirect immuno-staining and immuno-fluorescent assays, are expensive, time consuming, and do not allow for plaque purification of the virus. In order to enhance research efforts for studying these two viruses, direct plaque assays for both hMPV and aMPV have been developed. By optimizing the chemical components of the agarose overlay, it was found that both hMPV with a trypsin-independent F cleavage site and aMPV formed clear and countable plaques in a number of mammalian cell lines (such as Vero-E6 and LLC-MK2 cells) after 5 days of incubation. The plaque forming assay has similar sensitivity and reliability as the currently used immunological methods for viral quantitation. The plaque assay is also a simpler, rapid, 52

and economical method compared to immunological assays, and in addition allows for plaque purification of the viruses. The direct plaque assay will be a valuable method for the quantitation and evaluation of the biological properties of some metapneumoviruses.

2.2. Introduction

Human metapneumovirus (hMPV) is a non-segmented negative-sense RNA virus belonging to the family of Paramyxoviridae, the subfamily Pneumovirinae, and the genus Metapneumovirus (van den Hoogen et al., 2001). There are at least two lineages of hMPV circulating in human population, designated A and B, which can be further divided into A1, A2, B1, and B2, based on their surface glycoproteins and antigenicity

(van den Hoogen et al., 2004). Since its discovery in 2001, hMPV has been recognized worldwide as one of the leading causes of lower respiratory infections, second only to human respiratory syncytial virus (hRSV) (van den Hoogen, de Jong et al. 2001; Kahn

2006). hMPV infections are observed in all age groups with a high prevalence and severity among infants, children, the elderly, and immunocompromised patients

(Stockton, Stephenson et al. 2002; Bastien, Normand et al. 2003; Boivin, De Serres et al.

2003; Falsey, Erdman et al. 2003; van den Hoogen, van Doornum et al. 2003; Williams,

Harris et al. 2004; Falsey 2008; Falsey, Hennessey et al. 2010). Clinical symptoms associated with hMPV infection are similar to those caused by hRSV, ranging from asymptomatic infection to severe bronchiolitis and pneumonia (Boivin et al., 2002;

Esper et al., 2003). To date, no vaccines or antiviral drugs are available against this 53

important pathogen.

Avian metapneumovirus (aMPV) is the only other member in the

Metapneumovirus genus. Discovered in the 1970s, aMPV has been recognized as an important pathogen for the poultry industry worldwide (Buys, du Preez et al. 1989;

Buys, du Preez et al. 1989). aMPV is the causative agent of tract reproaccute rhinotracheitis (TRT) in turkeys, and is associated with swollen head syndrome (SHS) in chickens (Buys, du Preez, and Els, 1989). It causes respiratory tract and reproductive infections with low mortality but high morbidity in turkeys and chickens, resulting in economic losses in both egg and poultry production for the turkey and chicken industries. Based on genetic and serological homology, aMPV is divided into four antigenic subtypes, namely, A, B, C, D (Bayon-Auboyer, Jestin et al. 1999;

Bayon-Auboyer, Arnauld et al. 2000). Interestingly, aMPV subtype C is more closely related to hMPV than the other aMPV subtypes.

One of the major challenges in metapneumovirus research is the lack of a convenient, economic, and reliable quantitation method, which is due mainly to the fact that both hMPV and aMPV grow poorly in cell culture. It usually takes more than 10 days for hMPV to develop significant cytopathic effects (CPE) in mammalian cells.

The growth of some hMPV strains is trypsin-dependent and, in general, it has been shown that aMPV grows better than hMPV (van den Hoogen et al., 2001). However, it takes more than 5 days for aMPV subtype C strains to develop typical CPE.

Currently, indirect immunostaining or an immunofluorescence assay (IFA) followed by TCID50 calculations are used to determine the titer of hMPV and aMPV

(van den Hoogen et al., 2001; Herfst et al., 2004; Ebihara et al., 2005). Unfortunately, this methodology is relatively complicated and costly. Although hMPV forms small spots in immunostaining-based assays, it requires fixation of the cells followed by antibody straining. The viruses are inactivated by fixation and therefore purification of the virus post-staining cannot be executed. Agarose overlay plaque assays have been used to quantify, separate, and propagate a great number of viruses. However, no plaque assay for tittering hMPV has been reported to date. As for aMPV, only one study has shown that aMPV subtype C can form visible plaques in the Japanese quail fibrosarcoma cell line (QT-35) after 6 days incubation (Sabara and Larence 2002;

Sabara, Larence et al. 2003). It is not known whether aMPV can form plaques in other cell lines.

In this study, an agarose overlay plaque assay is developed for titer determination and plaque purification of hMPV strains with trypsin-independent cleavage sites in F protein and aMPV using African green monkey kidney cells (Vero) and rhesus monkey kidney cells (LLC-MK2). The results indicate that plaque formation can occur in 5 days with sensitivity comparable to that of the conventional indirect immunostaining assays.

In addition, the plaque assay is more convenient and economical than other immunological methods. Hence, the direct plaque assay developed in this study could provide a valuable method for detection, isolation, purification, and quantitation of 55

aMPV and some hMPV strains with self-cleavable F protein.

2.3. Materials and methods

2.3.1. Cell culture

African green monkey kidney epithelial cells (Vero ATCC no. CCL-81 and Vero

E6 ATCC no. CRL-1586) were cultured in Dulbecco’s modified Eagle’s medium

(DMEM; Invitrogen, Carlsbad, CA, USA) supplemented with 10% fetal bovine serum

(FBS; Invitrogen) in a humidified incubator at 37 °C and 5% CO2. Rhesus monkeys kidney cell (LLC-MK2) cells were maintained in OPTI-MEM reduced serum medium

(Invitrogen) supplemented with 2% FBS under the same incubation conditions.

2.3.2. Virus recovery and characterization

An infectious cDNA clone of hMPV lineage A strain NL/1/100 was kindly provided by Dr. Ron A. M. Fouchier at Department of Virology, Erasmus Medical

Center, Rotterdam, The Netherlands. Recombinant hMPV (rhMPV) was isolated using a reverse genetics system (Herfst et al., 2004; Biacchesi et al., 2004). Briefly, rhMPV was recovered by co-transfection of a plasmid encoding the full-length genomic cDNA of hMPV NL/1/00 (phMPV) and support plasmids encoding viral N (pCITE-N), P

(pCITE-P), L (pCITE-L), and M2-1 (pCITE-M2-1) proteins into BHK.SR19T7pac cells (kindly provided by Apath LLC, Brooklyn, NY, USA) which express stably the T7

RNA polymerase. Six days post-transfection, the cells were subjected to three freeze-thaw cycles followed by centrifugation at 3000×g for 10 min. The supernatant 56

was used subsequently to infect LLC-MK2 cells. Since hMPV requires trypsin to grow,

TPCK-trypsin was added to the media to the final concentration of 0.1µg/ml at day 2 post-infection. Cytopathic effect (CPE) was observed 5 days post-infection and the recovered viruses were amplified further in LLC-MK2 cells.

The F cleavage site (99RQSR102) in the full-length genome of wild type hMPV

NL/1/100 was mutated into 99RRRR102 (Fig. 1A) using the QuikChange Site-Directed

Mutagenesis Kit (Strategene, La Jolla, CA, USA) with the following primers:

Forward:

5’-GAGAGGAGCAAATTGAAAATCCCAGACGACGTAGATTCGTTCTAGGAG

CAATAGC -3’;

Reverse:

5’-GCTATTGCTCCTAGAACGAATCTACGTCGTCTGGGATTTTCAATTTGCTC

CTCTC-3’.

Recombinant hMPV carrying the trypsin-independent F cleavage site (rhMPV-F) was recovered as described above. Recombinant rhMPV-F exhibited trypsin-independent growth, and was propagated subsequently in LLC-MK2 cells in the absence of trypsin.

aMPV subtype C Minnesota (MN) strain was a generous gift from Mo Saif at the

Ohio Agricultural Research and Development Center (OARDC, Wooster, OH) and was propagated in LLC-MK2 cells.

2.3.3. Cytopathogenic effect(CPE)-based TCID50 assay for aMPV

LLC-MK2 cells were seeded in 96 well plates (Corning, Lowell, MA, USA) at a density of 105 cells per well and were grown at 37 °C for 18 h. Upon infection, the medium was removed and 0.2 ml of tenfold serial virus dilutions in Opti-MEM was added to each well. Eight wells containing a monolayer of cells were infected with 50

µl of each virus dilution and the cytopathogenic effect (CPE) was examined under a microscope daily for 10 days post infection. The CPE was recorded and the virus titer was calculated as the tissue culture infection dose (TCID50) using the Reed-Muench method (1938).

2.3.4. An indirect immunostaining assay for rhMPV

LLC-MK2 cells were seeded and then infected with serial dilutions of rhMPV or rhMPV-F as described in Section 2.3. At day 6 post-infection, the supernatant was removed and cells were fixed in a pre-chilled acetone: methanol solution (at the ratio of

3:2) at room temperature (RT) for 15 min. Cells were permeablized in a phosphate saline buffer (PBS) containing 0.4% Triton X-100 at RT for 10 min, and blocked at

37 °C for 1 h using 1% bovine serum albumin (BSA) in PBS. The cells were then labeled with an anti-hMPV N protein primary monoclonal antibody (Millipore,

Billerica, MA) at dilution of 1:1,000, followed by incubation with horseradish peroxidase (HRP)-labeled rabbit anti-mouse secondary antibody (Thermo Scientific,

Waltham, MA, USA) at dilution of 1:5,000. After incubation with AEC substrate

chromogen (Sigma, St. Louis, MO, USA), positive cells were then visualized under the microscope. Viral titer was calculated as TCID50 using the Reed-Muench method

(1938).

2.3.5. Direct plaque assays for rhMPV-F and aMPV

Vero or Vero-E6 cells were seeded in 6-well plates (Corning) at the density of

2×106 cells per well. After incubation for 18 h, the medium was removed and cell monolayers were infected with 400 µl of a 10-fold dilution series of each virus. After incubation at 37°C for 1 h with agitation every 10 min, the cells in each well were overlaid with 2.5 ml of Eagle minimum essential medium (MEM) containing 1%

agarose, 1% FBS, 0.075% sodium bicarbonate (NaHCO3), 20mM HEPES (pH 7.7), 2 mM L-Glutamine, 12.5 mg/mL of penicillin, 4 mg/mL of streptomycin, and 4 mg/mL of kanamycin. The plates were incubated at 4 oC for 30 min to solidify the overlay media.

o Cells were then grown at 37 C and 5% CO2 to allow for plaque formation. Where indicated, the overlay medium was supplemented with actinomycin-D (Sigma) (0.1 to

0.6 µg/ml) or TPCK-trypsin (0.1 to 0.6 µg/ml). After incubation for 4-10 days, the cells were fixed in 10% (v/v) formaldehyde for 2 h, and the plaques were visualized by staining with 0.05% (wt/vol) crystal violet.

2.3.6. Growth kinetics of rhMPV-F

Confluent LLC-MK2 cells in 35-mm dishes were infected by rhMPV-F at a multiplicity of infection (MOI) of 0.1. After 1 h of adsorption, fresh DMEM 59

(supplemented with 2% FBS) was added, and infected cells were incubated at 37°C. At different time points post infection, the cells were harvested by three freeze-thaw cycles followed by centrifugation at 1,500 × g, RT for 15 min. Virus titer was determined using indirect immunostaining assays or direct plaque assays on LLC-MK2 cells as described above.

2.3.7. Statistical analyses

All experiments were carried out in triplicate. Statistical analysis was performed by one-way multiple comparisons using SPSS 8.0 statistical analysis software (SPSS

Inc., Chicago, IL). A value of p <0.05 was considered statistically significant.

2.4. Results

2.4.1. aMPV, but not rhMPV, formed plaques in an agarose overlay plaque

assay.

First, the ability of aMPV to form plaques in Vero, Vero E6, and LLC-MK2 cells was investigated, because these cell lines are susceptible to metapneumovirus infection.

As shown in Fig. 2.1A, aMPV subtype C MN strain formed clear and countable plaques in all three cell lines tested. Plaques produced in Vero cells were round and much clearer than those in Vero E6 or LLC-MK2 cells (Fig.2.1A). Further microscopic examination showed that the edge between the intact and lysed cells was clear in Vero cells (Fig. 2.1B). In contrast, the cells at the edge of the plaque were not completely detached in LLC-MK2 and Vero E6 cells, leading to the irregular-shaped plaques. 60

Further optimization of the experimental conditions was established by including 0.1

µg/ml of actinomycin-D, an inhibitor of mammalian cell proliferation, in the agarose overlay media. The addition of actinomycin-D produced plaques that were visible and countable at day 4 post-infection in all three cell lines. As shown in Fig 2.1 C and D, plaques formed in the presence of actinomycin-D were significantly larger than those in the absence of actinomycin-D. Inhibition of cell growth with actinomycin-D significantly improved the ability of aMPV to replicate in cell culture and improved the quality of the plaque assay results.

Figure 2.1 Plaque formation of aMPV subtype C MN strain in Vero, Vero-E6, and LLC-MK2 cells. (A) Plaque morphology of aMPV in different cell lines. Agarose overlay plaque assay was performed in six-well plates as described in Materials and Methods. After incubation for 6 days, the plates were fixed in 10% formaldehyde, and the plaques were visualized by staining with 0.05% crystal violet. (B) Microscopic examination of viral plaques in different cell lines. Plaques were examined under at 100× magnification under light microscope. Digital photographs of plaque were taken under a Nikon TS100 inverted phase-contrast microscope mounted with a Nikon Coolpix995 camera. (C) Plaque morphology of aMPV in Vero cells without actinomycin-D. Plaque assay was performed without actinomycin-D. Plaques were developed at day 4 post-infection. (D) Plaque morphology of aMPV in Vero cells with actinomycin-D. The agarose overlay contains 0.1 μg/mL of actinomycin-D. Plaques were developed at day 4 post-infection.

Subsequently, an attempt was made to develop a similar plaque assay for hMPV. hMPV is known to grow much poorer compared to aMPV in most of the cell lines tested. Also, some hMPV strains require trypsin for growth, so in order to optimize the plaque assay for hMPV growth the assay was conducted as described above with the following modifications. First, the overlay medium was supplemented with 0.2

µg/ml of trypsin-TPCK but no FBS. Second, the time for plaque development was

extended to 3 weeks. Third, due to the high sensitivity of Vero cells to trypsin at concentrations required for hMPV growth, only Vero E6 and LLC-MK2 cells were chosen for this study. Unfortunately, no visible plaques were formed by hMPV in

LLC-MK2 or Vero E6 cells after incubation for up to three weeks post-infection (data not shown). In addition, increasing the concentration of trypsin and/or including actinomycin-D in the overlay medium did not led to any observable plaques, indicating that rhMPV is not able to form plaques in Vero E6 or LLC-MK2 cells although both cell lines are permissive for hMPV replication.

2.4.2. Recombinant hMPV with a trypsin-independent F cleavage site

(rhMPV-F) formed plaques in both Vero E6 and LLC-MK2 cells.

It has been reported that the growth of hMPV clinical isolates was improved in the presence of trypsin in cell culture medium. For paramyxoviruses, the F protein cleavage site sequence is the major determinant for trypsin (Collins, Bashiruddin et al.

1993; Lamb 1993). The F protein of hMPV NL/1/100 strain contains a trypsin-dependent cleavage site (99RQSR102 motif) (Biacchesi, Pham et al. 2006).

Hence, it was speculated that the inability of hMPV NL/1/100 strain to form plaques in

Vero E6 and LLC-MK2 cells could be associated with its putative cleavage site as well as trypsin dependence. To address this premise, the 99RQSR102 (Biacchesi, Pham et al.

2006) motif was mutated to 99RRRR102 in the F cleavage site in an infectious clone of hMPV NL/1/100, and successfully recovered the recombinant hMPV carrying the trypsin-independent F cleavage site (rhMPV-F). As expected, rhMPV-F displayed a 63

trypsin-independent growth pattern in LLC-MK2 cells (Fig. 2.2). Next, the evaluation of the ability of rhMPV-F to form plaques in Vero, Vero E6, and LLC-MK2 cells was carried out using an approach similar to that for aMPV. Remarkably, visible plaques were observed in Vero, Vero E6, and LLC-MK2 cells infected with rhMPV-F after 8 days incubation (Fig. 2.3). In addition, plaques formed in Vero and Vero E6 cells had larger and more pronounced zones of clearing and were easier to count than those in

LLC-MK2 cells. Taken together, these results demonstrated trypsin-independent rhMPV-F was able to form plaques in an agarose overlay plaque assay, suggesting that a direct agarose plaque assay is applicable for some (if not all) hMPV strains with trypsin-independent F cleavage sites.

Figure 2.2 Trypsin dependence of rhMPV-F and rhMPV. LLC-MK 2 cells were infected by rhMPV-F or rhMPV and incubated with DMEM with or without 0.2μg/mL of trypsin-TPCK. Indirect immunostaining assay was performed at day 6 post-infection. After removal of cell culture medium, the cells were fixed in acetone: methanol (at the ratio of 3:2) and permeablized in PBS containing 0.4% Triton X-100. The cells were then probed with anti-hMPV N monoclonal antibody, followed by incubation with HRP-conjugated rabbit anti-mouse antibody. After incubation with AEC substrate chromogen, positive cells were visualized under light microscope.

Figure 2.3 Plaque formation of rhMPV-F in Vero, Vero-E6, and LLC-MK2 cells. (A) Plaque morphology of rhMPV-F in different cell lines. Agarose overlay plaque assay was performed in six-well plates as described in Materials and Methods. Plaques were developed at day 6 post-infection. (B) Microscopic examination of viral plaques in different cell lines. Plaques were examined under at 100× magnification under light microscope.

2.4.3. Optimization of a direct plaque assay for metapneumoviruses

Modification of the parameters of the direct plaque assay was then optimized so that plaques could be visualized and detected in the shortest possible time. First, the effect of growth temperature on the plaque formation was evaluated. It was found that rhMPV-F formed significantly bigger plaques at 37 oC than at 30 oC after seven days of infection, indicative of a preference for the higher temperature. The average size of a total of twenty plaques formed at 37 oC at day 7 post-infection was 1.0 ± 0.12 mm, whereas the plaque size formed at 30 oC was 0.44 ± 0.11 mm. Secondly, the effect of cell confluence on the plaque formation was tested. Vero cells were seeded at different 66

densities to achieve 50%, 80%, and 100% confluency after incubation at 37 oC for 18 h, and plaque assays were performed as described previously. There was a reduction in the size of plaques observed as the cell confluency increased. The average plaque size in cells of 80% confluency at day 7 post-infection (1.1 ± 0.2 mm) was significantly larger than that in cells of 100% confluency (0.9 ± 0.2 mm) (p < 0.05). In addition, larger plaques were observed when the cell confluency was reduced to 50%. However, cells at 50% confluency made it challenging to count the plaques due to the light background after staining with crystal violet. Therefore, it was concluded that a cell culture of 80% confluence was the most suitable for the direct plaque assay taking into account the plaque size and clear background. Thirdly, the impact of actinomycin-D on the plaque formation of rhMPV-F was investigated. The plaque assay was performed in Vero cells with 80% confluency as described previously except that the agarose overlay medium contained various amounts of actinomycin-D and the plaque was developed at day 7 after incubation. As shown in Fig. 2.4, actinomycin-D increased the plaque size of rhMPV-F in a dose dependent manner. However, high concentrations of actinomycin-D

(0.5- 0.6 g/mL) brought about significant damage to the monolayer of cells. In contrast, 0.1-0.2 µg/mL of actinomycin-D was found to be the optimal concentration, as it increased significantly the plaque size (P<0.05) but did not cause any noticeable damage to the cell monolayer. Fourth, the shortest time for forming visible plaques was determined, as well as the dynamics of the plaque size formation for rhMPV-F and aMPV (Fig. 2.5). Plaque assays were carried out in the presence of 0.1 µg/mL of

actinomycin-D, plates were incubated at 37oC, and fixed from days 4-10 post-infection.

For both rhMPV-F and aMPV, the plaque size became bigger as the incubation time increased (Fig. 2.5). However, aMPV appeared to be more efficient in forming plaques than rhMPV-F. For rhMPV-F, visible plaques were observed at day 5 post-infection, and became countable at day 7 post-infection. In contrast, aMPV plaques could be visualized at day 4 and became countable at day 5 post-infection.

Finally, the effect of actinomycin-D on the accuracy of the direct plaque assay was evaluated. The stock of aMPV and hMPV was titrated using the direct plaque assay with or without 0.2 µg/mL of actinomycin-D. As shown in Fig. 2.6A, the plaques in actinomycin-D treated plates (developed at day 7) were bigger than those in non-treated plates (developed at day 10). As shown in Fig.2.6B, there was no significant difference between the titers determined by plaque assay with or without actinomycin-D (P>0.05).

Therefore, actinomycin-D increased the size of the plaques without altering the number of the plaques.

Figure 2.4 The effect of actinomycin-D on plaque formation of rhMPV-F. The plaque assay was performed in Vero cells with 80% confluency. Vero cells in six-well plates were infected with 400 µl of a 10-fold dilution series of rhMPV-F. The agarose overlay medium contained various amounts of actinomycin-D and the plaques were developed at day 7 post-infection. One representative well (dilution range from105 to107) from each six-well plate was shown. The size of individual plaque was measured by Photoshop software. The average size of a total of twenty plaques was shown.

Figure 2.5 Dynamics of plaque formation of rhMPV-F and aMPV in Vero cells. Direct plaque assay were carried out in the presence of 0.1 µg/mL of actinomycin-D and plates were incubated at 37oC and fixed from day 3 to 11 post-infection. One representative well (dilution range from105 to107) from each six-well plate was shown. The average size of a total of twenty plaques was shown.

Figure 2.6 Effect of actinomycin-D on the number and size of viral plaques. (A) Effect of actinomycin-D on the size of aMPV and rhMPV-F plaques. Plaque assay with or without 0.2 µg/ml of actinomycin-D was developed at days 7 and 10 post-infection, respectively. (B) Effect of actinomycin-D on the number of aMPV and rhMPV-F plaques. The titer of aMPV and rhMPV-F stocks was determined by direct plaque assay with or without 0.2 µg/ml of actinomycin-D. Titers were the average of three independent experiments.

Taken together, the optimized direct plaque assay for rhMPV-F was determined as the following: Vero or Vero E6 cells in six-well plates with 80% confluency infected by rhMPV-F; after 1 hour incubation at 37oC, the agarose overlay medium containing

0.1µg/mL of actinomycin-D is added to the cells; and the plates are incubated at 37oC for at least 5 days. 71

2.4.4. Sensitivity of the direct plaque assay

The sensitivity of the optimized direct plaque assay was compared to other conventional methods, such as TCID50 and indirect immunostaining assays. Briefly,

Vero E6 cells were infected by aMPV and rhMPV-F at MOI of 1, and viruses were harvested at day 7 post-infection. The titer of aMPV and rhMPV was determined by direct plaque assay, TCID50, or indirect immunostaining assay. Overall, the virus titer from direct plaque assays was consistent with the corresponding values determined by

TCID50 (aMPV) and indirect immunostaining (rhMPV-F). While the viral titer of

7 rhMPV-F determined by the direct plaque assay is 1.7 ± 0.2×10 PFU/mL, the

7 corresponding values determined by immunostaining assay is 3.8 ± 1.4×10 TCID50/mL.

6 For aMPV, the viral titer determined by direct plaque assay was 2.16 ± 0.29×10 , the

6 corresponding values determined by TCID50 was 3.76 ± 0.78×10 . The correlation in plaque formation between indirect immunostaining and direct plaque assays was also compared. Both methods were able to detect the presence of rhMPV-F at the highest virus dilution tested in this study (1:107), suggesting that the direct plaque assay has a comparable sensitivity to that of the conventional immunostaining method.

Figure 2.7 Growth kinetics of rhMPV-F determined by direct plaque assay and immunostaining assay. Confluent LLC-MK2 cells in 35-mm dishes were infected with rhMPV-F at an MOI of 0.1. After a 1-h adsorption, the inoculum was removed, the cells were washed with DMEM, and fresh medium (containing 2% FBS) was added, followed by incubation at 37°C. At the indicated intervals, the cells were harvested by three freeze-thaw cycles followed by centrifugation at 1,500 × g for 15 min. Virus titers were determined by direct plaque assay, indirect immunostaining assay or TCID50 assay in Vero cells. Titers were the average of three independent experiments.

Finally, the direct plaque assays were used to study the growth kinetics of rhMPV-F in LLC-MK2 cells. As shown in Fig. 2.7, the results reveal an exponential phase of rhMPV-F replication until day 6 post-infection whereas the virus titer starts to drop after day 8 post-infection. The growth curve generated from direct plaque assay was indistinguishable from those of TCID50 and indirect immunostaining assays, suggesting that the direct plaque assay is as reliable as other conventional methods.

2.5. Discussion

aMPV and hMPV are the only two known virus members in the genus 73

Metapneumovirus within the subfamily Pneumovirinae and family Paramyxoviridae. It has been a challenge to work with these viruses because they grow poorly in cell culture.

Although aMPV was discovered forty years ago, quantitation of this virus has been limited to TCID50 assay by observation of cytopathic effect (CPE) in permissive cell lines. hMPV was first discovered in 2001 and was found to grow much poorer than aMPV in cell culture. Quantitation of hMPV is limited to indirect immunostaining assay by examination of viral antigen expression using antibody staining. In general,

RNA viruses have a high mutation rate and it has been reported that both aMPV and hMPV can easily undergo gene truncation and/or mutation during passing in mammalian cells (Govindarajan et al., 2004; Agrawal et al., 2001). Therefore, there is an urgent need to develop a direct plaque assay that allows for plaque purification of virus. In this study, direct plaque assays for aMPV and trypsin-independent hMPV strains was developed. The plaque assay exhibited sensitivity and reliability comparable to conventional quantitation methods such as TCID50 and indirect immunostaining assays, but is more convenient and economical. The direct plaque assay takes only 5 days to develop plaques whereas conventional quantitation methods usually take more than 10 days to produce results. Most importantly, the direct plaque assay does not inactivate the viruses, providing a valuable tool to isolate and purify aMPV and hMPV.

Previously, it has been reported that aMPV subtypes A, B and C were able to form visible plaques in agarose overlay plaque assays using a Japanese quail 74

fibrosarcoma (QT-35) cell line (Sabara and Larence, 2003). However, it took 13 days to develop visible plaques in QT-35 cells. Since Vero, Vero E6, and LLC-MK2 cells are typically used for propagation of aMPV and hMPV, it is beneficial to determine whether the metapneumoviruses are also able to form plaques in these cell lines. By optimizing the chemical components of the agarose overlay, it was found that aMPV and certain hMPV strains formed clear and countable plaques in these permissive cell lines after 5 days of incubation. The size of the plaques was dependent on cell confluency, and incubation temperature and time. Furthermore, the addition of actinomycin D in the agarose overlay significantly increased the size of the plaques.

This compound inhibits the proliferation of cells in nonspecific ways by forming a stable complex with double-stranded DNA or causing single-strand breaks in DNA, thus inhibiting DNA-primed RNA synthesis (Mizutani et al., 2000). It is likely that addition of actinomycin D to the overlay medium inhibits cell proliferation which in turn supports viral infection and cell-cell spread. More importantly, the number of plaques is not significantly altered by actinomycin D treatment (P>0.05) (Fig.6), demonstrating that actinomycin D did not affect the accuracy and sensitivity of direct plaque assay.

The main limitation of the direct plaque assay is that it can be only applied to hMPV strains with self-cleavable F protein. The paramyxovirus F protein is

synthesized as an inactive precursor (F0), which is cleaved at the F cleavage site by

cellular proteases to produce a fusion-competent, disulfide-linked F2-F1 complex

(Lamb 993). It is well known that F protein, especially the F protein cleavage site sequence, is the major determinant for trypsin dependence and viral virulence of paramyxoviruses, such as Newcastle Disease Virus (NDV) and Sendai virus (SeV)

(Collins et al., 1993; Lamb 1993). While the trypsin-dependent hMPV strains contain a

99RQSR102 motif at the putative cleavage site of F gene, it has been reported that recombinant hMPV with a 99RRRR102 motif at the putative cleavage site (rhMPV-F) exhibits trypsin-independent growth (Schickli, Kaur et al. 2005; Biacchesi, Pham et al.

2006). It was found that wild type hMPV with a 99RQSR102 cleavage site was not able to form visible plaques whereas recombinant hMPV with a 99RRRR102 cleavage site efficiently formed plaques. Despite this limitation, it should not minimize the significance of the development of the direct plaque assay for hMPV. One striking characteristic of hMPV pathogenesis is that the F cleavage site is not the major determinant of virulence (Schickli et al., 2005; Biacchesi et al., 2006). It has been shown that rhMPV-F improves significantly viral growth in cell culture but does not affect the viral replication, distribution, and pathogenesis in hamster and African green monkeys as compared to wild type hMPV (Schickli et al., 2005; Biacchesi et al., 2006).

Therefore, rhMPV-F could serve as an appropriate surrogate to study the biology of hMPV. The direct plaque assay will be useful for the study of laboratory adapted hMPV strains and live vaccine candidates. Additionally, it has been documented that a number of hMPV isolates mutated spontaneously to self-cleavage F gene after multiple passages in cell culture (Schickli et al., 2005). Specifically, it was found that the F cleavage site mutation S101P was associated with trypsin-independent viral growth in vitro (Schickli et al., 2005). Interestingly, this mutation has been documented in hMPV isolated from clinical samples. Yang et al., (2009) identified two hMPV subgroup A1 76

clinical isolates and one subgroup B1 clinical isolate containing naturally occurring

S101P mutations which have trypsin-independent F cleavage sites. Presumably, the direct plaque assay would be applicable to these clinical isolates. Finally, it should also be emphasized that self-cleavable F protein does not necessarily require all four basic amino acids (99RRRR102) in the F cleavage site. The F cleavage sites of aMPV subtypes A, B, C are 99RRRR102, 99RKKR102, and 99RKAR102 respectively, all of which possess self-cleavable characteristics (Alkhalaf et al., 2002; Govindarajan et al., 2004).

Also, hMPV F protein with 99RQPR102 (S101P mutation) is self-cleavable, demonstrating that self-cleavable hMPV F protein can occur without all four basic amino acids (Schickli et al., 2005). Therefore, the direct plaque assay may be applicable to hMPV strains containing any F cleavage sequence other than 99RQSR102, which has been shown to be non-permissive to plaque formation in this study. Metapneumovirus research has been limited by the inability to plaque purify the virus and the expense of immunology-based assays for quantitation, the direct plaque assay will be a valuable tool for the quantitation and evaluation of the biological properties of metapneumoviruses.

Chapter 3.

Development of a small animal model for human

metapneumovirus: cotton rat is more permissive than hamster

and mouse

3.1. Abstract

Human metapneumovirus (hMPV) is the second most prevalent causative agent of pediatric respiratory infections worldwide. Currently, there are no vaccines or antiviral drugs against this virus. One of the major hurdles in hMPV research is the difficulty to identify a robust small animal model that can accurately evaluate the efficacy and safety of vaccines and therapeutics. In this study, we compared the replication and pathogenesis of hMPV in BALB/c mice, Syrian golden hamsters, and cotton rats. It was found that BALB/c mice are not permissive for hMPV infection despite a high dose

(6.5 log10TCID50) of virus was used for intranasal inoculation. In hamsters, hMPV had an efficient replication in nasal turbinate but had a limited replication in lungs. In contrast, hMPV replicated efficiently in both nasal turbinate and lung when intranasally

administered with three doses (4, 5, and 6 log10 PFU) of hMPV. Lungs of cotton rats infected by hMPV developed histological changes including interstitial pneumonia,

mononuclear cells infiltrates, and increased lumen exudates. Immunohistochemistry examination found that viral antigens were expressed at the luminal surfaces of the bronchial epithelium cells in lungs. Collectively, these results demonstrated that cotton rat is a robust small animal model for hMPV infection.

3.2. Introduction

Human metapneumovirus (hMPV) is the second most commonly detected virus in children suffering from acute respiratory tract infection. Currently, there is no vaccine or antiviral drug to combat this virus. A major hurdle for hMPV vaccine development and antiviral discovery is the difficulty of identifying a suitable animal model that can faithfully reproduce human infectious diseases and accurately evaluate the efficacy and safety of vaccines and therapeutics. Since the first discovery of hMPV in 2001, several small animal models (such as mice, cotton rats, hamsters, guinea pigs, and ferrets) and non-human primate models (such as chimpanzees, rhesus macaques, and

African green monkeys) have been reported to study virus-host interaction, viral pathogenesis, and antiviral immunity. To date, all these animal models don’t mimic the signs of human disease except for chimpanzees. However, hMPV has a variable degree of replication in upper and lower respiratory tracts in these animal models. Because of the accessibility and availability of reagents and tools, most studies have been focused on mouse, hamster, and cotton rat models. In fact, many contradictious results have been reported using these animal models. Therefore, it is necessary to re-evaluate the permissiveness of animal models for hMPV infection. 79

Hamelin et. al, (2004) and Alverez et. al, (2004) reported that hMPV infection in

BALB/c led to an appearance of signs of illness, including weight loss, ruffled coat, huddling and respiratory clinical symptoms (such as heavy breathing). The peak titers in lung tissue occurred at days 7 post-infection (8 log10PFU/g) and lung viral titer at day

14 post-infection was 7 log10PFU/g of tissue. This unusual high viral replication has not been observed for any of paramyxoviruses (such as RSV). In fact, many researchers found that mice are not permissive for hMPV infection. Later, Syrian golden hamsters

(Mesocricetus auratus) were found to be more permissive for hMPV infection than mice. Although no clinical symptoms were observed, hMPV infection caused pulmonary histological changes. In 2004, cotton rats were first tested for replication of hMPV. Unfortunately, it was found that less than 2 log10PFU/g of virus detected in lung tissue when cotton rats was inoculated with 6 log10PFU of hMPV. Later, several other groups reported that hMPV replicates efficiently in cotton rats. These controversial results may be due to the differences in hMPV strains, methodology, and environmental factors in the animal experiments.

The major goal of this thesis is to evaluate the safety and efficacy of hMPV vaccine candidates. Thus, it is necessary to identify a robust animal model for hMPV. In this study, we directly compared the replication of rhMPV in the upper and lower respiratory tracts in three animal models, mice, hamsters and cotton rats. . Our results showed that (i) BALB/c mice were not permissive for hMPV infection; (ii) hamsters supported viral replication in upper respiratory tract but not in lower respiratory tract; 80

and (iii) cotton rats is a robust animal model for hMPV and efficient viral replication was observed in the upper and lower respiratory tracts .

3.3. Material and Methods

3.3.1. Cell lines

LLC-MK2 (ATCC No. CCL-7) cells were maintained in Opti-MEM (Life

Technologies, Bethesda, MD) supplemented with 2% fetal bovine serum (FBS). Vero

E6 cells (ATCC No. CRL-1586) was grown in Dulbecco's modified Eagle's medium

(DMEM; Life Technologies) supplemented with 10% FBS.

3.3.2. Purification of recombinant hMPV

Wild-type rhMPV carrying the trypsin-independent mutations in the F cleavage site was purified and used in animal studies. Briefly, LLC-MK2 cells were infected with rhMPV at MOI=0.01 and incubated for 6 days at 37oC and harvested by scraping.

The cell suspension was clarified by low-speed centrifugation at 1,500 rpm for 20 min at 4 oC in a Beckman Coulter Allegra 6R centrifuge. The supernatants were collected and the cell pellet was resuspended in 2 ml of DMEM and subjected to three freeze and thaw cycles. After centrifugation at 5,000 rpm for 10 min, supernatants were collected and combined. The virus was pelleted by ultracentrifugation at 28,000 rpm in a

Beckman Ty 50.2 rotor for 2 h. The virus pellet was resuspended in Opti-MEM and stored at -80 oC. Viral titer was determined by an immunostaining assay

3.3.3. Replication of hMPV in mice

Ten four-week-old specific-pathogen-free (SPF) female BALB/c mice were purchased from Charles River Laboratories(Malvern, PA). These animals were housed within ULAR facilities of The Ohio State University under approved Institutional

Laboratory Animal Care and Use Committee (IACUC) guidelines. The animals were randomly divided into 2 groups (5 per group). Animals in group 1 were mock infected with Opti-MEM, and animals in group 2 were inoculated intranasally with 6.5 log10TCID50 of rhMPV under isoflurane anesthesia. After inoculation, the animals were evaluated on a daily basis for weight loss and the presence of any respiratory symptoms of hMPV. At day 4 post-infection, animal were sacrificed and their nasal turbinates and lungs were removed for virus titration and pathological examination. (i) Virus titer in lung and nasal turbinate. Nasal turbinate and left lung from each animal were weighed and homologized in 1 ml of phosphate-buffered saline (PBS) using Dounce homogenizer. Viral titers were determined by immunostaining assay, argarose overlay plaque assay, and TCID50 assay. (ii)Total viral RNA was extracted and detected by

RT-PCR. (iii) Pulmonary histology. Left lung from each mouse was fixed with 10% neutral buffered formalin for histology as described below.

3.3.4. Replication of hMPV in hamsters

Twenty four-week-old SPF female Golden Syrian Hamsters were purchased from

Charles River Laboratories. Animals were randomly divided into four groups.

Hamsters in groups 1-3 were intranasally inoculated with 0.1 mL of Opti-MEM containing 4, 5, and 6.5 log10PFU of rhMPV, respectively. Hamsters in group 4 were mock infected with 100 µl of Opti-MEM. After inoculation, the animals were evaluated on a daily basis for possible weight loss and the presence of any respiratory symptoms of hMPV. At day 4 post-infection, animal were sacrificed and their nasal turbinates and lungs were removed for virus titration and pathological examination as described in

Section 3.3.6. 3.3.7.

3.3.5. Replication of hMPV in cotton rats

Table 3.1 Experiment design of hMPV replication study in cotton rats

Dose Harvest time (day)

(log10PFU) 3 4 5 4.0 √ 5.0 √ √ √ 6.0 √

Twenty five SPF cotton rats (kindly provided by Dr. Stefan Niewiesk, College of

Veterinary Medicine, The Ohio State University) were randomly divided into five groups (5 rats per group). These cotton rats were housed within ULAR facilities of The

Ohio State University under approved Institutional Laboratory Animal Care and Use

Committee (IACUC) guidelines. Each inoculation group was separately housed in rodent cages under BSL-2 conditions. Experimental design was summarized in Table

3.1. Rats were anaesthetized under isofluorane and either mock infected with 0.1 ml of

Opti-MEM as uninfected control or 0.1 ml of Opti-MEM containing different amounts of rhMPV. After inoculation, the animals were evaluated on a daily basis for possible weight loss and the presence of any respiratory symptoms. At designated days post-infection, cotton rats were sacrificed, and their lungs and nasal turbinate were collected for virus isolation and histological analysis.

3.3.6. Pulmonary histology

Right lung of each animal was removed, inflated and fixed by 4% formaldehyde

(pH 7.0). Fixed tissues were embedded in paraffin, and sectioned at 5 microns. Slides were then stained with hematoxylin and eosin (H&E). The pulmonary histological changes were examined by three independent pathologists. Histological changes were scored based on the extent of inflammation (focal or diffuse), the pattern of inflammation (peribronchilolar, perivascular, interstitial, and alveolar), and the nature of the cells making up the infiltrate (neutrophils, eosinophils, lymphocytes, and macrophages).

3.3.7. Immunohistochemical staining (IHC)

Right lung of animal were fixed in 10% formalin and embedded in paraffin.

5-micron sections were cut and placed onto positively charged slides. After deparafinization, sections were incubated with target retrieval solution (Dako,

Carpinteria, CA) for antigen retrieval. After blocking, a primary mouse anti-hMPV monoclonal antibody (Virostat, Portland, ME) was incubated for 30 min at room 84

temperature followed by incubation with a biotinylated horse anti-mouse secondary antibody (Vector Laboratories, Burlingame, CA). Slides were further incubated with

ABC Elite complex to probe biotin (Vector Laboratories) and slides were then developed using a 3,3'-diaminobenzidine (DAB) chromogen Kit (Dako) and hematoxylin used as a counterstain. Lung sections from hMPV infected and uninfected samples were used as positive and negative controls respectively.

3.3.8. Determination of viral titer in lung and nasal turbinate

Nasal turbinate and the left lung from each cotton rat were removed, weighed, and homogenized in 1 ml of phosphate-buffered saline (PBS) solution using Precellys 24 tissue homogenizer (Precellys, Bertin, MD) following manufacturer’s recommendations. The homogenized tissues were clarified by centrifugation at 6,000 rpm for 5 min at 4 oC. The presence of infectious virus was determined by immunostaining plaque assay in Vero E6 cells. Briefly, Vero E6 cells were seeded in

24-well-plates and then infected with 0.2 ml of 10 fold serial dilutions of supernatant from homogenized tissues. After 1 adsorption, 0.5 ml of fresh culture medium containing 2% FBS and 0.75% methylcellulose (Sigma, St. Louis, MO) was added to each well and incubated in humidified 5% CO2 incubator at 37◦C. At day 6 post-infection, the supernatant was removed and cells were fixed in a pre-chilled acetone: methanol solution (at the ratio of 3:2) at room temperature (RT) for 15 min.

Cells were permeablized in a phosphate saline buffer (PBS) containing 0.4% Triton

X-100 at RT for 10 min, and blocked at 37 ◦C for 1 hour using 1% bovine serum 85

albumin (BSA) in PBS. The cells were then labeled with an anti-hMPV N protein primary monoclonal antibody (Millipore, Billerica, MA) at dilution of 1:1000, followed by incubation with horseradish peroxidase (HRP)-labeled rabbit anti-mouse secondary antibody (Thermo Scientific, Waltham, MA) at dilution of 1:5,000. After incubation with AEC substrate chromogen (Sigma), positive cells were then visualized under the microscope. Viral titer was calculated as plaque forming unit (PFU) per gram tissue.

3.3.9. Agarose overlay plaque assay.

Vero or Vero-E6 cells were seeded in 6-well plates (Corning) at the density of

o Cells were then grown at 37 C and 5% CO2 to allow for plaque formation. Where indicated, the overlay medium was supplemented with actinomycin-D (Sigma) (0.1 to

3.3.10. Reverse transcription polymerase chain reaction (RT-PCR).

Lung tissues from cotton rats infected by rhMPVwere homogenized in 1 ml of

PBS as described above. Total RNA was extracted from 200 µl of lung homogenate using an RNeasy mini-kit (Qiagen, Valencia, CA) following the manufacturer's recommendation. RT-PCR was performed using a One Step RT-PCR kit (Qiagen) with two hMPV specific primers spanning the MTase region of the L gene,

hMPV-L-11759-Foward: 5’-TATATAGGGTTTAAGAATTGG-3’

hMPV-L-13199-Reverse: 5’-ATCATTTTTTACTTACAAGC-3’

The reaction mixture was prepared as described in Table 3.2. For the cycling parameters, a holding stage at 50°C was maintained for 30 min for reverse transcription.

Temperature was raised to 95 °C to activate polymerase prior to cycling, followed by 35 cycles of 94°C for 1 min for denaturation, 60 °C for 1 min for annealing, and 72 °C for

1 min for extension. The PCR products were gel purified and sequenced at The Ohio

State University Plant Microbe Genetics Facility to confirm the presence of the introduced mutations using a sequencing primer:

hMPV-L-12113-Forward: 5’-GCTAAAGGAAAGCTAAC-3’

Table 3.2 Reaction components for one-step RT-PCR

Component Volume RNase-free water 12.8 µL 5×Qiagen OneStep RT-PCR Buffer 5.0 µL dNTP Mix (10 mM) 1.0 µL hMPV-L-11759-Forward (25 µM) 0.6 µL hMPV-L-13199-Reverse (25 µM) 0.6 µL Qiagen OneStep RT-PCR Enzyme Mix 1.0 µL Template RNA 4.0 µL

Total Volume 25.0 µL

2.3.11. Statistical analysis

Statistical analysis was performed by one-way multiple comparisons using SPSS

8.0 statistical analysis software (SPSS Inc., Chicago, IL). A P value of <0.05 was considered statistically significant.

3.4. Results

3.4.1. Replication of rhMPV in mice

Previously, several researchers reported that mouse is a permissive model for hMPV. However, others reported that hMPV had a poor replication in mice. We first aimed to examine the infectivity of hMPV in mice. Briefly, mice were inoculated intranasally with 6.5 log10TCID50 of rhMPV strain NL/1/00, and weight loss and

presence of respiratory symptoms were monitored on daily basis No respiratory symptoms (such as cough, increased mucus production, rhinitis, heavy breathing, and other respiratory illness) were observed in the animals infected by rhMPV. No significant change in body weight in rhMPV infected mice compared to mock infected control groups (Figure 3.1). No mortality, ruffled fur, or behavior changes were observed.

Figure 3.1 Body weight changes of BLAB/c mice infected by rhMPV. Body weight of each mouse was recorded on a daily basis. An average body weight of five mice was shown.

At day4 post-infection, mice were sacrificed and nasal turbinate and lung tissues were homogenized for virus detection by immunostaning, plaque assay, and TCID50 assays. None of the animals had detectable viruses in nasal turbinate and lung tissues using these three detection methods. To determine the presence of viral RNA in lung, one-step RT-PCR targeting conserved region VI (CR VI) of hMPV L gene was also performed. Unfortunately, no viral RNA was detected by RT-PCR (Data not shown). 89

These observations showed that BALB/c mice are not permissive models for hMPV.

3.4.2. Replication of rhMPV in Syrian golden hamster

Syrian golden hamsters were randomly divided into four groups. Hamsters in groups 1-3 were inoculated intranasally with 4, 5, and 6.5 log10TCID50 of rhMPV, respectively. Hamsters in group 4 served as mock infected control. No body weight losses or clinical symptoms were observed after virus inoculation (Figure 3.2). At day 4 post-infection, hamsters were euthanized and virus titers in nasal turbinate and lung tissues were measured by an immunostaining assay. The average virus titer in nasal turbinate in groups 1-3 was 5.43±0.21, 5.61±0.21, and 4.29±0.48 log10TCID50/g, respectively. However, no infectious virus was detected in lung tissues of hamsters challenged with all three doses (Figure 3.3).

Figure 3.2 Body weight changes of hamsters infected by rhMPV. Body weight of each hamster was recorded on a daily basis. An average body weight of five mice was shown. 90

Figure 3.3 Virus titer in nasal turbinates and lungs of Syrian golden hamsters infected by rhMPV. In the experiment 1, five hamsters in each group were intranasally infected with 6.5 log10TCID50 of rhMPV in 0.1 ml Opti-MEM. In the second experiment, 4 and 5 log10TCID50 of rhMPV was used. Four days post-infection, nasal turbinate and lung tissues were harvested, viral titer was measured by TCID50 assay. Mean virus titer in nasal turbinate and lung tissues was shown.

Figure 3.4 Histological changes in lungs of hamsters infected by rhMPV. A: Lungs from mock infected hamsters. B-C: Lungs from hamsters infected by rhMPV-at dose of 6.5 log10 TCID50. Arrows in B indicate lymphoid infiltrates, and arrows in C show severe infiltrates with prominent acute inflammation involving bronchiole and adjacent parenchyma. 91

Subsequently, lung histology of hamsters infected by 6.5 log10TCID50 was examined. It was found that lungs from rhMPV-infected hamsters displayed mild histological changes including lymphoplasmacytic infiltrates around bronchovascular bundles with extension into interlobular septae and interstitium and venules as well as acute inflammation involving bronchioles and alveolar parenchyma (Figure 3.4).

Therefore, these results showed that hamsters are a permissive model for rhMPV.

However, rhMPV replicates efficiently in upper but not lower respiratory tracts under our experimental conditions.

3.4.3. Replication of rhMPV in cotton rats

Table 3.3 Replication of rhMPV in upper and lower respiratory tracts in cotton rats

Nasal Wash Lung

Inoculum Numbers of Day of % of infected Mean titer % of infected Mean titer

(log10PFU) animals termination animals (log10PFU/ml) animals (log10PFU/g) 5.0 4 3 0 ND 100 4.43±0.27 5.0 4 4 25 2 100 5.05±0.47 5.0 4 5 50 1.94 100 4.87±0.40 6.0 4 4 50 1.66 100 5.79±0.12 4.0 4 4 25 1.7 100 3.58±0.51

ND: not detected

We sought to determine if cotton rat model is permissive for rhMPV infection. An experiment was designed to determine the optimum infection dose and best timing to harvest nasal turbinate and lung tissues (Table 3.1). Twenty five animals were divided into five groups. The first three groups were intranasally challenged by 5 log10PFU of

rhMPV and terminated on 3, 4, and 5 days post infection. The other two groups were infected by 4 and 6 log10PFU of viruses respectively and terminated on 4 days post infection. No body weight losses or clinical symptoms were observed after virus inoculation. The average virus titers in lung tissues of animals infected by 4 and 6 log10PFU and harvested on day 3, 4, and 5 post infection were 4.43, 5.05, and 4.87 log10PFU/g, respectively (Table 3.3). The average virus titer in lung tissues from cotton rats infected by 4.0 and 6.0 log10PFU of viruses harvested 4 days post-infection were

3.58 and 5.79 log10PFU/g, respectively (Table 3.3). To test the viral replication in upper respiratory tract, we performed nasal wash to isolate rhMPV in the nasal turbinate. No virus was detected in the nasal wash samples in the 5 log10PFU group at day 3 post-infection. In other groups the mean titers in nasal wash are much lower than lung, ranging from 1.7 to 2 log10PFU/ml. This experiment showed that rhMPV replicated efficiently in cotton rat.

Histological examination found that lungs associated with rhMPV infection exhibited mild to moderate pathological changes including peribronchial and perivascular inflammation of mononuclears, airway remodeling, as well as interstitial pneumonia and increased bronchial exudate containing epithelial cells debris, neutrophils, macrophages (Figure 3.5).

Figure 3.5 Histological changes in lungs of cotton rats infected by rhMPV. A: Lungs from mock infected cotton rats. Lungs were harvested at day 4 after mock infection. B, C, and D: Lungs from cotton rats intranasally infected by 5 log10PFU of rhMPV and harvested at days 3, 4, and 5 post-infection, respectively. Arrow indicates histopathological changes including lymphoid infiltrates, bronchial luminal exudate, and airway remodeling. E-F: Lungs from cotton rats intranasally infected by

6 or 4 log10PFU of rhMPV, respectively, and harvested at day 4 postinfection. Arrows indicate histopathological changes including lymphoid infiltrates, bronchial luminal exudate, and airway remodeling.

To further confirm the replication of hMPV, lung tissue sections were subjected to immunohistochemistry analysis using monoclonal antibody against hMPV matrix protein. It was found that hMPV antigens were observed in bronchiole epithelium cells in a discontinuous pattern (Figure 3.6). In addition, luminal exudates including some visible macrophage cells were also positive for hMPV antigen.

Figure 3.6 Immunohistochemical staining of lung tissues from cotton rats infected by rhMPV. A-B: Lungs from mock infected cotton rats. Lungs were harvested at day 4 after mock infection. No hMPV antigen was detected. C-D: Lungs from cotton rats infected by 6 log10PFU of rhMPV and were sacrificed at day 4 post infection. Arrow indicates the hMPV antigen expression in the infected bronchial epithelial cells.

Collectively, these experiments demonstrated that (i) rhMPV replicates efficiently in lungs of cotton rats; (ii) rhMPV causes histological changes in lungs; and (iii) a large number of rhMPV antigens were expressed in bronchiole epithelium cells. Therefore, cotton rat is a robust animal model for rhMPV infection.

3.5. Discussion

Our ultimate goal is to evaluate the safety and efficacy of hMPV vaccine candidates. Thus, it is critical to identify a robust small animal model for hMPV. We compared the replication and pathogenesis of rhMPV strain NL/1/00 in three rodent models, BALB/c mice (Mus musculus), Syrian golden hamsters (Mesocricetus auratus), and cotton rats (Sigmodon hispidus). We found that the cotton rat is an ideal model for hMPV whereas the hamster is less permissive and the mouse model is not permissive.

Currently, many controversial results have been observed using mouse as an animal model for hMPV infection. Alvarez, et. al., (2004) reported that hMPV causes clinical signs and breathing difficulties in mice. Hamelin et.al, (2004) reported that hMPV infection causes modest mononuclear cell infiltration in interstitium, airway remodeling, increased mucus production and bronchial infiltration in lungs. However, many other studies found that hMPV did not cause any clinical symptoms or viral replication in mice. Williams et al., (2005) compared the ability of hMPV replication in different strains of mouse (Mus musculus) 129, AKR, BALB/c, C3H, C57BL/10, CBA,

DBA/1, DBA/2, and SJL, all of which showed extremely low levels of viral replication.

In our study, we are not able to detect any infectious virus or viral RNA from nasal turbinate and lung tissue of any animal. Our results support that BALB/c mice are not permissive for hMPV infection.

MacPhail et al., (2004) first reported that hamsters are permissive for hMPV

infection. In their study, Syrian golden hamsters were administered intranasally in a 0.1 ml volume containing 6 log10PFU of hMPV. They found that 5.3 and 4.3 log10PFU /g tissue of virus was detected in nasal turbinate and lung, respectively, at day 4 post-inoculation. Williams et al., (2005) also found that Syrian golden hamsters are permissive for hMPV replication in upper and lower respiratory tracts, but the efficiency of viral replication is lower than that in cotton rats. In our study, no infectious virus was detected in lung when lower infection doses (4 and 5 log10TCID50) of virus were used for inoculation. When hamsters were infected intranasally with 6.5 log10TCID50 of rhMPV NL/1/00, we only found that 4.29 log10TCID50/g of virus was detected in nasal turbinate but not in the lung. Despite the low level of hMPV detected in lungs, hMPV infection caused significant histological changes in lungs. In our study, the replication efficiency in lungs of hamsters was much lower than those reported by

MacPhail et al., (2004) and Williams et al., (2005). Perhaps, this difference may result from the different virus strains used for infection and/or environmental factors (such as humidity and temperature) in animal facilities. Therefore, hamsters may not be robust enough to support hMPV replication under our experimental conditions.

The cotton rat S. hispidus is a small rodent model susceptible to a large variety of human pathogens. To date, this model was found to be the best model for respiratory viruses. Previously, it was found that permissiveness of cotton rats to infection with human respiratory syncytial virus (hRSV) was over 100-fold higher than that of mice.

In fact, cotton rats has not only been used as an animal model to study the pathogenesis 97

and efficacy of hRSV vaccine, but also used for evaluating the efficacy and safety of prophylactic treatment for severe hRSV disease (antibodies RespiGam and Synagis), bypassing the need for testing in primates. The cotton rat model also accurately predicted the dose of the drug currently being used in human infants. Recent years, many tools and reagents (such as cytokines, chemokines, cell surface markers and regulatory molecules) have been developed for cotton rats that accelerated the use of this model for human pathogens.

The first hMPV replication in cotton rats was reported by MacPhail et al., (2004).

Five cotton rats were intranasally inoculated with 6 log10PFU of hMPV. Unfortunately, less than 1.7 and 1.8 log10PFU/g tissue of virus was detected in nasal turbinate and lung in cotton rats, respectively. In contrast, Williams et al., (2005) determined the kinetics of hMPV replication in cotton rats and found that hMPV replicates efficiently in cotton rats. In their study, cotton rats were inoculated with 5 log10PFU of hMPV and were sacrificed at 2, 4, 6, 8, 10, or 14 days postinfection. It was found that the replication of hMPV in the lung tissues peaked on day 4 postinfection at a mean titer of 5.26 log10PFU/g and declined gradually. Virus was not detected in the lung after day 6.

Similarly, Wyde et al., (2005) found cotton rats are permissive for hMPV replication in nose and lungs, with a peak titer of 3.6 log10PFU/g in the nose and 4.4 log10PFU/g in the lung on day 4 post-infection. In our study, we compared hMPV replication in cotton rats using three different inoculation dosages at different time points. We found that hMPV replicates efficiently in lungs of cotton rats, but it was dose-dependent. At 98

inoculation dose of 4 log10PFU per rat, 3.58 log10PFU/g of virus was found in lungs.

When inoculation dose increased to 5 or 6 log10PFU per rat, virus titer in lungs increased to 5.05 and 5.79 log10PFU/g, respectively (P<0.05). Day 4 post-infection appears to be the best time to terminate the study as the viral titer in lungs at this time point was higher than that in days 3 and 5 post-infection although they were not significantly different (P>0.05). Furthermore, we demonstrated that rhMPV caused significant histological changes in lungs although no respiratory symptoms were observed. Our results were consistent with those from Williams et al., (2005) and Wyde et al., (2005). Also, it should be pointed out that we recovered less infectious viruses using nasal washing. However, we found that virus titer significantly increased when we homogenized the nasal turbinate (see Chapter 5 for detail).

Given the fact that a robust replication of other paramyxoviruses (such as RSV,

PIV3, and measles virus) in cotton rats was also observed, we concluded that cotton rat is an ideal model to study hMPV and other paramyxoviruses.

Chapter 4.

Recovery and characterization of methyltransferase-defective

recombinant human metapneumovirus

4.1. Abstract

The 5’ end of the messenger RNA (mRNA) of paramyxoviruses possesses a unique methylated cap structure that is essential for viral gene expression and, subsequently, viral replication. Available evidence suggests that mechanism of mRNA cap addition and methylation in paramyxoviruses is distinct from their hosts. This difference can be potentially used as a target for the development of antiviral drugs and vaccines. In this study, we performed a mutagenesis analysis in the putative mRNA cap methyltransferase (MTase) catalytic site and S-adenosyl methionine (SAM) binding site located in the conserved region VI (CR-VI) of large (L) polymerase protein.

Alanine substitutions to the amino acid residues involved in MTase catalysis and SAM binding diminished gene expression in a minigenome replication assay. Using a reverse genetic system, we recovered three recombinant hMPVs carrying mutations in the SAM binding site. Trans-methylation assay showed that these rhMPV mutants were specifically defective in ribose 2’-O, but not guanine N-7 (G-N-7) methylation. These 100

MTase-defective rhMPVs showed delayed growth kinetics, reduced viral genome replication and mRNA synthesis, and formed smaller plaques in cell culture compared to wildtype rhMPV. Collectively, our results indicate that (i) amino acid residues in the

SAM binding site in hMPV L protein are essential for 2’-O methylation, and (ii) inhibition of 2’-O methylation can serve as a new approach to rationally attenuate hMPV in cell culture.

4.2. Introduction

In eukaryotic cells, mRNA cap formation is an early posttranscriptional event that is essential for subsequent processing, nuclear export, stability, and translation of mRNA (Both, Furuichi et al. 1976; Furuichi, LaFiandra et al. 1977). Cap formation is mediated by a series of enzymatic reactions. First, the 5’ triphosphate end of the nascent mRNA chain (5’pppN-RNA) is hydrolyzed by an RNA triphosphatase (RTPase) to yield the diphosphate 5’ ppN-RNA. Second, an RNA guanylyltransferase (GTase) reacts with GTP to form a covalent enzyme-GMP intermediate and transfers GMP to

5’ppN-RNA via a 5′-5′ triphosphate linkage to yield 5’ GpppN-RNA. Third, the capping guanylate is methylated by a guanine N-7 (G-N-7) methyltransferase (MTase) to yield 7mGpppN-RNA (cap 0). Finally, the G-N-7 methylated cap structure can then be further methylated by a ribose-2’-O (2’-O) MTase to yield 7mGpppNm-RNA (cap 1)

(Furuichi and Shatkin 2000). In this conventional methylation reaction, G-N-7 methylation occurs prior to 2’-O methylation, and the two methylation events are carried out by two separate enzymes, each containing a unique binding site for the 101

methyl donor, S-adenosyl methionine (SAM). This capping and methylation mechanism is conserved in all cell types (Hakansson and Wigley 1998; Lima, Wang et al. 1999; Fabrega, Hausmann et al. 2004).

Figure 4.1 Conserved regions (CR) in the L proteins of NNS RNA viruses. Amino acid sequence alignment identified six CRs numbered I to VI (CRs I–VI) in L proteins. The function of CR III, V, and VI has been mapped in VSV L protein.

Cap formation in non-segmented negative-sense (NNS) RNA viruses has evolved independently of the host mechanism of mRNA cap formation. In early 1980, it was found that, for vesicular stomatitis virus (VSV) (Moyer and Banerjee 1975; Hercyk,

Horikami et al. 1988), human respiratory syncytial virus (hRSV) (Barik 1993), and spring viremia of carp virus (Gupta and Roy 1980), the two italicized phosphates of the

5′Gppp5′NpNpN triphosphate bridge are derived from a GDP donor, rather than GMP.

In recent years, our understanding of the mechanism of mRNA cap formation is largely shaped by the studies from VSV, a prototypical NNS RNA virus. It was found that capping of VSV mRNA is achieved by a novel polyribonucelotidyltransferase

(PRNTase) which transfers a monophosphate RNA onto a GDP acceptor through a covalent L-RNA intermediate. First, a GTPase associated with the VSV L protein removes the γ-phosphate group of GTP to generate GDP, an RNA acceptor. Second, a

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PRNTase activity of the L protein specifically transfers a 5′-monophosphorylated (p-)

RNA moiety to the GDP acceptor to yield a GpppA mRNA. In addition, methylation of the VSV mRNA cap structure is also unique in that mRNA cap is methylated by a dual

MTase within CR VI of L protein, and the two methylases share a single SAM binding site. First, the mRNA cap is methylated by a ribose 2’-O MTase to yield GpppAm-RNA.

The mRNA is further methylated by a G-N-7 MTase to yield 7mGpppAm-RNA

(Grdzelishvili, Smallwood et al. 2005; Li, Fontaine-Rodriguez et al. 2005). In this type of methylation, ribose 2′-O methylation precedes and facilitates subsequent G-N-7 methylation which is in contrast to all other known mRNA cap methylation reactions

The large (L) polymerase protein of NNS RNA viruses contains enzymatic activities for nucleotide polymerization, mRNA cap addition, cap methylation, and polyadenylation. Although no structural data for L is available, amino acid sequence alignment between the L proteins of representative members of each family within

NNS RNA viruses has identified six conserved regions numbered I to VI (CRs I–VI)

(Figure 4.1). Thus, it is generally thought that the enzymatic activities of L protein are located in these conserved regions. Previous studies using VSV have mapped the L enzymatic functions to the CRs. For example, the nucleotide polymerization activity has been identified in CR III and this region is also required for polyadenylation (Sleat and Banerjee 1993). The mRNA capping enzyme has been mapped to CR V whereas the mRNA cap MTase is located in CR VI.

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A detailed characterization of paramyxovirus capping and methylase activities and the mechanism involved in these reactions have not been described. This is due in large part to the fact that paramyxoviruses lack an in vitro mRNA reconstitution assay and it is technically challenging to purify a functional L protein in vitro. However, sequence alignment showed that the capping motif (GxxT[n]HR) and methylation signature motif are conserved in CR V and CR VI of L proteins of all paramyxoviruses, suggesting the general mechanism of capping and methylation found in VSV may be conserved in the paramyxoviruses. To date, it has been shown that the mRNAs of many paramyxoviruses (such as measles, Sendai virus, and RSV) are methylated at positions

G-N-7 and 2’-O. Interestingly, the mRNA of Newcastle disease virus (NDV), an avian paramyxovirus, lacks the 2’-O methylation (Colonno and Stone 1976). The capping and methylation status is not known for any member of the genus Metapneumovirus.

It appears that the mRNA capping and methylation machinery in NNS RNA viruses is distinct from that of their hosts. This difference, coupled with the fact that replication of NNS RNA viruses occurs in the cytoplasm, makes mRNA cap formation an excellent target for anti-viral drug discovery and a new target for generating attenuated vaccine strains. Inhibition of mRNA capping would likely be lethal to the virus since the mRNA cap is required for viral protein translation. However, inhibition of the viral mRNA cap methylation would likely reduce the binding affinity to eIF4E, the cellular cap-binding protein, and diminish the rate of viral protein translation, which in turn downregulates viral replication, viral spread, and ultimately result in viral 104

attenuation. Thus, we hypothesize that MTase-defective human metapneumovirus

(hMPV) can be recovered and will be highly attenuated in cell culture, and will be good live vaccine candidates for hMPV. We expect that ablating viral mRNA cap methylation would provide a new strategy to rationally attenuate hMPV for development of live attenuated vaccines and their exploitation as viral vectors for vaccines, oncolytic therapy, and gene delivery.

4.3. Materials and Methods

4.3.1. Cells and viruses

LLC-MK2 (ATCC No. CCL-7) cells were maintained in Opti-MEM (Life

Technologies, Bethesda, MD) supplemented with 2% fetal bovine serum (FBS). Vero

E6 cells (ATCC No. CRL-1586) and BHK-SR19-T7 cells (kindly provided by Apath,

L.L.C, Brooklyn, NY) were grown in Dulbecco's modified Eagle's medium (DMEM;

Life Technologies) supplemented with 10% FBS. The medium of the BHK-SR19-T7 cells was supplemented with 10 µg/ml puromycin (Life Technologies) during every other passage to select for T7 polymerase-expressing cells.

4.3.2. Plasmids

Plasmids encoding the hMPV minigenome, the full-length genomic cDNA of hMPV, and support plasmids expressing hMPV N (pCITE-N), P (pCITE-P), L

(pCITE-L), and M2-1 (pCITE-M2-1) were kindly provided by Dr. Ron A. M. Fouchier at Department of Virology, Erasmus Medical Center, Rotterdam, The Netherlands. The 105

methods for construction of these plasmids are described elsewhere. Briefly, (a) The minigenome plasmid was constructed using pSP72 as the backbone. A fragment containing the T7 terminator, HDV-ribozyme, hMPV leader, gene start, green fluorescent protein (GFP), gene end, hMPV trailer, and the T7 promoter was inserted between the NdeI and HpaI restriction enzyme sites in pSP72 (Figure 4.2). (b) Support plasmids. The N, P, and M2-1 genes were cloned into the pCITE vector between the

NcoI and XhoI restriction enzyme sites to yield pCITE-N, pCITE-P, and pCITE-M2-1, respectively. L gene was inserted into the pCITE vector between the NcoI and SmaI restriction enzyme sites. Gene expression of all cassettes was under the control of the

T7 promoter in the pCITE vector. (c) Full length genome plasmid. The full length hMPV NL/1/00 plasmid was constructed using pSP72 as the backbone. The cleavage site in the genome of hMPV NL/1/00 was modified to a trypsin-independent F cleavage site as described in Chapter 2. Full-length hMPV NL/1/00 was flanked by the

T7 promoter and hMPV leader at 5’ end, and flanked by HDV ribozyme, hMPV trailer and T7 terminator at the 3’ end and the cassette was inserted between NdeI and HpaI sites.

Figure 4.2 Schematic representation of hMPV minigenome construct. T-T7: T7 terminator, δ: HDV ribozyme, Le: hMPV leader, GS: hMPV gene start, GE: hMPV gene end, Tr: hMPV trailer, P-T7: T7 promoter.

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4.3.3. Site-directed mutagenesis

The L CR VI mutations were introduced into the full-length genome of wild type hMPV NL/1/100 carrying the F cleavage site mutations. A QuikChange Site-Directed

Mutagenesis Kit (Strategene, La Jolla, CA) was utilized following manufacturer’s recommendations. Mutagenesis primers are listed in Table 1 and positions of mutagenesis are underlined. Mutations were confirmed by DNA sequencing using the primers described below.

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4.3.4. Minigenome assay

Figure 4.3 Schematic diagram of minigenome assay.

The minigenome assay was performed in BHK-SR19-T7 cells which stably express the T7 RNA polymerase. Briefly, confluent BHK-SR19-T7 cells were transfected with 1.0 μg of the minigenome plasmid together with 0.8 μg of pCITE-N,

0.4 μg of pCITE-P, 0.4 μg of pCITE-M2.1, and 0.4 μg of pCITE-L or pCITE-L mutations using Lipofectamine 2000 (Invitrogen) (Figure 4.4). Transfections were performed overnight following the manufacturer’s recommendations. At 24 h post-transfection, the medium was replaced with DMEM containing 5% FBS. At day 3

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post-transfection, the expression of GFP in the transfected cells was visualized by a fluorescent microscope.

4.3.5. Recovery of recombinant hMPVs from the full-length cDNA clones

Recombinant hMPVs (rhMPVs) were rescued using a reverse genetics system as described previously. Briefly, BHK-SR19-T7 cells (kindly provided by Apath LLC) were transfected with 5.0 μg of phMPV full length genomic plasmid, 2.0 μg of pCITE-N, 2.0 μg of pCITE-P, 1.0 μg of pCITE-L, and 1.0 μg of pCITE-M2-1 using

Lipofectamine 2000 (Invitrogen). At day 6 post-transfection, the cells were harvested using scrapers and were co-cultured with 50-60% confluent LLC-MK2 cells. When an extensive cytopathic effects (CPE) was observed, the cells were subjected to three freeze-thaw cycles followed by centrifugation at 3,000×g for 10 min. The supernatant was used subsequently to infect new LLC-MK2 cells. The successful recovery of the rhMPVs was confirmed by immuno-staining, agarose overlay plaque assay, and

RT-PCR.

4.3.6. Reverse transcription polymerase chain reaction (RT-PCR) and

sequencing

All plasmids and viral stocks were sequenced. Viral RNA was extracted from 200

µL of each recombinant virus using an RNeasy mini-kit (Qiagen, Valencia, CA) following the manufacturer's recommendation. A 1.5 kb DNA fragment spanning CR

VI of hMPV L gene was amplified by RT-PCR using primers designed to anneal to

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nucleotide positions 11759 and 13199 (numbers are based on the genome sequence of hMPV strain NL/1/00), respectively.

hMPV-L-11759-Foward: 5’-TATATAGGGTTTAAGAATTGG-3’

hMPV-L-13199-Reverse: 5’-ATCATTTTTTACTTACAAGC-3’

The PCR products were purified and sequenced at The Ohio State University Plant

Microbe Genetics Facility to confirm the presence of the designed mutations using a sequencing primer: hMPV-L-12113-Forward: 5’-GCTAAAGGAAAGCTAAC-3’

4.3.7. Immuno-staining plaque assay

Vero E6 cells were seeded in 24 well plates and infected with serial dilutions of rhMPV. At day 6 post-infection, the supernatant was removed and cells were fixed in a pre-chilled acetone: methanol solution (at the ratio of 3:2) at room temperature (RT) for

15 min. Cells were permeablized in a phosphate saline buffer (PBS) containing 0.4%

Triton X-100 at RT for 10 min, and blocked at 37 ◦C for 1 h using 1% bovine serum albumin (BSA) in PBS. The cells were then labeled with an anti-hMPV N protein primary monoclonal antibody (Millipore, Billerica, MA) at a dilution of 1:1000, followed by incubation with horseradish peroxidase (HRP)-labeled rabbit anti-mouse secondary antibody (Thermo Scientific, Waltham, MA) at a dilution of 1:5,000. After incubation with 3-Amino-9-ethylcarbazole (AEC) substrate chromogen (Sigma, St.

Louis, MO), positive cells were then visualized under the microscope. Viral titer was calculated as plaque forming unit (PFU) per mL. 110

4.3.8. Viral replication kinetics in LLC-MK2 cells

Confluent LLC-MK2 cells in 35-mm dishes were infected by rhMPV or mutant rhMPV at a multiplicity of infection (MOI) of 0.1. After 1 h of adsorption, the inoculum was removed, and cells were washed three times with PBS. Fresh DMEM

(supplemented with 2% FBS) was added, and the infected cells were incubated at 37 oC.

At different time points post infection, the supernatant and cells were harvested by three freeze–thaw cycles followed by centrifugation at 1500×g, RT for 15 min. Virus titer was determined by an immunostaining assay in LLC-MK2 cells.

4.3.9. Analysis of hMPV N protein expression by Western blot

Confluent Vero E6 cells were infected with each rhMPV mutant at an MOI of 1.0.

At day 1, 3, 5, and 7 post-infection, the cell culture supernatant was removed and the cells were lysed in 200 µl of RIPA lysis buffer (25mM Tris-HCl (pH 7.6), 150mM NaCl,

1% NP-40, 1% sodium deoxycholate, 0.1% SDS). 20 µl of the cell lysate was denatured at 99°C for 5 min and analyzed on a 12% polyacrylamide Bis-Tris gel. The separated protein was transferred to a Hybond-P polyvinylidene difluoride membrane

(Amersham Biosciences, Pittsburgh, PA) using Trans-Blot SD Semi-Dry Transfer Cell

(Bio-Rad). Membranes were blocked with 5% skim milk in PBS, and subsequently probed with anti-hMPV N monoclonal antibody (Milipore) diluted 1:200 in PBS-milk, followed by incubation with horseradish peroxidase-conjugated anti-mouse IgG monoclonal antibody diluted to 1:2,000 in PBS-milk. Membranes were developed with

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a chemiluminescence substrate (Amersham Biosciences) and exposed to Biomax MR film (Kodak) for visualization of hMPV N protein.

4.3.10. Qiantification of viral genomeic RNA replication and mRNA synthesis

by real-time RT PCR

Confluent LLC-MK2 cells were infected with each rhMPV mutant at an MOI of

1.0. At day 1, 3, 5, and 7 post-infection, total RNA was isolated from virus-infected cells using Trizol reagent (Invitrogen). Viral genomic RNA copies were quantified by real-time RT-PCR using two primers specifically targeting hMPV leader sequence and

N gene. Poly (A)-containing viral mRNA was isolated from total RNA using

Dynabeads mRNA isolation Kit (Invitrogen) according to the manufacturer’s recommendations. Using the viral mRNA as the template, the N mRNA copies were quantified by real-time RT-PCR using two primers targeting viral N gene.

4.3.11. Genetics stability of rhMPV mutants in cell culture

Confluent LLC-MK2 cells in 150 mm dishes were infected by each mutant rhMPV at MOI of 0.1. At day 3 post-infection, the cell culture supernatant was harvested and used for the next passage in LLC-MK2 cells. Using this method, each rhMPV mutant was repeatedly passed 10 times in LLC-MK2 cells. At each passage, the

CR VI of L gene was amplified by RT-PCR and sequenced.

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4.3.12. In vitro trans methylation assay

Confluent Vero E6 cells in 150 mm dishes were mock infected or infected by wild-type or mutant rhMPVs at MOI of 0.1. At day 3 post-infection, total RNA was isolated from virus-infected cells using Trizol reagent (Invitrogen) and dissolved in 10 mM Tris-HCl buffer (pH 7.5). Subsequently, poly (A)-containing RNA was isolated from total RNA using Dynabeads mRNA isolation Kit (Invitrogen) according to the manufacturer’s recommendations. To determine whether rhMPV mutants were defective in G-N-7 methylation, 500 ng of mRNA were incubated with 10 units of vaccinia virus G-N-7 MTase supplied by m7G Capping System (Cellscript, Madison,

WI) in the presence of 15 μCi [3H]-SAM (85 Ci/mmol, PerkinElmer) for 4 h. After the methylation reaction, RNA was purified using the RNeasy Mini Kit (Qiagen), and the methylation of the mRNA cap structure was measured by 3H incorporation using a

1414 series scintillation counter (PerkinElmer). Similarly, trans ribose 2’-O methylation assay was performed by incubating 500 ng of mRNA with 10 units of vaccinia virus 2’-O MTase supplied by vaccinia 2'-O-Methyltransferase Kit (Cellscript,

Madison, WI) in the presence of 15 μCi [3H]-SAM (85 Ci/mmol, PerkinElmer). After the methylation reaction, RNA was purified, and the level of 2’-O methylation was measured by 3H incorporation using a scintillation counter.

4.3.13. Statistical analysis.

Statistical analysis was performed by one-way multiple comparisons using SPSS

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8.0 statistical analysis software (SPSS Inc., Chicago, IL). A P value of <0.05 was considered statistically significant.

4.4. Results

4.4.1. Sequence analysis in the putative MTase domain in hMPV L protein

To begin to utilize MTase as the target to attenuate hMPV, we first performed a detailed bioinformatics analysis of the CR-VI within the hMPV L protein. Our aim was to predict critical amino acids that may be essential for mRNA cap methylation by comparing the amino acid sequences and structural homology with available crystal structures of MTases such as vaccinia virus VP39 and E.coli RRMJ. The S-adenosyl methionine (SAM)-dependent MTase superfamily contains six motifs involved in either SAM binding (motifs I, III, IV) or in the catalytic reaction (motifs IV, VI, VIII,

X). Sequence alignment and structural modeling identified these motifs in the CR VI of paramyxovirus L proteins. Typically, an mRNA cap MTase contains a K-D-K-E tetrad that functions as the catalytic residues for methylation (Figure 4.4). As predicted, this

K-D-K-E tetrad is highly conserved in CR-VI of the L proteins of all known NNS RNA virus with the exception of Borna disease virus, which replicates in the nucleus. Amino acid sequence alignments suggest that residues K1673, D1779, K1817, and E1848 of the hMPV L protein correspond to the catalytic K-D-K-E tetrad. In methylation reactions, a G-rich motif and an acidic residue (D/E) are involved in binding the methyl donor SAM. Indeed, this GxGxG…..D/E motif is conserved in all paramyxoviruses and

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other NNS RNA viruses. Sequence alignments suggested that the SAM binding site residues of hMPV L protein include G1696, G1698, G1700, and D1755 (Figure 4.4).

Figure 4.4 Amino acid sequence alignment of conserved region VI (CR VI) of NNS RNA viruses L proteins and modeling with two known 2’-O MTase structures, VP39 and RRMJ. STR: structure of RRMJ and VP39. Predicted or known alpha-helical regions are shown as cylinders and the ß-sheet region. HMPV, human metapneumovirus; AMPVC, avian metapneumovirus subtype C; HRSV, human respiratory syncytial virus; BRSV, bovine respiratory syncytial virus; PVM, pneumonia virus of mice; PIV3, parainfluenza virus type 3; NDV, Newcastle disease virus; EBOM, Ebola virus, VSIV, vesicular stomatitis virus Indiana strain; RRMJ, E. coli, heat shock methyltransferase; VP39, vaccinia virus methyltransferase VP 39.

4.4.2. Examination of the function of L mutants in MTase region using a

minigenome assay.

Next, we performed an alanine scanning mutagenesis in the MTase region in hMPV L protein. Amino acids in the predicted KDKE motif and the SAM binding site were individually mutated to alanine in the pCITE-L plasmid using a site-directed

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mutagenesis. These L gene mutations were designated pCITE-L-K1673A, D1779A,

K1817A, E1848A, G1696A, G1698A, G1700A, and D1755A. The presence of these mutations in the L gene was confirmed by sequencing. A minigenome assay was then performed to examine the function of the L protein. In the hMPV minigenome

(pSP72-hMPV-GFP), the genome of hMPV was replaced by a negative sense open reading frame (ORF) of green fluorescent protein (GFP) gene which was flanked by hMPV leader (Le) and trailer (Tr) sequences (Figure 4.3). Thus, the expression of GFP is solely dependent on the ability of the RNA dependent RNA polymerase (RdRp) to replicate the minigenome. To do this, BHK-SR19T-7pac cells, which stably express T7

RNA polymerase, were transfected with the hMPV minigenome (pSP72-hMPV-GFP) together with support plasmids pCITE-N, pCITE-P, pCITE-M2-1 and pCITE-L or L mutants. At 48 h post infection, the cells expressing GFP were observed by fluorescence microscopy. As shown in Figure 4.5, a strong GFP expression was observed when wild type pCITE-L was used for transfection. No GFP signal was detected when pCITE-L was omitted from the transfection. Co-transfection of pCITE-L carrying mutations in the predicted MTase catalytic and SAM binding sites significantly decreased GFP expression. This result suggests that mutations to the predicted MTase catalytic and SAM binding sites confer a defect in replication, gene expression, or both.

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Figure 4.5 Mutations to the CR VI of hMPV L gene diminished GFP expression in a minigenome assay. Minigenome assay was performed as described above. The GFP expression was visualized by a fluorescent microscope at 48 h post-infection.

4.4.3. Recovery of recombinant hMPVs carrying mutations in the MTase

catalytic and SAM binding sites.

Since L mutants were functional in the minigenome replication assay, we then generated recombinant hMPVs carrying mutations in the MTase catalytic and SAM binding sites. Using QuikChange mutagenesis methodology, the amino acids in the predicted KDKE tetrad and GxGxG…..D/E motif were individually mutated to alanine in the full-length genomic cDNA of hMPV lineage A strain NL/1/00 (phMPV). The presence of the desired mutation was verified by sequencing. These plasmids were designated phMPV-K1673A, D1779A, K1817A, E1848A, G1696A, G1698A, G1700A, 117

and D1755A. Using a reverse genetics system, a panel of recombinant hMPVs (rhMPV) carrying mutations in the KDKE tetrad and GxGxG…..D/E motif were generated.

Briefly, rhMPV was recovered by co-transfection of phMPV (or phMPV-mutations) and support plasmids encoding viral N (pCITE-N), P (pCITE-P), L (pCITE-L), and

M2-1 (pCITE-M2-1) proteins into BHK-SR19-T7 cells. Six days post-transfection, the cells were harvested and co-cultured with 50-60% confluent LLC-MK2 cells. When extensive cytopathic effect (CPE) was observed, cell supernatant was harvested and the recovered viruses were further amplified in LLC-MK2 cells (Figure 4.6).

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Figure 4.6 Recovery of rhMPV from an infectious cDNA clone by reverse genetics. Plasmid encoding the full-length hMPV genomic cDNA and four support plasmids encoding N, P, M2-1, and L genes were co-transfected into BHK-SR19-T7 cells. At day 4 post transfection, cells were harvested and co-cultured with LLC-MK2 cells for another 7 days. The recovered rhMPV was further amplified in LLC-MK2 cells. The successful recovery of the virus was confirmed by plaque assay and RT-PCR.

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Figure 4.7 rhMPVs carrying mutations in the SAM binding site formed smaller plaques in Vero E6 cells. Top panel: immuno-spots formed by rhMPV mutants. Plaques in immunostaining assay were developed at day 4 post infection. Lower panel: viral plaques formed by rhMPV mutants. Plaques in agarose overaly plaque assay were developed at day 6 post infection.

The successful recovery of recombinant viruses was initially detected by an immuno-staining assay using a monoclonal antibody against N protein. As shown in

Figure 4.7, all rhMPV mutants were positive for viral N protein expression in the immuno-staining assay. However, the immune-spots formed by these rhMPV mutants were much smaller than wild type rhMPV, suggesting that these rhMPV mutants had a defect in cell-to-cell spread and/or viral gene expression. Therefore, these recombinant viruses were continuously passed in LLC-MK2 cells, and the viral titer was determined by an immune-staining assay. At passage 4, we found that rhMPVs carrying mutations in KDKE tetrad had titers of 3-4 log10PFU/ml. In contrast, titers of rhMPV carrying mutations in SAM binding site (rhMPV-G1696A, G1700A, and D1755A) reached

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approximately 5-6 log10PFU/ml. Thus, we decided to focus on rhMPV carrying mutations in the SAM binding site.

To further confirm the phenotype of these recombinant hMPVs, a direct agarose overlay plaque assay was performed. As shown in Figure 4.7, the plaque sizes of rhMPV-G1696A, G1700A and D1755A were 0.6±0.1 mm, 0.5±0.1 mm and 0.7±0.2 mm respectively, which were significantly smaller than that of wildtype rhMPV

(1.2±0.1 mm). This suggests that rhMPV carrying mutations in the SAM binding site had defects in cell-cell spread and/or viral replication.

Figure 4.8 RT-PCR and sequencing analysis of rhMPVs carrying mutations in the SAM binding site. A. RT-PCR of CR VI of L gene: a 1.44 kb fragment targeting CR VI of hMPV L gene was amplified from each recovered rhMPV mutant. B. Sequencing of RT-PCR products: RT-PCR products were gel purified and sequenced. All rhMPV mutants contained the desired mutations in the CR VI of the L gene.

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To confirm the mutant rhMPVs contained the desired mutations, viral RNA was extracted followed by RT-PCR using primers specific to CR VI of hMPV L gene

(Figure 4.8 A). A 1.4 kb DNA fragment spanning CR VI of L gene was amplified from each rhMPV mutant. The PCR products were gel purified and sequenced. It was found that all rhMPV mutants contained the desired mutation in CR VI of L gene (Figure 4.8

B).

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4.4.4. Recombinant hMPVs carrying mutations in the SAM binding site

were defective in viral replication.

The replication kinetics of these rhMPV mutants in LLC-MK2 cells was determined. Briefly, LLC-MK2 cells were infected with each of recombinant virus at an

MOI of 0.01. At the indicated time points, the amount of virus in the supernatant of infected LLC-MK2 cultures were determined by an immunostaining assay. Figure 4.9 shows the growth curves of these rhMPV mutants. Recombinants rhMPV-G1696A,

G1700A, and D1755A were significantly delayed in viral replication as compared to rhMPV. The peak viral titer of hMPV-G1755A was 4.6log10PFU/ ml, which was 1.4 log lower than wildtype rhMPV. The peak titers of rhMPV-G1696A and D1700A were

5.5 and 5.6 log10PFU/ ml at day 6 post-infection, respectively, which was only 0.5 log less than wildtype rhMPV (Figure 4.9).

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Figure 4.9 Growth kinetics of rhMPVs carrying mutations in the SAM binding site. LLC-MK2 cells were infected by each recombinant virus at an MOI of 0.01. Supernatant aliquots (1 mL out of a total volume of 10 mL) were taken on the indicated days post infection and replenished with same volume of fresh medium. Viral titer was determined by an immunostaining plaque assay. Each experiment was performed in duplicate.

The progression of the cytopathic effects (CPE) was recorded in LLC-MK2 cells.

Wildtype rhMPV had an extensive CPE at day 3 post-inoculation. The CPE caused by rhMPV mutants were significantly delayed. The onset of CPE caused by rhMPV-G1696A was delayed for 3 days whereas rhMPV-G1700A and D1755A were delayed for 1 day (Figure 4.10). Collectively, these results demonstrated that rhMPVs carrying mutations in the SAM binding site had defects in viral replication and were attenuated in cell culture.

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Figure 4.10 rhMPV mutants had delayed cytopathic effects (CPE). LLC-MK2 cells were infected by rhMPV or rhMPV mutants at an MOI of 0.1 and incubated at 37 °C for the indicated time period. CPE was visualized by inverted light microscope.

Since rhMPV-D1755A had 1.5 log defects in titer, we further optimized the growth of this rhMPV mutant by infecting LLC-MK2 cells at different MOIs. As shown in

Table 4.1, the viral titer significantly increased when LLC-MK2 cells were infected by rhMPV-D1755A at a higher MOI. At an MOI of 1.0, the titer of rhMPV-D1755A was approximately 1.3 log less than that of rhMPV.

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Table 4.1 Optimize the growth of rhMPV-D1755A

MOI 0.01 0.1 1.0 Harvest time (hours) 96 72 72

Titer (log10PFU/mL) 4.51±0.24 5.09±0.27 5.81±0.15

4.4.5. Recombinant hMPVs carrying mutations in SAM binding site had

defects in viral protein synthesis

Next, we determined whether these recombinant viruses had defects in viral protein synthesis by measuring N protein expression using Western blot. Figure 4.13 shows the kinetics of viral N protein synthesis. Recombinant rhMPV-G1700A and

D1755A had a significant delay in N protein synthesis. Quantitative analysis of protein bands showed that rhMPV-G1700A and D1755A had approximately 30, 50% and 20%,

40% decrease in N protein expression compared with rhMPV at days 3, 5, 7 post-infection respectively. In comparison, rhMPV-G1696A showed moderate

(70-90%) decreases in N protein expression compared to rhMPV(Figure 4.13).

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Figure 4.13 Analysis of hMPV N protein expression by Western blot. Vero E6 cells were infected with rhMPV or rhMPV mutant at an MOI of 1. The cells were harvest to prepare total cell lysate at indicated time and were subsequently subjected to SDS-PAGE, followed by Western blotting using a monoclonal antibody against hMPV N protein.

4.4.6. Recombinant hMPVs carrying mutations in the SAM binding site had

defects in viral genomic RNA replication and mRNA synthesis

We also determined viral genomic RNA replication and mRNA synthesis for each recombinant virus. Since we used a primer annealing to the hMPV leader sequence, viral genomic RNA can be directly quantified by real time RT-PCR using total RNA isolated from virus-infected cells. Since viral genomic RNA was not polyadenylated, we separated the mRNA from genomic RNA by using a poly (A) binding assay. As shown in Figure 4.11A, all three rhMPV mutants had significant defects in viral mRNA synthesis and genomic RNA replication (P<0.05). At day 5 post-infection, N mRNA of rhMPV-G1696A, G1700A, and D1755A was 1.1, 0.5, and 3.3 fold less than rhMPV, respectively (Figure 4.11A). At the same time point (day 5 post-infection), genomic

RNA of rhMPV-G1696A, G1700A, and D1755A was 1.6, 0.8, and 1.7 fold less than rhMPV, respectively (Figure 4.11B). 127

Figure 4.11 Quantification of viral N mRNA and genomic RNA by real-time RT-PCR. LLC-MK2 cells were infected with rhMPV or rhMPV mutant at an MOI of 0.1 for various periods of time as indicated. Total RNA was isolated using TRizol reagent (Invitrogen, CA). (A) Viral genomic RNA: Using total RNA as templates, genomic RNA was quantified by real-time RT-PCR using specific primers annealing to hMPV leader sequence and N gene. (B) N mRNA: Viral mRNA was further purified using Dynabeads mRNA isolation kit and subsequently quantified by real-time PCR using primers annealing to the N gene. The results are the average of three independent experiments and are expressed as means ± SEs of copy numbers of the transcribed N mRNA or viral genomic RNA.

4.4.7. Recombinant hMPV carrying mutations in the SAM binding site were

specifically defective in 2’-O, but not G-N-7 methylation

Messenger RNAs of paramyxoviruses are typically methylated at G-N-7 and ribose 2’-O positions. Unlike eukaryotic mRNA cap methylation, G-N-7 and ribose

2’-O methylations of NNS RNA viruses are catalyzed by a single peptide, the CR VI of the L protein. In methylation reactions, the SAM binding site of an MTase is involved in binding SAM molecules which donate the methyl group for methylation.

Bioinformatics analysis showed that only one single SAM binding site in CR VI of L protein, suggesting that G-N-7 and ribose 2’-O MTase of NNS RNA viruses share a

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same SAM binding site. Thus, mutations to this SAM binding site may potentially affect G-N-7, ribose 2’-O, or both methylations. An in vitro mRNA cap methylation assay was performed to determine if the mutant rhMPVs were defective in a specific mRNA cap methylation. Briefly, LLC-MK2 cells were mock infected or infected by wild-type or mutant rhMPVs at MOI of 0.1, and viral mRNAs for each recombinant virus were harvested. To determine whether rhMPV mutants were defective in G-N-7 methylation, 500 ng of mRNA was incubated with 10 units of vaccinia virus G-N-7

MTase in the presence of 15 μCi [3H]-SAM. After the methylation reaction, RNA was purified, and the methylation of mRNA cap structure was measured by 3H incorporation using a scintillation counter. As shown in Figure 4.12A, rhMPV mRNAs were not methylated by vaccinia virus G-N-7 MTase, which is consistent with the fact that rhMPV produces G-N-7 methylated mRNA.

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Similarly, mRNAs produced by rhMPV-G1696A, G1700A, and D1755A were not methylated by vaccinia virus G-N-7 MTase, suggesting that these recombinant viruses are not defective in G-N-7 methylation.

Figure 4.12 In vitro trans methylation assay. A. Trans G-N-7 methylation assay: Vaccinia virus G-N-7 MTase was used for trans G-N-7 methylation as described in Materials and Methods. The ratio of H3 incorporation of viral mRNA between rhMPV mutant and rhMPV was calculated. Average of four independent experiments was shown. . B. Trans 2’-O methylation assay: Vaccinia virus 2’-O MTase was used for trans 2’-O methylation as described in Materials and Methods. The ratio of H3 incorporation of viral mRNA between rhMPV mutant and rhMPV was calculated. Average of four independent experiments was shown. (**:p≤0.01, *:p≤0.05)

A similar strategy was used to determine the 2’-O methylation in mRNA. 500 ng of viral RNA were incubated with 10 units of vaccinia virus 2’-O MTase in the presence of 15 μCi [3H]-SAM. After the methylation reaction, RNA was purified, and the level of 2’-O methylation was measured by 3H incorporation using a scintillation counter. As shown in Figure 4.12B, rhMPV mRNAs were not methylated by vaccinia virus 2’-O

MTase, which is consistent with the fact that rhMPV produces 2’-O methylated mRNA.

The H3 incorporation of rhMPV-G1696A was approximately 10 fold higher than that of 130

wildtype rhMPV, suggesting that rhMPV-G1696A had a significant defect in 2’-O methylation. Also, the H3 incorporation of rhMPV-G1700A and D1755A was approximately 5 and 4 fold higher than that of wildtype rhMPV respectively, suggesting that rhMPV-G1700A and D1755A had moderate defects in 2’-O methylation. (Figure 4.12B). Therefore, rhMPVs carrying mutations in the SAM binding site were specifically defective in 2’-O, but not G-N-7 methylation.

4.4.8. Genetics stability

To determine whether rhMPV mutants were genetically stable in cell culture, each rhMPV mutant was passed repeatedly 10 times in LLC-MK2 cells. Virus from each passage was sequenced. It was found that all rhMPV mutants retained the desired mutation in CR VI of L gene. No additional mutation was found this this region. This result suggests that these rhMPV mutants were genetically stable.

4.5. Discussion

A growing body of evidence suggests that mRNA cap methylation in NNS RNA viruses evolved a unique mechanism in that both G-N-7 and 2’-O methylations are catalyzed by a single peptide (CR VI of L protein) and the two methylases share a single SAM binding site. In this study, we performed a mutagenesis analysis in the

SAM binding site of hMPV L. We showed that mutations to the SAM binding site diminished gene expression in a minigenome replication assay. Using a reverse genetics system, we successfully recovered three recombinant hMPVs carrying

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mutations in the SAM binding site (rhMPV-G1696A, G1700A, and D1755A). These recombinant viruses were specifically defective in 2’-O, but not G-N-7 methylation, and were highly attenuated in cell culture as judged by viral plaque size, replication kinetics, and viral RNA and protein synthesis. Our results demonstrated that inhibition of viral 2’-O methylation can serve as a new approach to attenuate hMPV.

The mechanism of mRNA cap methylation in NNS RNA viruses is still poorly understood. Our current understanding of mRNA cap methylation in NNS RNA viruses comes from the studies of VSV. First, VSV mRNA cap G-N-7 and 2’-O methylations are catalyzed by CR VI of L protein whereas cellular mRNA cap G-N-7 and 2’-O methylations are carried out by two separate proteins. Second, VSV mRNA cap methylation is achieved in a sequential order in which the 2’-O MTase precedes and facilitates the G-N-7 MTase. However, all known cellular mRNA caps are first methylated at G-N-7 position followed by 2’-O position. Third, mutations to the MTase active site and cap binding site in CR VI of the VSV L protein abolished both G-N-7 and 2’-O methylations whereas mutations to the SAM binding site specifically diminished G-N-7, but not 2’-O methylation. However, it is not known whether this unique mechanism is utilized by other NNS RNA viruses. Using an in vitro methylation assay, we found that hMPVs carrying mutations in the SAM binding site were specifically defective in 2’-O, but not G-N-7 methylation. In contrast, recombinant

VSVs carrying mutations in the SAM binding site (rVSV-G1670A, G1672A, S1673A, and D1735A) were specifically defective in G-N-7, but not 2’-O methylation. None of 132

VSV mutants in CR VI of L had specific defects in 2’-O, but not G-N-7 methylation.

This is consistent with the order of VSV mRNA cap methylation in which 2’-O methylation is required for G-N-7 methylation. Collectively, our results suggest that the order of mRNA cap methylation in hMPV is different from that of VSV. Specifically,

2’-O methylation is not required for G-N-7 methylation in hMPV whereas 2’-O methylation occurs prior to G-N7 methylation in VSV. Interestingly, several other studies also support that the order of mRNA cap methylation in other paramyxoviruses differs from VSV. In early 1980, it was found that the mRNA of RSV lacks 2’-O methylation. However, more recent studies found that RSV mRNA is G-N-7 methylated at a low SAM concentration but could be doubly methylated cap at higher

SAM concentration, providing evidence that mRNA cap methylation of RSV first occurs at position G-N-7 followed by position 2’-O. Recently, a fragment of Sendai virus L protein that includes CR-VI was able to methylate short Sendai virus-specific

RNA sequences in vitro at the G-N-7 position (Ogino, et al. 2005). However, 2’-O methylation was not detectable in their trans methylation assay. This result suggests that G-N-7 methylation of Sendai virus mRNA cap can occur without the pre-methylation of 2’-O position. Therefore, these evidences suggest that the order of cap methylation in paramyxoviruses is the same as cellular mRNAs, but differs from that of VSV.

An important finding in this study is that recombinant hMPVs carrying mutations in the SAM binding were highly attenuated in cell culture, which can potentially be 133

used as vaccine candidates. These recombinant viruses formed much smaller immune-spots and plaques, had significantly delayed CPE and replication kinetics, and were defective in viral RNA and protein synthesis compared to wild-type rhMPV.

Importantly, these recombinant viruses were genetically stable when passing in cell culture. By optimizing the growth, we found that the yield of these recombinant viruses in cell culture had only approximately 0.3 log defects compared to rhMPV. These characteristics suggest that targeting mRNA cap methylation is a new approach to attenuate hMPV and perhaps other paramyxoviruses for vaccine purposes. Previously, a variety of approaches have been employed to attenuate paramyxoviruses including deleting nonessential genes, and engineering mutations in surface glycoproteins. We now add the inhibition of mRNA cap methylation to this list. There are many advantages of using mRNA cap methylation as a target for virus attenuation. First, viruses lacking MTase would not affect immunogenicity since the MTase is located in L protein, which is not a neutralizing antibody target. Second, mRNA cap methylation requires a different cluster of amino acids residues for catalysis, SAM binding, and

RNA substrate binding site. Mutations to any of these critical residues would lead to the impairment of one common function, mRNA cap MTase. Since most NNS RNA viruses possess two MTases, recombinant viruses specifically defective in G-N-7 and/or ribose 2’-O can be generated. Furthermore, different amino acid substitutions in the MTase domain will have variable impacts on mRNA cap methylation which will lead to generating a number of recombinant viruses with different levels of attenuation.

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By combining multiple substitutions in the MTase region of L protein, it should be possible to generate an attenuated virus that is genetically stable, because reversion to wild type at any single amino acid should not provide fitness gain. For example, rVSVs carrying mutations in the MTase active site (approximately 2 log reduction in virus titer) were more attenuated than rVSVs carrying mutations in the SAM binding site and substrate binding site (1 log reduction in virus titer). In addition, rVSV carrying three or four combined mutations in the SAM binding site can also be generated (Li,

Fontaine-Rodriguez et al. 2005; Li, Wang et al. 2006). In this study, we also found that rhMPV carrying mutations in MTase active site had more defect in growth compared to those hMPV mutants in the SAM binding site. Our future work will focus on characterization of the phenotype of rhMPVs carrying mutations in the MTase active site and RNA substrate binding site.

The successful recovery of rhMPV mutants specifically defective in 2’-O methylation also provides an essential tool to study the biological function of 2’-O methylation in NNS RNA viruses. It is firmly established that G-N-7 methylation is essential for mRNA stability as well as efficient translation. However, the role(s) of ribose 2’-O methylation have proved more elusive although it has been discovered more than 4 decades. Recent studies on West Nile virus (WNV) suggest that the 2′-O methylation of the 5′ cap of viral RNA functions to evade innate host antiviral responses through escape of the suppression of interferon-stimulated genes, tetratricopeptide repeats (IFIT). Specifically, mutant WNV (E218A) defective in 2′-O 135

MTase activity was attenuated in wild-type C57BL/6 mice, but remained pathogenic in knockout mice that lacked the type I interferon (IFN) signaling pathway. In addition, a vaccinia virus mutant (J3-K175R) and mouse hepatitis virus mutant MHV-D130A, both of which lacked 2′-O MTase activity, exhibited enhanced sensitivities to the antiviral actions of IFN mediated by IFIT proteins. Interestingly, it was also reported that 2’-O methylation of mouse and human coronavirus RNA facilitates evasion from detection by the cytoplasmic RNA sensor Mda5 (Daffis, Szretter et al. 2010; Habjan,

Hubel et al. 2013). Taken together, these studies suggest that 2’-O methylation of viral

RNA provides a molecular signature for the discrimination of self and non-self mRNA.

However, it is necessary to determine whether 2’-O methylation plays a similar biological function in NNS RNA viruses. NNS RNA viruses include a wide range of human, animal, and plant pathogens, causing a diverse type of diseases. For example,

VSV can cause infection in many cell lines including yeasts, insect cells, worms, and mammalian cells. Interestingly, the order of mRNA cap methylation is reversed compared to all known mRNA cap methylation (Rahmeh, Li et al. 2009). It will also be interesting to understand the mechanism by which mRNAs of avian paramyxovirus

(Newcastle disease virus, NDV) naturally lacks 2’-O methylation (Colonno and Stone

1976).

In conclusion, we have generated a panel recombinant hMPVs carrying mutations in the SAM binding of L protein. These mutant rhMPVs were highly attenuated in cell culture and were specifically defective in 2’-O, but not G-N-7 methylation. Our results 136

demonstrated that targeting viral MTase is a novel approach to rationally attenuate hMPV and perhaps other paramyxoviruses for vaccine purpose.

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Chapter 5.

Pathogenicity and immunogenicity of

methyltransferase-defective human metapneumovirus in cotton

rats

5.1. Abstract

Human metapneumovirus (hMPV) is the second leading causative agent of pediatric respiratory tract disease worldwide. Despite major efforts, there are no vaccines or antiviral drugs to combat this virus. Since the discovery of hMPV, approaches to generate vaccines employing viral proteins or inactivated vaccines have failed either due to a lack of immunogenicity or the potential for causing enhanced pulmonary disease upon natural infection with the same virus. It was suggested that a live attenuated vaccine is the most promising vaccine for hMPV. However, it has been a challenge to identify a vaccine strain that has an optimal balance between attenuation and immunogenicity. Therefore, exploration of new approaches to attenuate hMPV is urgently needed. Here, we found that the viral mRNA cap methyltransferase is a novel target to generate live attenuated vaccines for hMPV. Recombinant hMPVs lacking

2’-O methylation (rhMPV-G1696A, G1700A, and D1755A) were not only highly

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attenuated in cell culture but also were attenuated in viral replication in upper and lower respiratory tracts in cotton rats. In addition, these recombinant viruses caused minimal lung histological changes and viral antigen expression in bronchial epithelium cells.

Importantly, cotton rats vaccinated with these MTase-defective rhMPVs triggered a high level of neutralizing antibody and were completely protected from challenge with wildtype rhMPV. Together, these results demonstrated that MTase-defective rhMPVs were sufficiently attenuated but retained high immunogenicity in cotton rats. Therefore,

MTase-defective rhMPVs are excellent live attenuated vaccine candidates for hMPV.

5.2. Introduction

Human metapneumovirus (hMPV) is an emerging human paramyxovirus causing upper and lower respiratory tract infections. It is estimated that hMPV is responsible for

5-15% of pediatric respiratory infections requiring hospitalization and virtually all children have been infected by the virus by the age of 5 (van den Hoogen, de Jong et al.

2001). Despite major efforts, there is no FDA approved vaccine or antiviral drug against this pathogen.

It has been a challenge to develop vaccines for the human paramyxoviruses.

Generally, inactivated and live attenuated vaccines are the two most common strategies used in vaccine development. For safety reasons, an inactivated vaccine is often preferred. However, an inactivated vaccine developed for human paramyxoviruses resulted in serious complications when tested in human clinical trials. In the early

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1960s, the vaccination of infants with a formalin-inactivated (FI) RSV vaccine not only failed to protect against RSV disease during the following RSV season, but some vaccine recipients developed enhanced disease upon infection with RSV, resulting in increased rates of severe pneumonia and 2 deaths. Since the discovery of hMPV, several vaccine strategies have been explored, including FI hMPV vaccine, live attenuated vaccines, vectored vaccines, and subunit vaccines. In 2007, Yim KC, et al. reported that the FI vaccine induced hMPV-specific immune responses but resulted in enhanced lung damage upon reinfection in cotton rats, similar to what had been previously described in the hRSV clinical trial. Enhanced lung damage was also observed using an FI vaccine for human parainfluenza type 3 (PIV3), another important human paramyxovirus. These results demonstrated that an FI vaccine is not advised for hMPV, RSV, and PIV3, all of which cause respiratory tract infection in the same populations: infants, children, the elderly, and immunocompromised individuals.

Subunit vaccine candidates have also been tested targeting the two major surface glycoproteins, G and F. It has been reported that a G protein-based subunit vaccine was immunogenic in cotton rats but was incapable of eliciting protective antibodies against hMPV challenge. In comparison, F protein-based subunit vaccines were able to induce neutralizing antibodies against hMPV and conferred protective immunity in animal models (Herfst, de Graaf et al. 2007; Liu, Zheng et al. 2009). Specifically, cotton rats immunized with a DNA vaccine encoding F gene and/or a soluble F protein lacking the transmembrane domain triggered neutralizing antibody responses and provided partial 140

protection against virus shedding in the lungs compared to controls. In a separate study, immunization of Syrian golden hamsters with the soluble F proteins with adjuvant induced high virus-neutralization titers against both homologous and heterologous virus challenge. Upon challenge, the viral titer in the nasal turbinate of immunized animals was significantly reduced compared to the unimmunized challenge control.

Live attenuated vaccines are the most promising vaccine candidates for human paramyxoviruses since enhanced lung damage has not been observed in animal models or human clinical trials. Importantly, live attenuated vaccines are capable of inducing robust and prolonged immune responses since they mimic a natural virus infection.

Soon after the discovery of hMPV, cold-passage (cp) temperature-sensitive (ts) hMPV strains have been isolated by randomly passing the virus in cell culture at “cold” temperatures. These cpts-hMPVs showed attenuation in replication in the upper and lower respiratory tracts and protected against challenge with hMPV strains (Herfst, de

Graaf et al. 2008). Recently, a reverse genetic system has been developed and has made manipulation of the hMPV genome possible. A variety of live attenuated vaccine candidates have been generated by deleting nonessential genes such as G, M2-2, SH, and M2-1 (Biacchesi, Skiadopoulos et al. 2004; Biacchesi, Pham et al. 2005; Buchholz,

Biacchesi et al. 2005; Schickli, Kaur et al. 2008). In addition, chimeric viruses of hMPV and aMPV were also successfully recovered (Pham, Biacchesi et al. 2005).

These vaccine candidates were attenuated in cell culture as well as in animal models.

Animals vaccinated with these live vaccine candidates generated varying levels of 141

neutralizing antibody and protection against hMPV challenge. Despite the promise of these live vaccine candidates, it has been a challenge to identify an hMPV vaccine strain that has an optimal balance between attenuation and immunogenicity. The genetic stability of these vaccine candidates is still not known and to date none of them have been tested in human clinical trials.

We found that the viral mRNA cap methyltransferase is a new and novel target to generate live attenuated vaccines for hMPV. In Chapter 4, we performed a detailed characterization of recombinant hMPVs carrying a mutation in the SAM binding site and found that these MTase-defective hMPVs were highly attenuated in cell culture.

The objectives of this chapter are to determine whether MTase-defective hMPVs are attenuated in replication in an animal model and to determine whether they can be used as live vaccine candidates for hMPV.

5.3. Materials and Methods

5.3.1. Cell lines

LLC-MK2 (ATCC No. CCL-7) cells were maintained in Opti-MEM (Life

Technologies, Bethesda, MD) supplemented with 2% fetal bovine serum (FBS). Vero

E6 cells (ATCC No. CRL-1586) were grown in Dulbecco's modified Eagle's medium

(DMEM; Life Technologies) supplemented with 10% FBS.

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5.3.2. Virus preparation

The rhMPV stocks used in animal studies were grown in LLC-MK2 cells and further purified by ultracentrifugation. Briefly, LLC-MK2 cells in confluent T150 flasks were infected with each rhMPV at a multiplicity of infection (MOI) of 0.01 in a volume of 2 ml of DMEM. After 1 h of adsorption with constant shaking, 20 ml of

Opti-MEM (supplemented with 2% FBS) was added to the cultures, and infected cells were incubated at 37°C for 6 days. When extensive cytopathic effect (CPE) was observed, cells were harvested by scraping. The cell suspension was clarified by low-speed centrifugation at 3,000 g for 20 min at 4 oC in a Beckman Coulter Allegra 6R centrifuge. The cell pellet was resuspended in 2 mL of Opti-MEM and was subjected to three freeze and thaw cycles. The mixture was clarified by low-speed centrifugation and the supernatants were combined. The virus was pelleted by ultracentrifugation at

28,000 g in a Beckman Ty 50.2 rotor for 2 h. The final virus pellet was resuspended in

0.3 ml of Opti-MEM, aliquotted, and stored at -80 oC freezer. One vial of virus was thawed and the titer was determined by an immuno-staining assay.

5.3.3. Pathogenesis of rhMPV in cotton rats

For the replication and pathogenesis study, twenty-five four-week-old female specific-pathogen-free (SPF) cotton rats (Harlan Laboratories, Indianapolis, IN) were randomly divided into five groups (5 rats per group). The cotton rats were housed within ULAR facilities of The Ohio State University under approved Institutional

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Laboratory Animal Care and Use Committee (IACUC) guidelines. Each inoculation group was separately housed in rodent cages under BSL-2 conditions.

Figure 5.1 Schematic diagram of rhMPV replication and pathogenesis study in cotton rats.

Prior to virus inoculation, rats were anaesthetized with isofluorane. Rats in group

1 were inoculated with 2.0 ×105 PFU of the wildtype rhMPV and served as a positive control. Rats in groups 2-5 were inoculated with 2.0×105 PFU of three MTase-defective rhMPV mutants (rhMPV-G1696A, D1755A, and D1700A). Rats in group 6 were mock infected with 0.1 ml of Opti-MEM and served as uninfected control. Each rat was inoculated intranasally with a volume of 0.1 mL. After inoculation, the animals were evaluated on a daily basis for mortality, and the presence of any respiratory symptoms.

At day 4 post-infection, cotton rats were sacrificed, and lungs and nasal turbinate were collected for both virus isolation and histological analysis as described below.

5.3.4. Immunogenicity of rhMPVs in cotton rats

For the immunogenicity study, thirty cotton rats (Harlan Laboratories,

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Indianapolis, IN) were randomly divided into six groups (5 rats per group). Rats in group 1 were mock infected with Opti-MEM as an uninfected control, groups 2 to 6 were intranasally inoculated with 2.0 ×105 PFU of wildtype rhMPV or MTase-defective hMPVs in 0.1 mL Opti-MEM. Rats in group 7 were inoculated with DMEM and served as the unimmunized challenged control. After immunization, cotton rats were evaluated daily for mortality and the presence of any symptoms of hMPV infection. Blood samples were collected from each rat weekly by facial vein retro-orbital bleeding, and serum was isolated for neutralizing antibody detection. At week 4 post-immunization, rats in groups 2-7 were challenged intranasally with wildtype rhMPV at a dose of 1.0

×106 PFU per rat. After challenge, the animals were evaluated twice every day for mortality and the presence of any symptoms of hMPV infection. At day 4 post-challenge, all rats from each group were euthanized by CO2 asphyxiation. The lungs and nasal turbinate from each rat were collected for virus isolation and histological evaluation. The immunogenicity of the MTase-defective hMPVs was evaluated using the following methods: (i) humoral immunity was determined by a virus-serum neutralization assay using an end-point dilution plaque reduction assay. (ii)

Viral titer in the nasal turbinate and lungs was determined by an immunostaining plaque assay and viral genomic RNA was quantified by real-time RT-PCR. (iii) Pulmonary histopathology and viral antigen distribution was determined using the procedure described in Chapter 3. The protection was evaluated with respect to viral replication, antigen distribution, and pulmonary histopathology.

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5.3.5. Pulmonary histology.

After sacrifice, the right lung of each animal was removed, inflated, and fixed by 4% neutral buffered formaldehyde. Fixed tissues were embedded in paraffin, and sectioned at 5 microns. Slides were then stained with hematoxylin and eosin (H&E stain) for the examination of histological changes by light microscopy. The pulmonary histopathological changes were reviewed by 2-3 independent pathologists.

Histopathological changes were scored to include the extent of inflammation (focal or diffuse), the pattern of inflammation (peribronchilolar, perivascular, interstitial, alveolar), and the nature of the cells making up the infiltrate (neutrophils, eosinophils, lymphocytes, macrophages).

5.3.6. Immunohistochemical staining (IHC)

Right lung of animal was fixed in 10% neutral buffered formaldehyde and embedded in paraffin. 5-micron sections were cut and placed onto positively charged slides. After deparaffinization, sections were incubated with target retrieval solution

(Dako, Carpinteria, CA) for antigen retrieval. After antibody block, a primary mouse anti-hMPV monoclonal antibody (Virostat, Portland, ME) was incubated for 30 min at room temperature followed by incubation with a biotinylated horse anti-mouse secondary antibody (Vector Laboratories, Burlingame, CA). Slides were further incubated with ABC Elite complex to probe biotin (Vector Laboratories) and slides were then developed using a 3,3'-diaminobenzidine (DAB) chromogen Kit (Dako) and

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hematoxylin used as a counterstain. Lung sections from hMPV infected and uninfected samples were used as positive and negative controls respectively.

5.3.7. Determination of hMPV neutralizing antibody

hMPV-specific neutralizing antibody titers were determined using a plaque reduction neutralizing assay. Briefly, cotton rat sera were collected by retro-orbital bleed weekly until prior to challenge. The sera samples were heat inactivated at 56 °C for 30 min. Two-fold dilutions of serum samples were mixed with an equal volume of

DMEM containing approximately 100 PFU/well hMPV NL/1/00 in a 96 well plate and incubated at room temperature for 1 h with constant rotation. The mixtures were then transferred to confluent Vero E6 cells in a 96-well plate in triplicate. After 1 h incubation at 37 °C, the virus-serum mixtures were removed and the cells were overlaid with 0.75% methylcellulose in DMEM and incubated for another 4 days before virus plaque titration as described previously. Plaques were counted and 50% plaque reduction titers were calculated as hMPV-specific neutralizing antibody titers.

5.3.8. Determination of viral titer in lung and nasal turbinate.

Nasal turbinate and the left lung from each cotton rat were removed, weighed, and homogenized in 1 ml of phosphate-buffered saline (PBS) solution using Precellys 24 tissue homogenizer (Bertin, MD) following manufacturer’s recommendations. The presence of infectious virus was determined by an immuno-staining plaque assay in

Vero cells as described previously. 147

5.3.9. Immunostaining of recombinant hMPV

Viral titer in lung and nasal turbinate was determined by an immunostaining assay.

Briefly, Vero E6 cells (at the confluency of 90%) in a 24-well plate were inoculated with 10-fold serial dilutions of recombinant hMPV mutants and incubated at 37°C for 1 h. Infected cells were cultured in fresh Opti-MEM media (0.5 ml per well) at 37°C. At day 4 post-infection, the supernatant was removed and cells were fixed in a pre-chilled acetone: methanol solution (at the ratio of 3:2) at room temperature (RT) for 15 min.

Cells were permeablized in a phosphate saline buffer (PBS) containing 0.4% Triton

X-100 at RT for 10 min, and blocked at 37 °C for 1 h using 1% bovine serum albumin

(BSA) in PBS. The cells were then labeled with an anti-hMPV N protein primary monoclonal antibody (Millipore, Billerica, MA) at dilution of 1:1,000, followed by incubation with horseradish peroxidase (HRP)-labeled rabbit anti-mouse secondary antibody (Thermo Scientific, Waltham, MA) at dilution of 1:5,000. After incubation with AEC substrate chromogen (Sigma, St. Louis, MO), positive cells were then visualized under the microscope. Positive plaques were counted to determine viral titers.

5.3.10. Quantification of hMPV genomic RNA by real-time RT-PCR

Viral genomic RNA copy in lung and nasal homogenate was determined by real-time RT-PCR. First strand cDNA was synthesized by SuperScriptase III (Life

Technologies, Carlsbad, CA) using the following primer targeting the hMPV leader

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sequence: hMPV-leader-F-10:

5’-AAAACGCGTATAAATTAGATTCCAAAA-3’

A fragment covering partial leader sequence and partial N gene was quantified by

SYBR Green real-time PCR (Life Technologies)) using the following primers: hMPV-leader-10-forward:

5’-AAAACGCGTATAAATTAGATTCCAAAA-3’ hMPV-N-157-reverse:

5’-TGCAGTTGTTGTACCACATCTCTTT-3’

Real-time PCR was performed on a StepOne real-time PCR machine (Applied

Biosystems, Foster City, CA). PCR reaction and cycling parameters were set following the manufacturer’s recommendations. Briefly, the reaction mixture was setup as described in Table 5.1. For cycling parameters, a holding stage at 95°C was maintained for 10min to activate polymerase prior to cycling, followed by 50 cycles of 95°C for 15 s for denaturation and 60°C for 1 min for annealing and extension. Standard curves and

StepOne Software v2.1 were used to quantify genomic RNA copies. Viral RNA was expressed as mean Log10 genomic RNA copies per gram tissue ± standard deviation.

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Table 5.1 Reaction components for real-time PCR

Components Volume Power SYBR Green PCR Master Mix 10.0 µL Template 4.0 µL hMPV-leader-10-forward (25 µM) 0.6 µL hMPV-N-157-reverse (25 µM) 0.6 µL

ddH2O 4.8 µL Total 20.0 µL

5.3.11. Genetic stability of MTase-defective rhMPV in vivo.

Lung tissues from cotton rats infected by wildtype or mutant rhMPV were homogenized in 1 ml of PBS as described in Chapter 3. Total RNA was extracted from

200 µl of lung homogenate using an RNeasy mini-kit (Qiagen, Valencia, CA) following the manufacturer's recommendation. RT-PCR was performed using a One Step RT-PCR kit (Qiagen) with two hMPV specific primers spanning the MTase region of the L gene,

hMPV-L-11759-Foward: 5’-TATATAGGGTTTAAGAATTGG-3’

hMPV-L-13199-Reverse: 5’-ATCATTTTTTACTTACAAGC-3’

The reaction mixture was prepared as described in Table 5.2. For the cycling parameters, a holding stage at 50°C was maintained for 30min for reverse transcription.

Temperature was raised to 95 °C to activate polymerase prior to cycling, followed by 35

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cycles of 94°C for 1 min for denaturation, 60 °C for 1 min for annealing, and 72 °C for

1 min 30 s for extension. The PCR products were gel purified and sequenced at The

Ohio State University Plant Microbe Genetics Facility to confirm the presence of the introduced mutations using a sequencing primer:

hMPV-L-12113-Forward: 5’-GCTAAAGGAAAGCTAAC-3’

Table 5.2 Reaction components for one-step RT-PCR

Total Volume 25.0 µL

5.3.12. Statistical analysis.

Statistical analysis was performed by one-way multiple comparisons using SPSS

8.0 statistical analysis software (SPSS Inc., Chicago, IL). A P value of <0.05 was considered statistically significant. 151

5.4. Results

5.4.1. Replication and pathogenesis of MTase-defective rhMPVs in cotton

rats

(1) Viral replication in cotton rats

To determine whether MTase-defective rhMPVs were attenuated in vivo, all of

the recombinant viruses were inoculated in cotton rats and viral replication and

pathogenesis were determined. None of the cotton rats inoculated with recombinant

viruses exhibited detectable clinical symptoms of respiratory tract infection. No

distinguishable body weight change was observed in any treatment group. At day 4

post-infection, cotton rats were sacrificed, the viral replication in nasal turbinates and

lungs, and pulmonary histology were determined.

Table 5.3 Replication of MTase-defective rhMPVs in cotton rats

Nasal turbinate Lung

Mean titer Mean titer

% of % of log (genomic log log (genomic infected log(pfu/g) infected RNA copies/g) (pfu/g) RNA copies/g) animals animals WT 100 5.24±0.35 7.54±0.32 100 3.02±0.46 8.54±0.34

G1696A 0 ND 6.34±0.72 0 ND 6.72±0.24

G1700A 80 2.69±0.34 6.95±0.18 60 2.27±0.28 7.46±1.10

D1755A 0 ND 5.27±0.77 0 ND 6.07±1.55

ND: not detected

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As shown in Table 5.3, wildtype rhMPV replicated efficiently in the nasal turbinate and lungs of all five cotton rats. An average viral titer of log 5.24±0.35 log10PFU/g and log 3.02±0.46 log10PFU/g was found in nasal turbinate and lung, respectively. All MTase-defective rhMPVs had defects in viral replication in the nasal turbinate and lung. For recombinant rhMPV-G1700A, 4 out of 5 cotton rats had detectable virus in nasal turbinate with an average titer of log 2.69±0.34 log10PFU/g, and only 1 out of 5 rats had detectable virus in lung tissue with a titer of log 2.02 log10PFU/g. For recombinants rhMPV-G1696A and G1755A, there was no detectable infectious virus in either nasal turbinate or lung tissue. This result demonstrated that recombinants rhMPV-G1696A and G1755A were completely defective in viral replication in both the upper and lower respiratory tracts in cotton rats, whereas recombinant rhMPV-G1700A had low levels of viral replication in the upper respiratory tract. This result also demonstrated that MTase-defective hMPVs were attenuated in viral replication in vivo.

Since no infectious virus was detected in some of the MTase-defective rhMPV inoculated animals, real time RT-PCR was performed to determine whether viral genomic RNA was present in the nasal turbinate and lung tissues. Our results demonstrated that all virus-infected cotton rats had detectable viral genomic RNA in their tissues although they were negative for infectious virus. In cotton rats inoculated with wildtype rhMPV, 7.54 and 8.54 log10genomic RNA copies/g tissue were detected in nasal turbinate and lung tissues, respectively. The viral RNA level was significantly 153

reduced in the upper and lower respiratory tracts of all animals infected by the

MTase-defective rhMPVs. 6.34, 6.95, and 5.27 log10genomic RNA copies/g tissue were detected in the nasal turbinate of animals infected by rhMPV-G1696A, G1700A, and

D1755A, respectively. In addition, 6.72, 7.46, and 6.07 log10genomic RNA copies per gram of tissue were detected in lungs of animals infected by rhMPV-G1696A,G1700A, and D1755A, respectively, which is approximately 10~100 fold lower than that of wildtype rhMPV. Therefore, all MTase-defective rhMPVs had significant defects in genome RNA replication in cotton rats.

(2) Pulmonary histopathology

Half of the lung tissue was inflated by formaldehyde and subjected to HE staining and histopathological examination. Each lung tissue was scored on a scale of 0 (no change) to 3 (severe change) and is summarized in Table 5.4. Wild-type rhMPV caused moderate pulmonary histopathological changes including interstitial pneumonia, peribronchial lymphoplasmocytic infiltrates, mononuclear cell infiltrates, and edematous thickening of the bronchial submucosa. Recombinant rhMPV-G1700A induced similar interstitial pneumonia but showed less epithelium change, bronchial exudate, and mononuclear infiltration. Recombinants rhMPV-G1696A and D1755A caused similar mononuclear inflammation as wildtype rhMPV but had significantly less pulmonary histopathological changes. No pathological changes were found in the lungs of cotton rats inoculated with DMEM. These results similarly indicate that

MTase-defective hMPVs are attenuated in cotton rats. 154

Figure 5.2 Lung histological changes in cotton rats infected by MTase-defective rhMPVs. (A) Lungs from mock infected cotton rats. No histological changes were observed. (B) Lungs from cotton rats infected by rhMPV. Arrows indicates significant histological changes including interstitial pneumonia, lymphoid infiltrates, bronchial luminal exudate and airway remodeling. (C) Lungs from cotton rats infected by rhMPV-G1696A. (D) Lungs from cotton rats infected by rhMPV-G1700A. (E) Lungs from cotton rats infected by rhMPV-D1755A. Significantly less histological changes (indicated by arrow) were observed in panels C, D, and E compared to panel B.

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Table 5.4 Pulmonary histological changes in cotton rats vaccinated with MTase defective rhMPVs and challenged with rhMPV

Interstitial Bronchial epithelium Bronchial Peribronchial/perivascular Groups pneumonia change exudate Mononuclears inflammation rhMPV 0.9 1.45 0.30 0.70 G1696A 0.0 0.50 0.07 0.30 G1700A 0.4 0.69 0.07 0.53 D1755A 0.3 0.53 0.03 0.37

(3) Viral antigen distribution in lung tissues

To determine viral antigen distribution in lung tissues, IHC was performed using an antibody against the hMPV matrix protein. As shown in Figure 5.3, a large number of viral antigens were detected at the luminal surfaces of the bronchial epithelium cells in the lungs of wild-type rhMPV infected rats. The pattern of viral antigen was discontinuous and appeared to occur in clusters of adjacent cells. In some cases, luminal cellular debris that may include both degenerated epithelial cells and macrophages also stained positive for hMPV antigen. Significantly less viral antigen-positive cells were detected in the bronchial epithelium cells of the rhMPVP-G1700A infected group. No or little viral antigen was detected in lung tissues from the rhMPV-G1696A and D1755A groups. Consistently, this data demonstrated that MTase-defective hMPVs had significant defects in viral replication and antigen expression in the lower respiratory tract.

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Figure 5.3 Immunohistochemical (IHC) staining of lungs from cotton rats infected by MTase-defective rhMPVs. (A) Lungs from mock infected cotton rats. (B) Lungs from cotton rats infected by rhMPV. A large number of hMPV antigens were detected at the luminal surfaces of bronchial epithelial cells. (C) Lungs from cotton rats infected by rhMPV-G1696A. No or little antigen was detected. (D) Lungs from cotton rats infected by rhMPV-G1700A. Significantly fewer antigens were detected compared to rhMPV-infected lungs. (E) Lungs from cotton rats infected by rhMPV-D1755A. No or little antigen was detected.

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5.4.1. Immunogenicity of MTase-defective hMPVs in cotton rats.

Since MTase-defective rhMPVs were attenuated in vitro and in vivo, we subsequently determined their immunogenicity in cotton rats. Briefly, the rhMPV mutants were inoculated intranasally in cotton rats. Serum samples were collected weekly for the detection of humoral immune response. At week 4 post-inoculation, animals were challenged with 106 PFU of rhMPV. At day 4 post-challenge, all the animals were sacrificed and nasal turbinate and lung samples were collected for virus detection and pathological examination.

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Figure 5.4 Schematic diagram of immunogenicity study in cotton rats.

(1) Serum-virus neutralizing antibody

Serum antibody was determined by a plaque reduction neutralizing assay. As shown in

Figure 5.5, all of MTase-defective hMPVs elicited high levels of neutralizing antibody

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in cotton rats. Antibody was detectable at week 1 post-immunization and gradually increased during weeks 2-4. Overall, the antibodies generated by MTase-defective hMPVs were comparable to rhMPV immunization (P>0.05). In contrast, no hMPV-specific antibody was detected in the unvaccinated control. This result demonstrated that MTase-defective rhMPVs were not only sufficiently attenuated but also capable of triggering high levels of antibody.

Figure 5.5 Serum neutralizing antibody titers in cotton rats vaccinated with MTase-defective rhMPVs. Cotton rats were immunized by each recombinant hMPV intranasally at a dose of 2.0×105 PFU per rat. Serum samples were collected from each cotton rat weekly after immunization. Serum neutralizing antibodies were determined using plaque reduction neutralization assay. The antibody titers were calculated as 50% plaque reduction titer.

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(2) MTase-defective rhMPVs provide complete protection against viral replication in upper and lower respiratory tracts in cotton rats.

Viral replication in the nasal turbinate and lungs of cotton rats after challenge was determined by an immunostaining assay. As shown in Table 5.5, cotton rats vaccinated with wildtype rhMPV or MTase-defective rhMPVs did not have any detectable infectious virus particles in either the nasal turbinate or lungs after challenge with rhMPV. In contrast, unvaccinated challenge controls had average titers of log

5.59±0.49 and log 4.76±0.29 log10PFU/g in the nasal turbinate and lung, respectively.

These results demonstrated that rhMPV-G1696A, G1700A, and D1755A provided complete protection against viral replication in both upper and lower respiratory tracts after challenge with wildtype rhMPV.

Table 5.5 Immunogenicity of MTase-defective rhMPVs in cotton rats

Nasal Turbinate Lung

Mean titer % of infected Mean titer % of infected Log (genomic RNA Log (PFU/g) 10 animals 10 animals Log10(PFU/g) copies/g) DMEM 100% 5.59±0.49 100% 4.76±0.29 5.65±0.22

rhMPV 0 ND 0 ND 2.54±0.35

G1696A 0 ND 0 ND 2.68±0.22

G1700A 0 ND 0 ND 2.50±0.28

D1755A 0 ND 0 ND 2.46±0.23

ND: not detected

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(3) Pulmonary histopathology

After challenge, lung histology was evaluated for each cotton rat. As expected, unvaccinated challenged control had moderate pulmonary histological changes characterized by interstitial pneumonia, mononuclear cell infiltration, and edematous thickening of the bronchial submucosa, and bronchial epithelium changes. In contrast, significantly less histological changes were found in the lungs of cotton rats vaccinated with rhMPV-G1696A, G1700A, and D1755A (Table 5.6). No enhanced lung damage was observed in vaccinated challenged groups. No histologic changes were found in the unvaccinated unchallenged controls. These results demonstrated that MTase-defective rhMPVs provided protection against lung damage from virulent challenge.

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Figure 5.6 Lung histological changes in cotton rats vaccinated with MTase-defective rhMPVs followed by challenge with rhMPV. (A) Lungs from mock infected cotton rats. (B) Lungs from unimmunized challenge control group. Arrows indicate significant histological changes including interstitial pneumonia, lymphoid infiltrates, bronchial luminal exudate, and airway remodelling. (C) Lungs from cotton rats immunized by rhMPV and challenged with the same virus. (D) Lungs from cotton rats immunized by rhMPV-G1696A and challenged with rhMPV. (E) Lungs from cotton rats immunized by rhMPV-G1700A and challenged with rhMPV. (F) Lungs from cotton rats immunized by rhMPV-D1755A and challenged with rhMPV. Significantly less histological changes (indicated by arrow) were observed in panels C, D, and E compared to panel B. No enhanced lung damage was observed in panels C, D, and E.

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Table 5.6 Pulmonary histological changes in cotton rats infected by MTase-defective rhMPVs

Interstitial Bronchial Bronchial Peribronchial/perivascular Groups pneumonia epithelium change exudate Mononuclears inflammation DMEM 1.60 1.0 0.5 1.75 rhMPV 0.60 0.5 0.2 1.4 G1696A 1.13 0.3 0.4 1.0 G1700A 0.56 0.4 0.4 1.8 D1755A 0.10 0.4 0.0 0.8

(4) Viral antigen distribution after challenge.

For lung tissues from unvaccinated challenged controls, a large number of viral antigens were found at the luminal surface of the bronchial epithelial cells. Interestingly, lung tissues from vaccinated challenged groups exhibited different antigen distribution patterns. A significant amount of antigens were found inside of bronchial tissue, but not on the luminal surface of the bronchial epithelial cells. No antigen was found in the unvaccinated unchallenged group.

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Figure 5.7 Immunohistochemical (IHC) staining of lungs from cotton rats vaccinated by MTase-defective rhMPVs followed by rhMPV challenge. (A) Lungs from mock infected cotton rats. (B) Lungs from unimmunized challenge control group. A large number of hMPV antigens were detected at the luminal surfaces of bronchial epithelial cells. (C) Lungs from cotton rats immunized by rhMPV and challenged with the same virus. (D) Lungs from cotton rats immunized by rhMPV-G1696A and challenged with rhMPV. (E) Lungs from cotton rats immunized by rhMPV-G1700A and challenged with rhMPV. (F) Lungs from cotton rats immunized by rhMPV-D1755A and challenged with rhMPV. For panels C, D, E and F, hMPV antigens were found inside of bronchial tissue, but not on the luminal surface of the bronchial epithelial cells.

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5.4.2. Genetic stability of MTase-defective rhMPVs in vivo.

To determine whether MTase-defective rhMPVs were genetically stable in cotton rats, we amplified the CR-VI of L gene from total RNA extracted from each lung tissues.

Sequencing of these PCR products showed that all MTase-defective rhMPVs retained the desired mutation in the CR VI of L gene. No additional mutation was found this region. This result suggests that MTase-defective rhMPVs were genetically stable in cotton rats.

5.5. Discussion

Despite the significant health and economic burdens hMPV causes, no vaccine is currently available against this virus. In this study, we found that MTase-defective rhMPVs were sufficiently attenuated but retained high immunogenicity in cotton rats.

These results demonstrated that MTase-defective rhMPVs are excellent live vaccine candidates.

A live attenuated vaccine is the most promising vaccine for human paramyxoviruses such as RSV, hMPV, and PIV3. Live attenuated vaccines offer many advantages: (i) enhanced lung damage has not observed either after vaccination with live attenuated viruses or after natural reinfection; (ii) intranasal administration of live vaccines induces balanced immune responses that closely resemble natural virus infection; and (iii) intranasal vaccination induces better local immunity compared with intramuscular injection of subunit vaccines. However, it has been technically

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challenging to identify vaccine strains that have an optimal balance between attenuation and immunogenicity. Previously, cold passage (cp) temperature sensitive

(ts) live attenuated hMPV, nonessential gene-deleted hMPVs, and chimeric hMPVs have been generated. Many of these approaches either affect immunogenicity, genetic stability, or lead to insufficient attenuation or poor virus growth in vitro. To date, only one live vaccine candidate, which is a chimeric rhMPV carrying aMPV P gene, has been approved for phase I clinical trial. The safety and efficacy of this live attenuated hMPV vaccine in humans is not known. However, in case of RSV, none of cpts live attenuated vaccines were found to have satisfactory attenuation and immunogenicity when they were tested in human clinical trials. In fact, the current lead RSV vaccine candidate (MEDI-559) which encodes a number of cpts mutations and a deletion of the

SH gene induced detectable antibody responses in only 44% of human vaccine recipients, even after two doses. These results suggest that the exploration of new approaches to attenuate human paramyxoviruses is urgently needed.

We hypothesize that viral mRNA cap MTase is an excellent target to rationally attenuate hMPV for the development of live attenuated vaccines. The rationale for this hypothesis is that methylation of viral mRNA is essential for efficient translation of viral proteins. Therefore, disruption of the viral MTase will downregulate viral protein expression and in turn viral replication, leading to the attenuation of hMPV. There are several advantages of using MTase as the target for virus attenuation. First, viruses lacking MTase would likely not affect immunogenicity, since the MTase is located in L 167

protein, which is not a neutralizing antibody target. Second, mRNA of paramyxoviruses typically possesses two MTases, thus recombinant viruses specifically defective in

G-N-7 and/or ribose 2’-O can be generated. Third, different amino acid substitutions in the MTase domain will have variable impacts on mRNA cap methylation which will lead to generation of a number of recombinant viruses with different degrees of attenuation. This will allow us to identify vaccine strains that have a proper balance between attenuation and immunogenicity. Finally, the MTase domain is highly conserved in L proteins of all non-segmented negative sense (NNS) RNA viruses with the exception of the family Bornaviridae. Thus, this novel attenuation strategy can be employed for other NNS RNA viruses.

Using a reverse genetics approach, we generated rhMPVs carrying mutations in the SAM binding site and showed that these recombinant viruses were specifically defective in 2’-O but not G-N-7 methylation. These MTase-defective rhMPVs showed smaller plaques, delayed growth kinetics, lower peak titer and less protein synthesis in cell culture compared to wildtype rhMPV. In cotton rats, MTase-defective rhMPVs were found to replicate less efficiently compared to wildtype rhMPV.

Specifically, rhMPV-G1696A and rhMPV-D1755A were highly attenuated in virus replication in vivo. No infectious virus particles were found in the upper or lower respiratory tracts in cotton rats inoculated with these two rhMPV mutants.

Recombinant rhMPV-G1700A was moderately attenuated based on the evidence that 4 out of 5 animals had low levels of virus replication in nasal turbinate and 1 out of 5 168

animals had low virus replication in lungs. These MTase-defective rhMPVs were also defective in viral genomic RNA replication in vivo compared to rhMPV. Consistent with infectious virus replication data, rhMPV-G1700A had higher genomic RNA copies in tissues compared to the rhMPV-G1696A and rhMPV-D1755A groups. In addition,

MTase-defective rhMPVs induced significantly less lung histopathological changes and had significantly less viral antigen expression in the epithelial cells on the surface of bronchi, correlating with the degree of attenuation of viral replication in the lower respiratory tract of the animals. Collectively, these data demonstrated that

MTase-defective rhMPVs were attenuated in vitro and in vivo.

Excitingly, despite the sufficient attenuation, all MTase-defective rhMPVs elicited a high titer of neutralizing antibody that was comparable to those induced by wildtype rhMPV. All immunized animals were completely protected from viral replication in the upper and lower respiratory tracts after challenge with wildtype rhMPV. Viral genomic

RNA in lower respiratory tract of the immunized animals showed at least a 1000-fold reduction.

One of the major concerns in paramyxovirus vaccine development is the potential for enhanced disease upon re-infection, a lesson learned from a formalin-inactivated (FI)

RSV vaccine clinical trial in the 1960s. To date, the enhanced disease has not been found in live attenuated (cold-adapted, gene-deleted, and chimeric viruses) and subunit vaccine candidates for hMPV. In our study, we also evaluated the histopathological

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changes and viral antigen distribution in lungs of the immunized animals upon challenge (re-infection) with wildtype rhMPV. No enhanced lung damage was found after vaccination of MTase-defective hMPV followed by challenge. In fact, vaccination protected lungs from histologic changes caused by wildtype challenge. Interestingly, a large number of viral antigens were found at the luminal surfaces of the epithelial cells of bronchi in unvaccinated challenged groups. This is consistent with the previous observation that viral antigens were localized almost exclusively at the apical surfaces of ciliated respiratory epithelial cells (Kuiken et al., 2004). However, viral antigens were distributed inside but not on the luminal surface of bronchi in vaccinated challenged groups. One possibility is that alveolar macrophage cells and/or dendritic cells were activated by vaccination with live vaccines, and migrated to the respiratory surfaces to capture viral antigens upon virus challenge. It is also possible that alveolar mucosal antibody binds and neutralizes the virus, which was further captured by immune cells. Further study is needed to elucidate the mechanism of viral clearance.

Collectively, MTase-defective rhMPVs have a satisfactory balance between attenuation and immunogenicity in cotton rats, thus can serve as excellent live vaccine candidates for hMPV.

Recently, the concept of using MTase-defective viruses as vaccine candidate has been also demonstrated for vesicular stomatitis virus (VSV), another NNS RNA virus.

Based on the level of mRNA methylation, the MTase-defective VSVs were classified into one of two groups: (i) viruses completely defective in both G-N-7 and 2’-O 170

methylation in vitro (rVSV-K1651A, D1762A, and E1833Q; catalytic site mutants), or

(ii) viruses defective in G-N-7, but not 2’-O methylation (rVSV-G1670A and G1672A;

SAM binding site mutants). It was found that VSV mutants carrying mutations in the predicted MTase catalytic site were highly attenuated in the mouse model. Mice inoculated with these viruses did not exhibit weight loss or clinical signs of VSV infection, and did not show any of the classic histologic lesions in the lung or brain.

Mice inoculated with VSV mutants carrying mutations in the predicted SAM binding site exhibited mild weight loss, clinical signs (i.e. ruffled fur), and histologic lesions in the lung and brain, indicating that VSV mutants defective in G-N-7 methylation alone retained low virulence. In vivo studies showed that both groups (i and ii) of recombinants stimulated high levels of VSV-specific antibody and provided full protection against challenge with wild type VSV, demonstrating that viruses which are defective in mRNA methylation are excellent vaccine candidates.

Besides safety and efficacy, an ideal live vaccine must be genetically stable. In this study, all MTase-defective rhMPVs were blindly passed 10 times in cell culture and

MTase domain in L gene was sequenced. No reversion was found for each passage.

During the animal study, we amplified the MTase domain of L gene using total RNA purified from lung tissues. Sequence results showed all the fragments retained the desired mutation. This suggests that MTase-defective rhMPVs are genetically stable in vitro and in vivo. We will continuously pass these viruses in cell culture and pass the viruses repeatedly in cotton rats. If reversion occurs, we will build double or triple 171

mutations in the MTase region of L protein. By combining multiple substitutions, it should be possible to generate an attenuated virus that is genetically stable, because reversion to wild type at any single amino acid should not provide a fitness gain.

The outcome of this study will also provide rationale to design MTase-defective hMPV as a vector to deliver other antigens since the genome of rhMPV, like other NNS

RNA viruses, has the capacity to harbor foreign genes. This attenuated rhMPV can be used as a vector to express glycoproteins of other important respiratory pathogens, e.g. hRSV F and G, and hPIV3 HN and F.

In conclusion, this study highlights a new and novel strategy for the development of live attenuated hMPV vaccines. Our results showed that single amino acid substitutions in the SAM binding site of hMPV L protein significantly reduced viral replication and pathogenesis in cotton rats while retaining wildtype levels of immunogenicity. Since amino acid residues essential for MTase activity are highly conserved, this vaccine strategy can be applied to human paramyxoviruses .

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Chapter 6.

Future directions

6.1. Determine the genetic stability of MTase-defective rhMPV vaccine candidates

In this study, we showed that all MTase-defective rhMPVs were genetically stable when passed repeatedly in cell culture for 10 times. In addition, viral RNA isolated from lungs of cotton rats retained the desired mutation in the L gene. These results suggest that MTase-defective rhMPVs were genetically stable. However, we will continue to pass these recombinant viruses in cell culture and sequence the entire viral genome. In addition, we plan to pass these viruses repeatedly in cotton rats. If a revision is found, we will engineer double or triple mutations in the SAM binding site to further stabilize the virus. By combining multiple substitutions within the MTase region in the

L protein, it should be possible to generate an attenuated virus that is genetically stable, because reversion to wild type at any single amino acid should not provide a fitness gain.

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6.2. Optimize the vaccination strategy for MTase-defective rhMPV vaccine candidates

In this study, we showed that a single vaccination (at dose of 105 pfu) of

MTase-defective rhMPV vaccine candidates triggered a high level of protective immunity that was comparable to wildtype rhMPV vaccination. However, it is necessary to determine the optimal vaccination dose, effect of booster vaccination, and the duration of antibody response and protection. (i) The effects of vaccine dosage.

Four different dosages (104,105,106, and 107 PFU) will be chosen for vaccination of cotton rats. This will allow us to determine the optimal vaccination dosage. (ii) The effects of booster vaccination. Once the optimized vaccination dosage has been established, we will compare the efficacy of single vaccination to a booster vaccination regime. (iii) The duration of protection conferred by hMPV vaccine candidates.

Cotton rats will be vaccinated with an optimal dosage of the hMPV vaccines, and antibody titer will be monitored for a one year time period. The duration of antibody titer of the single vaccination group will be compared to the booster group.

6.3. Evaluate the cross protection of MTase-defective rhMPV vaccine candidates

In this study, we showed that MTase-defective rhMPVs provided complete protection against challenge with a homologous hMPV (subgroup A strain). It is not known whether they can confer cross-protection against heterologous hMPV strains. To do this, we will select a representative strain of hMPV subgroup B as a challenge virus.

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If little or no cross-protection is observed, we will generate MTase-defective rhMPV subgroup B virus by engineering mutations in the SAM binding site. Subsequently, we will develop multivalent live attenuated vaccine candidates by mixing MTase-defective rhMPV subgroups A and B viruses and examine their cross-protection against both subgroups A and B hMPV.

6.4. Determine the mechanism of protective immune responses induced by MTase-defective rhMPVs.

Although we showed that MTase-defective rhMPVs generated a high level of neutralizing antibody and provided complete protection against challenge with wildtype rhMPV, the mechanism of the protective immunity is still poorly understood.

For example, what are the roles of T cell immune response and potential local immune response (such as IgA) in protection? What are the roles of macrophage and dendritic cells in immune responses? What is the mechanism of viral clearance? What is the cytokine level after vaccination? Does MTase-defective rhMPVs trigger a high level of interferon? Is innate immunity correlated with adaptive immunity? A better understanding of these questions will facilitate the future clinical trials of the vaccine candidates.

6.5. Improve the growth of MTase-defective rhMPV vaccine candidates

In this study, we found that the yield of MTase defective rhMPVs in LLC-MK2 cells was approximately 0.4-0.8 log lower than wildtype rhMPV. To reduce the cost of vaccine production, it is necessary to improve the growth of these vaccine strains. First, 175

we will test whether MTase-defective rhMPVs have a host range phenotype. Previous studies showed that MTase-defective vesicular stomatitis virus (VSV) can grow well in baby hamster kidney (BHK) cells and chicken embryo fibroblasts, but are restricted in growth in most cell lines of human origin, such as HEp-2 cells. By comparing different cells, we may be able to identify a cell line that can improve the growth of

MTase-defective rhMPVs. An alternative strategy is to supplement cap methylation activity by expressing a ribose 2’-O MTase in cell culture. Since hMPV replicates in cytoplasm and doesn’t have access to nucleus where host capping and methylation machinery is located, we can transient express a 2’-O MTase in LLC-MK2 cells or establish LLC-MK2 cell line stably expressing a 2’-O MTase. Presumably, 2’-O methylation of these recombinant viruses can be restored during their replication in cytoplasm which may result in an enhanced viral gene expression and replication so that the growth capability and virus yield can be improved in cell culture.

6.6. Use MTase-defective rhMPV as a vector to deliver antigens from other human paramyxoviruses

Since MTase-defective rhMPV is attenuated in vitro and in vivo, it can be an excellent vaccine vector to express the major surface glycoproteins from other respiratory tract pathogens, such F and G proteins of hRSV, and F and HN proteins of hPIV3. Previously, it has been reported that a bovine/human chimeric PIV3 (b/hPIV3) expressing hRSV F protein stimulated protective immune responses against both hPIV3 and hRSV in hamsters and African green monkeys (Haller AA, et al, 2003, Tang

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RS, et. al, 2004). Similarly, Sendai virus expressing RSV F protein elicited hRSV-specific B and T cell responses in cotton rats (Zhan X, et al, 2007). Clearly,

MTase-defective rhMPV could be a promising vector for the development of multivalent vaccine candidates against pediatric respiratory tract pathogens.

6.7. Recover and characterize rhMPVs carrying mutations in MTase catalytic site and RNA binding site.

The SAM-dependent MTase superfamily contains at least three key elements essential for mRNA cap methylation: a K-D-K-E tetrad that functions as the catalytic residues of the MTase, a SAM binding site that is involved in binding the methyl donor

SAM, and an RNA substrate binding site that binds and interacts with mRNA cap. In this study, we found that mutations to the SAM binding site yielded recombinant viruses that were defective in 2’-O methylation and were highly attenuated and immunogenic in cotton rats. We will continue to recover and characterize recombinant hMPVs carrying mutations in the MTase catalytic site and RNA binding site. Since mRNAs of hMPV contain G-N-7 and 2’-O methylation, we should be able to generate rhMPVs that are specifically defective in G-N-7, 2’-O, or both methylations. These recombinant hMPVs will exhibit different degrees of attenuation characteristics dependent on the specific methyl group involved and the degree of the defect in mRNA cap methylation. We anticipate that we will identify a number of hMPV mutants that are sufficiently attenuated and genetically stable, but maintain wild type levels of immunogenicity.

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6.8. Facilitate the clinical trials of the MTase-defective rhMPV vaccine candidates in nonhuman primates

Although MTase-defective rhMPV vaccine candidates were highly attenuated and immunogenic in cotton rats, it is not known whether they have similar attenuation characteristics and immunogenicity in nonhuman primates. The results generated from our study will facilitate the future clinical trials of these hMPV vaccine candidates in nonhuman primates and humans. It will be exciting to evaluate the safety and efficacy of these vaccine candidates in humans.

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