Soluble Respiratory Syncytial Virus Fusion Protein in the Fully Cleaved

Soluble Respiratory Syncytial Virus Fusion Protein in the Fully Cleaved,

Pretriggered State, a Tool to Study Protein Triggering

DISSERTATION

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

Supranee Chaiwatpongsakorn

The Comparative and Veterinary Medicine Graduate Program

The Ohio State University

2011

Dissertation Committee:

Mark Peeples

Michael Oglesbee

Stefan Niewiesk

Jianrong Li

Supranee Chaiwatpongsakorn

2011

ABSTRACT

Respiratory syncytial virus (RSV), a member of the Paramyxoviridae family,

Pneumovirinae subfamily is the most significant respiratory pathogen in infants and second only to influenza virus in the elderly. Despite extensive efforts, no vaccines or small molecule antiviral drugs are available. The RSV fusion (F) glycoprotein has been a major target for vaccine and antiviral drug development because of its importance in the viral replication cycle, its conserved sequence and structure, its exposed position in the virion, and its strong immunogenicity. Like other paramyxoviruses, the RSV F protein is anchored in the virion membrane in a metastable, pretriggered form. Once triggered, the F protein undergoes a dramatic conformational extension that inserts its hydrophobic fusion peptide into the target cell membrane, then folds back on itself to bring the membranes together and initiate fusion.

However, the Pneumovirinae F protein is unique in that it, alone, is sufficient to mediate membrane fusion and virus infection. It is, therefore, the simplest F protein to study. It likely has the ability to attach to target cells from which position it is triggered. Neither the trigger site on the F protein nor the triggering molecule/event has been identified. To begin to study the triggering mechanism of the RSV F protein biochemically, we have generated a soluble

F (sF) protein by replacing the transmembrane and cytoplasmic tail domains

ii with a 6His tag. This sF protein is secreted efficiently from 293T cells in a fully cleaved form. It is recognized by neutralizing monoclonal antibodies, appears spherical by electron microscopy, and is not aggregated, all consistent with a native, pretriggered trimer. The sF protein was purified on a Ni2+ column and eluted with 50 mM phosphate buffer containing 500 mM NaCl and 250 mM imidazole. Dialysis against 10 mM buffer caused the sF protein to trigger, forming “hatpin” shaped molecules that aggregated as rosettes, characteristic of the posttriggered form. Further dialysis experiments indicated that the efficiency of triggering correlated well with the reduction of buffer molarity.

Reduction of buffer molarity by dilution also resulted in exposure of the fusion peptide as detected by liposome association, confirming sF protein triggering.

Mutation of the furin cleavage site adjacent to the fusion peptide prevented liposome association, confirming that association is via the fusion peptide.

Although it is not clear whether reduction in molarity can serve as a physiological trigger of the intact F protein during the natural infection of RSV, our study has revealed a novel, surrogate method for triggering a viral fusion protein. The availability of pretriggered RSV sF protein capable of being triggered and transformation into its posttriggered conformation enables studies of its mechanism of attachment, triggering, and refolding, a protein vaccine for adults, assays to quantify antibodies against F, discovery of the mechanism of action of drugs known to target F, and high throughput screens to identify new and better drugs against F.

iii

DEDICATION

This dissertation is dedicated to my family and the loved ones. The first one to thank was my dad, for being a big inspiration and a role model of my life. His unequivocal support and encouragement always made me feel confident to overcome every difficulty I encountered. He always let me know that he was so proud of me, which motivates me to work harder and do my best. Thanks again for believing in me. Even though you are not here today, I always know how happy you would be to see my success. And you should know that you are a big part of it, always and will be. For my mom, this dissertation cannot be done and I would not have come this far without your unconditional love and support. You are truly my best friend who lives just a phone call away.

This work is also dedicated to my brother, P‟Kob, who always understands and shares the feeling I have had from study and research when no one else could.

I also dedicate this dissertation to everyone in Chaiwatpongsakorn family for their loving support. I thank them for being there to help out and comfort my parents when I and my brother were so far away in another country pursuing our doctoral degrees. Lastly, I also want to dedicate this work to P‟Jump for a wonderful time we have had together. Thanks for cheering me up, for trying to understand the person I am, and for believing in me when I lost faith in myself.

ACKNOWLDEGEMENTS

First of all, I would like to thank my advisor, Dr. Mark E. Peeples for his important support throughout this work. His wide knowledge and logical way of thinking have been a great value to me since the first day I joined the lab. I have learned a lot from him and realized for all these years how fortunate I am to have him as my mentor. In the last phase of my dissertation, he helped me to correct grammar mistakes and suggested possible improvements.

Without his help I could not have finished my dissertation successfully. I am also thankful to him for allowing me to go back home several times in the past few months to take care of my dad and to perform a religious ceremony after he passed away. Thank you so much again for your generous understanding during this difficult time.

I also thank Dr. Sunee Techaarpornkul for giving me a great opportunity and recommending me to Dr. Peeples. Laboratory training from her and Dr.

Nusara Piyapolrungroj in cell culture and molecular biology gave me tools that turned out to be essential in my PhD research. I own a sincere gratitude to both of them.

I would like to thank Drs. Richard and Raquel Epand for teaching me how to make and use liposomes and their valuable discussions and helpful

v comments on my project. I am also thankful for their warm hospitality and financial support during my training in Canada.

I wish to thank all past and present members of the Peeples lab for their technical support, scientific guidance, and great time we shared. It was my pleasure to share doctoral studies and life with wonderful people like Steve,

Anna, Olga, and Heather. Especially, Heather, I really appreciate your assistance on my F model project, thank you so much again for your kind friendship, for sharing the glory and sadness of our grant submission deadlines and day-to-day research.

I also would like to express my gratitude to my committee members, Dr.

Michael Oglesbee, Dr. Stefan Niewiesk, Dr. Jianrong Li, and Dr. Joan Durbin for providing me their valuable suggestions throughout my graduate study.

I would also like to extend my appreciation to many colleges from the

Veterinary Biosciences graduate program and Center for Vaccines and

Immunity who have assisted me in my studies and project for all these years.

And for Dr. Will Ray, I thank him for his amazing work on the RSV F computational models and for allowing me to present his work in my dissertation.

VITA

1998…………………………………………….The demonstration school of

Silpakorn University, Thailand

2003…………………………………………….B. Pharmacy, Silpakorn

University, Thailand (first class

honor)

2004 to present ……………………………….Graduate Research Associate,

Department of Veterinary

Biosciences, The Ohio State

University

PUBLICATIONS

Chaiwatpongsakorn, S., et al., Soluble Respiratory Syncytial Virus Fusion

Protein in the Fully Cleaved, Pretriggered State Is Triggered by Exposure to

Low-Molarity Buffer. J Virol, 2011. 85:3968-3977.

FIELDS OF STUDY

Major Field: The Comparative and Veterinary Medicine Graduate Program

Minor Fields: Molecular Virology

vii

TABLE OF CONTENTS

Abstract…...... ii

Dedication ...... iv

Acknowledgements ...... v

Vita……...... vii

List of Figures ...... xi

Abbreviations ...... xiv

Chapter 1: Respiratory Syncytial Virus ...... 1

Introduction ...... 1

Taxonomy and classification...... 2

RSV organization and morphology ...... 3

The RSV life cycle ...... 4

Attachment and entry ...... 4

Transcription, translation, and replication ...... 6

Assembly and budding ...... 8

The RSV proteins ...... 9

The nucleocapsid and polymerase complex ...... 9

viii

The nonstructural proteins NS1, NS2 ...... 10

The matrix (M) protein ...... 12

The attachment (G) protein ...... 12

The small hydrophobic (SH) protein ...... 15

The fusion (F) protein ...... 16

The mechanism of F protein-mediated membrane fusion ...... 26

Reverse genetics ...... 30

Vaccines...... 32

Treatment and preventions ...... 38

Chapter 2: Soluble RSV Fusion Protein ...... 40

Introduction ...... 40

Materials and methods ...... 44

Results…...... 48

Discussion ...... 70

Chapter 3: Relevance, Additional Studies and Future Studies ...... 76

Stabilization of the RSV soluble F protein in its pretriggered form ...... 76

Maximizing production of the RSV soluble F protein for crystalization: ...... 91

Application of the soluble F protein in screeing libralies of the compounds for

their ability to target the F protein ...... 99

Determine if reduction in molarity is the physiological cause of the F protein

triggering ...... 100 ix

Analysis of the MαH on the RSV F protein for its role in fusion function ...... 103

References ...... 120

LIST OF FIGURES

Fig. 1.1. Cartoon of the RSV F protein domains and cleavage sites ...... 18

Fig. 1.2. Cartoon of F protein refolding mediated viral-cell membranes fusion ... 27

Fig. 2.1. The wild-type RSV F protein and the cDNA-expressed SC-2 sF

protein generated from it ...... 49

Fig. 2.2. Western blot analysis of the SC-2 sF protein produced from

transfected 293T cells at 48 h posttransfection ...... 51

Fig. 2.3. MAb immunoprecipitation of the SC-2 sF protein after incubation at

4°C or 50°C for 1 h ...... 53

Fig. 2.4. Electron micrographs of the SC-2 sF protein ...... 55

Fig. 2.5. Analysis of the SC-2 sF protein aggregation state by velocity sucrose

gradient centrifugation ...... 56

Fig. 2.6. Sucrose velocity gradients and electron micrographs of the SC-2 sF

protein following dialysis with six different buffers ...... 60

Fig. 2.7. Association of the RSV sF protein with POPC-POPE-cholesterol

(8:2:5) large unilamellar liposomes ...... 63

Fig. 2.8. EM of SC-2 sF protein inserted into liposomes ...... 65

Fig. 2.9. Efficient secretion of sF protein cleavage mutants and the

requirement of cleavage at the furin site adjacent to the fusion

peptide for liposome association ...... 68

Fig. 3.1. Analysis of the SC-2 sF protein aggregation state by velocity

sedimentation on a sucrose gradient ...... 77

Fig. 3.2. Constructed RSV sF proteins ...... 79

Fig. 3.3. Western blot analysis of the sF proteins produced from transfected

293T cells at 48 h posttransfection ...... 81

Fig. 3.4. MAb immunoprecipitation of the SC-2 sF protein after incubation at

4°C or 50°C for 1 h ...... 84

Fig. 3.5. Analysis of the MP-A sF protein aggregation state by velocity

sedimentation on a sucrose gradient ...... 85

Fig. 3.6. Alignment of the F protein amino acid sequences of strain D53 and

A2 ...... 89

Fig. 3.7. Secretion efficiency and sucrose gradient analysis of SC-2 sF protein

and mutants containing one, two or all three differences with A2 sF

protein ...... 90

Fig. 3.8. Coomassie Blue staining analysis ...... 92

Fig. 3.9. Western blot analysis of MP-A sF protein expressed from two

different plasmids ...... 94

Fig. 3.10. Generation of the codon-optimized RSV Long strain F protein from

synthetic oligonucleotides and analysis of its fusion activity ...... 96

xii

Fig. 3.11. Models of the pretriggered (A) and posttriggered (B) RSV F protein

monomer ...... 106

Fig. 3.12. Model of the pretriggered RSV F protein trimer ...... 108

Fig. 3.13. Intracellular processing and cell surface expression of F protein

mutants in MαH ...... 114

Fig. 3.14. Comparison of protein production, cell surface expression and cell-

cell fusion activity of F protein mutants ...... 115

Fig. 3.15. Summary of critical residues that form the MαH motif (A) and

stereoimage of the MαH in side (B) and top (C) views highlighting

the critical amino acids ...... 118

xiii

ABBREVIATIONS

A Alanine (Ala)

AA Amino acid

ALSV Avian leukemia and sarcoma virus

ARE Apical recycling endosome

AUG Start codon sequence

BRSV Bovine respiratory syncytial virus

Ca2+ Calcium cDNA Complementary deoxyribonucleic acid

CMV Cytomegalovirus

CO2 Carbon dioxide

CTD Cytoplasmic tail

Cys Amino acid cysteine

C terminus Carboxy terminus

°C Degrees Celsius

D Aspartic acid, Asp

DMEM Dulbecco‟s modified Eagle medium xiv

DMPC Dimyristoylphosphatidylcholine

DNA Deoxyribonucleic acid dsRNA double stranded ribonucleic acid

E Glutamic acid, Glu

EM Electron microscopy

Endo H Endonuclease H

ER Endoplasmic reticulum

F Fusion protein

F Phenylalanine, Phe

F0 Precursor form of the fusion protein

F1 Larger subunit of the fusion protein formed after furin

cleavage

F2 Smaller subunit of the fusion protein formed after furin

cleavage

FBS Fetal bovine serum fcs Furin cleavage site

FDA Food and Drug Administration

Fig Figure

FI-RSV Formalin inactivated-RSV

FP Fusion peptide

xv g Gravity (unit of force)

G Respiratory syncytial virus attachment protein

GAG Glycosaminoglycan

GE Gene end

GFP Green fluorescent protein gp Glycoprotein

GS Gene start h Hour

HA Hemaglutinin

HB Helix bundle

HAE Human airway epithelium

HBSS Hank's Buffered Salt Solution

HEK Human kidney epithelial

His Histidine

HIV Human immunodeficiency virus

HMPV Human metapneumovirus

HRA Heptad repeat A

HRB Heptad repeat B

HRP Horseradish peroxide

HS Heparan sulfate xvi

I Isoleucine, Ile

ICAM Inter-cellular adhesion molecule

IFN Interferon

IG Intergenic region

IL Interleukin

IKKε IкB kinase

IP Immunoprecipitation

IRF3 Interferon regulatory factor 3

JAK Janus kinase

K Lysine, Lys kDa Kilodalton kV Kilovolt

L Large polymerase protein

L Leucine, Leu

Le Leader sequence at the 3‟ end of the paramyxovirus

genome

M Respiratory syncytial virus matrix protein

M2-1 Respiratory syncytial virus protein

M2-2 Respiratory syncytial virus protein

MAb Monoclonal antibody

xvii

Met Amino acid methionine

Mg2+ Magnesium min Minute ml Milliliter mm Millimeter mM Millimolar mRNA Messenger RNA

MV Measles virus

N Nucleoprotein

N Asparagine, Asn

NaCl Sodium Chloride

NDV Newcastle disease virus

NF-kB Nuclear Factor-Kappa B

Ni2+ Nickel nm Nanometer

NS1 Respiratory syncytial virus nonstructural protein 1

NS2 Respiratory syncytial virus nonstructural protein 2 nt Nucleotide

N terminus Amino terminus

ORF Open reading frame xviii

P Respiratory syncytial virus phosphoprotein

PBS Phosphate buffered saline

PC Phosphatidylcholine

PCR Polymerase chain reaction pep 27 Peptide fragment excised by the furin cleavage of the

RSV F protein

PFP Purified F protein pH Hydrogen ion concentration

PIV3 Parainfluenza virus type 3

PIV5 Parainfluenza virus type 5

PLL Poly-L-Lysine

PO4 Phosphate

Poly (A) Polyadenylate

POPC 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine

POPE 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphoethanolamine

PVM Pneumovirus of mice

R Arginine, Arg

RIG-I Retinoic acid inducible gene rgRSV Recombinant green fluorescent protein expressing RSV

RNA Ribonucleic acid

xix

RPM Revolutions per minute

RSV Respiratory syncytial virus

S Serine, Ser

SDS-PAGE Sodium dodecyl sulfate-polyacrylamide gel

electrophoresis

SeV Sendai virus sF Soluble F protein sG Soluble G protein

SH Respiratory syncytial virus small hydrophobic protein siRNA Small interfering RNA

SNAREs Soluble N-ethylmaleimide-sensitive factor attachment

proteins

STAT Signal Transducer and Activator of Transcription

SV5 Simian virus 5 (also known as PIV5)

T Tryptophan, Trp

T7 T7 bacteriophage

T7 polymerase Bacteriophage RNA polymerase

TCA Trichloroacitic acid

TEV protease Tobacco etch virus protease

TM Transmembrane

TNF Tumor necrosis factor

Tr Trailer sequence at the 5‟ end of the paramyxovirus

genome

TrC Trailer complement sequence at the 3‟ end of the

paramyxovirus antigenome

WHO World health organization wt Wild type

V Valine, Val

Vero Immortalized cell line derived from African green monkey

kidney cell vol Volume

Y Tyrosine, Tyr

α Alpha

β Beta

λ Lambda

µCi Microcurie (unit of radioactivity)

µl Microliter

µg Microgram

µm Micrometer

µM Micromolar

xxi

CHAPTER 1: RESPIRATORY SYNCYTIAL VIRUS

INTRODUCTION

Respiratory syncytial virus (RSV) was originally isolated in 1956 from a chimpanzee with cold-like symptoms and named Chimpanzee Coryza Agent

(143). It was also suspected to be a human pathogen, and is now recognized as a major pathogen in infants and young children, worldwide. RSV infection causes severe lower respiratory tract disease, bronchiolitis and pneumonia, in

1%-5% of this population (188) resulting in 90,000 emergency hospitalizations of infants in the U.S. and 160,000 deaths worldwide each year (43, 81, 109).

Although it is primarily known as pediatric pathogen, it is capable of repeatedly infecting and causing disease in individuals of all ages. For elderly and immunocompromised patients, RSV disease is also a significant cause of mortality, second to influenza virus in non-epidemic years (37, 62). In developed countries, a humanized monoclonal antibody (MAb) against the fusion (F) glycoprotein, Synagis, is used to protect premature infants who are at highest risk for severe RSV disease (163). Despite extensive efforts, a vaccine is not yet available.

RSV infection primarily begins in the epithelial cells of the upper respiratory tract but can spread to the lower respiratory tract where it often causes

1 severe respiratory disease. HRSV infection is restricted to the cells of respiratory tract in vivo. The virus obtained its name from the fact that expression of the RSV F protein on the surface of an infected cell in culture causes the cell membrane to merge with the cell membrane of adjacent cells, forming a syncytia which is an oversized multinucleated cell.

TAXONOMY AND CLASSIFICATION

RSV is a negative strand RNA virus, a member of the Mononegavirales, in the Paramyxoviridae family (37). Paramyxoviridae contains two subfamilies,

Paramyxovirinae and Pneumovirinae that are distinguishable by the degree of similarity between amino acid sequences, number of protein encoding mRNAs, the type of accessory proteins, and differences in the attachment protein properties.

The Pneumovirinae subfamily is further subdivided into two genera,

Pneumovirus which includes RSV (human, bovine and ovine) and pneumovirus of mice (PVM), and Metapneumovirus (avian and human).

According to an analysis of antigenic differences (2), RSV is a single serotype with 2 subgroups, A and B. The two subgroups differ in their hypervariable G protein while the F protein is antigenically similar (37).

RSV ORGANIZATION AND MORPHOLOGY

The single strand negative-sense RNA genome of RSV is composed of a helical nucleocapsid packaged in a lipid-bilayer membrane derived from a host cell membrane during viral budding. Viral particles are pleomorphic and appear as a spherical or filamentous shape ranging from 100 – 350 nm in length (175). The envelope contains three viral transmembrane glycoproteins, the attachment glycoprotein (G), the fusion (F) protein, and the small hydrophobic (SH) protein. The G and F proteins project as spikes from the viral particle membrane. Underneath the envelope lies the matrix (M) protein which is mainly involved in viral particle formation and budding.

The RSV genome is 15,222 nucleotides (nt) long encoding 11 proteins from the 10 open reading frames (ORFs). The RSV nucleocapsid is comprised of the RNA genome encapsidated by the nucleocapsid N protein in a helical form, and has three associated viral proteins, the phosphoprotein (P), the large polymerase subunit (L), and the antitermination factor, M2-1. These nucleocapsid-associated proteins form a polymerase complex and function in viral mRNA transcription and genome replication, using the nucleocapsid rather than free genomic RNA, as template.

THE RSV LIFE CYCLE

Attachment and Entry

RSV, like all enveloped viruses, must attach to a target cell and fuse its membrane with a cell membrane to deliver the viral genome to the cytoplasm and initiate infection. To accomplish these two functions, all members of the

Paramyxoviridae family express two glycoproteins, one to attach to the target cell and one to fuse the virion membrane with the host cell membrane. The

RSV G and F proteins, respectively, are responsible for these functions.

To gain entry, RSV must use the G protein to bind to a cellular receptor, glycosaminoglycans (GAGs), mostly heparan sulfate (HS) and possibly condroitin sulfate B, in immortalized cells in culture (89). In cells lacking HS due to mutation, RSV is 10-fold less infectious. However, in vivo infection may be different, as its target organ, human airway epithelial cells, do not express

HS on their apical surface, the entry point for RSV (224), suggesting that RSV uses a different receptor for its natural infection. Other cell surface molecules, annexin II, L-selectin, and the fractilkine receptor, have also been investigated as potential G protein receptors (133, 204).

To penetrate a target cell, the RSV F protein must cause the virus and the cell membranes to merge. The RSV F protein, alone, expressed transiently in cultured cells causes cells to fuse. RSV expressing F as its only glycoprotein is infectious and causes plaques to form by cell-cell fusion (7, 103, 196). This is a unique characteristic of the pneumovirus F protein; the F protein of members of the Paramyxovirinae requires the attachment protein to function

4 in fusion. RSV syncytia caused by virus expressing the F protein but no G protein are smaller than those produced by RSV that is also expressing the G protein (196), suggesting that while the F protein is the only glycoprotein necessary to mediate the viral entry step, the G protein enhances entry by binding a virion or an F protein-expressing cell to a target cell more tightly without playing a direct role in membrane fusion.

Virions expressing the RSV F protein as its only glycoprotein are infectious and appear to bind equally to either HS or another unidentified molecule(s) on the cell surface (197). Several peptides derived from the RSV F protein have been shown to bind to HS (47). RhoA and ICAM-1 have been shown to interact with the F protein and are potential receptors for the RSV F protein

(10, 85, 152). Taken together, the RSV F protein has the ability to attach to a target cell surface and to trigger, causing membrane fusion, without the aid of an attachment protein.

Even though the RSV G protein appears to be dispensible for viral entry in vitro, it is important in vivo. In the mouse and primate model, RSV lacking the

G protein is attenuated resulting in a significant reduction in viral titer (48,

103, 200).

Viral entry in primary well differentiated human airway epithelial (HAE) cultures, and therefore likely in vivo, occurs exclusively at the apical surface of ciliated cells (224). In that study, recombinant green fluorescent protein- expressing RSV (rgRSV) infected cultures only after they had been allowed to differentiate at the air-liquid interface for several weeks, developing ciliated

5 cells. RSV exclusively infected the ciliated columnar cells when added to the apical surface of the cultures (corresponding to intraluminal in vivo), not when added basolaterally.

Transcription, translation, and replication

Upon membrane fusion, the nucleocapsid is released into the cytoplasm and becomes the template for mRNA transcription and genome replication. The

RSV replication cycle is cytoplasmic and appears to be independent of the cell nucleus as the virus can grow in enucleated cells and in the presence of actinomycin D (71, 150). RSV encodes for 10 major subgenomic mRNAs.

Each contains a single ORF except for the M2 gene that has two ORFs. Like other viruses in the Paramyxoviridae family, RSV transcription initiates at only one position, the 3‟ promoter and occurs in a start-stop fashion. The genes located proximal to the promoter are more efficiently transcribed than those downstream because a proportion of the polymerase molecules fall off the template at each intergenic (IG) region, leading to a polar gradient of gene expression (34, 42, 90, 116, 118). The genes are terminated by a gene-end

(GE) signal, followed by an IG region of 1-52 nucleotides, and a gene-start

(GS) signal. These signaling elements direct the polymerase to initiate and terminate mRNA transcription. The 5‟ terminus of the mRNA is capped and the cap is methylated by the viral polymerase. The 3‟ terminus is polyadenylated by a repeated copying of the poly U tract of the GE signal before the mRNA is released (117, 119).

As the viral M2-2 protein accumulates, it reduces mRNA transcription, allowing the polymerase to switch to genome replication. Propagation of the viral genome occurs via RNA antigenome synthesis starting at the 3‟ terminus of the genome template. The leader (Le) sequence at the 3‟ genome end and complement of the 5‟ end trailer (Tr) sequence are necessary for polymerase binding and replication by serving as a replication promoter, thus they both are copied and included in the antigenome and genome molecules. By using the negative sense genomic RNA wrapped up in the N protein in a helical nucleocapsid as the template, the polymerase complex comprising of the L,

P, and N proteins ignores all transcription signals, synthesizing the antigenome, a complete positive sense complementary copy of the viral genome. This antigenome is used as the template to generate multiple copies of the negative sense viral RNA that will be encapsidated and assembled into new virion progeny (46). Both the genome and antigenome are tightly encapsidated with N protein as they are synthesized and neither is capped or polyadenylated (37).

To control the balance between transcription and genome replication, RSV employs the M2-2 protein. As the level of M2-2 protein accumulates, it inhibits transcription and favors RNA replication (11). The cellular cytoskeletal elements actin and profilin enhance in vitro transcription of the RSV genome suggesting that they are involved in activating RSV transcription in vivo (24,

25).

Assembly and Budding

The M protein is thought to orchestrate assembly and budding of the virion.

However, incorporation of the viral glycoproteins into the lipid raft may be the nucleation event that initiates viral assembly. When expressed alone, the

RSV F protein localizes to lipid rafts suggesting that it might be responsible for this initial step (70). Lipid rafts are microdomains in the plasma membrane that is enriched in sphingolipids and chloresterol. Assembly occurs on these rafts as the viral components gather there (18, 79, 136, 174). The M protein binds to the cytoplasmic tails of one or both of the viral glycoproteins (C- terminus of F and N-terminus of G). The M protein also binds to the nucleocapsid with its associated polymerase proteins and budding is initiated.

The forces that drive budding are not clear, nor is the mechanism by which the virion membrane pinches off. The site where the M protein is localized and organizes this assembly and budding characteristically appears to have multiple cytoplasmic inclusions (79).

In polarized cells, RSV budding takes place exclusively at the apical surface.

The host proteins controlling apical recycling endosome (ARE) pathway have been shown to regulate apical transport of virions, thus assisting directional budding from the polarized epithelial cells (17). In addition, cellular actin in the microfilament form, and microtubules elements promote viral assembly and release (24, 102).

THE RSV PROTEINS

The Nucleocapsid and Polymerase Complex

There are four proteins associated with the nucleocapsid inside the virion.

These proteins mediate replication and transcription of the RSV genome: the nucleocapsid protein (N) the phosphoprotein (P), the large polymerase subunit (L), and the anti-termination factor M2-1. The N protein is an RNA binding protein comprised of 391 amino acids. It encapsidates the genome in a helical nucleocapsid structure and remains associated with RNA genome during genome transcription and replication process as a required component of the template. The N protein binds to the entire length of the RSV genome and antigenome, protecting them from RNAses. Prior to binding to the genomic RNA, the N protein is maintained in a soluble state by the interaction of its C-terminus with the C-terminus of the P protein which functions as a chaperone (6, 68, 74, 189). However, the essential residues for its nucleocapsid assembly activity are located at the N-terminus of the N protein, residues 1-91 (146).

The P protein, 241 amino acids in length, is also responsible for the formation of stable polymerase complexes. Phosphorylation of the P protein which is required for the protein to function as a polymerase cofactor mainly occurs at the serine 232 (181). The C terminus of the P protein not only binds to the N protein, but also to the L protein attaching it to the nucleocapsid.

The L protein, 2165 amino acids in length, carries all of the enzymatic activities required for propagation of the viral genome including genome

9 replication, mRNA transcription, mRNA capping and cap methylation, and polyadenylation (158). Its polymerase activity was suggested by the high degree conservation of 6 amino acid motifs with other paramyxovirus polymerases. Its mRNA capping activity is in the central domain (domain II) of the molecule (130) and its polymeration and methylation catalytic sites are located in domains III and VI, respectively (159, 192).

The M2-1 protein is translated from the first ORF of the M2 gene. It is comprised of 194 amino acids and functions as the anti-termination factor of the polymerase during mRNA transcription. It keeps the whole enzyme complex associated with the nucleocapsid during viral transcription. It also appears to form a complex with the P protein (137).

The M2-2 protein is produced from the second ORF of the M2 gene by ribosomal termination-reinitiation, it contains 84-90 amino acids as a result of having three alternative AUG translation start sites. M2-2 functions as a negative regulator of mRNA transcription, favoring replication of viral genome

(11).

The Nonstructural Proteins NS1 and NS2

The NS1 and NS2 proteins, containing 139 and 124 amino acids, respectively, form homooligomers (61). Even though these two proteins have been found only in small amounts in the virion, they appear to be the two most abundant RNAs in RSV-infected cells, as expected from their promoter- proximal location (42, 61). 10

All paramyxoviruses encode accessory proteins that counteract the IFN response, the most important defensive mechanism of the innate immune response to viral infection. These proteins either inhibit IFN generation or block IFN signaling (84, 94). For the RSV, the NS1 and NS2 proteins are responsible for the inhibition of the IFN response induction (16, 190). They impede IFN induction by inhibiting the phosphorylation and nuclear localization of IFN regulatory factor (IRF3) (16, 191). This interference that greatly reduces the level of the kinase enzyme (IKKε) is primarily mediated by the NS1 protein, with the greatest effect achieved in conjunction with NS2.

The NS2 protein works primarily by targeting the STAT2, signal transducer and transcription activator of a JAK/STAT signaling protein, for proteosome- mediated degradation (167). STAT2 participates in ISGF-3 complex formation to initiate IFN gene transcription. Its degradation results in reduced expression of IFN-responsive genes, and antiviral immunity. Also, the NS2 protein interferes with RIG-I (retinoic acid inducible gene), a cytosolic receptor that recognizes intracellular viral dsRNA, leading to a suppressed IFN-beta production (129).

Overexpression of the NS1 inhibits RSV minigenome transcription and replication (4). However, removal of the NS1 and NS2 genes from the virus attenuates virus production. Although some of this effect undoubtedly comes from the loss of the ability of the virus to inhibit the IFN system, virus replication is also partially inhibited in the Vero cell line (98, 198, 199). Vero cells lack the IFN genes, so reduced replication in Vero cells indicates that

11 the NS1 and NS2 proteins enhance viral replication. However, they are not essential.

The Matrix (M) Protein

The M protein is a nonglycosylated protein containing 256 amino acids. The

C-terminal half of the protein contains a hydrophobic domain that might mediate membrane interaction. The protein forms a layer beneath the envelope of the virion. During budding, it accumulates at the plasma membrane of the virus-infected cells and mediates virion assembly (91).

Early in the course of infection, at least some of the M protein is localized to the nucleus (77). Later it is exported by the cellular Crm-1 dependent pathway to the cytoplasm where it performs its virion assembly function (78). It interacts with the G and F proteins (18, 70) in the plasma membrane and with the viral nucleocapsid to organize viral morphogenesis (79). Association with the nucleocapsid appears to serve two functions: (1) to facilitate assembly of the virion (79, 136, 155); and (2) to inhibit the transcriptase activity of the nucleocapsid before packaging (79).

The Attachment (G) Glycoprotein

The RSV G protein is 282 to 319 amino acids in length, depending on the strain of virus. It is a type II integral membrane protein with its signal sequence-containing N-terminus anchored in the membrane and its C-

12 terminus exposed to the extracellular space. The G protein of RSV shares similarities with other members of the Pneumovirinae subfamily but no sequence or structural homology with the attachment proteins of the viruses in Paramyxovirinae subfamily. The G protein is 36 kDa, but 4 or 5 N-linked glycan chains are added cotranslationally, and during passage through the

Golgi, a large number of O-glycans are added, resulting in a glycoprotein of approximately, 90 kDa. However the degree of O-linked glycosylation on the

G protein is partially dependent on the cell type used to produce the virus

(75). In cultured immortalized cells, the G protein mediates attachment of the virus to a host cell by binding to glycosaminoglycans (GAGs), mainly those containing iduronic acid, such as heparan sulfate (HS) (47, 69, 89). However, in a natural infection, the receptor is likely to be different. HAE cultures do not express HS on their apical surface as shown by immunostaining (223), even though RSV enters by the apical domain (224) strongly suggesting that RSV uses a different receptor for infection in vivo.

The RSV G protein has also been reported to share homology with the CX3C chemokine, fractalkine, and to bind to the fractalkine receptor on leukocytes

(204). Fractalkine and its receptor play a role in chemotaxis, therefore the

RSV G protein may affect this process, but the mechanism is not yet clear.

Some of the G protein is also synthesized and secreted in a soluble form (sG) which is the result of translational initiation at the second start codon (AUG) instead of the first, followed by proteolytic cleavage once the sG reaches the cell surface (176). The sG protein may act as a decoy, binding up antibody to

13 help the virus escape neutralization (22). In one study, human lung epithelial cells, A549, infected by RSV lacking the sG protein displayed increased IL-8 and RANTES mRNA and secreted protein compared to the same cells infected with RSV-wt (3). As a result, the authors suggested that the sG protein might have an inhibitory effect on NF-kB activation and on the production of the proinflammatory response mediators.

As a result of differences in protein modification like glycosylation or cleavage, the predominant form of G protein incorporated into virions may differ, depending on the cell line that produces them. This variation contributes to differences in viral infection efficiency (75, 121). Cleavage of the G protein in Vero cells results in the loss of its C-terminus which greatly reduces its infectivity for primary HAE cells (121).

Despite the fact that most members of the Paramyxoviridae family require the function of their attachment protein to initiate virus infection and membrane fusion activity, RSV is unique in that its F protein, alone, can mediate membrane fusion and support viral replication and spread in immortalized culture cells (7, 103, 196). These results indicate that the G protein is not essential for viral infection of immortalized cells or for syncytia formation in these cells. However, RSV syncytia caused by virus expressing F as its only glycoprotein are smaller than those produced by RSV that is also expressing the G protein (196). While the F protein is the only glycoprotein required to mediate the viral entry step, these results suggest that the G protein enhances entry probably by binding to a target cell more tightly, without

14 playing a direct role in membrane fusion. In addition, RSV lacking the G protein is attenuated in the mouse model with a significant reduction in virus production (49, 105, 200), indicating a more essential role for the G protein in vivo.

The Small Hydrophobic (SH) Protein

The SH protein is the smallest anchored protein in the viral envelope at 64 amino acids. Like the G protein, it is a type II transmembrane glycoprotein with its short C-terminus projecting extracellularly and its N-terminus in the cytoplasm (40). In RSV infected cells, the SH protein is present in 4 different forms, SH0, SHG, SHP, and SHT, with SH0 and SHP being the most abundant forms on the virion surface. Due to differences in translation initiation and post-translational modification, these species vary in size and carbohydrate modification: SH0 is a full-length nonglycosylated species, SHG posses a single N-linked sugar chain, SHP has an N-linked sugar chain with a polylactosaminoglycan addition to the sugar chain, and SHT is a truncated, nonglycosylated form derived from translation that initiates at the second

AUG (1, 148). However the significance of these multiple forms is currently unknown.

The function of the SH protein is not yet clear since it appears to be dispensable for viral survival. Deletion of the SH gene from the virus did not attenuate infection of immortalized cells or the ability to cause syncytial.

Deletion had no effect on replication in the mouse lower respiratory tract but

15 caused a 10-fold reduction in the upper respiratory tract. And only a slight attenuating effect in chimpanzees (21, 214), The SH protein may play a role in impeding cellular apoptosis by inhibiting the TNF pathway, similar to the role played by the SH protein of parainfluenzavirus 5 (PIV5) because recombinant PIV5 lacking its SH gene is sensitive to apoptosis but the addition of the RSV SH gene again made the virus resistant to apoptosis (72).

Molecular modeling of the SH protein suggests that it might form an ion channel (113). When expressed in bacteria it incorporates into the membrane and increases cell permeability to small molecules (156). Whether the channel activity is related to its anti-apoptotic activity has yet to be determined.

The Fusion (F) Protein

The F protein is a type I membrane glycoprotein with its N-terminus being extracellular. It is 574 amino acids long and its N terminus contains a cleavable signal sequence that targets synthesis to the ER membrane. The F protein is responsible for the membrane fusion in two scenarios: (1) between viral and cell membranes to initiate viral infection; and (2) between the plasma membrane of a cell expressing the F protein and a neighboring cell. The result is a syncytium or multinucleated giant cell. Because of its importance in the viral replication cycle, its conserved structure, its exposed position in the virion, and its strong immunogenicity, the RSV F protein has been a major target for vaccine and antiviral drug development (43, 99, 226).

In all paramyxoviruses, the F protein shares sequence and structural homology including many disulfide bond positions. Like many other class I viral fusion proteins, the RSV F protein is both a trimer and is activated by cleavage. Each monomer is produced as a precursor, F0, in the ER where it is also modified with 5 N-linked glycans and assembled as a trimer. During passage through the trans Golgi network the glycans are matured, trimmed back and rebuilt, and the protein is cleaved by a cellular furin-like protease

(Fig. 1.1). Unique to RSV, this enzyme cleaves the F protein in two places, following a multibasic furin cleavage site (fcs) 1 and 2 (KKRKRR and RARR, respectively), releasing a 27 amino acid (AA) peptide, pep27, with two attached glycans (83, 228, 229). The bovine RSV (BRSV) pep27 includes the

FYGLM, five signature AAs of tachykinin and possesses tachykinin activity.

This activity may play a role in inflammation induction and smooth muscle contraction which may contribute to BRSV pathogenesis (230). However, the

HRSV pep27 does not have this activity.

Fig. 1.1. Cartoon of the RSV F protein domains and cleavage sites. The wild-type F0 precursor is cleaved intracellularly by a furin-like protease at two sites, releasing a 27 amino acid peptide (pep27). The resulting F protein is composed of the N-terminal F2 protein linked by one, and probably two, disulfide bonds to the F1 transmembrane protein. The highly hydrophobic fusion peptide resides at the cleavage-created N-terminus of the F1 protein.

This cleaved form of the RSV F protein is fully active and able to cause cell- cell fusion once it reaches the plasma membrane or to cause virion-cell fusion. The two heptad repeats, HRA and HRB, interact to complete the conformational changes that result in the final posttriggered F protein 6HB.

The mature F protein is composed of the N-terminal F2 protein linked by two disulfide bonds to the transmembrane (TM) F1 protein (51). One of these disulfide bonds is conserved among all paramyxoviruses but the second is unique to the pneumoviruses. The cleavage-created N-terminus of the F1

18 protein contains the highly hydrophobic fusion peptide (FP), which inserts into a target cell membrane to initiate fusion (37). This cleaved form of the RSV F protein is in a native, fusion competent state, able to cause cell-cell fusion once it reaches the plasma membrane, or to be incorporated into virions where it is able to cause virion-cell fusion.

On the surface of virions or virus-infected cells, the F protein exists in a highly metastable form, like a compressed spring waiting to be triggered. Upon triggering, it undergoes a dramatic and irreversible conformational change that pulls the two opposing membranes together to initiate membrane fusion.

Despite extensive research, what triggers the RSV F protein to function remains unclear. Previous work by Jose Melero‟s group proposed that cleavage is the trigger, based on studies of an anchorless version of the RSV

F protein generated by removal of the transmembrane and cytoplasmic domains from the F protein (9, 83), The resulting soluble F (sF) protein appeared by electron microscopy (EM) as hairpins, with a long slender body and a small head. Some of these were individual molecules but many were in rosette clusters, aggregated at their narrow ends. They separated the individual hairpins from the rosettes by sucrose gradient centrifugation. The individual, least aggregated sF molecules remained near the top of the gradient while the rosette aggregates migrated further into the gradient due to their increased size. The sF proteins in the rosette aggregates were fully cleaved and represented the posttriggered sF, with its highly hydrophobic FP exposed, which would aggregate with other hydrophobic FPs to avoid exposure to water. But the unaggregated sF proteins were only partially 19 cleaved. Addition of trypsin caused this population to form rosettes. Melero speculated that the final proteolytic cleavage in a trimer causes it to trigger and that this cleavage might occur as the virion enters its target cell.

However the crystal structures of two paramyxovirus sF proteins, from

Newcastle disease virus (32) and parainfluenza virus 3 (PIV3) (219) were both found to be in the posttriggered form. Both had been mutated to inactivate their cleavage sites to prevent triggering, as suggested by Melero‟s work. These results demonstrated that cleavage was not required for triggering. These were two separate events. Our recent study on the sF protein of RSV described in Chapter 2 confirmed the idea that the cleavage and triggering events were separable and did not have a causal relationship.

We showed that the RSV sF protein can be produced in its pretriggered conformation despite being fully cleaved (29). Clearly the Melero theory that cleavage is the F protein triggering event is not correct. Furthermore, it would not be logical for the virus to have the majority of its F protein triggered and therefore inactive before it reaches the target cell membrane.

The efficiency of proteolytic cleavage of an F precursor molecule plays an important role in viral tropism and pathogenesis in some paramyxoviruses, most notably Newcastle disease virus (NDV) (82, 203). Sequence analysis of the cleavage site of virulent and a virulent isolates of NDV revealed a strong correlation between virulence and the number of basic amino acids at the cleavage site. Virulent NDV strains contain a furin cleavage site (R-X-[K/R]-

R↓) with multiple basic residues located at the 1st. 3rd, and the 4th position. In

20 the virulent viruses, the furin cleavage site at the C-terminus of F2 enables cleavage by widely expressed furin-like, intracellular proteases. Once cleaved, the F protein enables NDV to spread to various tissues, conferring high pathogenicity. However, that is not the case for the RSV F protein: all strains produce F protein with two furin cleavage sites, enabling intracellular cleavage that produces a fusion active F molecule during transport through the Golgi. However, RSV infection is generally limited to the respiratory tract.

Apparently the host adaptive immune system is capable of controlling the spread of this relatively slowly replicating virus to prevent spread to other organs.

The F protein cleavage site of the avirulent NDV strains contains a single basic residue. These F proteins are not cleaved intracellularly. In vivo, once it reaches the cell surface, it is exposed to an extracellular protease produced by Clara cells that cleaves it to activate its fusion potential. In immortalized cell culture this protease is not produced, the F protein is not cleaved, there is no cell-cell fusion unless trypsin is added. Virus particles are produced by these avirulent viruses, but they are inactive until they are treated with trypsin to cleave at this site.

The pneumovirus F protein is able to induce membrane fusion without the aid of the viral attachment protein, but members of the other subfamily,

Paramyxovirinae, require their homologous attachment glycoprotein in addition to the F glycoprotein to initiate fusion between the virus membrane and the target cell membrane. In these viruses, the attachment protein not only binds to the target cell receptor, but also triggers the F protein, resulting 21 in fusion (52, 57, 95, 195). It was recently proposed that the second F protein cleavage, which is unique to pneumoviruses, may enable membrane fusion in the absence of the viral attachment protein. This proposal has come from studies of recombinant Sendai virus (rSeV). Like most paramyxoviruses, fusion mediated by SeV F protein requires coexpression of its homologous attachment, HN, protein. Interestingly, The rSeV expressing F protein containing both cleavage site(s) of RSV F protein exhibited an increased fusogenicity with less dependence on its attachment partner and receptor binding activity compared to the wild-type rSeV (170, 171). The authors‟ interpretation was that removal of a peptide separating F1 and F2 enabled the

Sendai virus F protein to cause fusion without the help of its attachment protein. Because this peptide removal appears similar to the removal of pep27 from the RSV F protein and since the RSV F protein is capable of causing membrane fusion without its attachment protein, it is possible that removal of an intervening peptide may be responsible. However, it is also possible that the artificial insertion of this foreign peptide and/or its removal destabilizes the SeV F protein, causing a higher rate of spontaneous triggering and fusion. These modified SeV F proteins have recently been incorporated into recombinant SeV and displayed a reduced dependency on sialic acid receptor recognition conferred by its homotypic HN attachment protein. Following the infection of the CHO cell line that is defective in sialic acid expression or another cells line that was pretreated with neuraminidase, this virus appeared to be more infectious and fusogenic compared to the rSeV containing the wt F protein (170). 22

The F protein contains two important heptad repeat (HR) α-helices, HRA following the N-terminal fusion peptide, and HRB preceding the C-terminal transmembrane domain. For all paramyxovirus F proteins, these regions are critical in the fusion reaction as mutations result in a fusion-deficient F protein

(20, 80, 173, 185). HRA and HRB helices both participate in an irreversible structural change that is strongly coupled to a membrane fusion process. As a result of protein refolding, HRA and HRB interact to form a 6-helix bundle

(6HB) structure (28, 58) which is extremely stable to heat (5). Interestingly, eukaryotic intracellular fusion processes in the secretory or endocytic pathways are mediated by vesicle (v) - and target (t) -SNARE proteins. They result in the formation of a 4 helix coiled coil bundle, where 3 helices are contributed by t-SNAREs and one helix is contributed by v-SNARE. This 4 helical core complex is the hallmark of a cellular membrane merging event

(161, 193).

Despite the diversity in structures of many viral fusion proteins, all have a

6HB trimer-of-hairpins as their final structure that creates the fusion pore. The fold-back step that brings HRA and HRB together to create the 6HB trimer also brings the two membrane inserted domains together: the HRA-adjacent

FP embedded in the target membrane and the HRB-adjacent transmembrane domain. It is thought that accumulation of 5 to 8 F proteins that trigger and form 6HBs in a circle is needed to cause a fusion pore to form. As a result, the two opposing membranes are brought into intimate contact with their bilayer structure distorted. Lipid mixing, first of the outer leaflets (hemifusion) followed by the inner leaflets, leads to the opening of a fusion pore (213). 23

This final core trimer is a feature common to all viral fusion proteins. In the class I fusion proteins of the paramyxoviruses, retroviruses and orthomyxoviruses, core trimer is mainly α-helical but in class II fusion proteins it is mainly a β–sheet structure, and in class III fusion proteins, there is a combination of both structures. Previous studies with class I viral fusion proteins including the HA of influenza virus (23), the TM of Moloney murine leukemia virus (67), the gp41 of HIV-1 (30, 211), the GP2 of Ebola (132, 210), and the F protein of PIV5 (5), revealed that when protein folding is complete,

HRA and HRB form a 6HB in which the central coiled coil trimer is formed by the N-terminal HRA, with the three C-terminal HRB packed into the grooves of the HRA trimer in an antiparallel orientation. The RSV 6HB structure, solved by NMR and by X-ray diffraction of the crystallized complex of the

HRA and HRB peptides, displays a similar structure (126, 138, 226). Although the structure of the complete RSV F protein has not yet been solved, its 6HB is very similar to other members of class I fusion protein, so it clearly functions similarly in bringing the two membranes together to initiate membrane fusion (58, 140, 178).

The model of 6HB-dependent viral membrane fusion is also supported in part by the fact that the HR-derived peptides have been shown to have potent inhibitory effects on fusion caused by type I fusion proteins, including RSV

(123, 208). The HRB peptides are the most potent probably because they competitive binding to the HRA trimer groove, preventing the completion of

6HB formation and thus fusion. In the case of HIV, the FDA approved C- terminal peptide inhibitor, T20 (Enfuvirtide), interacts with the trimer HR1 24 region (equivalent to HRA in the RSV F protein, i.e. FP proximal) of the HIV gp41 fusion protein during an intermediate stage of conformational change. It forms a dead-end T20 peptide-gp41 hybrid 6HB structure, thus aborting HIV infection (108, 216). With the extreme requirement for 6HB formation to achieve membrane fusion during viral entry, Peptides targeting the formation of this crucial 6HB have also been used to study the fusion process of HIV-1,

SV5 (PIV5), and avian lymphoma and sarcoma virus (ALSV; Rous sarcoma virus) (31, 59, 97, 147, 178, 215).

The complete X-ray crystal structures of two paramyxovirus F proteins, both in the soluble (sF) form have been solved. One is in the pretriggered form and one in the posttriggered conformation. As mentioned above, both had been mutated to destroy the furin cleavage site since cleavage was thought to be the cause of triggering. The first to be solved, the parainfluenza virus type 3

(PIV3) sF protein was unexpectedly in the posttriggered form (219). In an attempt to stabilize the pretriggered form, the C-terminus of the PIV5 sF protein was fused to a trimerization domain (220). The PIV5 sF protein was in a very different, apparently pretriggered form, suggesting that stabilization of the C-terminus of the sF protein is important for maintaining its metastable form.

In collaboration with Dr. Will Ray of The Research Institute at Nationwide

Children‟s Hospital, we have developed computer models of the RSV F protein in the pretriggered and posttriggered forms based on the crystal structures of the PIV5 and PIV3 F proteins and sequence alignment. Details

25 of these models and predictions of residues critical for the fusion function of the RSV F protein, ideas that we derived from these models, will be discussed later (Chapter 3).

THE MECHANISM OF F PROTEIN-MEDIATED MEMBRANE FUSION

The F protein is known to drive membrane fusion by using irreversible protein refolding to bring two membranes into close contact (96, 122). The sequence of refolding events is graphically illustrated in Fig. 1. Initially, the paramyxovirus F protein is in a metastable, pretriggered trimer form (226).

Three short α-helices near the top of the trimer, “pre-HRA”, are likely in a spring-loaded state (178), awaiting a triggering event (Fig. 1.2A). Upon triggering, by an as yet unidentified mechanism, the F protein undergoes a dramatic and irreversible conformational change, converting this pre-HRA into the long HRA α-helix, thereby projecting the highly hydrophobic fusion peptide at the N-terminus of F1 into the target cell membrane (28). The three HRAs likely trimerize at this point, because this peptide in solution trimerizes spontaneously (126). This extended structural state (Fig. 1.2B) is called the

“pre-hairpin intermediate”. The three F1 monomers then fold back on themselves (Fig. 1.2C), juxtaposing the HRA trimer, with its adjacent FP inserted in the target cell membrane, with the HRB monomers, adjacent to the transmembrane domain of F1, and allowing the HRBs to insert into the grooves of the HRA trimer to form the highly stable 6HB (28, 58) (Fig. 1.2D).

This large-scale structural transition not only brings HRA and HRB together,

26 but also brings the two membrane-inserted domains together, which in turn brings the two membranes together to initiate fusion (58).

Fig. 1.2. Cartoon of F protein refolding mediated viral-cell membranes fusion. The three N-terminal α-helices in pre-HRA are connected by two non- helical peptides in the pretriggered form. Upon triggering these non-helical peptides refold into α-helices, completing the long HRA α-helix (dark gray) and in the process thrusting the F1 N-terminal fusion peptide into the target cell membrane. After HRA trimerizes, the molecule folds in half, and the HRB

α-helices (light gray) insert into the grooves on the surface of the HRA trimer to complete formation of the 6-helix bundle. As a result, the virion and cell membranes are brought together and initiate membrane fusion.

At this final state, the F protein is at its most energetically stable conformation and membrane merger is thought to be driven by the large energy released from 6HB formation (35, 140, 178). As a result, the two membranes begin to merge. First, the two opposing outer leaflets mix their lipid contents in a

“hemifusion” event, followed by merger of the inner leaflets that results in the opening of a fusion pore. Accumulation of additional fusion proteins that trigger and form 6HBs in the same area causes the fusion pore to enlarge.

The channel eventually becomes wide enough that the viral nucleocapsid can pass through it and into the cytoplasm (213).

F protein triggering is the step in this refolding process that initiates membrane fusion. But, it is not clear what triggers the RSV F protein.

Triggering of other class I viral fusion proteins is induced by a specific mechanism for each proteins. For example, influenza virus enters the cell by endocytosis and exposure of its hemagglutinin (HA) glycoprotein to low pH in the late endosome causes triggering (23). The triggering mechanism of the

Ebola GP protein is apparently unique in that it is cleaved during entry by cathepsin B and L before it can be triggered by low pH to cause membrane fusion (183). In the case of human immunodeficiency virus (HIV) gp41 fusion subunit, sequential binding of gp120, the receptor binding subunit, with two cellular receptors, CD4 and CXCR4 or CCR5, is a critical step that triggers a conformational change in gp120. Change in gp120 structure repositions this binding subunit resulting in exposure of gp41 and insertion of its N-terminal fusion peptide into the target membrane, leading to 6HB formation and membrane fusion (73, 142, 160). The triggering mechanism for the ALSV 28 envelope protein (envA) requires a specific interaction with its receptor (Tva) that triggers the envA protein and results in FP exposure, detectable by liposome membrane binding at neutral pH (50, 93).

Most paramyxovirus F proteins can be triggered and cause fusion only when they are co-expressed with their homologous attachment protein (52, 57, 95,

195). The attachment protein from one virus will not support the fusion activity of the F protein from another virus, unless the viruses are closely related (15).

Therefore in these viruses, specific interaction of F protein with its homotypic attachment protein probably initiates F protein triggering. The F protein from certain strains of Paramyxovirinae or F proteins containing particular mutations have been shown to cause some cell-cell fusion in the absence of an attachment protein (127, 144, 171, 186, 187) perhaps by destabilizing the already metastable F protein. However, none of these F proteins has been shown to support virus infection. Therefore in this triggering pathway, it is thought that the viral attachment protein binds to its receptor and destabilizes the F protein in some way that results in F protein triggering (44). The junction between the HRB and the head region has been postulated to be the trigger site at which the homologous attachment protein must interact to destabilize the F protein (220).

For all triggering mechanisms discussed above, binding of the viral attachment protein to its cell receptor is a prerequisite for entry (213).

However the F protein of RSV is unique in that it is capable of causing membrane fusion and initiating virus infection in immortalized, cultured cells

29 without the need for its attachment, G, glycoprotein (7, 103, 196), suggesting that it has cell attachment activity in addition to its fusion activity. The human metapneumovirus (hMPV) F protein is also able, as the only viral glycoprotein, to support infection of cultured immortalized cells (13), and to cause cell-cell fusion when expressed alone, although in this case cell-cell fusion may require low pH (184). Since hMPV is also in the Pneumovirinae subfamily of the Paramyxoviridae, it is possible that cell binding activity and direct triggering of the F protein of these viruses is a subfamily characteristic.

Since neither the G attachment glycoprotein nor low pH is required for RSV F protein triggering, it remains unclear what triggers the RSV F protein.

However, without the requirement for its attachment protein to function, the

RSV F protein appears to be the simplest paramyxovirus fusion protein for studying the triggering process. And what we learn could be relevant to the more complex attachment and fusion protein pairs of other paramyxoviruses, as well as retroviruses and others. What we learn could also lead to the development of novel antiviral drugs against RSV or an F protein vaccine to protect against RSV disease.

REVERSE GENETICS

Reverse genetics involves the study of a gene with a specifically inserted mutation in its sequence for its function as opposed to forward genetics that examines a phenotype and determines its sequence. Reverse genetics requires a set of techniques that enable an RNA sequence to be converted

30 into DNA, mutagenesis of that DNA and the use of that DNA to produce RNA, in most cases mRNA that encodes a mutant protein. A reverse genetics system for the complete RSV was first reported in 1995 (39). In this system, the entire 15,222 nucleotides of the RSV genome had been copied into cDNA and inserted into a plasmid. From this plasmid, replicating RSV could be rescued. This system is widely used in negative strand RNA viruses, including in addition to RSV, other members of Paramyxoviridae family, such as measles virus (MV) (166), SeV (76, 106), HMPV (12, 19), and Nipah virus

(221). The power of this system is that the full-length cDNA can be manipulated with routine DNA techniques to remove or add genes or to change any viral control sequence or mutate any viral protein.

For RSV, rescue from the full-length cDNA involves transfecting cells with a plasmid expressing the antigenomic RNA, along with plasmids expressing the

N, P, L, and M2-1 proteins that are essential for nucleocapsid formation and polymerase complex activity (36). Transcription from all of these plasmids is driven by the bacteriophage T7 polymerase promoter in the cytoplasm of transfected cells provided by vaccinia virus expressing the T7 polymerase

(MVA-T7). Reverse genetics has been a useful tool to create several live- attenuated vaccines in the past decade (36, 41). This RSV rescue system has also led to the generation of a potential RSV-based gene therapy vector for the treatment of Cystic Fibrosis (120) and an RSV replicon (134) that is presently being used to screen libraries of compounds to identify those that inhibit the RSV polymerase, as well as many other modified viruses.

VACCINES

Despite the importance of RSV in the pediatric population and extensive research, no effective vaccine is yet available. Even after a natural infection,

RSV can reinfect throughout life. Adults can even be infected repeatedly with the same strain of RSV in an experimental setting (88). Therefore, the goal of a vaccine must be to prevent disease, not to prevent infection. The second and subsequent infections with RSV cause a common cold, but seldom lower respiratory tract infection or severe disease. Therefore, an attenuated virus vaccine might be able to protect newborns from lower respiratory tract infection which can lead to severe disease. In the elderly, immune senescence might explain the decreased immune protection that enables repeated infection (88, 141).

For healthy adults, reinfection of the upper respiratory tract appears to be due to the fact that previous natural infections do not induce long-lived immunity capable of preventing subsequent infections. The inhibition of solid adaptive immunity to RSV may be due to the ability of the viral NS-1 and NS-2 proteins to inhibit the IFN response (16, 190), which is also important for the development of adaptive immunity (56).

Development of a vaccine to prevent RSV disease is clearly needed for the very young and the elderly. As a result of the vaccine-enhanced disease incidence in the 1960‟s clinical trial caused by the original formalin-inactivated

(FI) RSV vaccine and in the mouse model similarly immunized (110, 209), vaccine development has been approached with more caution. This

32 inactivated vaccine not only failed to induce protective immunity upon subsequent infection, but also caused enhanced disease in most of the recipients and resulted in two infant deaths. The lungs of these infants contained a high level of pulmonary inflammation and infiltrating eosinophils

(110, 124). Further analysis suggested that FI-RSV vaccine primed vaccinees for a Th2 response upon RSV challenge. An enhanced Th2 response, associated with inflammation, increased mucus production and airway hyperresponsiveness, and appeared to be the cause of this fatal immunopathology (86). Alternatively or in addition, the deposition of immune complexes of anti-RSV antibodies in lung tissue upon subsequent viral challenge might also contribute to the development of this enhanced illness

(162).

Because of the disease enhancement induced by inactivated vaccine, vaccine research has focused primarily on live-attenuated viruses for children.

However, protein subunit vaccines containing one or more of the major RSV proteins are being considered for adults who already have an established immune response pattern to RSV. Clearly, live vaccine does not result in enhance disease upon challenge, while greatly enhancing both the local and systemic immune responses (41).

Several live attenuated RSV vaccines have been developed by conventional methods including chemical mutagenesis and/or serial passage at low or high temperatures. The goal of this approach was to create viral mutants that replicate less under physiological conditions so they will not cause disease in

33 the vaccinee, but will replicate well enough to stimulate an adaptive immune response (49, 55). One promising vaccine candidate, cpts248/404 was found to be immunogenic, however nasal congestion followed vaccination indicating a need for further attenuation (217).

The conventional approach has identified attenuating mutations, and the availability of a reverse genetics system has enabled combining these mutations as well as other specific modifications to the virus genome to develop alternative live-attenuated RSV vaccines (41). It allows the use of site-directed mutagenesis to design and precisely introduce desired mutations into the RSV genome with more flexibility. Using this methodology, an ideal candidate with the best balance between immunogenicity and attenuation might be achieved.

Reverse genetics has also enabled the development of chimeric viral vaccines. In this approach, one virus is used as a vaccine vector to express an immunogen from another virus. In this manner, protective immunity against both pathogens is plausible. Related paramyxoviruses, such as bovine (B)PIV3, NDV, or SeV, have been tested as viral vectors to carry and express HRSV proteins (135, 182, 194). The chimeric vaccine that has progressed the farthest is the HPIV3/BPIV3 chimera (MEDI-534). It has the

BPIV backbone and so is naturally attenuated in humans. Its attachment and fusion proteins have been replaced with their human PIV3 counterparts and the genes for the HRSV G and F proteins have been added. This vaccine candidate is currently in clinical trials (182).

In another approach, RSV attenuation was achieved by nonessential gene deletion using reverse genetics. Removal of the interferon antagonist genes

(NS1 or NS2), the SH gene, or the M2-2 gene that regulates transcription and replication, yielded viable viruses with restricted replication in nonhuman primates (199, 214). Due to the loss of the type I IFN inhibitory effect of the

NS proteins, the vaccine induced a greater innate and therefore adaptive immune response. Due to the loss of transcription suppression in the M2-2 deleted virus, the vaccine displayed enhanced transcription of viral proteins.

Therefore, vaccines containing these deletions with or without the attenuating point mutations in other genes found by traditional methods have become good candidates for further clinical study (41).

Production of enough attenuated virus vaccine to make such an economically viable vaccine has also been a problem. The attenuated RSV vaccine candidates are produced in Vero cells, the cell line approved by WHO for vaccine production. However, a recent study from our laboratory showed that virus produced in Vero cells has a significantly reduced infectivity for primary well differentiated human airway epithelial cells and therefore most likely for human airway epithelium in vivo. Further investigation found that during production in Vero cells the G protein is cleaved and its C-terminus is lost resulting in the loss of infectivity (121). These observations indicate the importance of the cell type used in vaccine manufacturing. Production of the vaccine virus in a different cell line would result in a vaccine with higher infectivity, a smaller dose requirement and therefore the vaccine would be more economically viable, a problem with the current Vero-grown vaccines. 35

Since RSV is capable of causing disease in the elderly and in children with cystic fibrosis, there is also a need for a vaccine to protect these groups. It is also possible that boosting the antibodies against RSV in nursing mothers might confer protective antibody to babies in breast milk. A protein vaccine may be ideal for the elderly and nursing mothers because this is not their initial encounter with the virus so they have an established response pattern, unlike infants. Furthermore, an attenuated vaccine would be less immunogenic in these previously RSV-infected subjects (104).

The two major neutralizing antigens, the G and F proteins, would be the ideal immunogens for this type of vaccine. The F protein appears to be the better candidate for protein vaccine development because it is much more highly conserved between the two RSV subgroups and it is strongly immunogenic

(38, 99, 226). Purified F protein (PFP) vaccine candidates have been extracted from the HRSV-infected cell cultures and tested in healthy elderly

(64, 65). The PFP vaccine was well tolerated and induced modest levels of neutralizing antibody (65). A study of pregnant women showed only a minimal increase in RSV neutralizing IgG titers so that a protective effect conferred to offspring through breast feeding would not be expected (145). Furthermore, in children with cystic fibrosis, the PFP-3 vaccine failed to prevent HRSV infection (157).

It is not clear whether an adequate dose of the F protein was given in these

PFP vaccines or whether the PFP was in the ideal conformation for immunization. The PFP was a full-length RSV F protein extracted from

36 infected cells and purified by ion exchange chromatography. We have recently shown that exposure to a low molarity buffer triggers a soluble version of the RSV F protein (29) (Chapter 2). It is therefore possible that the molarity of the buffers used to purify the PFP from infected cells affected the final conformation of the PFP in the vaccine. Specific buffer concentrations were not listed in these reports, but the ion exchange procedure requires buffers of varying molarities and may well have cause the PFP to trigger. If the PFP were in the posttriggered form, antibodies raised against it may not be as effective as antibodies raised against the pretriggered form. An sF protein stabilized in the native, pretriggered conformation might be the ideal molecule for development of an F protein vaccine (29).

A recombinant G protein peptide vaccine that contains the central-conserved domain of the G protein fused to the albumin-binding domain of streptococcal protein G, BBG2Na, has also been developed (164). Despite its promising results in terms of safety and immunogenicity, further study has been postponed due to its side effects. Purpura, a type III hypersensitivity reaction, appeared in a few vaccine recipients during the Phase II Clinical Trial (111).

Another subunit vaccine containing the F and G protein, along with the M protein was also studied for it efficacy in the elderly (125). Vaccinated adults,

65 and older, exhibited an increased level of neutralizing antibody against both RSV subgroups with no severe adverse effects. Currently this vaccine is in a Phase II Clinical Trial for a potential use in elderly (66).

TREATMENT AND PREVENTION

Treatment of individuals suffering from lower respiratory tract HRSV infection involves supportive care such as suction removal of secretions, administration of humidified oxygen, and respiratory assistance. These procedures are recommended to reduce symptoms and lower the mortality rate for children hospitalized with RSV disease (37). Bronchodilators have been used to improve the asthma-like symptoms, wheezing caused by a constriction of airway smooth muscle, found in infant bronchiolitis. However the benefits are considered modest (107). Since pathogenesis of HRSV infection has been shown to be a result of a robust inflammatory response, anti-inflammatory therapy with corticosteroids or montelukast is commonly used in treatment of acute bronchiolitis despite a lack of strong evidence to support efficacy (14,

153).

The only approved antiviral compound available for treating HRSV infection is ribavirin, a nucleoside analog which causes mutations in the viral genome, thus diminishing viral replication. It has potent activity against HRSV infection in vitro and appears to reduce clinical illness in patients. However there is no significant data to support its efficacy in pulmonary function improvement or reduction of mortality and it does have side effects. Therefore, it is currently not used in most clinics (168, 206). Several companies are developing new antiviral agents to treat RSV disease, but none has yet reached the market.

Prevention of RSV disease has been accomplished for at-risk pre-mature newborns by passive immunization with a monoclonal antibody. Initially it was

38 demonstrated that intraperitoneal administration of HRSV neutralizing antibodies to cotton rats protects them from infection (165). A humanized version of the murine MAb 1129 (8), palivizumab, targets the RSV F protein and neutralizes HRSV of both subgroups. Under the commercial name

Synargis, this MAb is licensed for prophylactic treatment of at-risk children

(100). Palivizumab has become the standard of care in developed countries for the treatment of at-risk newborns. However the high cost per dose and requirement for repetitive administration, 5 doses during the winter months, to achieve and maintain an adequate amount of MAb in the serum limits access to this therapy in developing countries. Motavizumab, an improved version of palivizumab with 10 times greater neutralizing activity, is under FDA review

(218).

Palivizumab treatment of patients and cultured cells can select F protein mutations and the viruses carrying these mutations are no longer sensitive to this MAb (225, 227). The crystal structure of motavizumab complexed with the peptide containing these mutations confirms that this is its binding site.

However, this peptide in the pretriggered PIV5 F trimer, is partially obscured between two monomers. Either the trimeric configuration of the pretriggered

RSV F glycoprotein is different from that of PIV5 F in this region or the MAb induces monomers of the pretriggered F protein trimer to separate (139). It is also possible that motavizumab binds to its epitope when the epitope is exposed in the pre-hairpin intermediate during the structural transition from the pretriggered to the posttriggered forms, thus inhibiting the membrane fusion function of the F protein (139). 39

CHAPTER 2: SOLUBLE RSV FUSION PROTEIN

INTRODUCTION

Respiratory syncytial virus (RSV) is a major pathogen in infants and young children worldwide, causing bronchiolitis and pneumonia in 1%-5% of this population (188). It is also a significant cause of mortality in the elderly and in immunocompromised patients (37, 63). A humanized monoclonal antibody

(MAb) is used to protect premature infants who are at highest risk for severe

RSV disease (163). Despite extensive efforts, a vaccine is not yet available.

RSV is a member of the Mononegavirales in the Paramyxoviridae family (37),

Pneumovirinae subfamily. Other viruses in the Paramyxovirinae subfamily require their homologous attachment glycoprotein in addition to the fusion (F) glycoprotein to initiate fusion between the virus membrane and the target cell membrane. In these viruses, the attachment protein not only binds to the target cell receptor, but also triggers the F protein, resulting in fusion (52, 57,

95, 195). Fusion allows the viral nucleocapsid to enter the cytoplasm and initiate infection.

The F protein of RSV is unique in that it is capable of causing membrane fusion and initiating virus infection in immortalized, cultured cells in the absence of its attachment, G, glycoprotein (101, 103, 196). The RSV F

40 protein has been a major target for vaccine and antiviral drug development because of its importance in the viral replication cycle, its conserved sequence and structure, its exposed position in the virion, and its strong immunogenicity (37, 99, 226).

The RSV F protein expressed in nonpolarized cultured cells causes cell-cell fusion at neutral pH, leading to the characteristic syncytia or multinucleated giant cells. Since low pH is not required for cell-cell fusion by the RSV F protein, unlike the influenza virus HA protein for instance, and since the G attachment glycoprotein is not required for F protein triggering (7, 196), it is not clear what triggers the F protein.

Like other class I viral fusion proteins, the RSV F protein is a trimer. Each monomer is produced in a precursor, F0, form that is modified by N-linked glycans. During passage through the Golgi apparatus, these glycans mature and the protein is cleaved by a furin-like protease in two positions, releasing a

27-amino-acid peptide (pep27) with attached glycans (228, 231). The resulting F protein is composed of the N-terminal F2 protein linked by one, and probably two, disulfide bonds to the F1 transmembrane protein (51). The highly hydrophobic fusion peptide resides at the cleavage-created N terminus of the F1 protein (37). This cleaved form of the RSV F protein is fully active and able to cause cell-cell fusion once it reaches the plasma membrane and to cause virion-cell fusion.

Initially, the paramyxovirus F protein is in a metastable, pretriggered trimer form (226). Upon triggering, it is thought to undergo a dramatic and

41 irreversible conformational change, extending the α-helical region (HRA) that follows the N-terminal fusion peptide, thereby inserting the fusion peptide into the target cell membrane (28). The three F1 monomers then fold back on themselves, juxtaposing HRA with a second α-helix (HRB) adjacent to the transmembrane domain of F1 and forming a highly stable 6-helix bundle (6-

HB) (28, 58). This action pulls the two membranes together to initiate membrane fusion (58).

The complete X-ray crystal structures of two paramyxovirus soluble F (sF) proteins, one in a pre- and one in a posttriggered conformation, have been solved. The first, the parainfluenza virus (PIV) type 3 F protein, which lacked the transmembrane and cytoplasmic domains, was in the posttriggered form

(219). In an attempt to stabilize the pretriggered form for crystallization, the

PIV5 sF protein was modified with a trimerization domain at its C terminus

(220). The PIV5 sF protein was in a very different, apparently pretriggered form, suggesting that stabilization of the C terminus of the sF protein is important for maintenance of this form. The PIV5 sF protein could be triggered by the addition of a surrogate trigger, heat (45). The natural mechanism by which the PIV5 F protein causes membrane fusion requires the homologous attachment protein. Following receptor binding, a conformational change within the attachment protein is thought to exert a destabilizing effect on the F protein that initiates its refolding (44).

In the present study, we have produced fully cleaved, pretriggered RSV sF protein by removing the transmembrane and cytoplasmic domains and

42 replacing them with tags for purification and detection. This sF protein had a spherical shape and was not aggregated, consistent with it being in the pretriggered form. Dialysis into a low-molarity buffer converted the sF protein to a “hat pin” shape that aggregated in rosettes, characteristic of the posttriggered form of the sF protein. Likewise, dilution of the sF protein with low-molarity buffer in the presence of liposomes resulted in similar hat pin spikes embedded in the lipid membranes. We conclude that low-molarity buffer functions as a surrogate, or perhaps physiological, trigger for the RSV

F protein.

MATERIALS AND METHODS

Cells. Human kidney epithelial 293T cells were grown in Dulbecco modified

Eagle medium (DMEM), 10% fetal calf serum (FCS) at 37°C in a humidified chamber with 5% CO2. During sF protein production, transfected 293T cells were grown at 33°C.

Construction of the sF protein and mutant genes. A codon-optimized version of the RSV D53 F protein gene, similar to that of strain A2, was used to construct the SC-2 soluble F protein gene. Codons 1 to 524 of the F open reading frame, encoding the ectodomain of the F protein were fused to the

FLAG tag followed by the 6His tag by inverted PCR mutagenesis (26), effectively deleting the transmembrane and cytoplasmic domains. sF mutant proteins with furin cleavage site 1 (fcs-1) (i.e., at the downstream side of pep27; parental sequence KKRKRR), fcs-2 (at the upstream side of pep27; parental sequence RARR), or both sites ablated were generated by inverted PCR mutagenesis. The fcs-1 site was replaced by the enterokinase cleavage site (DDDDK), and the fcs-2 site was replaced with a trypsin cleavage site (NATR) to create the SC-33 and SC-31 sF genes, respectively.

The SC-35 sF gene had both fcs sites replaced. The production of all sF proteins was driven by a cytomegalovirus (CMV) promoter from the pcDNA3.1 expression plasmid (Invitrogen).

Production and purification of the sF proteins. Plasmids containing the sF gene were transfected into 293T cells. The sF protein was purified from the medium harvested at 48 h posttransfection, by using passage over a Ni2+

44 column (Probond purification system; Invitrogen) and elution with 250 mM imidazole in either 50 mM phosphate buffer (pH 8.0) with 500 mM NaCl or 10 mM HEPES buffer (pH 8.0) without additional NaCl. Samples were reduced and analyzed by Western blotting with 5His MAb (Invitrogen), anti-mouse antibody–horseradish peroxidase (Kirkegaard & Perry Laboratories), and

Lumi-Light Western blotting substrate (Roche).

Velocity gradient centrifugation. Purified sF protein was layered over an

11-ml 20-to-55% (wt/wt) linear sucrose gradient (in HBSS buffer with Ca2+ and Mg2+) and centrifuged at 41,000 RPM for 20 h in an SW 41Ti rotor

(Beckman Instruments). When indicated below, purified sF protein was incubated at 50°C for 1 h prior to centrifugation. Gradients were collected from the bottom with a peristaltic pump and a fraction collector. Two micrograms of bovine serum albumin was were added as carrier, and the proteins were precipitated with 15% (vol/vol) trichloroacetic acid (TCA) and analyzed by Western blotting with anti-5His MAb.

Radioimmunoprecipitation. 293T cells were transfected with plasmids expressing the SC-2 protein and metabolically labeled with [35S]methionine- cysteine (MP Biomedical, Solon, OH) at 100 µCi/ml for 1 h at 37°C, beginning at 46 h posttransfection. The medium was removed, complete DMEM with

10% FCS was added, and incubated continued for another 1 h. Then the medium from these cells was harvested, clarified, and incubated for 1 h at

4°C or 50°C, followed by immunoprecipitation overnight with protein G Plus-

Sepharose beads (Calbiochem) precoated with neutralizing anti-RSV F

MAbs. The beads were washed six times with 0.01% Triton X-100 in 45 phosphate-buffered saline and resuspended in Laemmli protein loading buffer before boiling for 5 min. Proteins were separated by SDS-PAGE, and protein bands were visualized using a Typhoon Phosphoimager and quantified using

ImageQuant TL software (GE Healthcare).

Protein dialysis. One milliliter of freshly prepared and purified SC-2 sF protein in 50 mM phosphate buffer containing 500 mM NaCl and 250 mM imidazole was promptly dialyzed against six different buffers (pH 8.0) described in the figure legends for 18 h at 4°C using dialysis cassette units

(Slide-A-Lyzer; Thermo Scientific). Dialyzed protein samples were analyzed by sucrose velocity gradient and electron microscopy (EM).

Liposome association assay. Liposomes were prepared from an 8:2:5 molar ratio of 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine (POPC), 1- palmitoyl-2-oleoyl-sn-glycero-3-phosphoethanolamine (POPE), and cholesterol (Avanti Polar Lipids) dissolved in chloroform- methanol (2:1). Lipid films were deposited on glass tubes by removing most of the solvent with a stream of argon and removing any that remained by evaporation under vacuum (Freezone 2.5; Labconco). Lipid films were stored at -20°C under argon for up to 1 month. Before use, the dried lipid film was resuspended in 1 ml of buffer, then taken through five freeze-thaw cycles with vortexing at maximum speed between cycles. The suspension of multilayer liposomes was extruded 40 times through 100-µm filters with a miniextruder (Avanti

Polar Lipids). In each protein-liposome reaction mixture, 2.2 µg of sF protein was mixed with 20 µl of 20 mM liposome (4 mmole of total lipids), and the same buffer used in the liposome preparation was added to bring the volume 46 up to 80 µl. The samples were then incubated at 4°C for 30 min. As a control, protein alone was incubated at 4°C for 30 min. When indicated, 80 µl of 100 mM sodium carbonate (pH 11) was added at 21°C for 10 min before sucrose gradient flotation.

Each reaction mixture was combined with 840 µl of 60% sucrose made in the same buffer used in the liposome preparation step to obtain a final sucrose concentration of 50%, and the mixture was overlaid with 1 ml each of 40% sucrose, 30% sucrose, 20% sucrose, and 10 mM HEPES. Following centrifugation in an SW55Ti rotor at 55,000 rpm for 2 h, 20 min at 4ºC, the gradients were collected in 1.2-ml fractions as described above. Lipids were solubilized in 0.5% Triton X-100 and TCA precipitated as described above.

Proteins were analyzed by Western blotting with anti-5His MAb.

Electron microscopy. sF protein samples were adsorbed onto 300-mesh carbon-coated copper grids freshly treated by glow discharge and stained with fresh 1% sodium uranyl formate, pH 4.5 to 5. An FEI Technai G2 Spirit transmission electron microscope operating at 80 kV was used to examine the sample. For sF protein-liposome mixtures, the sF protein was mixed with liposome in 10 mM HEPES containing 0.8 mM lipid to a final protein concentration of 12.8 µg/ml and incubated at 4°C for 30 min. Samples were examined by EM as described above.

RESULTS

Production of RSV sF protein. We constructed a full-length, codon- optimized synthetic gene for the RSV F protein to enable plasmid expression from the nucleus (201). Transient expression in 293T cells resulted in syncytium formation (data not shown), demonstrating that the optimized F gene produced a functional product. To facilitate structural and functional studies of the RSV F protein, we modified the gene sequence by removing the transmembrane and cytoplasmic domains to produce an anchorless sF protein, SC-2 (Fig. 2.1). FLAG and 6His tags were linked to the C-terminus to facilitate purification, concentration, and detection. This sF protein sequence retained the entire F protein ectodomain, with its functional furin cleavage sites.

Fig. 2.1. The wild-type RSV F protein and the cDNA-expressed SC-2 sF protein generated from it. The wild-type F0 precursor is cleaved intracellularly by a furin-like protease at two sites to generate the mature F1 and F2 subunits that remain linked by probably two disulfide bonds. The two heptad repeats, HRA and HRB, interact to complete the conformational changes that result in the final post-triggered F protein 6-HB. To generate the cDNA-expressed sF protein, the cytoplasmic and transmembrane (TM) domains were replaced with the FLAG and 6His tags, and the sF protein was expressed from pcDNA3.1.

A plasmid containing the SC-2 sF gene, driven by a CMV promoter, was transfected into 293T cells and the sF protein was detected in the cell culture medium at 48 h posttransfection (Fig. 2.2). Samples from the cell lysate (C) and the medium (M) were reduced before loading onto the SDS- polyacrylamide gel for electrophoresis. Both the precursor, sF0, and the fully cleaved sF1 protein were detected intracellularly. However, only the sF1 protein was detected in the medium, indicating that only completely cleaved sF protein was secreted. Because the cell lysates were loaded at a 10-fold- greater level than the medium, and the total signals in these lanes were similar, it appears that at least 90% of the sF protein was released into the medium. Thus, the SC-2 sF protein was efficiently cleaved and released from

293T-transfected cells in its fully cleaved form.

Fig. 2.2. Western blot analysis of the SC-2 sF protein produced from transfected 293T cells at 48 h posttransfection. The sF protein from cell lysates (C) and media (M) were stained with the 5His MAb. The C lanes represent 10X more cell equivalents than the M lanes. pcDNA3.1 was included as the empty vector control.

The sF protein is produced in the pretriggered form. If the SC-2 sF protein represents the pretriggered form of the F protein, MAbs that recognize the RSV F protein and are capable of neutralizing RSV infectivity in cell culture should recognize it. To test this possibility, transfected cells were metabolically labeled with [35S]-methionine-cysteine and the medium was immunoprecipitated individually with 11 neutralizing MAbs representing four antigenic sites with multiple epitopes in each site. All 11 of these MAbs precipitated the sF protein. The findings with one MAb from each antigenic site are presented in Fig. 2.3. These results suggest that this sF protein is in its native conformation, at least relative to these antigenic sites and their epitopes. The reduced reactivity of these MAbs with heated (50°C) sF protein that is presented in Fig. 2.3, will be discussed below.

Fig. 2.3. MAb immunoprecipitation of the SC-2 sF protein after incubation at 4°C or 50°C for 1 h. 293T cells were transfected with plasmids expressing SC-2 sF protein and metabolically labeled with [35S]-methionine- cysteine. The medium from these cells was incubated for 1 h at 4°C or 50°C followed by immunoprecipitation with individual MAbs. The antigenic sites for the MAbs that have been mapped by competition (8) are indicated across the top.

Affinity-purified RSV SC-2 sF protein in 50 mM phosphate or 10 mM HEPES buffer was examined by negative staining with EM. It appeared primarily as a sphere with a diameter of 10.9-15.2 nm, often with a dark center (Fig. 2.4A).

This shape and the dark center were similar to that previously reported for the pretriggered PIV5 sF protein (44, 45). However, the PIV5 sF protein, whose

C-terminus was linked to a GCNt self-trimerizing domain, was reported to be somewhat smaller, 7-11 nm in diameter, and often included an attached stem.

The SC-2 sF protein did not include such a stem, probably because we did not link a trimerization domain to it. The dark center of these sF proteins

53 probably represents a collection of the negative stain in the hollow sF protein head.

A small number of rosette aggregates were detected in the RSV sF protein preparations (Fig. 2.4A-3, final panel with asterisk). The individual molecules in these rosettes have a “hat pin” shape with a much smaller head and a thin, long body compared to the majority of the sF molecules that appear as spheres. Similar hat pin or cone-shaped structures were previously observed for the posttriggered RSV sF protein (27), the PIV3 sF protein, and for the heat triggered PIV5 sF protein (45). These structures likely represent the posttriggered form of the sF protein, aggregated via their highly hydrophobic fusion peptides. This low level of triggering presumably occurred nonspecifically during preparation. The effect of heat and dialysis on the sF protein presented in this figure will be discussed below.

Fig. 2.4. Electron micrographs of the SC-2 sF protein. Ni2+ affinity- purified

SC-2 sF protein was analyzed by negative staining EM using 1% uranyl formate. (A) SC-2 sF protein eluted from the Ni2+ beads in either 50 mM phosphate buffer, 500 mM NaCl, 250 mM imidazole (images in section 1 and

3) or 10 mM HEPES buffer (section 2) with imidizole. (B) The SC-2 protein in

50 mM phosphate buffer from panel A, heated to 50°C for 30 min. (C) SC-2 protein released from Ni2+ beads in 50 mM phosphate buffer, 500 mM NaCl, and 250 mM imidizole and dialyzed against 10 mM HEPES buffer for 18 h at

4°C.

Test of mild heat as a surrogate trigger for the RSV sF protein. To determine whether mild heat would act as a surrogate trigger for the SC-2 sF protein as it had for the PIV5 sF protein (45), we compared SC-2 sF protein maintained at 4°C or at 50°C for 1 h by analysis on sucrose gradients. If the unheated SC-2 sF protein is in the pretriggered form, trimers would remain unaggregated and near the top of a velocity sedimentation sucrose gradient

(9, 45). Aggregates, including posttriggered trimers, would move further into the gradient. The SC-2 sF protein was found in fractions 2 and 3 (Fig. 2.5A), near the top of the gradient, indicating that it was not in an aggregated form.

This conclusion is consistent with the EM identification of individual pretriggered sF protein spheres.

Fig. 2.5. Analysis of the SC-2 sF protein aggregation state by velocity sucrose gradient centrifugation. Freshly prepared and purified SC-2 was incubated at 4°C or 50°C for 1 h before loading on top of a linear 25%-to-55% sucrose gradient for ultracentrifugation in an SW41 rotor at 41,000 rpm for 20 h. The protein in each fraction was TCA precipitated, separated by SDS-

PAGE and detected by Western blotting with 5His MAb. The top fraction of the gradient is indicated.

In contrast, the heated SC-2 sF protein migrated further into the gradient with a peak in fractions 5 and 6 (Fig. 2.5B). By EM, most of the heated sF protein appeared to be in larger aggregates with no obvious definition (Fig. 2.4B-1), probably due to heat induced denaturation and aggregation. A smaller portion of the heat-treated sF protein appeared as rosettes of hat pins with their thin ends pointing toward the center (Fig. 2.4B-2) indicative of posttriggered sF protein aggregated by their hydrophobic fusion peptides. These rosettes ranged from 36 - 43 nm in diameter (Fig. 2.4B-2).

Consistent with this interpretation, 50ºC treatment resulted in the loss of nearly all reactivity with the F-specific MAbs (Fig. 2.3, “+” lanes). Together, these results confirm that mild heating induced a dramatic change in the RSV sF protein conformation resulting in aggregation that destroys or hides all of the antigenic sites in most of the protein molecules rather than causing an orderly triggering process.

Effect of dialysis on protein triggering. With the exception of the immunoprecipitation experiment (Fig. 2.3), the sF protein was partially purified and concentrated by Ni2+ column chromatography. It was released from the column by 250 mM imidazole in 50 mM phosphate, 500 mM NaCl buffer. To reduce the salt concentration and remove the imidazole from the purified sF protein solution, we dialyzed against 10 mM HEPES buffer. Surprisingly, dialysis converted most of the sF protein spheres to rosettes (Fig. 2.4C).

We reasoned that the sF protein must have been triggered by one of three changes: exposure to the low molarity (10 mM HEPES) buffer; the loss of imidazole; or contact with the dialysis membrane. To distinguish among these possibilities, we dialyzed the purified sF protein (50 mM phosphate, 500 mM

NaCl, 250 mM imidazole) with three buffers, each with or without 250 mM imidazole for a total of 6 conditions. The three buffers were: the same buffer the sF protein was in; 10 mM phosphate with 100 mM NaCl; and 10 mM

HEPES. Dialyzed samples were examined by EM and by sucrose velocity sedimentation to determine which variable caused the change in protein morphology and aggregation.

When 50 mM phosphate, 500 mM NaCl buffer, with or without imidazole, was used for dialysis, the sF protein retained its spherical shape and was not aggregated (Fig. 2.6A and B), corresponding to the pre-dialysis unaggregated pretriggered protein (Fig. 2.5A). These results indicate that contact with the dialysis membrane did not cause the sF protein to trigger. Also, removal of

58 the imidazole failed to cause the sF protein to trigger, indicating that imidazole is not essential for maintaining the pretriggered form.

Dialysis against either 10 mM phosphate buffer with 100 mM NaCl, or 10 mM

HEPES buffer, both in the presence of 250 mM imidazole, converted a small amount of the sF protein spheres to rosettes, consistent with the migration of a small amount of the protein further into the gradient (Fig. 2.6C and D). In the absence of imidazole, more of the sF protein was converted to the posttriggered form when dialyzed against 10 mM phosphate buffer, 100 mM

NaCl (Fig. 2.6E), and nearly all of it was converted to the posttriggered form when dialyzed against 10 mM HEPES buffer (Fig. 2.6F), as previously shown in Fig. 2.4C. Since the removal of imidazole was ruled out as the single cause of triggering above, these results indicate that low buffer molarity is responsible for triggering the sF protein. Imidazole contributes to this molarity effect in conjunction with both the 10 mM phosphate, 100 mM NaCl, and the

10 mM HEPES buffers.

This posttriggered sF protein retained reactivity with all 5 of the anti-F MAbs

(data not shown) that were demonstrated in Fig. 2.3 to react with the pretriggered sF protein. Apparently none of these antigenic sites are altered by triggering. None of these MAbs bind to the sF protein in a western blot

(data not shown), indicating that their corresponding antigenic sites are conformational.

Fig. 2.6. Sucrose velocity gradients and electron micrographs of the SC-

2 sF protein following dialysis with six different buffers. A 1-ml aliquot of freshly prepared and purified SC-2 in 50 mM phosphate (PO4) buffer containing 500 mM NaCl and 250 mM imidazole was dialyzed against six different buffers for 18 h at 4°C before sucrose velocity gradient and EM analyses. The buffers and the net salt molarity are listed above each sucrose gradient panel. Proteins were detected by Western blotting as described in the legend to Fig. 2.5. 60

Association of the sF protein with liposomes. The classical method for detecting triggering of a viral fusion protein is by liposome association and co- floatation. A soluble fusion protein does not insert its fusion peptide into the liposomes, until an agent that triggers the fusion protein is added (50, 54, 92).

Liposome association is detected by co-flotation of the fusion protein with the liposomes. For instance, a mixture of trypsin-treated PIV5 sF protein and liposomes treated with a surrogate stimulus, mild heat (60°C for 30 min), causes triggering, liposome association and co-flotation (45).

To confirm that triggering is induced by low molarity, we prepared large unilamellar liposomes in each of the 6 buffers used above. We mixed 30 µl of sF protein with 50 µl of liposomes in each buffer, and incubated the mixtures at

4ºC for 30 min. To separate the free sF protein from the liposomes, we added sucrose to a final concentration of 50%, overlaid with solutions of 40%, 30%,

20% and 0% sucrose, and subjected the gradient to ultracentrifugation.

Protein that associates with liposomes will float with them into the two less dense sucrose fractions.

Without liposomes, the sF protein remained in the bottom two sucrose gradient fractions (Fig. 2.7G). With liposomes in the high molarity 50 mM phosphate, 500 mM NaCl buffer, most of the sF protein did not associate with the liposomes, remaining in the bottom two fractions regardless of whether imidazole was included (Fig. 2.7A and B), indicating that the high molarity buffer did not cause triggering and fusion peptide exposure. With liposomes in the intermediate molarity, 10 mM phosphate, 100 mM NaCl buffer or 10 mM

HEPES in the presence of the 250 mM imidazole, approximately half of the sF protein associated with the liposomes (Fig. 2.7C and D).

With liposomes in lower molarity, 10 mM phosphate, 100 mM NaCl without imidazole, most of the sF protein associated with the liposomes (Fig. 2.7E).

And at the lowest molarity, 10 mM HEPES buffer, nearly all of the sF protein associated with liposomes (Fig. 2.7F). The interactions between the sF protein and liposomes was not disrupted by pH 11 treatment (Fig. 2.7, bottom row), indicating that, in all cases, the sF protein was stably inserted into the liposome membrane rather than peripherally associated. As in the EM and velocity gradient assays in the previous section, the association with liposomes correlated closely with the molarity of the buffer, confirming that low buffer molarity triggers the sF protein.

Fig. 2.7. Association of the RSV sF protein with POPC-POPE-cholesterol

(8:2:5) large unilamellar liposomes. The SC-2 sF protein (30 µl, 2.2 µg) was mixed with liposomes (4.4 mmole) in the presence of six different buffers and incubated for 30 min at 4ºC (lane groups A to F) in two aliquots. One aliquot was treated with carbonate buffer at pH 11 for 10 min at 20°C to release any superficially bound sF protein. Both aliquots were adjusted to a final concentration of 50% sucrose, overlaid with steps of 40%, 30% and 20% sucrose, and ultracentrifuged in an SW55 Ti rotor at 55,000 rpm for 2.5 h. (G)

The sF protein was also incubated for 1 h at 4ºC without liposomes and analyzed on a sucrose flotation gradient. The protein in each fraction was precipitated and analyzed as described in the legend to Fig. 2.5. The top fraction of the gradient is indicated.

If the RSV sF protein was indeed triggered by low molarity buffer conditions and associates with the liposomes via its fusion peptide, as supposed, we should be able to visualize the posttriggered sF protein projecting from the liposomes like spikes. Such spikes have been demonstrated in class II membrane fusion proteins from Semliki Forest virus and Dengue virus in which triggering is low pH and cholesterol dependent (128, 180). Consistent with this expectation, when the sF protein was incubated at 4°C for 30 min with liposomes in 10 mM HEPES buffer, protein spikes were found inserted into the liposome membranes (Fig. 2.8). These molecules had the same hat pin shape and size (14-16 nm x 5.4 nm at the top) of the posttriggered sF protein found in the rosettes (Fig. 2.4), with the narrow end of the hat pin inserted in the membrane. This is the end that should contain the fusion peptide. These results again indicate that low molarity buffer causes the sF protein to trigger, exposing its fusion peptide and inserting into a lipid membrane.

Fig. 2.8. EM of SC-2 sF protein inserted into liposomes. The purified SC-2 protein was incubated with liposomes in 10 mM HEPES at 4°C for 30 min before negative staining and EM examination. A diagram resembling SC-2 spikes inserted into a liposome membrane is shown at the bottom right.

Involvement of the fusion peptide in liposome association. To confirm that the association between the SC-2 sF protein and the liposome membrane occurred specifically through insertion of the exposed hydrophobic fusion peptide, rather than by a hydrophobic patch on the sF molecule, we generated three RSV sF proteins with mutations in one or the other, or both furin cleavage sites. Specifically, in SC-31, the upstream cleavage sequence was changed from RARR to NATR; in SC-33, the downstream cleavage site

(at the F2-F1 junction, and thus proximal to the fusion peptide) was changed from KKRKRR to DDDDK; and in SC-35, both changes were made.

All three sF cleavage mutant proteins were cloned and produced in 293T cells by transient transfection, as described for the parental SC-2 sF protein in Fig.

2.2. Western blot analysis showed that all of these sF protein mutants were efficiently synthesized and released from 293T-transfected cells (Fig. 2.9A).

These results indicate that cleavage is not required for transit through the cell and release into the media. Under reducing conditions, the SC-33 and SC-35 mutants migrated with a higher molecular mass than the parental SC-2 sF protein because their F1 protein retains the pep27 fragment or the pep27+F2 fragment, respectively. All 5 of the anti-F MAbs shown in Fig. 2.3 recognize the three cleavage site mutants (data not shown) indicating that, at least as far as these antigenic sites are concerned, these sF mutant proteins are in the native conformation.

These cleavage site mutants were tested for liposome association in the most effective triggering condition identified above, 10 mM HEPES buffer. Only SC-

31, the sF protein mutant that retained cleavage site proximal to the fusion peptide associated with liposomes (Fig. 2.9Bii), like the parental SC-2 (Fig.

2.9Bi). The two mutants lacking the fusion peptide proximal cleavage site,

SC-33 and SC-35, did not associate with liposomes (Fig. 2.9Biii and iv).

These results confirm that the RSV sF protein associates with liposomes via its fusion peptide, but only after the fusion peptide is released by cleavage, becoming the N-terminus of the F1 protein.

Fig. 2.9. Efficient secretion of sF protein cleavage mutants and the requirement of cleavage at the furin site adjacent to the fusion peptide for liposome association. (A) Western blot analysis of the SC-2-derived cleavage site mutant produced from transfected 293T cells. The sF protein from cell lysates (C) and media (M) were stained with the 5His MAb. The C lanes represent 10-fold more cell equivalents than the M lanes. pcDNA3.1 is the empty vector control, and SC-2 is the positive-control parental sF protein.

(B) Association of furin cleavage site sF mutants with liposomes. Each sF protein was incubated with liposomes in 10 mM HEPES buffer without imidazole at the indicated temperatures for 30 min. The mixtures were then treated with carbonate buffer, pH 11, for 10 min at 20°C to release any superficially bound sF protein before the floatation analysis. The mixtures were analyzed by using a flotation gradient as described in the legend to Fig.

2.7. The top fraction of the gradient is indicated. As indicated in the cartoons to the left of each panel, SC-2 is the parental protein with both cleavage sites intact. In SC-3, the upstream cleavage site has been ablated, in SC-33, the 68 downstream, fusion peptide-proximal site has been ablated, and in SC-35, both sites have been ablated.

DISCUSSION

The RSV F protein appears to be the simplest paramyxovirus fusion protein for studying the triggering process because it does not need its attachment protein to cause membrane fusion. It is also the major target for vaccine development and for small molecule antiviral drug development in this medically important virus. To initiate studies on the RSV F protein triggering process, we have generated an sF protein that lacks the transmembrane and cytoplasmic domains. This sF protein was secreted in the fully-cleaved, pretriggered conformation, as confirmed by its MAb reactivity, spherical morphology, and unaggregated state. The addition of an artificial trimerization domain was not required, as had been found for the only other pretriggered paramyxovirus sF protein that had previously been produced, the PIV5 sF protein (220). We have identified a treatment that triggers the metastable

RSV sF protein: placement in a buffer of low molarity. It is not yet clear if this treatment initiates triggering in a manner similar to the physiological triggering of the F protein on the cell or virion surface.

In previous work by other investigators, the sF protein of a different strain of

RSV, the Long strain, was generated by removing the transmembrane and cytoplasmic domains (83) in a manner similar to the present study. However, most of that sF protein aggregated in posttriggered rosettes. That portion of the sF protein population was efficiently cleaved, whereas the sF population that remained unaggregated retained some uncleaved molecules. The addition of trypsin to the unaggregated, uncleaved sF molecules caused them to form rosettes. Those results led the authors to speculate that the final 70 proteolytic cleavage triggers the F protein trimers and that in the virion, this final cleavage might take place on the surface of the target cell during entry, initiating fusion and infection (9). However, both their supposed pretriggered and posttriggered forms had a very similar appearance by EM, a small round head with a relatively long stem, and both were very stable to heating, showing no evidence for conformational change until approximately 90°C

(177). Extreme heat stability is not a characteristic of metastable proteins like the F protein including the pretriggered form of the PIV5 sF protein (45).

However, the posttriggered, 6-HB form of the F protein is very heat stable (5).

In hindsight, it is likely that both the uncleaved and cleaved forms of their RSV sF protein were in fact already triggered, as discussed below for the PIV3 sF protein.

Their use of the F protein from the Long strain, with 6 amino acids differences from the D53 strain that we used, could have contributed to the differences in our results. It is also possible that the triggered sF protein observed in the previous study was due to a difference in the handling of the sF protein. The low pH of the Glycine-HCl buffer used to elute their purified sF protein from the MAb column or its molarity (100 mM) may have caused triggering. We have shown here that a 10 mM phosphate, 100 mM NaCl buffer converted a large proportion of the sF protein to posttriggered rosettes. In addition, it is possible that other differences in the treatment of the sF protein during isolation and storage were responsible. Their sF protein was frozen, thawed and precipitated with 65% ammonium sulfate prior to analysis (27). We have found that freezing and thawing causes the sF protein to trigger 71

(Chaiwatpongsakorn, S. and Peeples, M.E., unpublished data). In contrast, the pretriggered sF protein that we produced and rapidly purified was surprisingly stable when stored in 50 mM phosphate (pH 8.0), 500 mM NaCl,

250 mM imidazole buffer at 4ºC, converting to the posttriggered form with a half-life of approximately 3 weeks (Chaiwatpongsakorn, S. and Peeples, M.E., unpublished data).

Our ability to produce the RSV sF protein in the pretriggered form, despite complete proteolytic cleavage, indicates that cleavage does not cause triggering of the RSV F protein. This conclusion is consistent with the results of the three successful X-ray crystallographic studies of other paramyxovirus sF proteins, all of which started with mutant sF protein that could not be cleaved by furin during transport through the cell, to avoid triggering. The structures of both the Newcastle disease virus (33) and the PIV3 (219) sF proteins were, surprisingly, in the posttriggered form. Trypsin treatment of this already triggered PIV3 sF protein resulted in cleavage at the former furin site and rosette formation (45). Clearly, rosette formation was the result of fusion peptide exposure, not of triggering, because this PIV3 sF protein was already in the posttriggered form before trypsin treatment. Proteolytic cleavage clearly does not serve as a triggering mechanism as had previously been proposed

(83).

Removal of the membrane anchor of the PIV3 F protein led to its triggering

(219). To avoid this, Yin et al. (220) fused a trimerization domain from a foreign protein, GCNt, to the C-terminus of the PIV5 sF protein. This PIV5 sF

72 protein appeared by EM as a sphere with a short stem composed of its HRB and the GCNt α-helical extension (44, 45), similar to the shape found in its pretriggered crystal structure (220). Proteolytic cleavage of this sF protein at the site between F1 and F2 did not trigger a conformational change, reinforcing the idea that cleavage is not the trigger. However once it is cleaved, the PIV5 sF protein can be triggered by heat, adopting a different structure with a smaller head and a longer stem, similar to the posttriggered

6-HB form of the PIV3 sF protein (45) that forms rosettes. Therefore, the PIV5 sF protein was originally in the pretriggered form and heat served as a surrogate trigger. In membrane fusion caused by PIV3 and PIV5, engagement of the attachment protein with its receptor is thought to lead to a conformational change within the attachment protein that destabilizes and triggers the F protein resulting in refolding and leading to membrane fusion

(44). As reported here, neither the membrane anchor nor a substitute trimerization domain is necessary to maintain the RSV sF protein in its pretriggered form as found for the PIV3 and PIV5 sF proteins.

Here we have identified low molarity as a novel method for triggering the RSV sF protein. It is not clear why reduction in buffer molarity causes sF protein triggering, but buffers are known to play a major role in protein stability and conformational change (205). Modifying the ionic strength or molarity of a solution can affect the stability of a protein (172, 207) depending on its charge distribution (114). Similarly charged residues in a protein, which are screened in solutions containing salt, repel each other in low salt environments (53, 60,

169), thereby destabilizing it. 73

We found that the RSV sF protein association with liposomes was not temperature dependent, occurring equally well at 37°C and 4°C (Fig. 2.9). In contrast, the avian leukemia and sarcoma virus (ALSV) attachment glycoprotein associated with liposomes in a temperature dependent manner, with very poor association at 4°C (50, 92). The influenza virus HA also associated with liposomes in a temperature dependent manner if the liposomes are composed of dimyristoylphosphatidylcholine (DMPC).

However, with liposomes composed of egg phosphatidylcholine (PC) there was no temperature effect probably because egg PC melts below 0°C and therefore remains fluid at 4°C while DMPC melts at 24°C and therefore does not (54). From these earlier studies it appears that the melting temperature of the lipids is critical in liposome-fusion protein binding experiments. In other words, the lack of ALSV glycoprotein insertion into the liposomes at low temperature may have been caused by the state of the lipids rather than the lack of glycoprotein triggering. The major lipid in the liposomes that we used in the present studies was POPC, a PC derivative with a low melting temperature similar to PC. It would also be fluid at 4°C.

The addition of heat has been reported to be a surrogate trigger for many full- length paramyxovirus fusion proteins (45, 154, 178, 212, 222). The mechanism probably involves thermal destabilization of the metastable F protein, initiating its conformational change. Yunis et al. (222) found that heating RSV-infected cells to 45-55°C enhanced antibody recognition of the membrane-anchored F protein by antiserum specific for the posttriggered 6-

HB structure, indicating that elevated temperature can act as a surrogate 74 trigger for the transmembrane RSV F protein. Raising the temperature to

60°C has also been shown to act as a surrogate trigger for the PIV5 sF protein, resulting in liposome association (45). However, in our present study heating the sF protein to 50°C destroyed its ability to react with neutralizing

MAbs and caused it to form aggregates, both apparently due to denaturation rather than triggering. The difference between the RSV transmembrane F protein and the released sF protein is the membrane anchor, so it is possible that when mild heat is used as a surrogate stimulus, the membrane anchor regulates or stabilizes the re-folding of the F protein, enabling it to re-fold properly, resulting in the posttriggered 6-HB form, while without the anchor, the sF protein denatures.

Here, we have produced a fully cleaved paramyxoviral sF protein in its pretriggered form for the first time without the addition of a foreign trimerization domain. We also identified molarity of the solution containing the sF protein as an unexpected but critical component in the production of the pretriggered RSV sF protein and used it as a novel, surrogate method for triggering a viral fusion protein. Although it is not clear whether this triggering method mimics the physiological triggering of the intact F protein, it confirms that the SC-2 sF protein is produced in the pretriggered form. The availability of pretriggered sF protein should enable studies of the RSV sF protein attachment and triggering mechanisms.

CHAPTER 3: RELEVANCE, ADDITIONAL STUDIES AND FUTURE

STUDIES

STABILIZATION OF THE RSV SOLUBLE F PROTEIN IN ITS

PRETRIGGERED FORM

RSV is known as an unstable virus whose infectivity is easily lost by storage

(except below -70C) and manipulation. One or more of the viral glycoprotein genes is responsible for this instability since replacing them with the baculovirus GP64 glycoprotein results in a virus that is very stable to storage at 4C for 8 weeks, compared to RSV which lost 90% of its infectivity within 2 weeks (149). The F protein is likely to be the glycoprotein responsible for this virion instability because it is highly metastable, readily to undergo major conformational changes following a triggering event. If such triggering happens prematurely, the protein will change the conformation to the posttriggered form before it reaches the target membrane where it functions, leading to the loss of the ability to cause membrane fusion.

We showed in Chapter 2 that the RSV sF protein in the pretriggered form can be generated by simply replacing the TMD and CTD with a 6-his tag (131).

We have found that this, SC-2 sF protein is also somewhat unstable when stored in 800 mM buffer (pH 8.0) at 4ºC, having a half-life of approximately 3 weeks as determined by sucrose gradient analysis (data not shown). It was

76 also triggered by freezing (Fig. 3.1B). Stabilizing the protein in this pretriggered form would be of practical use for storage, shipping, and crystallization.

Fig. 3.1. Analysis of the SC-2 sF protein aggregation state by velocity sedimentation on a sucrose gradient. SC-2 protein was either freshly prepared (A) or kept frozen (B) at -20°C for 2 days prior to analysis. Both preparations were kept at 4°C before loading on a 25% to 55% linear sucrose gradient for ultracentrifugation in an SW41 rotor at 41,000 rpm for 20 h. The protein in each fraction was TCA precipitated, separated by SDS-PAGE and detected by western blot with 5His MAb. “Top” indicates the top fraction of the gradient.

A stabilized pretriggered sF protein may also be the ideal molecule for development as an RSV F protein vaccine for adults. Even though it is not being considered for infants, a protein vaccine would be acceptable for adults because it would not be their initial encounter with RSV. Adults contact RSV repeatedly throughout life and have an established response pattern that 77 could be boosted by a protein vaccine to protect against RSV disease. The alternative that is the prime candidate for infants, an attenuated vaccine, would likely be less immunogenic in these previously infected adults (104).

The F protein is much more conserved than the G protein, the only other neutralizing antigen, and the ideal candidate for that reason. The sF protein could be present in its posttriggered form. However, the sF protein in the pretriggered form would likely present novel antigenic sites found on the infectious virions that are not present in the posttriggered protein. Therefore it might induce significantly more or better quality Abs, more effectively reducing the burden of RSV disease in this population.

1. Stabilization by addition of the trimerization domain to the truncated protein.

The first pretriggered paramyxovirus sF protein, the PIV5 sF protein produced by another group, also lacked both TMD and CTD. However, this sF protein contained a C-terminal stabilizing domain, GCNt, that naturally trimerizes

(220). The PIV5 sF protein could be triggered by the addition of a surrogate trigger, heat (45). The PIV5 sF protein, with its foreign trimerization domain, appeared to be stable at 20°C during the crystallization period, indicating that the GCNt domain can stabilize the truncated protein (220). Despite the fact that addition of trimerization domain is not required for a production of our pretriggered SC-2 sF protein, it might be helpful in stabilizing its pretriggered structure, thereby extending its „shelf-life‟ and therefore its ease of use for some biochemical studies and its value for vaccine studies. We decided to follow the same approach and generated RSV sF proteins with the GCNt 78 domain attached. Two versions were created. The MP-B sF protein was designed to match as closely as possible the PIV5 sF protein, with the GCNt trimerization domain followed by epitope tags, FLAG, Factor Xa cleavage site and 6-his (Fig. 3.2). MP-A sF protein was designed similarly except that an additional protease site was included for future manipulation: a TEV protease site between the F and the GCNt domain.

Fig. 3.2. Constructed RSV sF proteins. 6-his and FLAG tags were included for detection and protein isolation. Factor Xa and TEV are specific protease sites. GCNt is a foreign trimerization domain that stabilizes the sF protein.

The pcDNA3.1 plasmid was used to express these proteins in 293T cells.

Plasmids containing each gene were transfected into 293T cells to analyze expression and secretion efficiency relative to the SC-2. All three sF proteins were detected in the cell lysate (C) and in the culture supernatant (S) at 48 h posttransfection using the 5-his MAb that recognizes the 6-his tag linked to the C-terminus of the F1 fragment (Fig. 3.3). As previously described in the

Chapter 2, the results indicate that the RSV sF proteins with the GCNt domain in both versions were produced, cleaved, and secreted as in a fully cleaved form, but somewhat less efficiently than the wt sF protein. As expected, the GCNt fused sF proteins migrated at a larger size compared to

SC-2 in both the F0 and F1 species due to their C-terminal extensions. The origin of the minor, slightly larger form of the MP-A and MP-B proteins is unknown.

Fig. 3.3. Western blot analysis of the sF proteins produced from transfected 293T cells at 48 h posttransfection. The sF protein from cell lysates (C) and supernatant (S) were stained with the 5His MAb. The C lanes represent 10X more cell equivalents than the S lanes. pcDNA3.1 is the empty vector control.

To determine the protein conformation of the MP-A and MP-B proteins, reactivity of these sF proteins against all 11 neutralizing RSV F MAbs was tested and compared to that of the SC-2 protein. As previous described, transfected cells were metabolically labeled with 35[S] -methionine/cysteine and the medium was immunoprecipitated individually with 11 representing 4 antigenic sites with multiple epitopes in each site. All 11 of these MAbs recognized each of the RSV sF proteins (Fig. 3.4, “+” lane), suggesting that the sF proteins, including those with the trimerization domain attachment, are 81 properly folded. The reduced signal for the MP-A and MP-B sF proteins compared to the SC-2 was also detected when the 5-his MAb was used to precipitate all 3 proteins, reflecting the somewhat lower yield of the MP-A and

MP-B protein, as found in Fig. 3.3.

In Chapter 2 we showed that heating at 50°C induced a dramatic conformational change in the SC-2 protein that resulted in the loss of nearly all reactivity by MAbs representing each antigenic site ((131); Fig. 3.4, “-” lane). To determine if heat has the same effect on the MP-A and MP-B protein, the sF proteins were radioactively labeled and the cell culture medium containing them was heated to 50°C for 1 h before immunoprecipitation with the same set of the F MAbs. We found that heating had little to no effect on the ability of these MAbs to recognize MP-A or MP-B while it destroyed nearly all reactivity of these MAbs with the SC-2 sF protein.

These results suggest that addition of the GCNt trimerization domain stabilizes the structure of the RSV sF protein. Consistent with this finding, when we analyzed the unheated and heated MP-A sF protein by sucrose gradient analysis, we found no difference in the protein migration pattern: both samples remained near the top of the gradient (Fig. 3.5). This result then confirms that following heat treatment, the MP-A and MP-B sF-GCNt protein molecules remain in the unaggregated, pretriggered form. This is surprising since the PIV5 sF protein with the same GCNt structural modification, after a similar heat treatment adopted the posttriggered conformation as determined by the gain of reactivity to a MAb specific to the posttriggered form and aggregation detected by sucrose gradient and by EM (44). Heat, then, serves 82 as a surrogate trigger for the PIV5 sF protein linked to GCNt, but not to the

RSV sF protein linked to the same peptide. It is possible that differences in the triggering mechanism may be responsible for this difference. However, it is also possible that the difference in buffer was responsible. The MP-A and

MP-B sF protein were in 800 mM buffer while PIV5 sF was in 210 to 500 mM buffer, depending on the report. The PIV5 sF protein was treated with trypsin in 100 mM phosphate buffer at 25°C for 30 min before heating, to cleave the sF protein, releasing the fusion peptide. Without trypsin treatment the PIV5 sF protein was not triggered by heat. It is possible that the trypsin treatment also cleaved one or more of sF monomers within the GCNt domain since there are three single basic residues within the first half of this domain. Such a cleavage might release that momomer from the stabilization of the GCNt domain and allow it to trigger.

Fig. 3.4. MAb immunoprecipitation of the SC-2 sF protein after incubation at 4°C or 50°C for 1 h. 293T cells were transfected with plasmids expressing SC-2 sF protein and metabolically labeled with 35S- methionine/cysteine. The medium from these cells was incubated for 1 h at

4°C or 50°C followed by immunoprecipitation with individual MAbs. The antigenic sites for the MAbs that have been mapped by competition (8) are indicated across the top.

Fig. 3.5. Analysis of the MP-A sF protein aggregation state by velocity sedimentation on a sucrose gradient. Freshly prepared MP-A protein was incubated at 4°C or 50°C for 1 h before loading on a 25% to 55% linear sucrose gradient for ultracentrifugation in an SW41 rotor at 41,000 rpm for 20 h. The protein in each fraction was TCA precipitated, separated by SDS-

PAGE and detected by western blot with 5His MAb. “Top” indicates the top fraction of the gradient.

Since MP-A and MP-B appear to be more stable than the SC-2 sF protein. To further characterize these two sF proteins, we will perform EM analysis to confirm their morphology. If it appears to be in the pretriggered form as we predict, we will expose them to low molarity and assess if this exposure causes triggering as it does with the SC-2 sF protein (Fig. 2.6), by EM and liposome association. If they are triggered by low molarity even though their

C-terminus is stabilized by the GCNt domain, it would suggest that triggering does not initiate at the bottom of the pretriggered head as has been suggested for the PIV5 sF protein (45, 179, 220). If they are not triggered by low molarity, then it is likely that low molarity triggers from the bottom of the

85 pretriggered head. Identifying the site of triggering is a critical step in understanding the physiological triggering function.

For cell attachment studies we may treat MP-A and MP-B with the factor Xa protease to remove the 6His tag from their C termini because the 6His tag has been reported to nonspecifically bind to cellular molecules. Factor Xa protease will not remove the FLAG tag which we can use to immunoprecipitate. These sF proteins may enable us to identify the receptor for the F protein on the surface of target cells. For MP-A, treatment with TEV protease would remove both the GCNt domain and the 6His tag. This sF protein would have been produced and secreted in its GCNt stablized, pretriggered conformation. Removal of GCNt may result in a more natural pretriggered sF protein than one produced without any anchor (SC-2). We will then be in an ideal position to examine the effect of low molarity buffer on the sF protein with and without the GCNt stabilizing domain. If the MP-A protein is not triggered by low molarity unless it is first digested with TEV protease, it would suggest that triggering initiates at the bottom of the pretriggered head as has been suggested for the PIV5 sF protein (45, 179,

220). This experiment may enable us to locate the site of triggering.

The MP-A and MP-B sF proteins, which are stabilized in the pretriggered form by the GCNt domain, may be good candidates for many additional studies, such as crystallization for X-ray structural determination, as vaccines for adults, and as immunogens to develop MAbs specific to pretriggered form of

RSV F protein that will enable more detailed studies of F protein triggering events. 86

2. Improving stabilization the sF protein by switching from strain D53 to

A2. The sF protein sequence that we have used in all of these studies is derived from the recombinant D53 strain of RSV. We have also used this

RSV strain for nearly all of the studies in our laboratory. D53 is identical to the more commonly studied A2 strain of RSV except for three AAs within the F protein. We have recently found that D53 virions are dramatically less stable to freezing at -80°C than A2 virions. Therefore, one or more of the AAs that differ in the F proteins of these strains must be responsible for this instability.

The 3 AAs that differ between the two F strains are K66E, Q101P, and

Y342F, in which the first AA is from D53 and the second is A2. AAs 66 and

101 are located within the F2 peptide and 342 is located near the middle of the F1 sequence (Fig. 3.6). By changing all 3 of these AAs in the SC-2 (D53- derived) sF protein, we have created the A2 sF protein. We have also mutated each of these three AAs individually, in two of the three possible combinations in the SC-2 gene for a total of 6 sF mutants. SM-1, SM-2, and

SM-3 are constructs with single AA changes in the first, second, and third positions, respectively. SM-5 and SM-6 represent two of the three pairs of AA changes. SM-7 contains all three of the AA changes and therefore represents the A2 sF protein.

All of the mutant sF genes transfected into 293T cells displayed efficient protein production and secretion, comparable to SC-2 (Fig. 3.7A). They were secreted as fully cleaved proteins. The SM-7 construct has also been tested for its aggregation state by sucrose gradient analysis compared to the SC-2 sF protein. Both the SC-2 and SM-7 sF proteins were found in fractions 2-4 87

(Fig. 3.7B), near the top of the gradient, consistent with the pretriggered form.

This result will be confirmed by EM.

Next, we will compare the D53 and the A2 sF proteins for their ability to withstand -80°C freezing, examining them after one freeze-thaw cycle by EM and sucrose velocity gradient. We will also test these sF proteins for their stability to triggering by heating for 1 h at various temperatures. The heating experiment will be performed in HBSS, a 150 mM physiological buffer, rather than in the 800 mM buffer that the sF protein is in when it is released from the

Ni2+ column. In the 800 mM buffer, the sF protein appears to be stabilized in the pretriggered form. After heating at 50°C, the SC-2 sF protein denatures

(Fig. 2.3 and 2.4B). We will also compare the stability of these two sF proteins during storage at 4°C in the 800 mM and in the HBSS buffers.

We will also compare the triggering effects of low molarity buffer on both sF proteins. If low molarity is the physiological trigger, following exposure to low molarity buffer we would expect both sF proteins to change their conformation, exposing their FP as a result of protein triggering and forming rosettes determined by EM and associating with liposomes, but the A2 sF protein may require a lower molarity or a longer time to trigger. If the two sF proteins differ in any of these tests, the single and double mutants will be used to identify the AA or AAs responsible.

If the A2 sF protein, or any of the intermediate sF protein mutants, are more stable than SC-2, these mutants may be more effective in the crystallization, biochemical triggering, or vaccination experiments described above.

Fig. 3.6. Alignment of the F protein amino acid sequences of strain D53 and A2. The AAs that differ between the 2 strains are in color.

Fig. 3.7. Secretion efficiency and sucrose gradient analysis of SC-2 sF protein and mutants containing one, two or all three differences with A2 sF protein. A. Western blot analysis of the sF mutant proteins produced from transfected 293T cells at 48 h posttransfection. The sF proteins from cell lysates (C) and culture supernatant (S) were stained with the 5-his MAb. The

C lanes represent 10X more cell equivalents than the S lanes. pcDNA3.1 is the empty vector control. B. Analysis of the SC-2 (D53) and SM-7 (A2) sF protein aggregation state by velocity sedimentation on a sucrose gradient as described in the legend to Fig. 2.5. “Top” indicates the top fraction of the gradient.

MAXIMIZING PRODUCTION OF THE RSV SOLUBLE F PROTEIN FOR

CRYSTALLIZATION

To facilitate the structural and functional studies of the RSV F protein, the crystal structure of the RSV F protein in both pre and posttriggered forms would be very useful. To accomplish that ultimate goal, there are several things to consider and optimize, including the stability, quantity, and purity of the sF protein. Two approaches to increase sF protein stability were discussed above. Here we will discuss the quantity of sF protein produced in our system and its purity.

We have used transient expression to produce the SC-2 sF protein by transfecting 293T cells in a 150 mm tissue culture dishes with the SC-2 plasmid. At 48 h posttransfection, the medium was collected and purified on a

Ni2+ column as described in the Materials and Methods section of the Chapter

2. The sF protein was readily detected by SDS-PAGE and Coomassie blue staining (Fig. 3.8). The reduced sF protein migrated at 50 kDa, consistent with the sF1 molecule. The non-reduced sF protein migrated at 70kDa, consistent with the sF1-F2 disulfide linked monomer. The total yield of partially purified sF protein obtained from three 150 mm plates was approximately 1.0 mg as determined by the BCA protein assay. A similar amount was obtained in a second harvest at 72 h posttransfection. However, in this sF preparation we also detected minor contaminants. One protein at 66 kDa is probably albumin, a major component of the fetal bovine serum added to the media. The minor proteins smaller than sF1 are likely breakdown products from the sF protein.

Fig. 3.8. Coomassie Blue staining analysis. Ni2+ column purified RSV SC-2 sF protein analyzed by SDS-PAGE in the presence and absence of 2- mercaptoethanol and stained with Coomassie Blue. Serial 2-fold dilutions of the sF protein were loaded.

The albumin must be removed as it can interfere with the crystallization process. We will attempt to remove it by several methods. We will grow cells in the absence of serum after transfection, replace serum with a non-serum supplement, purify the sF protein on the Ni2+ column twice to minimize this contaminant, or pass the purified sF protein over a Cibacron Blue 3GA agarose (Sigma Aldrich) which efficiently binds and removes the albumin.

And to avoid breakdown products, we will add protease inhibitors to buffers used in each purification step. However, if the breakdown occurs intracellularly, this may be difficult to avoid.

Our current ability to produce 1.0-1.5 mg of the sF protein from three p150 dishes will not be sufficient for the crystallization for which 10-20 mg of the sF protein at the concentration between 10-20 mg/ml is needed. We could increase the amount of sF protein produced by transfecting more cells, but maximizing protein production from each cell may make production more efficient. We have developed and tested several strategies to enhance sF protein production.

The MP-A sF protein is probably the best candidate for crystallization due to its GCNt stabilized, pretriggered conformation at least as determined by the mild heat treatment (Fig. 3.4, 3.5). We have also found that MP-A is stable at

4°C for many months which should facilitate crystallization attempts.

However, both GCNt versions of the sF protein were found to be produced and secreted in somewhat lower amounts, approximately 2-fold less, than

SC-2 that does not contain the GCNt domain (Fig. 3.3). First, to enhance the level of the sF protein expression from the transfected mammalian cells, we moved the MP-A sF gene from its original expression vector, pcDNA3.1, in which the expression of each gene is driven by a CMV promoter to the pCAGGs.mcs vector (112). This mammalian expression vector has a chicken

β-actin promoter fused with the core CMV promoter that drives high levels of gene expression.

We compared production of MP-A from pcDNA3.1 and pCAGGs.mcs, in transiently transfected 293T cells, harvesting at 48 h posttransfection (Fig.

3.9). MP-A sF protein was efficiently produced and secreted into the medium by both plasmids. Although the blots are over-exposed making quantification 93 difficult, we estimate that approximately twice as much MP-A sF protein was produced from pCAGGs.mcs. While this level of production remained below that of the SC-2 sF protein, a two-fold increase in MP-A sF protein production would reduce the number of plates needed 50%. Therefore, we will use this plasmid for MP-A sF production.

Fig. 3.9. Western blot analysis of MP-A sF protein expressed from two different plasmids. At 48 h posttransfection 293T cell lysates (C) and supernatants (S) were harvested and western blotted with the 5His MAb. The

C lanes represent 10X the cell equivalents of the S lanes.

In order to express the RSV F protein from the nucleus, we and others (201,

202) had found that the coding sequence had to be optimized to remove cryptic splice and polyadenylation sites. We were given the optimized D53 F gene by Peter Collins (NIH) and we used that gene for all of the experiments described above. Because we had observed that infection with the Long

94 strain of RSV caused more cell-cell fusion than infection with the D53 strain, we hypothesized that the Long strain F protein may be more active.

To test our hypothesis, we generated a completely synthetic, codon-optimized

RSV sF gene that would express the same AA sequence as the F protein of the RSV Long strain. We designed the optimized Long F gene by substituting the viral codons with codons preferentially found in mammalian cell (87). A total of 28 oligonucleotides, 80-120 nt in length, were created to match this sequence, each having at least a 17 nt overlap with the neighboring oligonucleotide. The oligonucleotides were designed such that their 3‟ termini were good PCR primers. Eight fragments of the F gene, indicated in Fig.

3.10A, were produced by mixing 6 to 8 overlapping oligonucleotides and performing PCR to fill in and amplifying the fragments. The correctly sized fragments were isolated by agarose gel electrophoresis, combined and subjected to PCR, including primers representing the ends of the genome to generate the complete synthetic gene. A full length template was created from the first round PCR, which became a template for amplification using two small flanking primers (Fig. 3.10A). We cloned this synthetic gene codon- optimized version of the Long strain F gene (SC-5) into pcDNA3.1 and confirmed its sequence.

Fig. 3.10. Generation of the codon-optimized RSV Long strain F protein from synthetic oligonucleotides and analysis of its fusion activity. A.

Diagram of the 6 overlapping F gene fragments, each produced from 4-6 synthetic oligonucleotides that were used to create the synthetic RSV Long stain F gene by PCR. B. Cell-cell fusion activity of the D53 (MP340) and Long

(SC-5) strains of the RSV F protein, examined at 23 h posttransfection. Each

F plasmid was co-transfected with the GFP-expressing plasmid pTracer-GFP to facilitate visualization.

To test the fusogenicity of the codon-optimized RSV Long strain synthetic F gene, its ability to cause cell-cell fusion was compared to that of the codon- optimized D53 gene. Each F protein construct was co-transfected into 293T cells with a GFP-expressing plasmid that enables visualization and photography of cells that were successfully transfected. Cells expressing the

F protein on their surface would fuse with the neighboring cells and appear as oversized, multinucleated cells, called syncytia. At 23 h posttransfection, it appeared that the RSV D53 F protein clearly caused a greater amount of fusion than the RSV Long F protein (Fig. 3.10B). In a separate, similar experiment, fusion of both sets of transfected cells was observed at 72 h posttransfection (data not shown), indicating a possible ability of the Long strain F protein to cause fusion, but with delayed kinetics. Because of its higher fusogenicity, the codon-optimized D53 F gene (MP340) was chosen for our studies and used as a template for the construction of the RSV sF protein.

It remains unclear why the Long strain F protein is more fusogenic than the

D53 strain when it is expressed in infected cells but less fusogenic when it is expressed from a plasmid. Immunostaining of these transfected cells demonstrated that both optimized F proteins were produced and localized to the cell surface (data not shown). However, we have not quantified the amount of F protein that is produced or that reaches the cell surface in the transfected cells or quantified fusion caused by both F proteins over time. It is possible that the codon optimization of the Long strain F gene was not as effective in enhancing translation of the F protein as the D53 strain. We used 97 the Genbank sequence for the Long strain F gene to produce this optimized F gene. Others in our laboratory have PCR-amplified the F gene from Long strain (American Type Culture Collection) virions and found minor sequence variations. One of these variants may have caused the efficient fusion noted in the Long strain-infected cells, but that sequence may not be the one in

Genbank. It is also possible that co-expression of another viral protein during infection, most likely the G attachment glycoprotein, might enhance the fusion activity of the Long strain F protein more effectively than the G protein from the D53 strain does.

The delay in cell-cell fusion caused by the Long strain F protein may suggest that it is more stable than the D53 F protein, a quality that may be useful for crystallization. We will produce a sF protein from this Long strain F protein and test its stability. If it is significantly enhanced, we will attempt to crystallize it for structural determinations.

APPLICATION OF THE SOLUBLE F PROTEIN IN SCREENING

LIBRARIES OF COMPOUNDS FOR THEIR ABILITY TO TARGET THE F

PROTEIN

Here we have produced a fully cleaved paramyxoviral sF protein in its pretriggered form for the first time without the addition of a foreign trimerization domain. We also identified molarity of the solution containing the sF protein as an unexpected but critical component in the production of the pretriggered RSV sF protein and used it as a novel, surrogate method for triggering a viral fusion protein. The availability of pretriggered sF protein in a form that can be triggered should enable studies to determine the functions of known anti-RSV F protein compounds, Compounds can be tested for the first time to determine how they block F protein function: by causing the F protein to trigger prematurely, by preventing triggering, or by preventing 6-HB formation. In addition, we may be able to use the sF protein to identify the position of antiviral drug binding and their effects on F protein structure by co- crystallization. In addition to the identification of the binding site for a drug and the mechanism by which it inactivates F protein function, these studies could lead to the identification the F protein trigger. Experiments that can detect changes in protein shape, aggregation state, and exposure of hydrophobic fusion peptide (Chapter 2) can be used to answer these questions.

DETERMINE IF REDUCTION IN MOLARITY IS THE PHYSIOLOGICAL

CAUSE OF F PROTEIN TRIGGERING

It is not clear what the trigger is for the full-length F protein in the virion membrane to initiate the dramatic conformational change, in which the protein converts to the posttriggered form while causing membrane fusion. We identified, for the first time, that exposure to the low molarity buffer can trigger the sF protein. We demonstrated that exposure to 110 mM or lower molarity buffer triggered the sF protein, causing nearly all molecules to convert from the spherical shape to hat-pins that aggregate into rosettes, However, the same triggering effect was minor when the protein was exposed to the 260 mM buffer, indicating that the SC-2 sF protein may be triggered by molarity that is higher than 110 mM but lower than 260 mM. Physiological molarity is

150 mM and our finding that triggering occurs around the physiological range suggests that reduction in buffer molarity may be the actual physiological trigger of the F protein in a natural infection. A reduction in buffer molarity would require an enclosed compartment, such as endosome. In one study, the clathrin-mediated endocytosis pathway and an early endosome compartment in particular, was shown to be essential for RSV infectivity

(115). This is quite surprising since RSV has always been thought to enter the cell by direct fusion between the viral envelope and the target cell membrane and the entry is pH independent. Because cell-cell fusion must take place at the plasma membrane, it has long been assumed that virus-cell fusion would take place in a similar manner and location.

100

Taken together, we propose a novel viral entry/ triggering pathway in which the virion enters the cell through the endocytic pathway. Following endosomal uptake, ion pumps and channels in the endosome membrane lower the overall molarity, or perhaps the molarity of a crucial ion, and the F protein is triggered. Once triggered, the F protein will fuse the viral envelope with the endosomal membrane and the viral nucleocapsid content is released into the cytoplasm where the viral genome is transcribed and replicates.

To test this hypothesis, we first should narrow the range of molarity that causes the obvious change in protein structure as a result of triggering. A series of 0.8X, 1.0X, and 1.2X of physiological molarity buffer (150 mM) will be used to test their triggering effect on the SC-2 protein. If the protein is triggered by 0.8X buffer only, it would indicate that when molarity is reduced below the physiological point, then the sF protein is triggered. If the sF protein is triggered in this manner, the full length F protein might also be, within an endosome. In addition, if a reduction in molarity is the physiological cause of

F protein triggering, it should enhance cell-cell fusion of the cells expressing the F protein. If F-expressing cells fuse more at the lower physiological molarity point, the molarity may serve as a physiological trigger. And such fusion activity should be dampened when cells are treated with 1.2X media, as a result of high molarity stabilizing the F protein, thus prevent triggering.

Viral infectivity should also increase as a result of enhanced membrane fusion activity of the F protein if low molarity medium is applied.

If endocytosis and ion pumps play some role in the RSV entry/ F protein triggering as we proposed, blockage of ion pumps by drugs would result in 101 decreased viral infectivity, indicating that the physiological change in ionic strength causes the F protein to trigger and it is needed to initiate infection.

To test the involvement of channels and pumps in the endocytic pathway in the initiation of RSV infection, we could also use siRNAs to knock down expression of critical genes in a manner similar to the study that demonstrated the crucial role for endocytosis (115). The siRNA should cause fewer side effects on the other cellular pathway than inhibitory drugs, therefore the role of the buffer molarity on the fusion activity of the F protein could be determined.

102

ANALYSIS OF THE MαH ON THE RSV F PROTEIN FOR ITS ROLE IN

FUSION FUNCTION.

Unlike most other paramyxoviruses, RSV and human metapneumovirus

(hMPV), members of the Pneumovirinae subfamily do not require their attachment glycoprotein (G) for fusion or infectivity in cell culture (7, 13, 101,

197). Their F proteins apparently have the ability to catalyze membrane fusion and support viral spread on its own, suggesting that it is capable of attachment and triggering. However, the sites responsible for these functions have not been identified. To cause membrane fusion, the pretriggered form of the F protein is converted to the posttriggered form. The crystal structures of one example of each form, the posttriggered form of the PIV3 F protein and the pretriggered PIV5 F protein, have been solved (219, 220). While the actual crystal structure of the RSV F protein in both forms are not yet available, to help us better understand the structural change during protein triggering and domain (s) involved in that critical step, the availability of the molecular models of the RSV F protein in both pre and posttriggered forms would be very useful.

Dr. Will Ray, our collaborator on this project, has used the two crystal structures of PIV3 and PIV5 F proteins and sequence alignment to develop molecular models of the RSV F protein in both conformations. Modeling was facilitated by the abundance of conserved cysteine residues and the constraints resulting from disulfide bonding. The monomer models of the pretriggered and posttriggered RSV F protein are presented here with his permission in Fig. 3.11A and 3.11B, respectively. 103

Our confidence in these models is enhanced by the fact that they not only placed the cysteines that are conserved between the RSV F and PIV3 F proteins appropriately, but that the 6 cysteines not present in the PIV F proteins are placed in appropriate proximity to form disulfide bonds in both models, and the 4 cysteines present in the parent proteins but not in the RSV

F protein are replaced in both models with amino acids that would form salt bridges, preserving a bond in those positions. In addition, the proposed structure of the 6HB in our posttriggered model was very similar to the previously solved crystal structure of the RSV F protein 6HB (126, 138, 226).

The pretriggered model suggests a critical domain. As shown in the pretriggered model, the central portion of the F protein (Fig. 3.11A) remains relatively constant during the transition (marked as the “head” because it becomes the top of the posttriggered form, Fig. 3.11B). In the pretriggered conformation at the upper left region, the pre-HRA region (blue in Fig. 3.11A) is composed of three short α-helices and the two loop regions (L1 and L2) that connect them. The pre-HRA is presented in blue that becomes lighter in tone with increasing distance from the fusion peptide (white). In the posttriggered structure, L1 and L2 have dramatically rearranged, changing from random coils and β-sheets into α-helices to complete the long HRA α- helix in the posttriggered form (blue in Fig. 3.11B). The formation of HRA would both expose the fusion peptide and thrust it upwards, into the target cell membrane. The long HRA α-helix would likely trimerize at this point, since the HRA peptide naturally forms a trimer in solution. The entire molecule would then fold in half, bringing its two ends together, like a jackknife. The 104

HRB helices would fit into the grooves of the HRA trimer to form the 6HB, bringing the two membranes into contact to initiate membrane fusion. The fusion peptide would be below the HRA in the posttriggered form shown in

Fig. 3.11B

105

Fig. 3.11. Models of the pretriggered (A) and posttriggered (B) RSV F protein monomer. In both, the cysteines and the disulfide bonds they form are yellow and the 2 N-linked glycosylation sites are pink. A third N-linked site would be at the N-terminus of F2 if the previous amino acid, Asn, had been included in the structure. The pretriggered form is based on the PIV5 F protein structure (220). The posttriggered form is based on the PIV3 F0 protein structure (219). These models were produced by Dr. Will Ray and are used with his permission.

106

A comparison of these structures clearly indicates that a dramatic refolding event takes place at the top of the molecule, the apically positioned pre-HRA in particular, that converts from three short α-helices into one long α-helix,

HRA. A trimer model of the RSV F protein in a pretriggered form has revealed additional interesting features of this region that might suggest its potential involvement in modulating protein triggering/ function (Fig. 3.12A).

The crown of the pretriggered F protein trimer contains a central pore where the three monomers converge (Fig. 3.12B) that leads into the apparently empty space in the trimer. The middle α-helix (MαH) of the pre-HRA lines this central pore (fishnet in Fig. 3.12C) and would seem to be in the ideal position to function as the F protein trigger for several reasons. 1) Its apical position would ensure that it would be the first surface of the F protein to contact the target cell. 2) The central position of the MαH would enable all three monomers to trigger at the same time. 3) MαH is situated between the two loop regions, L1 and L2, which must refold into α-helices in the posttriggered form (Fig. 3.11A). This central position is ideal for MαH to affect their rearrangement. If the MαH were pulled out of this pocket, it would tension L1 and L2 perhaps initiating their conformational change to α-helices. 4) The location of MαH within the central pore should protect it from antibody binding and the resulting evolutionary pressure. Therefore, the MαH appears to be in the ideal position to serve as the triggering domain that initiates the F protein refolding and fusion function.

107

Fig. 3.12. Model of the pretriggered RSV F protein trimer. One monomer in each is presented in multicolored-cartoon form showing secondary structures, the other two in the space-filling mode (pink and gray). (A) The pretriggered trimer is oriented such that the hole in the side of the head into which the fusion peptide probably slips after cleavage is visible. The stalk is composed of the three HRB domains. (B and C) The top view of the pretriggered F protein trimer. A central pore in the top of the trimer (B) leads to the interior of the head in this space-filling model. The MαH domain

108

(indicated by fishnet) lines the pore (C). The amino acids from this monomer that face the central pore of the trimer are highlight in red.

To determine the importance of the F protein MαH in mediating membrane fusion and to identify the most important amino acids, we separately mutated each of the 7 amino acids of the MαH, 195LKNYIDK, to Ala. Ala substitutions should not affect the α-helical structure of MαH but will reduce the side chain to the minimal, single methyl group, disrupting most contacts. The mutations were made in a codon-optimized version of the complete RSV F protein gene and tested in transfected cells for the ability to cause cell-cell fusion. Because mutants that do not reach the cell surface would also be defective in fusion activity, we also examined these mutants for defects in transport, processing and cell surface expression.

Effects of MαH mutations on transport and processing. To assess the effects of these Ala mutations on F protein processing, transfected cells were labeled with 35[S]-Met/Cys for 1 hr followed by a 1 hr chase with cold methionine to allow newly synthesized and labeled F protein to be processed.

The Ala mutants were all produced and immunoprecipitated with the L4 anti-F

MAb at an efficiency of greater than 60% that of wt F protein, with the exception of L195A which precipitated at 30% the level of wt F protein (Fig.

3.14A, striped bars). Because the antibody used for immunoprecipitation is conformation dependent (151), the fact that it did not precipitate L195A efficiently suggests that the majority of the L195A F protein molecules had not folded properly.

109

Greater than 90% of the wild-type (wt) F protein was in the F1 form (Fig. 3.13,

“-” lane). Because the F protein cleaved by a furin-like enzyme which resides in the trans Golgi, this result indicates that 90% of the wt F protein had reached the trans Golgi. F1 was the major species (70% to 93%) precipitated for all of these mutants (Fig. 3.14A, numbers across the bottom), indicating that the majority these F proteins had reached the trans Golgi. The one exception, again L195A, was cleaved at a much lower, 40% efficiency, indicating that this proportion of its molecules had reached the trans Golgi.

To determine whether the N-linked glycans on these F protein molecules were mature we treated immunoprecipitates with Endoglycosidase H (Endo

H) (Fig. 3.13 “+” lanes). The migration of the wt F1 protein, as well as the F1 protein from all of the mutants was unaffected by Endo H treatment, confirming that these molecules had reached the Golgi where N-linked glycans maturation takes place. The remaining F0 protein in the wt F protein and all mutants was digested by Endo H, resulting in a change in protein migration of the F0 specie. Therefore, it is clear that the glycans on F0 were not mature, confirming that these F0 molecules had not reached the Golgi.

This result was consistent with the uncleaved status of the F0 proteins and confirmed that its lack of cleavage was due to the lack of transport to the

Golgi rather than a direct effect on the process of F protein cleavage.

Effects of MαH mutations on cell surface expression. To determine whether these mutations altered the ability of the F protein to reach the cell surface, we biotinylated the surface of transfected cells in the radiolabeling experiment described above. Ala mutants N197A, Y198A and K201 reached 110 the cell surface with at least 90% that the efficiency of wt F protein (Fig.

3.14A, white bars). Cell surface expression of K196A, I199A and D200A was

40% to 50% that of wt F protein. L195A could not be detected at the cell surface. In an attempt to increase the level of cell surface of these mutant F proteins we increased the amount of transfected DNA by 2-fold. However that approach only slightly increased the cell surface expression of D200A without improving the others (data not shown) and was abandoned. Instead, we have attempted draw conclusions from a comparison of the level of cell surface expression for each mutant, described here, and the level of cell-cell fusion, described below.

Effects of Ala mutations in the MαH on cell-cell fusion. We used a luciferase assay to quantify the cell-cell fusion activity of the mutant F proteins in transfected cells. Two of the 7 Ala mutants, N197A and D200A, retained their fusion activity (Fig. 3.14A, gray bars) relative to their level of cell surface expression (Fig. 3.14A, white bars). D200A reached the cell surface only 45% the level of the wt F protein but was as just as fusogenic, if not more. N197A was either produced at a higher level or reacted with MAb L4 twice as efficiently as the wt F protein, reaching the cell surface even more efficiently and fusing cells yet more efficiently, 4 times as efficiently as wt F. Neither of these mutants are functionally inhibited, and therefore neither of these amino acids are critical for the fusion function of the F protein.

The five remaining F mutants displayed no or very little fusion activity, suggesting that their mutations had lost their ability to drive fusion. Mutants

Y198A and K201A displayed no or very low fusion activity, despite the fact 111 that they both reached the cell surface at 90% that the efficiency of the wt F protein (Fig. 3.14A). These amino acids are clearly required for the fusion function of the F protein. Mutants K196A and I199A reached the cell surface at 40% and 47% the level of the wt F protein but displayed no fusion activity, suggesting that K196 and I199 are also important amino acids in fusion. The lack of fusion by L195A is clearly explained by its lack of cell surface expression. L195 is clearly important for F protein folding or stability, but its importance in the fusion function of the F protein cannot be assessed by this mutant.

Important characteristics of the MαH AAs. We generated several additional mutants in the MαH in an attempt to identify important characteristics on the AAs in critical positions. L195V, unlike L195A, did reach the cell surface at an efficiency of 29% but caused no fusion, suggesting that

L195 may, in fact be critical for the fusion function in addition to protein structure. Another mutation at the same position, L195I reached the cell surface twice as efficiently as wt F, and supported fusion at a higher level than wt F, indicating that that this conservative change was not detrimental.

The central AA in the MαH, Y198, could be changed to Phe, a similar AA with a phenyl ring, with an actual gain in cell surface expression and an even greater increase in fusion activity (Fig. 3.14B), indicating that Phe is more effective in this position than the wt Tyr. Replacement of Y198 with Trp likewise caused enhanced cell fusion (data not shown). The following AA,

I199, could be changed to Tyr, a bulkier hydrophobic AA, with a slight reduction in cell surface expression but with a near complete loss of fusion 112 activity, confirming the importance of AA I199 in fusion that had been suggested by the I199A mutation.

The most conservative mutation at the following position, K201R, maintained the positive charge, actually enhanced cell surface expression to over 200% that of wt F protein and fusion to more than 600% that of wt F. To determine whether the positively charged K201 could be replaced with a negative charge, it was replaced with Asp and Glu. Both K201D and K201E reached the cell surface efficiently but displayed low fusion activity, suggesting that the type of charge is important. Replacement of K201 with Ser, a similarly sized

AA lacking a charge again had no effect on cell surface expression and reduced fusion activity, but only to 30% (Fig. 3.14B).Taken together, these results indicate that fusion is most efficient if AA 201 contain a polar side chain but a positive charge further enhances fusion while no charge (Ala or

Ser), or a negative charge (Asp or Glu) is less supportive of fusion, while retaining a low level of activity.

113

Fig. 3.13. Intracellular processing and cell surface expression of F protein mutants in MαH. 293T cells were radiolabeled with 100 mCi/ml 35[S]

-Met/Cys at 15 hr post-transfection for 1 hr and chased with cold methionine for 1 hr. The cell monolayers were then labeled with cell impermeable biotin.

Cells were lysed and immunoprecipitated with MAb L4. (A) One portion of the immunoprecipitated F protein was treated with Endo H, another was not. Both were separated by SDS-PAGE and detected by autoradiography. (B) A third portion of the same immunoprecipitate was separated by SDS-PAGE, transferred to Immobilon and probed with streptavidin-HRP to detect the biotinylated protein.

114

Fig. 3.14. Comparison of protein production, cell surface expression and cell-cell fusion activity of F protein mutants. All F proteins were expressed from the pcDNA3.1 in 293T cells. Any signal from cells transfected with pcDNA3.1 lacking an insert is subtracted as a background in order to obtain the absolute value of each sample for quantification. Protein production is quantified from absolute signal obtained from immunoprecipitation by MAb L4 after 1 hr of Tran35[S]-Met/Cys pulse-labeling and 1.3 hr chase. Total protein immunoprecipitated (striped bars) is the sum of the F0 and F1 bands after background subtraction. Cell surface expression is determined by

115 biotinylation (white bars). Cell-cell fusion activity is quantified by the luciferase assay (gray bars). All were analyzed in triplicate and normalized to the respective wt F protein activities. The average values are plotted with the standard error of the mean. Mean percentage of protein cleavage is also indicated at the bottom of graph with standard deviation value less than 2% in all cases. Percent of protein cleavage is calculated from the ratio of F1 to a total F0+F1 signal within the mutant itself.

116

The data summary of mutational analysis of residues in this MαH motif is shown in Fig. 3.15A. Red indicates the residues important for protein to function and black indicates unimportant residues. Mutations that resulted in defective protein function or production are located below the sequence and unaffected mutants are located above the sequence.

We have found that 3 of the 7 amino acids (underlined) in the MαH are critical for the fusion function of the F protein, independent of cell surface expression:

195LKNYIDK. Two of the mutants, N197A and D200A, caused efficient fusion relative to their cell surface expression indicating that these amino acids are not critical for F protein function. Mutations of L195 and K196 resulted in a defect in protein production and/ or transport that prevents us from a further examination of their significance in a fusion function. However, they both appear to be critical for the production of protein that can reach the cell surface.

The MαH appears as α-helical conformation in which each turn of the helix involves approximately 3.5 residues. Therefore the 3 residues (Y198, I 199, and K201) identified here as being required for function would line two neighboring faces of the MαH α-helix (Fig. 3.15B, red), while the two amino acids that are not critical would line the third face (Fig. 3.15B, green). One particular face containing the two critical residues, Y198 and K201, is oriented toward the pore in the top of the F protein (red in Fig. 3.15C).

117

Fig. 3.15. Summary of critical residues that form the MαH motif (A) and stereoimage of the MαH in side (B) and top (C) views highlighting the critical amino acids. Individual mutations in 3 (red) of these 7 amino acids destroy fusion activity without affecting protein processing or cell surface appearance. Mutations in the other 2 (red) residues are defective in protein production, processing, or surface transport, suggesting that they are essential in a production of the functional fusion unit. Mutations in the remaining two amino acids (green) did not inhibit fusion, suggesting that these amino acids are not involved in fusion. Note that both green amino acids are on the backside of this helix, while the red amino acids cover the two front faces of the helix. The side chains of the amino acids important for fusion (red) appear to form a groove. Panels B and C are included with the permission of Dr. Will Ray. 118

In summary, we use the PIV3 and PIV5 crystal structures to create computational models for RSV F protein pretriggered and posttriggered forms. From these models, we suggest that the middle of the three short α- helices (MαH) in pre-HRA is in the ideal position to serve as the triggering site that initiates the dramatic F protein refolding leading to membrane fusion. Our

Ala-mutational analysis of the MαH region confirms its importance in the fusion function of the F protein and identifies 3 amino acids critical to this process as measured by cell-cell fusion without affecting cell surface expression, indicating their critical role in protein function. Mutations in two other AAs predicted to be on the side of MaH facing away from the pore had no effect on fusion, suggesting that this face was not important for either structure or function. Mutation in remaining two AAs disrupted cell surface expression suggested a role in protein folding or stability. Additional mutation of these residues indicated an important characteristic required for fusion activity.

119

REFERENCES

1. Anderson, K., A. M. King, R. A. Lerch, and G. W. Wertz. 1992. Polylactosaminoglycan modification of the respiratory syncytial virus small hydrophobic (SH) protein: a conserved feature among human and bovine respiratory syncytial viruses. Virology 191:417-30. 2. Anderson, L. J., J. C. Hierholzer, C. Tsou, R. M. Hendry, B. F. Fernie, Y. Stone, and K. McIntosh. 1985. Antigenic characterization of respiratory syncytial virus strains with monoclonal antibodies. J Infect Dis 151:626-33. 3. Arnold, R., B. Konig, H. Werchau, and W. Konig. 2004. Respiratory syncytial virus deficient in soluble G protein induced an increased proinflammatory response in human lung epithelial cells. Virology 330:384-97. 4. Atreya, P. L., M. E. Peeples, and P. L. Collins. 1998. The NS1 protein of human respiratory syncytial virus is a potent inhibitor of minigenome transcription and RNA replication. J Virol 72:1452-61. 5. Baker, K. A., R. E. Dutch, R. A. Lamb, and T. S. Jardetzky. 1999. Structural basis for paramyxovirus-mediated membrane fusion. Mol Cell 3:309-19. 6. Barr, J., P. Chambers, C. R. Pringle, and A. J. Easton. 1991. Sequence of the major nucleocapsid protein gene of pneumonia virus of mice: sequence comparisons suggest structural homology between nucleocapsid proteins of pneumoviruses, paramyxoviruses, rhabdoviruses and filoviruses. J Gen Virol 72 ( Pt 3):677-85. 7. Barretto, N., L. K. Hallak, and M. E. Peeples. 2003. Neuraminidase treatment of respiratory syncytial virus-infected cells or virions, but not target cells, enhances cell-cell fusion and infection. Virology 313:33- 43. 8. Beeler, J. A., and K. van Wyke Coelingh. 1989. Neutralization epitopes of the F glycoprotein of respiratory syncytial virus: effect of mutation upon fusion function. J Virol 63:2941-50. 9. Begona Ruiz-Arguello, M., L. Gonzalez-Reyes, L. J. Calder, C. Palomo, D. Martin, M. J. Saiz, B. Garcia-Barreno, J. J. Skehel, and J. A. Melero. 2002. Effect of proteolytic processing at two distinct sites on shape and aggregation of an anchorless fusion protein of human respiratory syncytial virus and fate of the intervening segment. Virology 298:317-26. 10. Behera, A. K., H. Matsuse, M. Kumar, X. Kong, R. F. Lockey, and S. S. Mohapatra. 2001. Blocking intercellular adhesion molecule-1 on human epithelial cells decreases respiratory syncytial virus infection. Biochem Biophys Res Commun 280:188-95. 120

11. Bermingham, A., and P. L. Collins. 1999. The M2-2 protein of human respiratory syncytial virus is a regulatory factor involved in the balance between RNA replication and transcription. Proc Natl Acad Sci U S A 96:11259-64. 12. Biacchesi, S., M. H. Skiadopoulos, K. C. Tran, B. R. Murphy, P. L. Collins, and U. J. Buchholz. 2004. Recovery of human metapneumovirus from cDNA: optimization of growth in vitro and expression of additional genes. Virology 321:247-59. 13. Biacchesi, S., M. H. Skiadopoulos, L. Yang, E. W. Lamirande, K. C. Tran, B. R. Murphy, P. L. Collins, and U. J. Buchholz. 2004. Recombinant human Metapneumovirus lacking the small hydrophobic SH and/or attachment G glycoprotein: deletion of G yields a promising vaccine candidate. J Virol 78:12877-87. 14. Bisgaard, H. 2003. A randomized trial of montelukast in respiratory syncytial virus postbronchiolitis. Am J Respir Crit Care Med 167:379- 83. 15. Bossart, K. N., L. F. Wang, M. N. Flora, K. B. Chua, S. K. Lam, B. T. Eaton, and C. C. Broder. 2002. Membrane fusion tropism and heterotypic functional activities of the Nipah virus and Hendra virus envelope glycoproteins. J Virol 76:11186-98. 16. Bossert, B., S. Marozin, and K. K. Conzelmann. 2003. Nonstructural proteins NS1 and NS2 of bovine respiratory syncytial virus block activation of interferon regulatory factor 3. J Virol 77:8661-8. 17. Brock, S. C., J. R. Goldenring, and J. E. Crowe, Jr. 2003. Apical recycling systems regulate directional budding of respiratory syncytial virus from polarized epithelial cells. Proc Natl Acad Sci U S A 100:15143-8. 18. Brown, G., C. E. Jeffree, T. McDonald, H. W. Rixon, J. D. Aitken, and R. J. Sugrue. 2004. Analysis of the interaction between respiratory syncytial virus and lipid-rafts in Hep2 cells during infection. Virology 327:175-85. 19. Buchholz, U. J., S. Biacchesi, Q. N. Pham, K. C. Tran, L. Yang, C. L. Luongo, M. H. Skiadopoulos, B. R. Murphy, and P. L. Collins. 2005. Deletion of M2 gene open reading frames 1 and 2 of human metapneumovirus: effects on RNA synthesis, attenuation, and immunogenicity. J Virol 79:6588-97. 20. Buckland, R., E. Malvoisin, P. Beauverger, and F. Wild. 1992. A leucine zipper structure present in the measles virus fusion protein is not required for its tetramerization but is essential for fusion. J Gen Virol 73 ( Pt 7):1703-7. 21. Bukreyev, A., S. S. Whitehead, B. R. Murphy, and P. L. Collins. 1997. Recombinant respiratory syncytial virus from which the entire SH gene has been deleted grows efficiently in cell culture and exhibits site- specific attenuation in the respiratory tract of the mouse. J Virol 71:8973-82. 22. Bukreyev, A., L. Yang, J. Fricke, L. Cheng, J. M. Ward, B. R. Murphy, and P. L. Collins. 2008. The secreted form of respiratory 121

syncytial virus G glycoprotein helps the virus evade antibody-mediated restriction of replication by acting as an antigen decoy and through effects on Fc receptor-bearing leukocytes. J Virol 82:12191-204. 23. Bullough, P. A., F. M. Hughson, J. J. Skehel, and D. C. Wiley. 1994. Structure of influenza haemagglutinin at the pH of membrane fusion. Nature 371:37-43. 24. Burke, E., L. Dupuy, C. Wall, and S. Barik. 1998. Role of cellular actin in the gene expression and morphogenesis of human respiratory syncytial virus. Virology 252:137-48. 25. Burke, E., N. M. Mahoney, S. C. Almo, and S. Barik. 2000. Profilin is required for optimal actin-dependent transcription of respiratory syncytial virus genome RNA. J Virol 74:669-75. 26. Byrappa, S., D. K. Gavin, and K. C. Gupta. 1995. A highly efficient procedure for site-specific mutagenesis of full- length plasmids using Vent DNA polymerase. Genome Res 5:404-7. 27. Calder, L. J., L. Gonzalez-Reyes, B. Garcia-Barreno, S. A. Wharton, J. J. Skehel, D. C. Wiley, and J. A. Melero. 2000. Electron microscopy of the human respiratory syncytial virus fusion protein and complexes that it forms with monoclonal antibodies. Virology 271:122- 31. 28. Carr, C. M., C. Chaudhry, and P. S. Kim. 1997. Influenza hemagglutinin is spring-loaded by a metastable native conformation. Proc Natl Acad Sci U S A 94:14306-13. 29. Chaiwatpongsakorn, S., R. F. Epand, P. L. Collins, R. M. Epand, and M. E. Peeples. 2011. Soluble respiratory syncytial virus fusion protein in the fully cleaved, pretriggered state is triggered by exposure to low-molarity buffer. J Virol 85:3968-77. 30. Chan, D. C., D. Fass, J. M. Berger, and P. S. Kim. 1997. Core structure of gp41 from the HIV envelope glycoprotein. Cell 89:263-73. 31. Chan, D. C., and P. S. Kim. 1998. HIV entry and its inhibition. Cell 93:681-4. 32. Chen, L., P. M. Colman, L. J. Cosgrove, M. C. Lawrence, L. J. Lawrence, P. A. Tulloch, and J. J. Gorman. 2001. Cloning, expression, and crystallization of the fusion protein of Newcastle disease virus. Virology 290:290-9. 33. Chen, L., J. J. Gorman, J. McKimm-Breschkin, L. J. Lawrence, P. A. Tulloch, B. J. Smith, P. M. Colman, and M. C. Lawrence. 2001. The structure of the fusion glycoprotein of Newcastle disease virus suggests a novel paradigm for the molecular mechanism of membrane fusion. Structure (Camb) 9:255-66. 34. Cheng, X., H. Park, H. Zhou, and H. Jin. 2005. Overexpression of the M2-2 protein of respiratory syncytial virus inhibits viral replication. J Virol 79:13943-52. 35. Cohen, F. S., and G. B. Melikyan. 2004. The energetics of membrane fusion from binding, through hemifusion, pore formation, and pore enlargement. J Membr Biol 199:1-14.

122

36. Collins, P. L., E. Camargo, and M. G. Hill. 1999. Support plasmids and support proteins required for recovery of recombinant respiratory syncytial virus. Virology 259:251-5. 37. Collins, P. L., and J. E. J. Crowe. 2007. Respiratory syncytial virus and metapneumovirus, p. 1601-1646. In D. M. Knipe and P. M. Howley (ed.), Fields Virology, 5th ed, vol. 2. Lippincott Williams & Wilkins, Philadelphia. 38. Collins, P. L., and B. S. Graham. 2008. Viral and host factors in human respiratory syncytial virus pathogenesis. J Virol 82:2040-55. 39. Collins, P. L., M. G. Hill, E. Camargo, H. Grosfeld, R. M. Chanock, and B. R. Murphy. 1995. Production of infectious human respiratory syncytial virus from cloned cDNA confirms an essential role for the transcription elongation factor from the 5' proximal open reading frame of the M2 mRNA in gene expression and provides a capability for vaccine development. Proc Natl Acad Sci U S A 92:11563-7. 40. Collins, P. L., and G. Mottet. 1993. Membrane orientation and oligomerization of the small hydrophobic protein of human respiratory syncytial virus. J Gen Virol 74 ( Pt 7):1445-50. 41. Collins, P. L., and B. R. Murphy. 2005. New generation live vaccines against human respiratory syncytial virus designed by reverse genetics. Proc Am Thorac Soc 2:166-73. 42. Collins, P. L., and G. W. Wertz. 1983. cDNA cloning and transcriptional mapping of nine polyadenylylated RNAs encoded by the genome of human respiratory syncytial virus. Proc Natl Acad Sci U S A 80:3208-12. 43. Collins, P. L. a. J. E. C., Jr. 2007. Respiratory syncytial virus and metapneumovirus, p. 1601-1646. In D. M. Knipe, Howley, P.M. (ed.), Fields Virology, vol. 2. Lippincott Williams & Wilkins, Philadelphia. 44. Connolly, S. A., G. P. Leser, T. S. Jardetzky, and R. A. Lamb. 2009. Bimolecular complementation of paramyxovirus fusion and hemagglutinin-neuraminidase proteins enhances fusion: implications for the mechanism of fusion triggering. J Virol 83:10857-68. 45. Connolly, S. A., G. P. Leser, H. S. Yin, T. S. Jardetzky, and R. A. Lamb. 2006. Refolding of a paramyxovirus F protein from prefusion to postfusion conformations observed by liposome binding and electron microscopy. Proc Natl Acad Sci U S A 103:17903-8. 46. Cowton, V. M., D. R. McGivern, and R. Fearns. 2006. Unravelling the complexities of respiratory syncytial virus RNA synthesis. J Gen Virol 87:1805-21. 47. Crim, R. L., S. A. Audet, S. A. Feldman, H. S. Mostowski, and J. A. Beeler. 2007. Identification of linear heparin-binding peptides derived from human respiratory syncytial virus fusion glycoprotein that inhibit infectivity. J Virol 81:261-71. 48. Crowe, J. E., Jr., C. Y. Firestone, S. S. Whitehead, P. L. Collins, and B. R. Murphy. 1996. Acquisition of the ts phenotype by a chemically mutagenized cold- passaged human respiratory syncytial

123

virus vaccine candidate results from the acquisition of a single mutation in the polymerase (L) gene. Virus Genes 13:269-73. 49. Crowe, J. E., Jr., C. Y. Firestone, S. S. Whitehead, P. L. Collins, and B. R. Murphy. 1996. Acquisition of the ts phenotype by a chemically mutagenized cold-passaged human respiratory syncytial virus vaccine candidate results from the acquisition of a single mutation in the polymerase (L) gene. Virus Genes 13:269-73. 50. Damico, R. L., J. Crane, and P. Bates. 1998. Receptor-triggered membrane association of a model retroviral glycoprotein. Proc Natl Acad Sci U S A 95:2580-5. 51. Day, N. D., P. J. Branigan, C. Liu, L. L. Gutshall, J. Luo, J. A. Melero, R. T. Sarisky, and A. M. Del Vecchio. 2006. Contribution of cysteine residues in the extracellular domain of the F protein of human respiratory syncytial virus to its function. Virol J 3:34. 52. Deng, R., Z. Wang, A. M. Mirza, and R. M. Iorio. 1995. Localization of a domain on the paramyxovirus attachment protein required for the promotion of cellular fusion by its homologous fusion protein spike. Virology 209:457-69. 53. Dominy, B. N., D. Perl, F. X. Schmid, and C. L. Brooks, 3rd. 2002. The effects of ionic strength on protein stability: the cold shock protein family. J Mol Biol 319:541-54. 54. Doms, R. W., A. Helenius, and J. White. 1985. Membrane fusion activity of the influenza virus hemagglutinin. The low pH-induced conformational change. J Biol Chem 260:2973-81. 55. Dudas, R. A., and R. A. Karron. 1998. Respiratory syncytial virus vaccines. Clin Microbiol Rev 11:430-9. 56. Durbin, J. E., T. R. Johnson, R. K. Durbin, S. E. Mertz, R. A. Morotti, R. S. Peebles, and B. S. Graham. 2002. The role of IFN in respiratory syncytial virus pathogenesis. J Immunol 168:2944-52. 57. Ebata, S. N., M. J. Cote, C. Y. Kang, and K. Dimock. 1991. The fusion and hemagglutinin-neuraminidase glycoproteins of human parainfluenza virus 3 are both required for fusion. Virology 183:437-41. 58. Eckert, D. M., and P. S. Kim. 2001. Mechanisms of viral membrane fusion and its inhibition. Annu Rev Biochem 70:777-810. 59. Eckert, D. M., V. N. Malashkevich, L. H. Hong, P. A. Carr, and P. S. Kim. 1999. Inhibiting HIV-1 entry: discovery of D-peptide inhibitors that target the gp41 coiled-coil pocket. Cell 99:103-15. 60. Elcock, A. H., and J. A. McCammon. 1998. Electrostatic contributions to the stability of halophilic proteins. J Mol Biol 280:731-48. 61. Evans, J. E., P. A. Cane, and C. R. Pringle. 1996. Expression and characterisation of the NS1 and NS2 proteins of respiratory syncytial virus. Virus Res 43:155-61. 62. Falsey, A. R. 2005. Respiratory syncytial virus infection in elderly and high-risk adults. Exp Lung Res 31 Suppl 1:77. 63. Falsey, A. R., P. A. Hennessey, M. A. Formica, C. Cox, and E. E. Walsh. 2005. Respiratory syncytial virus infection in elderly and high- risk adults. N Engl J Med 352:1749-59. 124

64. Falsey, A. R., and E. E. Walsh. 1996. Safety and immunogenicity of a respiratory syncytial virus subunit vaccine (PFP-2) in ambulatory adults over age 60. Vaccine 14:1214-8. 65. Falsey, A. R., and E. E. Walsh. 1997. Safety and immunogenicity of a respiratory syncytial virus subunit vaccine (PFP-2) in the institutionalized elderly. Vaccine 15:1130-2. 66. Falsey, A. R., E. E. Walsh, J. Capellan, S. Gravenstein, M. Zambon, E. Yau, G. J. Gorse, R. Edelman, F. G. Hayden, J. E. McElhaney, K. M. Neuzil, K. L. Nichol, E. A. Simoes, P. F. Wright, and V. M. Sales. 2008. Comparison of the safety and immunogenicity of 2 respiratory syncytial virus (rsv) vaccines--nonadjuvanted vaccine or vaccine adjuvanted with alum--given concomitantly with influenza vaccine to high-risk elderly individuals. J Infect Dis 198:1317-26. 67. Fass, D., S. C. Harrison, and P. S. Kim. 1996. Retrovirus envelope domain at 1.7 angstrom resolution. Nat Struct Biol 3:465-9. 68. Fearns, R., M. E. Peeples, and P. L. Collins. 1997. Increased expression of the N protein of respiratory syncytial virus stimulates minigenome replication but does not alter the balance between the synthesis of mRNA and antigenome. Virology 236:188-201. 69. Feldman, S. A., S. Audet, and J. A. Beeler. 2000. The fusion glycoprotein of human respiratory syncytial virus facilitates virus attachment and infectivity via an interaction with cellular heparan sulfate. J Virol 74:6442-7. 70. Fleming, E. H., A. A. Kolokoltsov, R. A. Davey, J. E. Nichols, and N. J. Roberts, Jr. 2006. Respiratory syncytial virus F envelope protein associates with lipid rafts without a requirement for other virus proteins. J Virol 80:12160-70. 71. Follett, E. A., C. R. Pringle, and T. H. Pennington. 1975. Virus development in enucleate cells: echovirus, poliovirus, pseudorabies virus, reovirus, respiratory syncytial virus and Semliki Forest virus. J Gen Virol 26:183-96. 72. Fuentes, S., K. C. Tran, P. Luthra, M. N. Teng, and B. He. 2007. Function of the respiratory syncytial virus small hydrophobic protein. J Virol 81:8361-6. 73. Gallo, S. A., C. M. Finnegan, M. Viard, Y. Raviv, A. Dimitrov, S. S. Rawat, A. Puri, S. Durell, and R. Blumenthal. 2003. The HIV Env- mediated fusion reaction. Biochim Biophys Acta 1614:36-50. 74. Garcia-Barreno, B., T. Delgado, and J. A. Melero. 1996. Identification of protein regions involved in the interaction of human respiratory syncytial virus phosphoprotein and nucleoprotein: significance for nucleocapsid assembly and formation of cytoplasmic inclusions. J Virol 70:801-8. 75. Garcia-Beato, R., I. Martinez, C. Franci, F. X. Real, B. Garcia- Barreno, and J. A. Melero. 1996. Host cell effect upon glycosylation and antigenicity of human respiratory syncytial virus G glycoprotein. Virology 221:301-9.

125

76. Garcin, D., T. Pelet, P. Calain, L. Roux, J. Curran, and D. Kolakofsky. 1995. A highly recombinogenic system for the recovery of infectious Sendai paramyxovirus from cDNA: generation of a novel copy-back nondefective interfering virus. Embo J 14:6087-94. 77. Ghildyal, R., C. Baulch-Brown, J. Mills, and J. Meanger. 2003. The matrix protein of Human respiratory syncytial virus localises to the nucleus of infected cells and inhibits transcription. Arch Virol 148:1419- 29. 78. Ghildyal, R., A. Ho, M. Dias, L. Soegiyono, P. G. Bardin, K. C. Tran, M. N. Teng, and D. A. Jans. 2009. The respiratory syncytial virus matrix protein possesses a Crm1-mediated nuclear export mechanism. J Virol 83:5353-62. 79. Ghildyal, R., J. Mills, M. Murray, N. Vardaxis, and J. Meanger. 2002. Respiratory syncytial virus matrix protein associates with nucleocapsids in infected cells. J Gen Virol 83:753-7. 80. Ghosh, J. K., S. G. Peisajovich, M. Ovadia, and Y. Shai. 1998. Structure-function study of a heptad repeat positioned near the transmembrane domain of Sendai virus fusion protein which blocks virus-cell fusion. J Biol Chem 273:27182-90. 81. Glezen, P., and F. W. Denny. 1973. Epidemiology of acute lower respiratory disease in children. N Engl J Med 288:498-505. 82. Glickman, R. L., R. J. Syddall, R. M. Iorio, J. P. Sheehan, and M. A. Bratt. 1988. Quantitative basic residue requirements in the cleavage- activation site of the fusion glycoprotein as a determinant of virulence for Newcastle disease virus. J Virol 62:354-6. 83. Gonzalez-Reyes, L., M. B. Ruiz-Arguello, B. Garcia-Barreno, L. Calder, J. A. Lopez, J. P. Albar, J. J. Skehel, D. C. Wiley, and J. A. Melero. 2001. Cleavage of the human respiratory syncytial virus fusion protein at two distinct sites is required for activation of membrane fusion. Proc Natl Acad Sci U S A 98:9859-9864. 84. Gotoh, B., T. Komatsu, K. Takeuchi, and J. Yokoo. 2001. Paramyxovirus accessory proteins as interferon antagonists. Microbiol Immunol 45:787-800. 85. Gower, T. L., M. K. Pastey, M. E. Peeples, P. L. Collins, L. H. McCurdy, T. K. Hart, A. Guth, T. R. Johnson, and B. S. Graham. 2005. RhoA signaling is required for respiratory syncytial virus-induced syncytium formation and filamentous virion morphology. J Virol 79:5326-36. 86. Graham, B. S. 2011. Biological challenges and technological opportunities for respiratory syncytial virus vaccine development. Immunol Rev 239:149-66. 87. Haas, J., E. C. Park, and B. Seed. 1996. Codon usage limitation in the expression of HIV-1 envelope glycoprotein. Curr Biol 6:315-24. 88. Hall, C. B., E. E. Walsh, C. E. Long, and K. C. Schnabel. 1991. Immunity to and frequency of reinfection with respiratory syncytial virus. J Infect Dis 163:693-8.

126

89. Hallak, L. K., P. L. Collins, W. Knudson, and M. E. Peeples. 2000. Iduronic acid-containing glycosaminoglycans on target cells are required for efficient respiratory syncytial virus infection. Virology 271:264. 90. Hardy, R. W., and G. W. Wertz. 1998. The product of the respiratory syncytial virus M2 gene ORF1 enhances readthrough of intergenic junctions during viral transcription. J Virol 72:520-6. 91. Henderson, G., J. Murray, and R. P. Yeo. 2002. Sorting of the respiratory syncytial virus matrix protein into detergent-resistant structures is dependent on cell-surface expression of the glycoproteins. Virology 300:244-54. 92. Hernandez, L. D., R. J. Peters, S. E. Delos, J. A. Young, D. A. Agard, and J. M. White. 1997. Activation of a retroviral membrane fusion protein: soluble receptor- induced liposome binding of the ALSV envelope glycoprotein. J Cell Biol 139:1455-64. 93. Hernandez, L. D., R. J. Peters, S. E. Delos, J. A. Young, D. A. Agard, and J. M. White. 1997. Activation of a retroviral membrane fusion protein: soluble receptor-induced liposome binding of the ALSV envelope glycoprotein. J Cell Biol 139:1455-64. 94. Horvath, C. M. 2004. Silencing STATs: lessons from paramyxovirus interferon evasion. Cytokine Growth Factor Rev 15:117-27. 95. Hu, X. L., R. Ray, and R. W. Compans. 1992. Functional interactions between the fusion protein and hemagglutinin-neuraminidase of human parainfluenza viruses. J Virol 66:1528-34. 96. Jardetzky, T. S., and R. A. Lamb. 2004. Virology: a class act. Nature 427:307-8. 97. Jiang, S., K. Lin, N. Strick, and A. R. Neurath. 1993. Inhibition of HIV-1 infection by a fusion domain binding peptide from the HIV-1 envelope glycoprotein GP41. Biochem Biophys Res Commun 195:533-8. 98. Jin, H., H. Zhou, X. Cheng, R. Tang, M. Munoz, and N. Nguyen. 2000. Recombinant respiratory syncytial viruses with deletions in the NS1, NS2, SH, and M2-2 genes are attenuated in vitro and in vivo. Virology 273:210-8. 99. Johnson, P. R., Jr., R. A. Olmsted, G. A. Prince, B. R. Murphy, D. W. Alling, E. E. Walsh, and P. L. Collins. 1987. Antigenic relatedness between glycoproteins of human respiratory syncytial virus subgroups A and B: evaluation of the contributions of F and G glycoproteins to immunity. J Virol 61:3163-6. 100. Johnson, S., C. Oliver, G. A. Prince, V. G. Hemming, D. S. Pfarr, S. C. Wang, M. Dormitzer, J. O'Grady, S. Koenig, J. K. Tamura, R. Woods, G. Bansal, D. Couchenour, E. Tsao, W. C. Hall, and J. F. Young. 1997. Development of a humanized monoclonal antibody (MEDI-493) with potent in vitro and in vivo activity against respiratory syncytial virus. J Infect Dis 176:1215-24. 101. Kahn, J. S., M. J. Schnell, L. Buonocore, and J. K. Rose. 1999. Recombinant vesicular stomatitis virus expressing respiratory syncytial 127

virus (RSV) glycoproteins: RSV fusion protein can mediate infection and cell fusion. Virology 254:81-91. 102. Kallewaard, N. L., A. L. Bowen, and J. E. Crowe, Jr. 2005. Cooperativity of actin and microtubule elements during replication of respiratory syncytial virus. Virology 331:73-81. 103. Karron, R. A., D. A. Buonagurio, A. F. Georgiu, S. S. Whitehead, J. E. Adamus, M. L. Clements-Mann, D. O. Harris, V. B. Randolph, S. A. Udem, B. R. Murphy, and M. S. Sidhu. 1997. Respiratory syncytial virus (RSV) SH and G proteins are not essential for viral replication in vitro: clinical evaluation and molecular characterization of a cold- passaged, attenuated RSV subgroup B mutant. Proc Natl Acad Sci U S A 94:13961-6. 104. Karron, R. A., P. F. Wright, R. B. Belshe, B. Thumar, R. Casey, F. Newman, F. P. Polack, V. B. Randolph, A. Deatly, J. Hackell, W. Gruber, B. R. Murphy, and P. L. Collins. 2005. Identification of a recombinant live attenuated respiratory syncytial virus vaccine candidate that is highly attenuated in infants. J Infect Dis 191:1093- 104. 105. Karron, R. A., P. F. Wright, J. E. Crowe, Jr., M. L. Clements-Mann, J. Thompson, M. Makhene, R. Casey, and B. R. Murphy. 1997. Evaluation of two live, cold-passaged, temperature-sensitive respiratory syncytial virus vaccines in chimpanzees and in human adults, infants, and children. J Infect Dis 176:1428-36. 106. Kato, A., Y. Sakai, T. Shioda, T. Kondo, M. Nakanishi, and Y. Nagai. 1996. Initiation of Sendai virus multiplication from transfected cDNA or RNA with negative or positive sense. Genes Cells 1:569-79. 107. Kellner, J. D., A. Ohlsson, A. M. Gadomski, and E. E. Wang. 2000. Bronchodilators for bronchiolitis. Cochrane Database Syst Rev:CD001266. 108. Kilgore, N. R., K. Salzwedel, M. Reddick, G. P. Allaway, and C. T. Wild. 2003. Direct evidence that C-peptide inhibitors of human immunodeficiency virus type 1 entry bind to the gp41 N-helical domain in receptor-activated viral envelope. J Virol 77:7669-72. 109. Kim, H. W., J. O. Arrobio, C. D. Brandt, B. C. Jeffries, G. Pyles, J. L. Reid, R. M. Chanock, and R. H. Parrott. 1973. Epidemiology of respiratory syncytial virus infection in Washington, D.C. I. Importance of the virus in different respiratory tract disease syndromes and temporal distribution of infection. Am J Epidemiol 98:216-25. 110. Kim, H. W., J. G. Canchola, C. D. Brandt, G. Pyles, R. M. Chanock, K. Jensen, and R. H. Parrott. 1969. Respiratory syncytial virus disease in infants despite prior administration of antigenic inactivated vaccine. Am J Epidemiol 89:422-34. 111. Kneyber, M. C., and J. L. Kimpen. 2004. Advances in respiratory syncytial virus vaccine development. Curr Opin Investig Drugs 5:163- 70. 112. Kobasa, D., M. E. Rodgers, K. Wells, and Y. Kawaoka. 1997. Neuraminidase hemadsorption activity, conserved in avian influenza A 128

viruses, does not influence viral replication in ducks. J Virol 71:6706- 13. 113. Kochva, U., H. Leonov, and I. T. Arkin. 2003. Modeling the structure of the respiratory syncytial virus small hydrophobic protein by silent- mutation analysis of global searching molecular dynamics. Protein Sci 12:2668-74. 114. Kohn, W. D., C. M. Kay, and R. S. Hodges. 1997. Salt effects on protein stability: two-stranded alpha-helical coiled-coils containing inter- or intrahelical ion pairs. J Mol Biol 267:1039-52. 115. Kolokoltsov, A. A., D. Deniger, E. H. Fleming, N. J. Roberts, Jr., J. M. Karpilow, and R. A. Davey. 2007. Small interfering RNA profiling reveals key role of clathrin-mediated endocytosis and early endosome formation for infection by respiratory syncytial virus. J Virol 81:7786- 800. 116. Krempl, C., B. R. Murphy, and P. L. Collins. 2002. Recombinant respiratory syncytial virus with the G and F genes shifted to the promoter-proximal positions. J Virol 76:11931-42. 117. Kuo, L., R. Fearns, and P. L. Collins. 1997. Analysis of the gene start and gene end signals of human respiratory syncytial virus: quasi- templated initiation at position 1 of the encoded mRNA. J Virol 71:4944-53. 118. Kuo, L., R. Fearns, and P. L. Collins. 1996. The structurally diverse intergenic regions of respiratory syncytial virus do not modulate sequential transcription by a dicistronic minigenome. J Virol 70:6143- 50. 119. Kuo, L., H. Grosfeld, J. Cristina, M. G. Hill, and P. L. Collins. 1996. Effects of mutations in the gene-start and gene-end sequence motifs on transcription of monocistronic and dicistronic minigenomes of respiratory syncytial virus. J Virol 70:6892-901. 120. Kwilas, A. R., M. A. Yednak, L. Zhang, R. Liesman, P. L. Collins, R. J. Pickles, and M. E. Peeples. 2010. Respiratory syncytial virus engineered to express the cystic fibrosis transmembrane conductance regulator corrects the bioelectric phenotype of human cystic fibrosis airway epithelium in vitro. J Virol 84:7770-81. 121. Kwilas, S., R. M. Liesman, L. Zhang, E. Walsh, R. J. Pickles, and M. E. Peeples. 2009. Respiratory syncytial virus grown in Vero cells contains a truncated attachment protein that alters its infectivity and dependence on glycosaminoglycans. J Virol 83:10710-8. 122. Lamb, R. A. 1993. Paramyxovirus fusion: a hypothesis for changes. Virology 197:1-11. 123. Lambert, D. M., S. Barney, A. L. Lambert, K. Guthrie, R. Medinas, D. E. Davis, T. Bucy, J. Erickson, G. Merutka, and S. R. Petteway, Jr. 1996. Peptides from conserved regions of paramyxovirus fusion (F) proteins are potent inhibitors of viral fusion. Proc Natl Acad Sci U S A 93:2186-91.

129

124. Lampinen, M., M. Carlson, L. D. Hakansson, and P. Venge. 2004. Cytokine-regulated accumulation of eosinophils in inflammatory disease. Allergy 59:793-805. 125. Langley, J. M., V. Sales, A. McGeer, R. Guasparini, G. Predy, W. Meekison, M. Li, J. Capellan, and E. Wang. 2009. A dose-ranging study of a subunit Respiratory Syncytial Virus subtype A vaccine with and without aluminum phosphate adjuvantation in adults > or =65 years of age. Vaccine 27:5913-9. 126. Lawless-Delmedico, M. K., P. Sista, R. Sen, N. C. Moore, J. B. Antczak, J. M. White, R. J. Greene, K. C. Leanza, T. J. Matthews, and D. M. Lambert. 2000. Heptad-repeat regions of respiratory syncytial virus F1 protein form a six-membered coiled-coil complex. Biochemistry 39:11684-95. 127. Li, J., V. R. Melanson, A. M. Mirza, and R. M. Iorio. 2005. Decreased dependence on receptor recognition for the fusion promotion activity of L289A-mutated newcastle disease virus fusion protein correlates with a monoclonal antibody-detected conformational change. J Virol 79:1180-90. 128. Liao, M., C. Sanchez-San Martin, A. Zheng, and M. Kielian. 2010. In vitro reconstitution reveals key intermediate states of trimer formation by the dengue virus membrane fusion protein. J Virol 84:5730-40. 129. Ling, Z., K. C. Tran, and M. N. Teng. 2009. Human respiratory syncytial virus nonstructural protein NS2 antagonizes the activation of beta interferon transcription by interacting with RIG-I. J Virol 83:3734- 42. 130. Liuzzi, M., S. W. Mason, M. Cartier, C. Lawetz, R. S. McCollum, N. Dansereau, G. Bolger, N. Lapeyre, Y. Gaudette, L. Lagace, M. J. Massariol, F. Do, P. Whitehead, L. Lamarre, E. Scouten, J. Bordeleau, S. Landry, J. Rancourt, G. Fazal, and B. Simoneau. 2005. Inhibitors of respiratory syncytial virus replication target cotranscriptional mRNA guanylylation by viral RNA-dependent RNA polymerase. J Virol 79:13105-15. 131. Lorusso, V., A. Paradiso, M. Guida, F. Berardi, and M. De Lena. 1991. Ifosfamide plus mitoxantrone as salvage treatment in non- Hodgkin's lymphomas. Am J Clin Oncol 14:492-5. 132. Malashkevich, V. N., D. C. Chan, C. T. Chutkowski, and P. S. Kim. 1998. Crystal structure of the simian immunodeficiency virus (SIV) gp41 core: conserved helical interactions underlie the broad inhibitory activity of gp41 peptides. Proc Natl Acad Sci U S A 95:9134-9. 133. Malhotra, R., M. Ward, H. Bright, R. Priest, M. R. Foster, M. Hurle, E. Blair, and M. Bird. 2003. Isolation and characterisation of potential respiratory syncytial virus receptor(s) on epithelial cells. Microbes Infect 5:123-33. 134. Malykhina, O., M. A. Yednak, P. L. Collins, P. D. Olivo, and M. E. Peeples. 2011. A respiratory syncytial virus replicon that is noncytotoxic and capable of long-term foreign gene expression. J Virol 85:4792-801. 130

135. Martinez-Sobrido, L., N. Gitiban, A. Fernandez-Sesma, J. Cros, S. E. Mertz, N. A. Jewell, S. Hammond, E. Flano, R. K. Durbin, A. Garcia-Sastre, and J. E. Durbin. 2006. Protection against respiratory syncytial virus by a recombinant Newcastle disease virus vector. J Virol 80:1130-9. 136. Marty, A., J. Meanger, J. Mills, B. Shields, and R. Ghildyal. 2004. Association of matrix protein of respiratory syncytial virus with the host cell membrane of infected cells. Arch Virol 149:199-210. 137. Mason, S. W., E. Aberg, C. Lawetz, R. DeLong, P. Whitehead, and M. Liuzzi. 2003. Interaction between human respiratory syncytial virus (RSV) M2-1 and P proteins is required for reconstitution of M2-1- dependent RSV minigenome activity. J Virol 77:10670-6. 138. Matthews, J. M., T. F. Young, S. P. Tucker, and J. P. Mackay. 2000. The core of the respiratory syncytial virus fusion protein is a trimeric coiled coil. J Virol 74:5911-20. 139. McLellan, J. S., M. Chen, A. Kim, Y. Yang, B. S. Graham, and P. D. Kwong. 2010. Structural basis of respiratory syncytial virus neutralization by motavizumab. Nat Struct Mol Biol 17:248-50. 140. Melikyan, G. B., R. M. Markosyan, H. Hemmati, M. K. Delmedico, D. M. Lambert, and F. S. Cohen. 2000. Evidence that the transition of HIV-1 gp41 into a six-helix bundle, not the bundle configuration, induces membrane fusion. J Cell Biol 151:413-23. 141. Meyer, K. C. 2010. The role of immunity and inflammation in lung senescence and susceptibility to infection in the elderly. Semin Respir Crit Care Med 31:561-74. 142. Moore, J. P., and R. W. Doms. 2003. The entry of entry inhibitors: a fusion of science and medicine. Proc Natl Acad Sci U S A 100:10598- 602. 143. Morris, J. A. J., R. E. Blount, and R. E. Savage. 1956. Recovery of cytopathogenic agent from chimpanzees with coryza. Proc. Soc. Exp. Biol. Med. 92:544-550. 144. Morrison, T., C. McQuain, T. Sergel, L. McGinnes, and J. Reitter. 1993. The role of the amino terminus of F1 of the Newcastle disease virus fusion protein in cleavage and fusion. Virology 193:997-1000. 145. Munoz, F. M., P. A. Piedra, and W. P. Glezen. 2003. Safety and immunogenicity of respiratory syncytial virus purified fusion protein-2 vaccine in pregnant women. Vaccine 21:3465-7. 146. Murphy, L. B., C. Loney, J. Murray, D. Bhella, P. Ashton, and R. P. Yeo. 2003. Investigations into the amino-terminal domain of the respiratory syncytial virus nucleocapsid protein reveal elements important for nucleocapsid formation and interaction with the phosphoprotein. Virology 307:143-53. 147. Netter, R. C., S. M. Amberg, J. W. Balliet, M. J. Biscone, A. Vermeulen, L. J. Earp, J. M. White, and P. Bates. 2004. Heptad repeat 2-based peptides inhibit avian sarcoma and leukosis virus subgroup a infection and identify a fusion intermediate. J Virol 78:13430-9. 131

148. Olmsted, R. A., and P. L. Collins. 1989. The 1A protein of respiratory syncytial virus is an integral membrane protein present as multiple, structurally distinct species. J Virol 63:2019-29. 149. Oomens, A. G., and G. W. Wertz. 2004. The baculovirus GP64 protein mediates highly stable infectivity of a human respiratory syncytial virus lacking its homologous transmembrane glycoproteins. J Virol 78:124-35. 150. Pal, S. R., and K. C. Das. 1976. Respiratory syncytial virus-specific RNA synthesis in primary monkey kidney cell cultures. Acta Virol 20:253-6. 151. Paradiso, P. R., B. T. Hu, R. Arumugham, and S. Hildreth. 1991. Mapping of a fusion related epitope of the respiratory syncytial virus fusion protein. Vaccine 9:231-7. 152. Pastey, M. K., J. E. Crowe, Jr., and B. S. Graham. 1999. RhoA interacts with the fusion glycoprotein of respiratory syncytial virus and facilitates virus-induced syncytium formation. J Virol 73:7262-70. 153. Patel, H., R. W. Platt, G. S. Pekeles, and F. M. Ducharme. 2002. A randomized, controlled trial of the effectiveness of nebulized therapy with epinephrine compared with albuterol and saline in infants hospitalized for acute viral bronchiolitis. J Pediatr 141:818-24. 154. Paterson, R. G., C. J. Russell, and R. A. Lamb. 2000. Fusion protein of the paramyxovirus SV5: destabilizing and stabilizing mutants of fusion activation. Virology 270:17-30. 155. Peeples, M. 1991. Paramyxovirus M proteins: Pulling it all together and taking it on the road. Plenum Press, New York. 156. Perez, M., B. Garcia-Barreno, J. A. Melero, L. Carrasco, and R. Guinea. 1997. Membrane permeability changes induced in Escherichia coli by the SH protein of human respiratory syncytial virus. Virology 235:342-51. 157. Piedra, P. A., S. G. Cron, A. Jewell, N. Hamblett, R. McBride, M. A. Palacio, R. Ginsberg, C. M. Oermann, and P. W. Hiatt. 2003. Immunogenicity of a new purified fusion protein vaccine to respiratory syncytial virus: a multi-center trial in children with cystic fibrosis. Vaccine 21:2448-60. 158. Poch, O., B. M. Blumberg, L. Bougueleret, and N. Tordo. 1990. Sequence comparison of five polymerases (L proteins) of unsegmented negative-strand RNA viruses: theoretical assignment of functional domains. J Gen Virol 71 ( Pt 5):1153-62. 159. Poch, O., I. Sauvaget, M. Delarue, and N. Tordo. 1989. Identification of four conserved motifs among the RNA-dependent polymerase encoding elements. EMBO J 8:3867-74. 160. Pohlmann, S., and J. D. Reeves. 2006. Cellular entry of HIV: Evaluation of therapeutic targets. Curr Pharm Des 12:1963-73. 161. Poirier, M. A., W. Xiao, J. C. Macosko, C. Chan, Y. K. Shin, and M. K. Bennett. 1998. The synaptic SNARE complex is a parallel four- stranded helical bundle. Nat Struct Biol 5:765-9.

132

162. Polack, F. P., M. N. Teng, P. L. Collins, G. A. Prince, M. Exner, H. Regele, D. D. Lirman, R. Rabold, S. J. Hoffman, C. L. Karp, S. R. Kleeberger, M. Wills-Karp, and R. A. Karron. 2002. A role for immune complexes in enhanced respiratory syncytial virus disease. J Exp Med 196:859-65. 163. Pollack, P., and J. R. Groothuis. 2002. Development and use of palivizumab (Synagis): a passive immunoprophylactic agent for RSV. J Infect Chemother 8:201-6. 164. Power, U. F., T. N. Nguyen, E. Rietveld, R. L. de Swart, J. Groen, A. D. Osterhaus, R. de Groot, N. Corvaia, A. Beck, N. Bouveret-Le- Cam, and J. Y. Bonnefoy. 2001. Safety and immunogenicity of a novel recombinant subunit respiratory syncytial virus vaccine (BBG2Na) in healthy young adults. J Infect Dis 184:1456-60. 165. Prince, G. A., V. G. Hemming, R. L. Horswood, and R. M. Chanock. 1985. Immunoprophylaxis and immunotherapy of respiratory syncytial virus infection in the cotton rat. Virus Res 3:193-206. 166. Radecke, F., P. Spielhofer, H. Schneider, K. Kaelin, M. Huber, C. Dotsch, G. Christiansen, and M. A. Billeter. 1995. Rescue of measles viruses from cloned DNA. Embo J 14:5773-84. 167. Ramaswamy, M., L. Shi, S. M. Varga, S. Barik, M. A. Behlke, and D. C. Look. 2006. Respiratory syncytial virus nonstructural protein 2 specifically inhibits type I interferon signal transduction. Virology 344:328-39. 168. Randolph, A. G., and E. E. Wang. 2000. Ribavirin for respiratory syncytial virus infection of the lower respiratory tract. Cochrane Database Syst Rev:CD000181. 169. Rao, J. K., and P. Argos. 1981. Structural stability of halophilic proteins. Biochemistry 20:6536-43. 170. Rawling, J., O. Cano, D. Garcin, D. Kolakofsky, and J. A. Melero. 2011. Recombinant Sendai viruses expressing fusion proteins with two furin cleavage sites mimic the syncytial and receptor-independent infection properties of respiratory syncytial virus. J Virol 85:2771-80. 171. Rawling, J., B. Garcia-Barreno, and J. A. Melero. 2008. Insertion of the two cleavage sites of the respiratory syncytial virus fusion protein in Sendai virus fusion protein leads to enhanced cell-cell fusion and a decreased dependency on the HN attachment protein for activity. J Virol 82:5986-98. 172. Record, M. T., Jr., W. Zhang, and C. F. Anderson. 1998. Analysis of effects of salts and uncharged solutes on protein and nucleic acid equilibria and processes: a practical guide to recognizing and interpreting polyelectrolyte effects, Hofmeister effects, and osmotic effects of salts. Adv Protein Chem 51:281-353. 173. Reitter, J. N., T. Sergel, and T. G. Morrison. 1995. Mutational analysis of the leucine zipper motif in the Newcastle disease virus fusion protein. J Virol 69:5995-6004. 174. Rixon, H. W., G. Brown, J. Aitken, T. McDonald, S. Graham, and R. J. Sugrue. 2004. The small hydrophobic (SH) protein accumulates 133

within lipid-raft structures of the Golgi complex during respiratory syncytial virus infection. J Gen Virol 85:1153-65. 175. Roberts, S. R., R. W. Compans, and G. W. Wertz. 1995. Respiratory syncytial virus matures at the apical surfaces of polarized epithelial cells. J Virol 69:2667-73. 176. Roberts, S. R., D. Lichtenstein, L. A. Ball, and G. W. Wertz. 1994. The membrane-associated and secreted forms of the respiratory syncytial virus attachment glycoprotein G are synthesized from alternative initiation codons. J Virol 68:4538-46. 177. Ruiz-Arguello, M. B., D. Martin, S. A. Wharton, L. J. Calder, S. R. Martin, O. Cano, M. Calero, B. Garcia-Barreno, J. J. Skehel, and J. A. Melero. 2004. Thermostability of the human respiratory syncytial virus fusion protein before and after activation: implications for the membrane-fusion mechanism. J Gen Virol 85:3677-87. 178. Russell, C. J., T. S. Jardetzky, and R. A. Lamb. 2001. Membrane fusion machines of paramyxoviruses: capture of intermediates of fusion. Embo J 20:4024-34. 179. Russell, C. J., K. L. Kantor, T. S. Jardetzky, and R. A. Lamb. 2003. A dual-functional paramyxovirus F protein regulatory switch segment: activation and membrane fusion. J Cell Biol 163:363-74. 180. Sanchez-San Martin, C., H. Sosa, and M. Kielian. 2008. A stable prefusion intermediate of the alphavirus fusion protein reveals critical features of class II membrane fusion. Cell Host Microbe 4:600-8. 181. Sanchez-Seco, M. P., J. Navarro, R. Martinez, and N. Villanueva. 1995. C-terminal phosphorylation of human respiratory syncytial virus P protein occurs mainly at serine residue 232. J Gen Virol 76:425-30. 182. Schmidt, A. C., D. R. Wenzke, J. M. McAuliffe, M. St Claire, W. R. Elkins, B. R. Murphy, and P. L. Collins. 2002. Mucosal immunization of rhesus monkeys against respiratory syncytial virus subgroups A and B and human parainfluenza virus type 3 by using a live cDNA-derived vaccine based on a host range-attenuated bovine parainfluenza virus type 3 vector backbone. J Virol 76:1089-99. 183. Schornberg, K., S. Matsuyama, K. Kabsch, S. Delos, A. Bouton, and J. White. 2006. Role of endosomal cathepsins in entry mediated by the Ebola virus glycoprotein. J Virol 80:4174-8. 184. Schowalter, R. M., S. E. Smith, and R. E. Dutch. 2006. Characterization of human metapneumovirus F protein-promoted membrane fusion: critical roles for proteolytic processing and low pH. J Virol 80:10931-41. 185. Sergel-Germano, T., C. McQuain, and T. Morrison. 1994. Mutations in the fusion peptide and heptad repeat regions of the Newcastle disease virus fusion protein block fusion. J Virol 68:7654-8. 186. Sergel, T. A., L. W. McGinnes, and T. G. Morrison. 2000. A single amino acid change in the Newcastle disease virus fusion protein alters the requirement for HN protein in fusion. J Virol 74:5101-7. 187. Seth, S., A. Vincent, and R. W. Compans. 2003. Mutations in the cytoplasmic domain of a paramyxovirus fusion glycoprotein rescue 134

syncytium formation and eliminate the hemagglutinin-neuraminidase protein requirement for membrane fusion. J Virol 77:167-78. 188. Shay, D. K., R. C. Holman, R. D. Newman, L. L. Liu, J. W. Stout, and L. J. Anderson. 1999. Bronchiolitis-associated hospitalizations among US children, 1980-1996. Jama 282:1440-6. 189. Slack, M. S., and A. J. Easton. 1998. Characterization of the interaction of the human respiratory syncytial virus phosphoprotein and nucleocapsid protein using the two-hybrid system. Virus Res 55:167- 76. 190. Spann, K. M., K. C. Tran, B. Chi, R. L. Rabin, and P. L. Collins. 2004. Suppression of the induction of alpha, beta, and lambda interferons by the NS1 and NS2 proteins of human respiratory syncytial virus in human epithelial cells and macrophages [corrected]. J Virol 78:4363-9. 191. Spann, K. M., K. C. Tran, and P. L. Collins. 2005. Effects of nonstructural proteins NS1 and NS2 of human respiratory syncytial virus on interferon regulatory factor 3, NF-kappaB, and proinflammatory cytokines. J Virol 79:5353-62. 192. Stec, D. S., M. G. d. Hill, and P. L. Collins. 1991. Sequence analysis of the polymerase L gene of human respiratory syncytial virus and predicted phylogeny of nonsegmented negative-strand viruses. Virology 183:273-87. 193. Sutton, R. B., D. Fasshauer, R. Jahn, and A. T. Brunger. 1998. Crystal structure of a SNARE complex involved in synaptic exocytosis at 2.4 A resolution. Nature 395:347-53. 194. Takimoto, T., J. L. Hurwitz, C. Coleclough, C. Prouser, S. Krishnamurthy, X. Zhan, K. Boyd, R. A. Scroggs, B. Brown, Y. Nagai, A. Portner, and K. S. Slobod. 2004. Recombinant Sendai virus expressing the G glycoprotein of respiratory syncytial virus (RSV) elicits immune protection against RSV. J Virol 78:6043-7. 195. Tanabayashi, K., and R. W. Compans. 1996. Functional interaction of paramyxovirus glycoproteins: identification of a domain in Sendai virus HN which promotes cell fusion. J Virol 70:6112-8. 196. Techaarpornkul, S., N. Barretto, and M. E. Peeples. 2001. Functional analysis of recombinant respiratory syncytial virus deletion mutants lacking the small hydrophobic and/or attachment glycoprotein gene. J Virol 75:6825-34. 197. Techaarpornkul, S., P. L. Collins, and M. E. Peeples. 2002. Respiratory syncytial virus with the fusion protein as its only viral glycoprotein is less dependent on cellular glycosaminoglycans for attachment than complete virus. Virology 294:296-304. 198. Teng, M. N., and P. L. Collins. 1999. Altered growth characteristics of recombinant respiratory syncytial viruses which do not produce NS2 protein. J Virol 73:466-73. 199. Teng, M. N., S. S. Whitehead, A. Bermingham, M. St Claire, W. R. Elkins, B. R. Murphy, and P. L. Collins. 2000. Recombinant respiratory syncytial virus that does not express the NS1 or M2-2 135

protein is highly attenuated and immunogenic in chimpanzees. J Virol 74:9317-21. 200. Teng, M. N., S. S. Whitehead, and P. L. Collins. 2001. Contribution of the respiratory syncytial virus G glycoprotein and its secreted and membrane-bound forms to virus replication in vitro and in vivo. Virology 289:283-96. 201. Ternette, N., D. Stefanou, S. Kuate, K. Uberla, and T. Grunwald. 2007. Expression of RNA virus proteins by RNA polymerase II dependent expression plasmids is hindered at multiple steps. Virology Journal 4:51. 202. Ternette, N., B. Tippler, K. Uberla, and T. Grunwald. 2007. Immunogenicity and efficacy of codon optimized DNA vaccines encoding the F-protein of respiratory syncytial virus. Vaccine 25:7271- 9. 203. Toyoda, T., T. Sakaguchi, K. Imai, N. M. Inocencio, B. Gotoh, M. Hamaguchi, and Y. Nagai. 1987. Structural comparison of the cleavage-activation site of the fusion glycoprotein between virulent and avirulent strains of Newcastle disease virus. Virology 158:242-7. 204. Tripp, R. A., L. P. Jones, L. M. Haynes, H. Zheng, P. M. Murphy, and L. J. Anderson. 2001. CX3C chemokine mimicry by respiratory syncytial virus G glycoprotein. Nat Immunol 2:732-8. 205. Ugwu, S. O., S. P. Apte. 2004. The effects of buffers on protein conformational stability. Pharmaceutical Technology:86-113. 206. Ventre, K., and A. Randolph. 2004. Ribavirin for respiratory syncytial virus infection of the lower respiratory tract in infants and young children. Cochrane Database Syst Rev:CD000181. 207. Vonhippel, P. H., and K. Y. Wong. 1964. Neutral Salts: The Generality of Their Effects on the Stability of Macromolecular Conformations. Science 145:577-80. 208. Wang, E., X. Sun, Y. Qian, L. Zhao, P. Tien, and G. F. Gao. 2003. Both heptad repeats of human respiratory syncytial virus fusion protein are potent inhibitors of viral fusion. Biochem Biophys Res Commun 302:469-75. 209. Waris, M. E., C. Tsou, D. D. Erdman, S. R. Zaki, and L. J. Anderson. 1996. Respiratory synctial virus infection in BALB/c mice previously immunized with formalin-inactivated virus induces enhanced pulmonary inflammatory response with a predominant Th2-like cytokine pattern. J Virol 70:2852-60. 210. Weissenhorn, W., A. Carfi, K. H. Lee, J. J. Skehel, and D. C. Wiley. 1998. Crystal structure of the Ebola virus membrane fusion subunit, GP2, from the envelope glycoprotein ectodomain. Mol Cell 2:605-16. 211. Weissenhorn, W., A. Dessen, S. C. Harrison, J. J. Skehel, and D. C. Wiley. 1997. Atomic structure of the ectodomain from HIV-1 gp41. Nature 387:426-30. 212. Wharton, S. A., J. J. Skehel, and D. C. Wiley. 2000. Temperature dependence of fusion by sendai virus. Virology 271:71-78.

136

213. White, J. M., S. E. Delos, M. Brecher, and K. Schornberg. 2008. Structures and mechanisms of viral membrane fusion proteins: multiple variations on a common theme. Crit Rev Biochem Mol Biol 43:189- 219. 214. Whitehead, S. S., A. Bukreyev, M. N. Teng, C. Y. Firestone, M. St Claire, W. R. Elkins, P. L. Collins, and B. R. Murphy. 1999. Recombinant respiratory syncytial virus bearing a deletion of either the NS2 or SH gene is attenuated in chimpanzees. J Virol 73:3438-42. 215. Wild, C., J. W. Dubay, T. Greenwell, T. Baird, Jr., T. G. Oas, C. McDanal, E. Hunter, and T. Matthews. 1994. Propensity for a leucine zipper-like domain of human immunodeficiency virus type 1 gp41 to form oligomers correlates with a role in virus-induced fusion rather than assembly of the glycoprotein complex. Proc Natl Acad Sci U S A 91:12676-80. 216. Wild, C., T. Greenwell, and T. Matthews. 1993. A synthetic peptide from HIV-1 gp41 is a potent inhibitor of virus-mediated cell-cell fusion. AIDS Res Hum Retroviruses 9:1051-3. 217. Wright, P. F., R. A. Karron, R. B. Belshe, J. Thompson, J. E. Crowe, Jr., T. G. Boyce, L. L. Halburnt, G. W. Reed, S. S. Whitehead, E. L. Anderson, A. E. Wittek, R. Casey, M. Eichelberger, B. Thumar, V. B. Randolph, S. A. Udem, R. M. Chanock, and B. R. Murphy. 2000. Evaluation of a live, cold- passaged, temperature-sensitive, respiratory syncytial virus vaccine candidate in infancy. J Infect Dis 182:1331-42. 218. Wu, H., D. S. Pfarr, S. Johnson, Y. A. Brewah, R. M. Woods, N. K. Patel, W. I. White, J. F. Young, and P. A. Kiener. 2007. Development of motavizumab, an ultra-potent antibody for the prevention of respiratory syncytial virus infection in the upper and lower respiratory tract. J Mol Biol 368:652-65. 219. Yin, H. S., R. G. Paterson, X. Wen, R. A. Lamb, and T. S. Jardetzky. 2005. Structure of the uncleaved ectodomain of the paramyxovirus (hPIV3) fusion protein. Proc Natl Acad Sci U S A 102:9288-93. 220. Yin, H. S., X. Wen, R. G. Paterson, R. A. Lamb, and T. S. Jardetzky. 2006. Structure of the parainfluenza virus 5 F protein in its metastable, prefusion conformation. Nature 439:38-44. 221. Yoneda, M., V. Guillaume, F. Ikeda, Y. Sakuma, H. Sato, T. F. Wild, and C. Kai. 2006. Establishment of a Nipah virus rescue system. Proc Natl Acad Sci U S A 103:16508-13. 222. Yunus, A. S., T. P. Jackson, K. Crisafi, I. Burimski, N. R. Kilgore, D. Zoumplis, G. P. Allaway, C. T. Wild, and K. Salzwedel. 2010. Elevated temperature triggers human respiratory syncytial virus F protein six-helix bundle formation. Virology 396:226-37. 223. Zhang, L., A. Bukreyev, C. I. Thompson, B. Watson, M. E. Peeples, P. L. Collins, and R. J. Pickles. 2005. Infection of ciliated cells by human parainfluenza virus type 3 in an in vitro model of human airway epithelium. J Virol 79:1113-24.

137

224. Zhang, L., M. E. Peeples, R. C. Boucher, P. L. Collins, and R. J. Pickles. 2002. Respiratory syncytial virus infection of human airway epithelial cells is polarized, specific to ciliated cells, and without obvious cytopathology. J Virol 76:5654-66. 225. Zhao, X., F. P. Chen, and W. M. Sullender. 2004. Respiratory syncytial virus escape mutant derived in vitro resists palivizumab prophylaxis in cotton rats. Virology 318:608-12. 226. Zhao, X., M. Singh, V. N. Malashkevich, and P. S. Kim. 2000. Structural characterization of the human respiratory syncytial virus fusion protein core. Proc Natl Acad Sci U S A 97:14172-7. 227. Zhao, X., and W. M. Sullender. 2005. In vivo selection of respiratory syncytial viruses resistant to palivizumab. J Virol 79:3962-8. 228. Zimmer, G., L. Budz, and G. Herrler. 2001. Proteolytic activation of respiratory syncytial virus fusion protein. Cleavage at two furin consensus sequences. J Biol Chem 276:31642-50. 229. Zimmer, G., K. K. Conzelmann, and G. Herrler. 2002. Cleavage at the furin consensus sequence RAR/KR(109) and presence of the intervening peptide of the respiratory syncytial virus fusion protein are dispensable for virus replication in cell culture. J Virol 76:9218-24. 230. Zimmer, G., M. Rohn, G. P. McGregor, M. Schemann, K. K. Conzelmann, and G. Herrler. 2003. Virokinin, a bioactive peptide of the tachykinin family, is released from the fusion protein of bovine respiratory syncytial virus. J Biol Chem 278:46854-61. 231. Zimmer, G., I. Trotz, and G. Herrler. 2001. N-glycans of F protein differentially affect fusion activity of human respiratory syncytial virus. J Virol 75:4744-51.

138